1. health and fitness
2. disease
3. cholesterol

The Metabolism of Androstenone and Other Steroid Hormone
Conjugates in Relation to Boar Taint
by
Heidi M. Laderoute
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 with Toxicology
Guelph, Ontario, Canada
© Heidi M. Laderoute, April, 2015
ABSTRACT
THE METABOLISM OF ANDROSTENONE AND OTHER STEROID HORMONE
CONJUGATES IN RELATION TO BOAR TAINT
Heidi M. Laderoute
University of Guelph, 2015
Advisor:
Dr. E.J. Squires
Increased public interest in the welfare of pigs reared for pork production has led
to an increased effort in finding new approaches for controlling the unpleasant odour and
flavour from heated pork products known as boar taint. Therefore, this study investigated
the metabolism of androstenone and the enzymes involved in its sulfoconjugation in
order to further understand the pathways and genes involved in the development of this
meat quality defect. Leydig cells that were incubated with androstenone produced 3-ketosulfoxy-androstenone, providing direct evidence, for the first time, that sulfoconjugation
of this steroid does occur in the boar. In addition, human embryonic kidney cells that
were overexpressed with porcine sulfotransferase (SULT) enzymes showed that
SULT2A1, but not SULT2B1, was responsible for sulfoconjugating androstenone. These
findings emphasize the importance of conjugation in steroid metabolism and its relevance
to boar taint is discussed.
ACKNOWLEDGEMENTS
I would like to gratefully and sincerely thank my advisor, Dr. E. James Squires,
for providing me with the opportunity to be a graduate student and for introducing me to
the world of boar taint. This project would not have been possible without your guidance,
encouragement, and patience over the last few years. I would also like to thank Dr. James
Raeside and Dr. John Cant for serving on my committee and taking the time to provide
me with valuable advice, information, and support on numerous aspects of my project.
I am very grateful to Yanping Lou for her technical expertise and friendship. I
wouldn’t have finished my studies without her ability to troubleshoot and keep the HPLC
‘Frankenstein monster’ alive. I am also grateful to Heather Christie for the time she spent
assisting with early morning Leydig cell isolations and for her advice on different
experimental approaches. I would also like to thank Dr. Dyanne Brewer from the Mass
Spectrometry Facility at the University of Guelph for her help with sample analysis and
interpretation.
My thanks and appreciation also goes to my friends and colleagues in the
Department of Animal and Poultry Science. To Dr. Matthew Gray, thank you for your
patience in answering my many questions, for helping to isolate hepatocytes, and for
teaching me how to run a Western blot. To my lab mates, both past and present, thank
you for the well-needed distractions and for reminding me that it was okay to take breaks.
Finally, thank you to my family and my amazing husband Chris. Your love and
support were vital in keeping me sane throughout the writing process.
iii
TABLE OF CONTENTS
ACKNOWLEDGEMENTS ........................................................................................... III
TABLE OF CONTENTS ............................................................................................... IV
LIST OF FIGURES ...................................................................................................... VII
LIST OF TABLES .......................................................................................................... IX
LIST OF ABBREVIATIONS ......................................................................................... X
CHAPTER 1: LITERATURE REVIEW ....................................................................... 1
1.1 INTRODUCTION ........................................................................................................... 1
1.2 ANDROSTENONE BIOSYNTHESIS ................................................................................. 4
1.2.1 Steroidogenesis ................................................................................................... 4
1.2.2 Biological Function ............................................................................................ 8
1.2.3 Accumulation in Adipose Tissue ....................................................................... 10
1.3 ANDROSTENONE METABOLISM ................................................................................ 11
1.3.1 Phase I Metabolism .......................................................................................... 11
1.3.2 Phase II Metabolism ......................................................................................... 13
1.3.2.1 Sulfoconjugation ........................................................................................ 13
1.3.2.2 Glucuronidation ......................................................................................... 20
1.3.3 Enterohepatic Circulation ................................................................................ 24
1.4 CONTROLLING BOAR TAINT ..................................................................................... 25
1.4.1 Early Slaughter ................................................................................................. 25
1.4.2 Sperm Sorting ................................................................................................... 26
1.4.3 Immunocastration ............................................................................................. 27
1.4.4 Genetic Selection .............................................................................................. 28
iv
CHAPTER 2: HYPOTHESIS AND RESEARCH OBJECTIVES ............................ 30
2.1 HYPOTHESIS ............................................................................................................. 30
2.2 RESEARCH OBJECTIVES ............................................................................................ 30
CHAPTER 3: METABOLISM OF ANDROSTENONE IN PRIMARY CULTURED
PORCINE LEYDIG CELLS AND HEPATOCYTES............................................. 32
3.1 ABSTRACT ................................................................................................................ 32
3.2 INTRODUCTION ......................................................................................................... 33
3.3 MATERIALS AND METHODS ...................................................................................... 35
3.3.1 Reagents ........................................................................................................... 35
3.3.2 Research Animals ............................................................................................. 35
3.3.3 Leydig cell isolation ......................................................................................... 35
3.3.4 Hepatocyte isolation ......................................................................................... 36
3.3.5 Biosynthesis studies .......................................................................................... 37
3.3.6 High-Performance Liquid Chromatography (HPLC) ...................................... 37
3.3.7 Steroid Conjugate Analyses .............................................................................. 38
3.3.8 Mass Spectrometry ........................................................................................... 39
3.3.9 Data Analysis.................................................................................................... 40
3.4 RESULTS ................................................................................................................... 40
3.5 DISCUSSION .............................................................................................................. 49
CHAPTER 4: THE SULFOCONJUGATION OF ANDROSTENONE AND
DEHYDROEPIANDROSTERONE BY HUMAN AND PORCINE SULT2A1
AND SULT2B1 ENZYMES ....................................................................................... 56
4.1 ABSTRACT ................................................................................................................ 56
v
4.2 INTRODUCTION ......................................................................................................... 57
4.3 MATERIALS AND METHODS ...................................................................................... 59
4.3.1 Materials........................................................................................................... 59
4.3.2 Plasmid Constructs ........................................................................................... 60
4.3.3 Cell Culture Transfection and Enzyme Expression .......................................... 61
4.3.4 HPLC ................................................................................................................ 62
4.3.5 Steroid Conjugate Analyses .............................................................................. 63
4.3.6 Western Blot Analysis ....................................................................................... 63
4.3.7 Data Analysis.................................................................................................... 64
4.4 RESULTS ................................................................................................................... 65
4.5 DISCUSSION .............................................................................................................. 77
CHAPTER 5: GENERAL DISCUSSION AND CONCLUSIONS ............................ 83
5.1 DISCUSSION AND FUTURE CONSIDERATIONS ............................................................ 83
5.2 CONCLUSIONS .......................................................................................................... 89
LITERATURE CITED .................................................................................................. 90
vi
LIST OF FIGURES
Figure 1.1 Steroid hormone biosynthesis pathway. ............................................................ 6
Figure 1.2 Steroid hormone sulfoconjugation .................................................................. 14
Figure 1.3 Potential sulfoconjugation pathways of androstenone. ................................... 16
Figure 1.4 Steroid sulfonate synthesis pathway ................................................................ 19
Figure 1.5 Steroid hormone glucuronidation .................................................................... 21
Figure 3.1 Characteristic HPLC profiles of conjugated and unconjugated steroids from 8
hour incubations of radiolabelled [3H]-androstenone with cells from the testis and
liver ....................................................................................................................... 41
Figure 3.2 Mass spectra of conjugated metabolites of androstenone produced by the testes
and liver ................................................................................................................ 44
Figure 3.3 Relative peak areas, as determined by HPLC analysis, of metabolites produced
from incubations of radiolabelled [3H]-androstenone with Leydig cells over time.
............................................................................................................................... 45
Figure 3.4 Relative peak areas, as determined by HPLC analysis, of metabolites produced
from incubations of radiolabelled [3H]-androstenone with hepatocytes over time.
............................................................................................................................... 46
Figure 3.5 Relative peak areas, as determined by HPLC analysis, of metabolites produced
from incubations of radiolabelled [3H]-androstenone with Leydig cells over time
when considering only sulfoconjugated and unconjugated steroids. .................... 47
Figure 3.6 The production of 3-keto-sulfoxy androstenone and 3-enol sulfonate from
androstenone ......................................................................................................... 52
vii
Figure 4.1 Amino acid sequence alignment of SULT2A1 enzymes from human and pig
species ................................................................................................................... 66
Figure 4.2 Amino acid sequence alignment of SULT2B1 enzymes from human and pig
species ................................................................................................................... 67
Figure 4.3 Typical Western blot of human and porcine SULT proteins. ......................... 70
Figure 4.4 Characteristic HPLC profiles of conjugated and unconjugated steroids from
HEK cells expressing SULT enzymes and incubated with radiolabelled [3H ]androstenone (A) and [1,2,6,7-3H(N)]-DHEA (B). .............................................. 71
Figure 4.5 Relative peak areas, as determined by HPLC analysis, of androstenone
sulfonate (A) and DHEAS (B) production over time for human and porcine
SULTs. .................................................................................................................. 72
Figure 4.6 The production of androstenone sulfonate by porcine SULT2A1 (A) and
human SULT2A1 (B) with increasing androstenone concentrations. .................. 73
Figure 4.7 The production of androstenone sulfonate by human SULT2B1a (A) and
human SULT2B1b (B) with increasing androstenone concentrations. ................ 74
Figure 4.8 The production of DHEAS by porcine SULT2A1 (A) and human SULT2A1
(B) with increasing androstenone concentrations. ................................................ 75
Figure 4.9 The production of DHEAS by human SULT2B1a (A) and human SULT2B1b
(B) with increasing androstenone concentrations. ................................................ 76
Figure 4.10 Kinetic model for partial substrate inhibition by DHEA............................... 81
viii
LIST OF TABLES
Table 3.1 Absolute mean peak areas of metabolites produced by Leydig cells and
hepatocytes ............................................................................................................ 48
Table 4.1 Gene-specific primers used for PCR of human and porcine SULTs ................ 60
Table 4.2 Percent identities of human and porcine SULT amino acid sequences ............ 68
Table 4.3 Kinetic constants for porcine and human SULTs ............................................. 77
ix
LIST OF ABBREVIATIONS
AI
artificial insemination
AKR
aldo-keto reductase
APS
adenosine 5’-phosphosulfate
ATP
adenosine triphosphate
Aβ
andien-β-synthetase
BSA
bovine serum albumin
CAR
constitutive androstane receptor
CYP
cytochrome P450
CYParo
cytochrome P450 aromatase
CYPscc
cytochrome P450 side chain cleavage
DHEA
dehydroepiandrosterone
DHEAS
dehydroepiandrosterone sulfonate
DHT
dihydrotestosterone
DMSO
dimethyl sulfoxide
EGTA
ethylene glycol tetraacetic acid
ESI
electrospray ionization
FBS
fetal bovine serum
FXR
farnesoid X receptor
G1P
glucose-1-phosphate
GABAA
γ-aminobutyric acid type A
GnRH
gonadotropin releasing hormone
HEK
human embryonic kidney
HEPES
hydroxy ethyl piperazine ethane sulfonic acid
HPLC
high-performance liquid chromatography
HSD
hydroxysteroid dehydrogenase
x
LC-MS
liquid chromatography-mass spectrometry
LH
luteinizing hormone
NMDA
N-methyl-D-aspartate
NMR
nuclear magnetic resonance
OATP
organic anion transporting polypeptide
PAGE
polyacrylamide gel electrophoresis
PAPS
3’-phosphoadenosine 5’-phosphosulfate
PCR
polymerase chain reaction
PSE
pale, soft, exudative
PVDF
polyvinylidene fluoride
PXR
pregnane X receptor
QTL
quantitative trait loci
Q-TOF
quadrupole time-of-flight
RIA
radioimmunoassay
RIPA
radio-immunoprecipitation assay
SDR
short-chain dehydrogenase/reductase
SDS
sodium dodecyl sulfate
SLC
solute carrier
SNP
single nucleotide polymorphism
SOAT
sodium-dependent organic anion transporter
SULT
sulfotransferase
UDP
uridine diphosphate
UDPG
uridine diphosphate glucose
UDPGA
uridine diphosphate glucuronic acid
UGT
uridine diphosphate glucuronosyltransferase
UTP
uridine triphosphate
xi
CHAPTER 1: LITERATURE REVIEW
1.1 Introduction
In the swine industry, male piglets are surgically castrated within the first week of
life in order to reduce aggression and eliminate the unpleasant odour and flavour from
heated pork products known as boar taint. 5α-androst-16-en-3-one (androstenone) and 3methylindole (skatole) are the two main compounds responsible for this meat quality
defect which occurs in approximately 5 to 15% of uncastrated (intact) male pigs
(Patterson, 1968; Lundström et al., 1988; Babol and Squires, 1999; Diaz et al., 1999).
Although castration reduces the levels of these compounds, this method is less than ideal
for both the producer and consumer.
From an economic standpoint, uncastrated boars are more profitable because they
have better feed conversion efficiency, faster growth, and greater lean yields than
castrates (Gower, 1971; Babol and Squires, 1995; Andersson et al., 1997). In addition,
intact boars excrete less nitrogen and have a higher level of unsaturated fatty acids, which
directly benefits the environment and the health of consumers, respectively (Wood et al.,
1986; Barton-Gade, 1987; Huber et al., 2013).
From an ethical standpoint, there is a growing concern for animal welfare since
the vast majority of male pigs are castrated without the use of anaesthesia and analgesia.
Consequently, this has been shown to induce physiological and behavioural reactions
indicative of pain (Giersing et al., 2006; Albrecht, 2013). During castration, pain-related
call types can be identified and it has been shown that piglets castrated without the use of
anaesthesia produce almost twice as many ‘screams’ as piglets castrated with anesthesia
1
(Marx et al., 2003). In addition, castrated piglets are less active while awake and show
more pain related behaviours such as trembling and tail wagging when compared to
sham-castrated piglets (Hay et al., 2003). Some of these behaviours last for at least four
days following castration. As a result of these welfare concerns, the European Union has
prohibited castration without anaesthesia since 2012, and intends to ban surgical
castration altogether by 2018 (EC Declaration, 2010). Therefore, there is increased
pressure placed on the pig industry worldwide to find alternative methods for reducing
boar taint without using castration. Before new methods can be developed, however,
factors affecting the synthesis and metabolism of skatole and androstenone have to be
well understood.
Androstenone is a 16-androstene steroid that is produced in the Leydig cells of the
testes together with androgens, estrogens, and other 16-androstene steroids (Gower,
1972). Its production is regulated by the hypothalamic-pituitary axis where luteinizing
hormone (LH) is released from the pituitary in response to gonadotropin releasing
hormone (GnRH) secreted from the hypothalamus (Zamaratskaia and Squires, 2009).
Since androstenone is produced in association with testicular steroids that stimulate
growth and fertility, the challenge is to reduce the level of androstenone without affecting
the other testicular steroids (Moe et al., 2007b). Due to its lipophilic properties,
androstenone readily accumulates in adipose tissue and the cut off levels used for sensory
perception of androstenone range between 0.5 μg/g and 1.0 μg/g in fat (Desmoulin et al.,
1982; Claus et al., 1994). However, consumer acceptability is highly variable and
depends on the intensity of boar taint in raw material, the manufacturing process, the
temperature at which eaten, and the consumers’ sensitivity (Desmoulin et al., 1982;
2
Annor-Frempong et al., 1997). While some consumers can detect androstenone at very
low concentrations, others are anosmic to it (Wysocki and Beauchamp, 1984; Keller et
al., 2007). Bremner et al. (2003) reviewed several studies on androstenone sensitivity and
found that between 7.6 and 75% of consumers are anosmic, while Weiler et al. (2000)
reported values between 48 and 70% as being anosmic, depending on geographic location
and gender. The ability to detect androstenone is genetically determined and is related to
the OR7D4 human odour receptor; however women are generally more sensitive
(Wysocki and Beauchamp, 1984; Weiler et al., 2000; Keller et al., 2007). While the
majority of sensitive consumers describe the odour of androstenone as urine/perspirationlike, it has been reported that up to 8% of highly sensitive consumers liked the smell of
androstenone, describing it as sweet, fruity, or perfume-like (Griffiths and Patterson,
1970; Font i Furnols et al., 2003; Andresen, 2006; Robic et al., 2008).
Unlike androstenone, most people are able to detect skatole and describe it as
having a fecal-like or naphthalene odour and bitter taste (Weiler et al., 1997, 2000;
Andresen, 2006). Its threshold value for consumer acceptance is between 0.20 μg/g and
0.25 μg/g in fat (Andresen, 2006). Skatole is produced when bacteria metabolize
tryptophan in the large intestine (Jensen et al., 1995). Some skatole is excreted with the
feces while the remaining amount is absorbed through the gut wall and is either
metabolized in the liver or absorbed into adipose tissue (Lundström and Zamaratskaia,
2006; Tajet et al., 2006; Zamaratskaia and Squires, 2009). Because the amount of skatole
produced is primarily dependent on the availability of tryptophan and the composition of
intestinal bacteria, it can be controlled through dietary means (Giersing et al., 2006;
Lundström and Zamaratskaia, 2006; Zamaratskaia and Squires, 2009). Including certain
3
types of fibers, such as raw potato starch or sugar beet pulp in the diet, for example, has
been shown to reduce skatole levels in the fat and plasma of boars (Van Oeckel et al.,
1998; Whittington et al., 2004; Zamaratskaia et al., 2005; Chen et al., 2007b). In addition,
undigested carbohydrates have been shown to increase fecal wet and dry weight and
decrease intestinal transit time, which reduces the amount of skatole absorbed through the
intestinal wall (Drochner, 1993; Wang et al., 2004; Zamaratskaia and Squires, 2009). In
addition to diet, fat skatole levels can be reduced by rearing pigs on slatted floors rather
than concrete, by using wet feeding instead of dry, by keeping pigs clean, and by
providing unlimited access to drinking water (as reviewed by Bonneau, 1998). Because
skatole can largely be controlled through diet and other environmental factors, and
because androstenone is strongly associated with testicular steroids and the occurrence of
boar taint, the remainder of this review will focus on biochemical and genetic factors
involved in the regulation of androstenone in boars.
1.2 Androstenone Biosynthesis
1.2.1 Steroidogenesis
The production of androstenone and other testicular steroids is controlled by the
neuroendocrine system. LH is secreted by the pituitary gland in response to GnRH and
binds to receptors on the Leydig cells to increase steroidogenesis (Claus et al., 1994;
Zamaratskaia et al., 2007). Consequently, androstenone, androgens, and estrogens are
secreted following a closely parallel pattern (Andresen, 2006). Although a temporary
activation of the hypothalamic-pituitary-gonadal axis occurs at approximately two to four
weeks of age, biosynthesis of androstenone and other testicular steroids is low in young
pigs (Gower, 1972; Bonneau, 1982; Kwan et al., 1985; Sinclair et al., 2001). As boars
4
reach sexual maturity, at approximately 14 to 15 weeks of age, there is a surge of
testosterone and androstenone production, which is quickly followed by an increase in
the concentration of androstenone in fat (Claus et al., 1994; Gunn et al., 2004;
Zamaratskaia et al., 2008a,b). These steroids continue to gradually increase and reach
their highest levels at approximately two years of age (Booth, 1975). Therefore,
steroidogenesis depends to a large extent on sexual maturity and consequently on the age
and weight of the animal (Bonneau, 1982).
All steroid hormones are synthesized from cholesterol and thus have very similar
structures (Figure 1.1). In addition, steroid synthesis is primarily carried out by
cytochrome P450 (CYP) enzymes, a family of heme-containing mono-oxygenases
located on intracellular membranes (Rose et al., 1997). The first step in steroidogenesis
involves the side chain cleavage of cholesterol to produce pregnenolone by the enzyme
cytochrome P450 side chain cleavage (CYPscc), also known as CYP11A1 (Payne and
Youngblood, 1995; Robic et al. 2008, 2014). This occurs in the inner mitochondrial
membrane (Hébert and Cooke, 1992; Payne and Youngblood, 1995). Pregnenolone is
then transported to the smooth endoplasmic reticulum where it can be converted into
progesterone, 17α-hydroxypregnenolone, or 5,16-androstadien-3β-ol (androstadienol)
(Robic et al., 2014). Both pregnenolone and progesterone are precursors for the three
major groups of steroid hormones, corticoids, androgens, and estrogens, as well as the
16-androstene steroids (Katkov and Gower, 1970; Saat et al., 1972; Brooks and Pearson,
1986; Robic et al., 2014).
Androgens and estrogens are necessary for spermatogenesis and sexual behaviour
(Gunn et al., 2004). In addition, they favour the development of lean tissue and influence
5
3α-androstenol
3α-HSD
5α-R
3β-HSD
Aβ
Aldosterone
3β-androstenol
CYP11
3β-HSD
Androstenone
Androstadienone
Androstadienol
Aβ
11β-HSD
CYP11
CYP21
3β-HSD
CYPSCC
11-dehydro
corticosterone
Corticosterone
11-DOC
Progesterone
CYP17
CYP17
Pregnenolone
Cholesterol
11β-HSD
CYP11
CYP21
3β-HSD
Cortisone
Cortisol
11-deoxycortisol
17α-OHprogesterone
CYP17
17α-OHpregnenolone
CYP17
CYParo
CYParo
3β-HSD
17β-HSD
Estrone
17β-HSD
19-OH-∆4-AD
Androstenedione
DHEA
17β-HSD
CYParo
CYParo
3β-HSD
Estradiol
19-OH-Testo
Testosterone
Androstenediol
Figure 1.1 Steroid hormone biosynthesis pathway. CYPscc, cytochrome P40 side chain cleavage; Aβ, andien β-synthetase; 3βAdapted
Brooks and Pearson,
1986 and Robic et5α-R,
al., 2014
HSD,from
5α-reductase; 3α-HSD, 3α-hydroxysteroid dehydrogenase; CYP21, cytochrome P450
3β-hydroxysteroid
dehydrogenase;
21; 11-DOC, 11-deoxycorticosterone; CYP11, cytochrome P450 11; 11β-HSD, 11β-hydroxysteroid dehydrogenase; CYP17,
cytochrome P450 17; CYParo, cytochrome P450 aromatase; 19-OH-∆4-AD, 19-hydroxy androstenedione; 17β-HSD, 17βhydroxysteroid dehydrogenase; 19-OH-Testo, 19-hydroxy testosterone (adapted from Brooks and Pearson, 1986; Robic et al., 2014).
6
metabolism, which helps to explain the improved carcass qualities seen in boars
compared to castrates (Babol et al., 1998; Gunn et al., 2004). Testosterone, the main
androgen in males, can be synthesized through two metabolic pathways. In the first
pathway, 17α-hydroxyprogesterone is converted to androstenedione by CYP17, which is
then transformed into testosterone via the reduction of the C-17 keto group to a hydroxyl
group (Cooke and Gower, 1977; Ruokonen, 1978; Shimizu, 1979; Brooks and Pearson,
1986; Robic et al., 2014). The alternate pathway involves the oxidation of 17αhydroxypregnenolone to dehydroepiandrosterone (DHEA), also by CYP17, which then
undergoes oxidation and isomerisation to yield androstenedione or androstenediol
(Brooks and Pearson, 1986). Cytochrome P450 aromatase (CYParo), also known as
CYP19A, expressed in Leydig cells, is responsible for converting testosterone to estradiol
(Payne and Youngblood, 1995).
The conversion of pregnenolone into the 16-androstene steroids can also occur
through two pathways; however, unlike androgen and estrogen synthesis, neither pathway
requires prior C-17 hydroxylation of pregnenolone or progesterone (Brophy and Gower,
1972; Gower, 1972; Shimizu, 1979). In the first pathway, pregnenolone is converted into
progesterone, which is then converted into androsta-4,16-dien-3-one (androstadienone).
Androstadienone is then metabolized to androstenone via 5α-reductase, which is further
converted into either 5α-androst-16-en-3α-ol (3α-androstenol) or 5α-androst-16-en-3β-ol
(3β-androstenol) (Brophy and Gower, 1972; Bonneau, 1982). In the second and more
predominant pathway, pregnenolone is irreversibly converted to androsta-5,16-dien-3β-ol
(androstadienol) by the andien-β-synthetase (Aβ) enzyme complex. Androstadienol is
further metabolized to androstadienone by 3β-hydroxysteroid dehydrogenation and
7
isomerization of the double bond from the B ring to the A ring (Hébert and Cooke, 1992;
Sinclair, 2004). The Aβ enzyme complex responsible for 16-androstene production
includes CYP17 and cytochrome b5 (CYB5) (Ahmad and Gower, 1968; Meadus et al.,
1993; Moe et al., 2007a; Robic et al., 2008, 2014). CYB5, a membrane bound protein,
stimulates the activity of CYP17 by acting as an allosteric modulator to promote the
association of CYP17 with its electron donor (Zamaratskaia et al., 2007). CYB5 is also
particularly important in the synthesis of androstenone, as pigs with high CYB5 activity
have high levels of androstenone in fat (Davis and Squires, 1999; Leung et al., 2010). In
addition, a single nucleotide polymorphism (SNP) has been found in the untranslated
region of the CYB5 gene which is associated with a lower fat androstenone level (Lin et
al., 2005).
1.2.2 Biological Function
Androstenone synthesized in the testis is released into the spermatic vein and then
passes into the systemic circulation where the majority is taken up by the submaxillary
glands and adipose tissue, and the remaining portion is metabolized (Katkov et al., 1972;
Bonneau, 1982; Lundström and Zamaratskaia, 2006; Zamaratskaia and Squires, 2009).
The androstenone taken up by the submaxillary glands becomes concentrated by binding
to a specific low molecular mass protein called pheromaxein (Booth, 1987; Austin et al.,
2004; Andresen, 2006). Once bound, androstenone is reduced to physiologically active
3α-androstenol and, to a lesser extent, 3β-androstenol (Katkov et al., 1972; Booth, 1987).
The function of pheromaxein is considered essential since the 16-androstenes are highly
lipophilic and occur in very high concentrations in submaxillary glands and saliva
8
(Booth, 1980a,b, 1987; Booth and von Glos, 1991; Austin et al., 2004; Andresen, 2006).
When a boar comes in contact with a female, he starts to salivate profusely and thus
excretes a large amount of androstenone and 3α-androstenol into the environment
(Bonneau, 1982; Booth, 1987; Sinclair, 2004). These androstenes consequently stimulate
the typical mating stance seen in sows that are in estrous, allowing the boar to mount and
copulate (Bonneau, 1982; Brooks and Pearson, 1986; Lundström and Zamaratskaia,
2006). Therefore, although the 16-androstenes do not possess any androgenic activity,
androstenone and 3α-androstenol do act as pheromones to stimulate reproductive
functions in the female pig (Andresen, 1976, 2006). In addition to being present in the
submaxillary glands, androstenone and 3α-androstenol have also been found in boar
sweat glands which are assumed to also play a role in boar courtship behaviour (Stinson
and Patterson, 1972; Perry et al., 1980; Bonneau, 1982).
Several other possible pheromonal actions of the 16-androstene steroids have
been investigated. For example, androstenone has been shown to suppress aggressive
behaviour in young pigs and it is thought that the level of androstenes in saliva may be
involved in establishing hierarchy (Andresen, 1976; McGlone et al., 1987). In addition, it
has been shown that androstenone induces the release of oxytocin in estrous pigs, which
may affect the fertility of the female by influencing the sperm transport rate (Mattioli et
al., 1986). There is also evidence suggesting that 3α-androstenol is involved as a priming
pheromone which acts to accelerate puberty in gilts (Hughes, 1982; Kirkwood et al.,
1983; Pearce et al., 1988).
9
1.2.3 Accumulation in Adipose Tissue
Androstenone is highly lipophilic and accumulates in adipose tissue in much
higher concentrations than the other testicular hormones (Brooks and Pearson, 1986;
Claus et al., 1994; Robic et al., 2008). However, storage in adipose tissue is reversible,
since fat androstenone levels decrease following castration (Bonneau, 1982; Brooks and
Pearson, 1986; Claus et al., 1994). The concentration of androstenone in adipose tissue is
directly related to the concentration in blood, which is consequently determined by a
boar’s genetic capacity to both synthesize and metabolize 16-androstenes (Zamaratskaia
et al., 2008a). In plasma, androstenone concentration is variable, ranging from a few
nanograms to at least 60 ng/ml, and levels greater than 15 ng/ml have been associated
with high androstenone concentrations in adipose tissue (Andresen, 1976; Sinclair et al.,
2001; Andresen, 2006). Therefore, increased production, decreased metabolism, or both
will lead to a higher amount of androstenone in the blood, which will consequently lead
to an increased concentration in adipose tissue (Sinclair et al., 2001).
In many species, steroids are transported to target tissues partly bound to specific
proteins such as sex hormone binding globulin (SHBG) (Andresen, 2006; Zamaratskaia
et al., 2008a). The binding of androstenone to plasma proteins would increase its polarity
and reduce the transfer of the steroid into adipose tissue (Andresen, 2006; Zamaratskaia
et al., 2008a). However, SHBGs have not been detected in porcine plasma, and it is
possible that androstenone only binds non-specifically to serum components such as αglobulins and albumin (Cook et al., 1977; Zamaratskaia et al., 2008a). Since
androstenone binding capacity does not vary greatly between animals, this non-specific
binding is unlikely to affect androstenone accumulation in adipose tissue (Zamaratskaia
et al., 2008a).
10
1.3 Androstenone Metabolism
The metabolism of androstenone and other steroids occurs primarily in the
kidney, liver, and testes through both phase I and phase II reactions (Hobkirk, 1985;
Sinclair, 2004). Phase I metabolism includes oxidation, reduction, and hydrolysis
reactions while phase II metabolism involves the conjugation of a hydrophilic group,
such as sulfonate or glucuronic acid, to hydroxyl groups that already exist on the steroid
or are added on during phase I metabolism (Sinclair, 2004; Rasmussen et al., 2012a,b).
Consequently, the biological properties of steroids and other compounds undergoing
these metabolic biotransformations are dramatically affected. Typically, phase I and
phase II reactions have been thought of as deactivation or detoxification mechanisms;
however the biological significance of these reactions is more complex than this
(Sinclair, 2004; Rasmussen et al., 2012b).
1.3.1 Phase I Metabolism
The key enzymes involved in the phase I metabolism of androstenone are
hydroxysteroid dehydrogenases (HSDs) (Doran et al., 2004). HSDs are oxidoreductases
that belong to two distinct protein superfamilies. The short-chain
dehydrogenase/reductase (SDR) family includes 3β-HSD, 11β-HSD, and 17β-HSD,
while the aldo-keto reductase (AKR) family includes 3α-HSD and 20α-HSD (Pawlowski
et al., 1991; Krozowski, 1994; Miura et al., 1994; Jörnvall et al., 1995; Penning, 2003).
The reduction and oxidation activities of HSDs allow isoforms to function as molecular
switches that will either inactivate a hormone or produce an active ligand, thus regulating
the amount of hormone that binds to a steroid hormone receptor (Penning, 1997, 2003).
11
In the case of androstenone, 3α- and 3β- HSD reduce the 3-keto group to a 3α- or 3βhydroxyl group to produce either 3α-androstenol or 3β-androstenol, respectively (Figure
1.1) (Doran et al., 2004; Sinclair et al., 2005a,b). Androstenone was also shown to be
transformed into 6-hydroxy androstenone in axillary microflora; however the enzyme
responsible for this is unknown (Austin and Ellis, 2003). Consequently, these compounds
are less likely to accumulate in adipose tissue due to the hydrophilic nature of the
hydroxyl groups (García-Regueiro and Diaz, 1989). Although hepatic and testicular
phase I metabolism differ in terms of the percentage of produced metabolites, the rate of
3β-androstenol formation, and therefore 3β-HSD activity, is three to six times higher than
that of 3α-androstenol, regardless of metabolic site (Brophy and Gower, 1972; Doran et
al., 2004, 2014; Sinclair et al., 2005a; Zamaratskaia and Squires, 2009).
3β-HSD is a particularly important enzyme as it is involved in the biosynthesis of
all steroid hormones and controls critical steroid hormone related actions in the adrenal
cortex, gonads, liver, fat, and other peripheral target tissues (Mason et al., 1998; Kershaw
and Flier, 2004; Rasmussen et al., 2013). Numerous 3β-HSD isoforms have been
identified in several species including three in humans, four in rats, and six in mice;
however only one isoform has been found in pigs (Simard et al., 1993; Abbaszade et al.,
1997; McBride et al., 1999; Von Teichman et al., 2001; Rasmussen et al., 2013). Several
investigations have shown that low expression and activity of 3β-HSD in liver and testis
is associated with reduced clearance and increased accumulation of androstenone in
adipose tissue (Doran et al., 2004; Nicolau-Solano et al., 2006; Chen et al., 2007a).
Conversely, high 3β-HSD expression and activity enhance androstenone metabolism and
thus decrease androstenone accumulation in adipose tissue (Chen et al., 2007a;
12
Rasmussen et al., 2012b). Nicolau-Solano and Doran (2008) established that
androstenone, as well as testosterone and estrone sulfate, induce expression of hepatic 3βHSD; however, this was dependent on substrate concentration and the age and weight of
the boar. Conversely, Chen et al. (2007a) suggested that the induction of 3β-HSD
expression may be a result of the absence of steroids following castration. External
factors, such as diet and environmental toxins, have also been shown to affect the activity
of 3β-HSD (Fink-Gremmels and Malekinejad, 2007; Hu et al., 2010; Rasmussen et al.,
2012b, 2013). The exact mechanisms regulating this enzyme in porcine liver and testis,
however, are still unclear.
1.3.2 Phase II Metabolism
Phase II conjugation reactions play an important role in the biotransformation of
hormones, neurotransmitters, and xenobiotic compounds (Falany, 1997). These reactions
generate metabolites that are more water-soluble and thus more readily excreted in the
urine and bile (Falany, 1997). Sulfoconjugation and glucuronidation are the primary
phase II reactions responsible for the metabolism of androstenes, androgens, and
estrogens.
1.3.2.1 Sulfoconjugation
Sulfoconjugation, or sulfonation, is a major phase II biotransformation reaction
that is important in the synthesis, transport, and metabolism of steroids (Falany, 1997;
Weinshilboum et al., 1997). Sulfonation of steroids involves the transfer of a sulfo group
(SO3-) from a donor molecule to the hydroxyl group of an acceptor molecule to form a
13
sulfonate conjugate (Figure 1.2) (Falany et al., 1995; Strott, 2002). Due to phase I
metabolism, hydroxyl groups may be present at multiple positions on the steroid nucleus;
however hydroxyl groups at positions 3, 17, and 21 are the most common sites for
sulfonation to occur (Adams and McDonald, 1981; Strott, 1996). The donor molecule
required for sulfonation is 3’-phosphoadenosine 5’-phosphosulfate (PAPS), and all
mammalian tissues are able to carry out its synthesis (Robbins et al., 1956; Falany et al.,
1995). PAPS is synthesized in the cytosol from inorganic sulfate and adenosine
triphosphate (ATP) via a two-step process involving the enzymes ATP sulfurylase and
adenosine 5’-phosphosulfate (APS) kinase (Falany, 1997; Strott, 2002). Sources of
inorganic sulfate required for PAPS synthesis are acquired through the diet or from the
catabolism of sulfur-containing amino acids (Falany, 1997).
Sulfonation reactions are mediated by a family of enzymes termed
sulfotransferases (SULTs) which are found in many tissues including liver, kidney, testis,
ovary, intestinal tract, lung, and brain (Roberts and Lieberman, 1970; Gasparini et al.,
1976; Hobkirk, 1985; Falany et al., 1995). Two broad classes of SULTs have been
identified in human and animal tissues: (1) membrane-bound SULTs that are localized in
the Golgi apparatus of most cells and are responsible for the sulfonation of
+
Steroid
+
SULT
PAPS
Steroid Sulfonate
Figure 1.2 Steroid hormone sulfoconjugation. PAPS, 3’-phosphoadenosine 5’phosphosulfate; SULT, sulfotransferase; PAP, 3’- phosphoadenosine 5’-phosphate.
14
PAP
macromolecular endogenous structures such as glycosaminoglycans, glycoproteins, and
tyrosines in proteins and peptides; and (2) cytosolic SULTs which metabolize relatively
small molecules including steroid hormones, xenobiotics, and neurotransmitters (Falany
et al., 1995; Falany, 1997; Glatt and Meinl, 2004). To date, six distinct cytosolic SULT
gene families have been identified in mammals; however only the SULT1 and SULT2
families are capable of sulfonating steroids (Strott, 2002; Blanchard et al., 2004; Neunzig
et al., 2014). The SULT1 family primarily conjugates phenolic molecules including
estrogens, thyroid hormones, catecholamines, and xenobiotics (Falany et al., 1989; Strott,
2002). The SULT2 family is primarily involved with the conjugation of hydroxyl groups
of steroids with saturated A rings, including 3α-androstenol and 3β-androstenol (Falany
et al., 1994; Falany, 1997; Falany and Rohn-Glowacki, 2013). Previously, it was thought
that since androstenone did not contain any hydroxyl groups, it could not undergo
sulfonation; however hydroxylated forms of androstenone have been produced by
axillary microflora and porcine hepatocytes (Austin and Ellis, 2003; Sinclair et al.,
2005b; Chen et al., 2007b). In addition, a large proportion of androstenone sulfonate has
been found in the plasma of boars (Sinclair and Squires, 2005). It has also been suggested
that androstenone may undergo enolisation of the 3-keto group prior to sulfoconjugation
(Sinclair et al., 2005a,b, 2006). Since steroid enols are relatively unstable,
sulfoconjugation may serve to stabilize this form of steroid (Sinclair et al., 2005a,b). The
potential pathways of androstenone sulfoconjugation are shown in Figure 1.3.
The SULT2 family consists of SULT2A1 and SULT2B1 subfamilies. In pigs,
SULT2A1 is thought to be the key enzyme in the hepatic and testicular phase II
metabolism of androstenone (Sinclair 2004, Sinclair et al., 2005b, 2006). This enzyme is
15
Androstenone
ENO
Hydroxy-androstenone
3-enol
SULT
SULT
SULT
3-enol sulfonate
Hydroxy-androstenone sulfonate
3-keto-4-sulfate
Figure 1.3 Potential sulfoconjugation pathways of androstenone. ENO, enolase;
SULT, sulfotransferase
(adapted
from
Sinclair2011
et al., 2005b; Desnoyer, 2011).
Adapted
from Sinclair et al.,
2005b and
Desnoyer,
capable of sulfonating both 3α- and 3β-androstenol and its expression and activity have
been negatively correlated with both fat and plasma androstenone concentrations in boars
(Falany et al., 1989, 1994; Sinclair and Squires, 2005; Sinclair et al., 2006). SULT2A1
activity may be affected by genetic polymorphisms, as several SNPs in humans have
resulted in decreased levels of enzyme activity and protein (Igaz et al., 2002; Thomae et
al., 2002). In addition, porcine SULT2A1 has been shown to be down-regulated by
various nuclear receptors including the constitutive androstane receptor (CAR), the
pregnane X receptor (PXR), and the farnesoid X receptor (FXR) (Gray and Squires,
2013). This has the potential to decrease steroid sulfoconjugation and increase
androstenone accumulation in adipose tissue. SULT2B1 has also been negatively
16
correlated to fat androstenone concentration and has shown selectivity for 3βhydroxysteroids (Moe et al., 2007a,b; Meloche and Falany, 2001). Therefore, SULT2B1
may also play an important role in the phase II metabolism of androstenone, although
direct evidence has yet to be generated. In pigs, only one SULT2B1 enzyme exists;
however in humans this gene encodes two isoforms, SULT2B1a and SULT2B1b, which
are generated by alternate splicing of the first exon (Her et al., 1998; Meloche and
Falany, 2001; Kohjitani et al., 2006).
In the blood, the concentration of sulfoconjugated steroids is several times higher
than the unconjugated or free form (Saat et al., 1972; Gasparini et al., 1976; Neunzig et
al., 2014). In fact, Sinclair and Squires (2005) showed that up to 69-72% of
androstenone, 3α-androstenol, and 3β-androstenol are present in their sulfoconjugated
forms in the peripheral and testicular vein plasma of mature boars. This may be
attributable to an increase in serum protein binding, which would dramatically increase
the half-life and decrease metabolic clearance rates of steroid sulfonates (Ruder et al.,
1972; Glatt and Meinl, 2004). Protein binding also provides a soluble inactive transport
mechanism to target tissues where the unconjugated steroid could be regenerated via
hydrolysis of the sulfonate group by the action of membrane bound sulfatase enzymes
(Falany, 1997; Glatt, 2000; Sinclair et al., 2005a). Therefore, high levels of
sulfoconjugated steroids in the blood could act as a reservoir that serves to regulate the
availability of active steroid hormones at both systemic and local levels (Hobkirk, 1993;
Glatt and Meinl, 2004). In this way, the level of free androstenone that accumulates in
adipose tissue can be controlled. These steroid sulfonates are also capable of acting as
substrates for steroidogenic enzymes in the steroid-sulfonate synthesis pathway (Figure
17
1.4) (Falany et al., 1995; Sinclair, 2004). This pathway allows steroid sulfonates to be
metabolized in the same way as unconjugated steroids, with compounds retaining their
sulfonate groups throughout the entire series of biotransformations, originating from
cholesterol sulfonate (Roberts et al., 1964; Gasparini et al., 1976). Cholesterol can be
sulfoconjugated at its 3β-hydroxyl group and subsequently converted into pregnenolone
sulfonate by side-chain cleavage via CYPscc (Hochberg et al., 1974; Tuckey, 1990).
Pregnenolone sulfonate can then be metabolized to 17-hydroxypregnenolone sulfonate
via CYP17 and to androstadienol sulfonate (Ruokonen, 1978; Gasparini et al., 1976;
Neunzig et al., 2014). Further, 17-hydroxypregnenolone sulfonate can be converted to
dehydroepiandrosterone sulfonate (DHEAS) (Lebeau et al., 1964). Since any of these
steroid sulfonates can be hydrolyzed to their unconjugated forms by sulfatase enzymes,
the steroid-sulfonate synthesis pathway may serve as an alternate route for the formation
of free, active steroid hormones.
Sulfonation typically increases water solubility and prevents steroid sulfates from
binding to and activating steroid receptors (Falany, 1997; Strott, 2002). In addition,
steroid sulfonates are unable to passively diffuse through cell membranes, since these
compounds remain ionized at physiological pH values (Falany, 1997; Glatt and Meinl,
2004). However, cell uptake of steroid sulfonates can occur via the membrane bound
solute carrier (SLC) group of transporter proteins found in a number of tissues including
kidney, liver, lung, and testis (Hagenbuch and Meier, 2003; Harteneck, 2013). For
example, the organic anion transporting polypeptide (OATP) family has been shown to
have broad and overlapping substrate specificities for steroid conjugates as well as bile
18
Sulfonate Pathway
Free Steroid Pathway
SULT
Cholesterol sulfonate
CYPscc
Cholesterol
CYPscc
Aβ
SULT
Androstadienol
sulfonate
Pregnenolone
sulfonate
Pregnenolone
CYP17
CYP17
SULT
17α-OH-pregnenolone
sulfonate
CYP17
17α-OH-pregnenolone
CYP17
SULT
DHEAS
DHEA
Figure
Steroid
sulfonate
synthesis
pathway.
SULT,
sulfotransferase;
CYPscc,
Figure
1.41.4
Steroid
sulfonate
synthesis
pathway.
SULT,
sulfotransferase;
CYPscc
,
cytochrome
P450
chain
cleavage;
andien-β-synthetase;
CYP17,
cytochrome
cytochrome
P450
sideside
chain
cleavage;
Aβ,Aβ,
andien-β-synthetase;
CYP17,
cytochrome
P450
17;17;
DHEAS,
dehydroepiandrosterone
sulfate;
DHEA,
dehydroepiandrosterone
P450
DHEAS,
dehydroepiandrosterone
sulfonate;
DHEA,
dehydroepiandrosterone
(adapted
from
Hobkirk,
1985;
Sinclair,
2004).
(adapted from Hobkirk, 1985; Sinclair, 2004).
19
salts, thyroid hormones, and xenobiotics (Meier et al., 1997; Kullak-Ublick et al., 2001;
Hagenbuch and Meier, 2003; Petzinger and Geyer, 2006). Further, one member of the
sodium-dependent organic anion transporter (SOAT) family appears to be specific for the
transport of a number of steroid sulfonates including pregnenolone sulfonate and DHEAS
(Geyer et al., 2006, 2007; Fietz et al., 2013). Although knowledge of the physiological
role of steroid sulfonates is limited, recent studies have shown that they are involved in
many different processes that do not require regeneration of the unconjugated steroid
(Sinclair, 2004; Neunzig et al., 2014). Cholesterol sulfonate has a stabilizing role in cell
membranes, supports platelet adhesion, and can regulate the activity of a variety of
functional proteins (as reviewed by Strott and Higashi, 2003); pregnenolone sulfonate has
been shown to regulate the γ-aminobutyric acid type A (GABAA) and the N-methyl-Daspartate (NMDA) receptors in the brain (Bowlby, 1993; Park-Chung et al., 1999); and
DHEAS induces a non-classical signalling pathway in spermatogenic cells (Shihan et al.,
2013). The physiological role of androstenone sulfonate has not been investigated.
1.3.2.2 Glucuronidation
Glucuronidation is another major phase II biotransformation process involved in
steroid metabolism. Steroid glucuronidation involves the transfer of glucuronic acid from
uridine diphosphate (UDP) glucuronic acid (UDPGA) to a hydrophobic steroid molecule
that has one or more electrophilic groups (Figure 1.5) (Bélanger et al., 1998; Shipkova
and Wieland, 2005). UDPGA is synthesized via a two-step process. First, uridine
diphosphate glucose (UDPG) is produced from glucose-1-phosphate (G1P) and uridine
triphosphate (UTP) by the action of UDPG pyrophosphorylase. Second, UDPG is
oxidized by UDPG dehydrogenase to produce UDPGA. Since glycogen is the ultimate
20
precursor of UDPGA, glucuronidation capacity is dependent on the levels of this
compound (Miettinen and Leskinen, 1970). Like sulfonation, glucuronidation can occur
at multiple sites on the steroid molecule due to functional groups added during phase I
metabolism. However, the most common sites for glucuronidation to occur are at the 3αand 21-hydroxyl groups of C21 steroids and the 3-(C18) or 3α-(C19) and 17β-hydroxyl
groups of C18 and C19 steroids (Makin and Trafford, 1972). Steroids with 3β-hydroxyl
groups such as DHEA are generally excreted as sulfonates (Mackenzie et al., 1992).
Glucuronidation reactions are catalyzed by uridine diphosphate
glucuronosyltransferase (UGT) enzymes found in the endoplasmic reticulum of a number
of tissues including liver, kidney, intestine, skin, and testes (Miettinen and Leskinen,
1970; Dutton, 1978; Burchell and Coughtrie, 1989). Mammalian UGTs have been
classified into four major families based on protein sequence similarity: UGT1, UGT2,
UGT3, and UGT8 (Mackenzie et al., 1997, 2005). The UGT1 family primarily
conjugates bilirubin, estrogens, bile acids and numerous other drugs and xenobiotics
(Ebner and Burchell, 1993; Senafi et al., 1994; Hum et al., 1999). The UGT2 family is
divided into three subfamilies: UGT2A, whose members are specific to the olfactory
UGT
+
Steroid
+
UDPGA
UDP
Steroid Glucuronide
Figure 1.5 Steroid hormone glucuronidation. UDPGA, uridine diphosphate glucuronic
acid; UGT, uridine diphosphate glucuronosyltransferase; UDP, uridine diphosphate.
21
epithelium and have glucuronidation activity towards several odorants (Lazard et al.,
1991; Jedlitschky et al., 1999); UGT2B, whose members conjugate steroids, bile acids,
fatty acids, carboxylic acids, phenols and carcinogens (Mackenzie et al., 1997; Turgeon
et al., 2001); and UGT2C, which has only been identified in rabbits (Parkinson, 2001).
The UGT3 family uses UDP-N-acetylglucosamine, UDP-glucose, or UDP-xylose instead
of UDPGA as a glycosyl source and conjugates steroid hormones, bile acids, and
xenobiotics (Mackenzie et al., 2008, 2011; Meech and Mackenzie, 2010). The UGT8
family contains only one member, UGT8A1, which uses UDP-galactose instead of
UDPGA as a glycosyl source to conjugate ceramides and bile acids (Sprong et al., 1998;
Meech et al., 2015).
The UGT enzymes have wide and overlapping substrate specificities; however
enzymes belonging to the UGT2B family are mainly responsible for the glucuronidation
of steroids (Mackenzie et al., 1992; Turgeon et al., 2001). In humans, UGT2B7,
UGT2B15, and UGT2B17 have shown a high capacity to conjugate androgens (Bélanger
et al., 2003). UGT2B7 conjugates 5α-reduced metabolites of mineralocorticoids,
glucocorticoids, progestins, and androgens, as well as 5β-reduced C19 and C21 steroids
(Bélanger et al., 2003; Girard et al., 2003). UGT2B15 specifically conjugates the 17hydroxy position of 5α-reduced androgens, while UGT2B17 conjugates both the 3- and
17-hydroxy positions (Beaulieu et al., 1996; Bélanger et al., 2003). Although
glucuronidation of androstenone, 3α-androstenol, and 3β-androstenol is possible, specific
UGT enzymes responsible for these conjugations have not been identified.
As with sulfoconjugation, glucuronidation results in an increase in polarity and
facilitates excretion via the bile or urine (Mackenzie et al., 1992). In addition, steroid
22
glucuronidation may regulate the availability of active steroid hormones by serving as a
storage pool from which the free steroid can be regenerated via glucuronidase enzymes
(Bradlow, 1970; Hobkirk, 1985; Guillemette et al., 1996). However, high levels of certain
steroid glucuronides have been associated with hepatotoxicity. 17β-estradiol glucuronide
and testosterone glucuronide, for example, have been shown to inhibit bile flow, thus
inducing cholestasis (Gosland et al., 1993; Clarke and Burchell, 1994). In contrast to
sulfonation, a glucuronide synthesis pathway does not seem to exist as steroid
glucuronides generally do not act as substrates for the enzymes involved in
steroidogenesis (Roy, 1970; Sinclair, 2004). However, Rittmaster (1993) has suggested
that the conversion of dihydrotestosterone (DHT) glucuronide to 3α-androstanediol 17glucuronide, and of 3α-androstanediol 3-glucuronide to androsterone glucuronide is
possible.
Although UGT enzymes are found in multiple extrahepatic tissues, the 16androstenes have not been detected as glucuronide conjugates in either the peripheral or
testicular vein plasma of boars (Sinclair and Squires, 2005; Sinclair et al., 2005a). This
led to the suggestion that the glucuronidation of 16-androstene steroids serves primarily
as a mechanism to facilitate metabolic clearance (Sinclair et al., 2005b). This is supported
by the high concentrations of androgen, estrogen, and androstene glucuronides found in
the urine and bile, as well as the high levels of UGT enzymes found in the liver and
kidney (Gower and Patterson, 1970; Sinclair et al., 2005b). In terms of biological roles in
relation to boar taint, steroid glucuronidation has received very little attention, and much
more research is needed.
23
1.3.3 Enterohepatic Circulation
Most steroids and other compounds conjugated in the liver are excreted in the bile
(Winter and Bokkenheuser, 1987). When these compounds reach the duodenum, they are
exposed to the action of deconjugating enzymes secreted by the intestinal wall as well as
by the intestinal microflora. The deconjugated compounds are then either reabsorbed in
the ileum and caecum or eliminated in the feces (Eriksson, 1971; Groh et al., 1993). The
intestinal wall only produces glucuronidase enzymes while intestinal bacteria, primarily
Bacteroides, Escherichia coli, and Clostridiae, synthesize both glucuronidases and
sulfatases (Adlercreutz and Martin, 1980; Macdonald et al., 1983). Gastrointestinal
disorders, antibiotic treatment, and diet can all influence the composition of the intestinal
microflora and thus impact the quantity of steroids that are reabsorbed (Van Eldere et al.,
1990; Groh et al., 1993). Consequently, there is a large interest in finding ways to reduce
androstenone concentrations through dietary additives that bind androstenone in the
intestine (for review see Jen, 2009). Increasing the conjugation of steroids by enhancing
the activation of SULTs and UGTs in the intestine, or decreasing the deconjugation
activity of the intestine and bacteria would potentially increase the amount of steroids
excreted in feces and decrease the amount of androstenone available to accumulate in
adipose tissue.
Following reabsorption, steroids are reconjugated by SULTs and UGTs present in
the intestinal wall and liver (Falany, 1997). From here, the reconjugated steroids may
once again undergo enterohepatic circulation or be sent to the kidney and excreted in the
urine (Groh et al., 1993). In addition, these steroids may also re-enter the systemic
circulation and become available to tissues that are capable of removing the conjugate
groups. Although deconjugation is considered essential for efficient reabsorption,
24
evidence has suggested that some conjugated steroids such as estrone sulfonate, estrone
glucuronide, pregnenolone sulfonate, and deoxycorticosterone sulfonate are capable of
being reabsorbed without cleavage of the conjugate group (Eriksson, 1971; Sim et al.,
1983; Sim and Back, 1985). However, much more research is needed to fully understand
and confirm these endocrinological processes.
1.4 Controlling Boar Taint
Surgical castration has been the traditional method used to control boar taint;
however economical and ethical drawbacks have prompted researchers to find feasible,
low-cost alternatives to castration that do not negatively affect other carcass
characteristics. Some of these alternatives include early slaughter, sperm sorting,
immunocastration, and genetic selection.
1.4.1 Early Slaughter
Early slaughter involves slaughtering boars at a younger age and at a lighter
weight before they have reached sexual maturity. Since androstenone is generally not
produced and accumulated in adipose tissue until puberty, early slaughter may reduce the
frequency of boar taint (Gunn et al., 2004). However, the correlation of fat androstenone
concentration and carcass weight is low and most studies have shown that slaughtering at
a lower weight does not entirely eliminate boar taint (Gunn et al., 2004; Zamaratskaia and
Squires, 2009). In addition, the carcasses of young pigs often do not have enough meat to
return a profit for producers. There are also welfare issues associated with the early
slaughter of boars. Entire males are more aggressive than castrates, particularly when
25
mixed with unfamiliar pigs on farms, during transport, and during pre-slaughter lairage,
which increases the risk for skin damage and leg injuries (Gunn et al., 2004; Giersing et
al., 2006). Further, the physical exertion associated with fighting may mobilise muscle
glycogen stores and lead to a higher frequency of pale, soft, exudative (PSE) meat which
has a lower economic value (Warriss and Brown, 1985; Gunn et al., 2004).
1.4.2 Sperm Sorting
Sperm sorting involves separating the X- and Y-chromosome bearing
spermatozoa using flow cytometry (Johnson et al., 2005). The X-chromosome bearing
spermatozoa may then be used in artificial insemination (AI) to produce predominately
female pigs for pork production. Since boar taint is not a problem in sows, sperm sorting
effectively reduces this issue (Gunn et al., 2004). In addition, the need for castration is
greatly reduced and animal welfare is potentially increased. However, current sorting
methods are not efficient enough to produce the high number of sperm needed for AI as
conventional techniques require approximately 3 billion sperm per dose of semen, and
the flow cytometry technique currently allows the sorting of only 10 to 15 million sperm
per hour (Johnson, 1991, 2000; Gunn et al., 2004). One procedure has been developed
that deposits spermatozoa deep into the uterine horn and reduces the number of sperm
required per dose (Martinez et al., 2001, 2002). However, the catheter used in this
technique may cause damage to the cervix and uterine mucosa, potentially compromising
subsequent fertility and reducing the welfare of the sows (Giersing et al., 2006; Vazquez
et al., 2008). In addition to welfare issues, the raising of only females for pork production
may not be economically advantageous to producers as female pigs have poorer feed
efficiency and produce less lean meat than intact boars (Thun et al., 2006).
26
1.4.3 Immunocastration
Immunocastration involves vaccination with Improvac against GnRH which
effectively castrates boars near the end of the grower-finisher phase of production
(Dunshea et al., 2001). Consequently, this vaccination inhibits the production of LH and
FSH from the pituitary, prevents testicular development, reduces the production of
testicular steroid hormones and accumulation of skatole, and decreases aggressive
behaviour (Dunshea et al., 2001; Metz et al., 2002; Cronin et al., 2003; Giersing et al.,
2006). Pigs are vaccinated twice, at least four weeks apart, with the second booster
injection given four to six weeks before slaughter. Prior to the second injection, pigs
retain the production benefits of entire males (Dunshea et al., 2001; Cronin et al., 2003;
Jaros et al., 2005). In addition, immunocastrates have been shown to grow faster to
slaughter weight and have higher feed conversion efficiency than surgical castrates
(Dunshea et al., 2001; Cronin et al., 2003; Zamaratskaia et al., 2008b). One US economic
model estimated that these benefits may result in an additional net income of $5.32 per
pig, after the cost and implementation of the vaccine (Buhr et al., 2013).
Although immunocastration can be used to reduce androstenone concentrations in
fat and decrease aggressive behaviour without negatively affecting performance, several
disadvantages exist. For example, handling of heavy pigs that need to be captured,
restrained, and injected twice is not practical for some pig producers (Einarsson, 2006;
Giersing et al., 2006). Assuming that no pigs are missed, some boars will still have fat
androstenone concentrations that are too high due to individual variation in
immunological response to the vaccine (Einarsson, 2006). Another disadvantage involves
the safety of humans. Since the Improvac vaccine is not species-specific, workers who
27
are accidentally self-injected may develop antibodies against GnRH and consequently
become sterile (Prunier et al., 2006; Squires, 2010); however, two injections are needed.
In addition, consumer acceptability of products from immunocastrated pigs may not be
favourable and needs to be further investigated (Lundström and Zamaratskaia, 2006;
Squires, 2010). Despite these drawbacks, Improvac has been approved for use in over 60
countries (Buhr et al., 2013).
1.4.4 Genetic Selection
Genetic selection involves identifying genetic factors that control boar taint and
implementing them in breeding programs to select pigs with low levels of androstenone
(Moe et al., 2009). Since there are large differences in androstenone concentrations
between breeds, with more Meishan, Duroc, and Piétrain boars showing higher
androstenone levels than Landrace, Large White, Hampshire, and Yorkshire boars, it was
suggested that there may be genetic differences in the tissue levels of androstenone (Xue
et al., 1996; Tajet et al., 2006; Zamaratskaia and Squires, 2009). In addition, the
heritability of fat androstenone is relatively high, ranging from 0.25 to 0.88, suggesting
that genetic selection for low boar taint is possible (Jonsson and Andresen, 1979; Willeke
et al., 1980; Sellier and Bonneau, 1988; Sellier et al., 2000). Previous attempts at
selection against high fat androstenone were successful, however they were also
associated with decreased growing performance in boars and delayed puberty in gilts due
to a lower production of androgens and estrogens (Bonneau, 1982, 1998; Willeke et al.,
1987; Sellier and Bonneau, 1988; Squires, 2006). Therefore, before starting selection,
animals that have a decreased genetic capacity to accumulate androstenone in fat while
maintaining normal levels of testicular steroids need to be identified using genetic
28
markers (Squires, 2006; Zamaratskaia and Squires, 2009). This would make it possible to
select intact boars with little or no boar taint that otherwise grow as normal boars.
Identification of quantitative trait loci (QLT) and investigation of SNPs in
candidate genes are two common approaches used for developing genetic markers
(Squires, 2006; Zamaratskaia and Squires, 2009). Research in this area has identified
multiple SNPs in the CYP members CYP2E1, CYP21, CYP2D6 and CYP2C49 that were
found to be significantly associated with fat androstenone but not testosterone or
estrogens (Moe et al., 2009). In addition, a SNP in CYB5 was found to be significantly
associated with lower fat androstenone concentrations (Lin et al., 2005). Further, Squires
et al. (2014) showed that numerous SNPs in Duroc, Landrace, and Yorkshire breeds were
significantly associated with androstenone and skatole levels. Therefore, these SNPs may
be used to reduce androstenone without negatively affecting phenotypes related to growth
and reproduction, however much more research is needed (Moe et al., 2009).
29
CHAPTER 2: HYPOTHESIS AND RESEARCH OBJECTIVES
The traditional method of surgical castration used to control boar taint has come
under scrutiny and economical, ethical, and legal issues are pushing for reliable
alternatives. In order to develop new and effective boar taint control methods, a complete
understanding and identification of the physiological mechanisms regulating fat
androstenone accumulation and the genes controlling these processes are needed. Since
androstenone accumulation is influenced by phase II metabolism, the focus of this
research will be on the biosynthesis of androstenone and other steroid hormones in the
liver and testis, with particular emphasis on sulfonate and glucuronide conjugates.
2.1 Hypothesis
It is generally thought that androstenone does not undergo sulfoconjugation since
it does not possess any hydroxyl groups; however, several researchers have provided
evidence suggesting otherwise. In addition, both porcine SULT2A1 and SULT2B1 have
been identified as important enzymes involved in steroid hormone sulfoconjugation.
Therefore, it is expected that both porcine Leydig cells and hepatocytes will
sulfoconjugate androstenone through the action of SULT2A1 and SULT2B1 enzymes.
2.2 Research Objectives
Objective 1: To investigate the phase I and phase II metabolism of androstenone in both
Leydig cells and hepatocytes.
Previous research has suggested that androstenone, 3α-androstenol, and 3βandrostenol undergo sulfoconjugation, but not glucuronidation, in porcine Leydig cells;
30
however these results were achieved indirectly (Sinclair et al., 2005a; Desnoyer, 2011).
In addition, 3β-androstenol was shown to be glucuronidated by porcine hepatocytes while
studies investigating the sulfoconjugation capabilities of these cells have been conflicting
(Sinclair et al., 2005b; Chen et al., 2015). Therefore, the conjugation abilities of Leydig
cells and hepatocytes towards androstenone and its metabolites were investigated.
Objective 2: To further characterize the roles of porcine SULT2A1 and SULT2B1 in
androstenone sulfoconjugation.
SULT2A1 has been identified as an important enzyme involved in the
sulfoconjugation of 16-androstene steroids and its activity has been negatively correlated
with adipose androstenone concentration (Sinclair et al., 2006). Similarly, decreased
expression of SULT2B1 in porcine liver and testes has been associated with high
androstenone concentrations (Moe et al., 2007a). Therefore, the sulfoconjugation abilities
of porcine SULT2A1 and SULT2B1 towards androstenone were further investigated.
Objective 3: To compare the sulfonation activity of human and porcine sulfotransferases
toward androstenone and other steroid hormones.
Pigs share many similar anatomical and physiological characteristics with humans
making them a valuable experimental model for multiple research applications (Patterson
et al., 2008; Swindle et al., 2012). Since many sulfotransferases are known to conjugate
drugs and xenobiotics, similarities between human and porcine enzymes may further
justify the increased use of pigs in research, particularly in preclinical toxicological
testing of pharmaceuticals (Swindle et al., 2012).
31
CHAPTER 3: METABOLISM OF ANDROSTENONE IN PRIMARY CULTURED
PORCINE LEYDIG CELLS AND HEPATOCYTES
3.1 Abstract
Increased public interest in the welfare of pigs reared for pork production has led
to an increased effort in finding alternatives to castration for controlling the unpleasant
odour and flavour from heated pork products known as boar taint. The purpose of this
study was to investigate the testicular and hepatic metabolism of androstenone in the
boar. Leydig cells and hepatocytes were isolated from five mature Yorkshire boars and
incubated with radiolabelled androstenone for 10 min, 1h, 4h, 8h, and 12h. Entire steroid
profiles were analyzed by high-performance liquid chromatography (HPLC) and liquid
chromatography-mass spectrometry (LC-MS). Sulfoconjugated steroids were produced
by the Leydig cells but not hepatocytes, while glucuronides were produced by
hepatocytes but not Leydig cells. Approximately 85% of androstenone was converted
into sulfoconjugate metabolites in Leydig cell incubations after 8 hours. This
sulfoconjugate fraction included androstenol sulfonate and 3-keto-sulfoxy androstenone.
These findings provided direct evidence of the testicular production of a sulfoconjugated
form of androstenone in the boar. In addition, the high proportion of sulfoconjugates
produced by the Leydig cells emphasizes the importance of steroid conjugation and
supports the theory that these steroids could act as a reservoir which serves to regulate the
amount of unconjugated steroid hormones available for accumulation in adipose tissue.
Keywords: androstenone, boar taint, glucuronidation, sulfoconjugation
32
3.2 Introduction
In the swine industry, male piglets are surgically castrated within the first week of
life in order to reduce aggression and eliminate the unpleasant odour and flavour from
heated pork products known as boar taint. However, increasing welfare concerns are
putting pressure on pork producers to find alternatives to castration. Before new methods
can be developed, factors controlling and contributing to this meat quality defect have to
be well understood.
5α-androst-16-en-3-one (androstenone), the primary compound responsible for
boar taint (Patterson, 1968), is a 16-androstene steroid that is produced in the Leydig cells
of the testes and readily accumulates in adipose tissue (Desmoulin et al., 1982; Claus et
al., 1994). Since androstenone is produced together with testicular steroids that stimulate
growth and fertility, the challenge is to reduce the level of androstenone without affecting
the other testicular steroids (Gower, 1972; Moe et al., 2007b). This may be accomplished
by either selectively reducing the rate of androstenone biosynthesis or increasing the rate
of its metabolism (Doran et al., 2004). Androstenone metabolism occurs primarily in the
kidney, liver, and testes through phase I and phase II reactions (Hobkirk, 1985; Sinclair,
2004). The phase I enzymes, 3α- and 3β-hydroxysteroid dehydrogenase (HSD), reduce
androstenone to 3α-androstenol or 3β-androstenol, respectively (Doran et al., 2004;
Sinclair et al., 2005a,b). These compounds may then be conjugated by the action of phase
II enzymes, such as sulfotransferases (SULTs) or uridine diphosphate
glucuronosyltransferases (UGTs), which result in the generation of metabolites that are
more water-soluble and thus more likely to be excreted via the urine and bile (Falany,
1997).
33
Previous research has indirectly shown that porcine Leydig cells produce a
relatively large amount of steroid sulfoconjugates, including androstenone sulfonate, but
do not produce steroid glucuronides (Sinclair and Squires, 2005; Sinclair et al., 2005a;
Desnoyer, 2011). The sulfoconjugation of androstenone likely occurs through prior
enolisation of the 3-keto group (Sinclair et al., 2005a,b, 2006). Previous studies have also
shown that porcine hepatocytes produce steroid glucuronides; however there is
conflicting data regarding the ability of these cells to produce sulfoconjugated steroids
(Sinclair et al., 2005b; Chen et al., 2015).
One of the problems with steroid metabolism studies is a lack of appropriate
analytical methods (Chen et al., 2015). Radioimmunoassays (RIAs), for example, can
give erroneous results if there is antibody cross-reactivity and antibodies may not exist
for all steroid metabolites of interest (Soldin and Soldin, 2009; Stanczyk and Clarke,
2010). In addition, gas chromatography-mass spectrometry (GC-MS) requires tedious
extraction and derivatization procedures for each sample and thus steroid conjugates
cannot be analyzed directly (Penning et al., 2010). The use of mass spectrometry with
electrospray ionization (ESI) can now avoid many of these problems (Penning et al.,
2010; Gouveia et al., 2013). In addition, mass spectrometry with a quadrupole time-offlight (Q-TOF) mass analyzer provides a relatively high sensitivity and accuracy for mass
detection and has a virtually unlimited mass range (Gouveia et al., 2013).
The purpose of this study was to investigate the metabolism of androstenone in
primary cultured porcine Leydig cells and hepatocytes using appropriate analytical
techniques that minimized sample processing.
34
3.3 Materials and methods
3.3.1 Reagents
Radiolabelled [3H]-androstenone (10 Ci/mmol) was obtained from Moravek
Biochemicals (Brea, CA, USA). Nonradioactive steroids were purchased from Steraloids
Inc. (Newport, RI, USA). Organic solvents of analytical grade were obtained from Fisher
Scientific (Toronto, ON, Canada). All other chemicals used for the isolation and
incubation of Leydig cells and hepatocytes were of analytical grade and were purchased
from Sigma–Aldrich Canada, Ltd. (Oakville, ON, Canada).
3.3.2 Research Animals
A total of five mature intact Yorkshire boars weighing between 120 and 190kg
were obtained from the Arkell Swine Research Facility at the University of Guelph and
were used in accordance with the guidelines of the Canadian Council of Animal Care and
the University of Guelph Animal Care Policy. Boars were electrically stunned and
exsanguinated, followed by immediate surgical excision of the testes and liver for further
processing.
3.3.3 Leydig cell isolation
Primary Leydig cells were isolated as previously described (Sinclair et al., 2005a),
with modifications. Briefly, testes were rinsed with cold water, decapsulated, and sliced
into pieces approximately 1 cm2 × 1 mm thick. Tissue (120 – 150 g) was digested at 37°C
for 45 – 60 minutes by constant stirring in 500 ml of culture medium (TC 199 without
phenol red) containing 0.1% L-glutamine, 2.2% sodium bicarbonate, 1% bovine serum
35
albumin (BSA), 1% glucose, 1% collagenase, 0.05% DNase, and 0.05% trypsin inhibitor.
Following digestion, tissue homogenate was passed through nylon mesh (155 µm and 80
µm) and the filtrate was centrifuged at 400 × g for 10 minutes at 4°C. The cell pellet was
then resuspended in approximately 80 ml of media and layered onto discontinuous
Percoll gradients consisting of 21, 26, 34, 40, and 60% interfaces, as previously described
(Sinclair et al., 2005a). Cells were counted using a hemocytometer and cell viability was
determined with a trypan blue exclusion test. Typical Leydig cell viability was 90%.
3.3.4 Hepatocyte isolation
Primary hepatocytes were isolated as previously described (Sinclair et al., 2005b)
with modifications. Briefly, the largest vein in one liver lobe was catheterized and
perfused with Hanks’ balanced salt solution, pH 7.4, (without Ca2+, Mg2+, HCO3--, and
phenol red) containing 1 mM ethylene glycol tetraacetic acid (EGTA) and 10mM
hydroxy ethyl piperazine ethane sulfonic acid (HEPES) for approximately 10 minutes
and then with Hanks’ balanced salt solution, pH 7.4, containing 10 mM HEPES for
approximately 5 minutes at 37°C with a flow rate of 25 ml/minute. The blanched lobe
was then digested with Williams’ Medium E, pH 7.4, containing 10 mM HEPES and 0.5
mg/ml Type I collagenase for approximately 30 minutes.
After digestion, the hepatic lobe was dissected under sterile conditions to liberate
the hepatocytes from the collagen matrix. Dissociated hepatocytes were collected in
Williams’ Medium E, pH 7.4, supplemented with 10 mM HEPES, 0.02 units/ml insulin,
1% penicillin/streptomycin, and 10% fetal bovine serum (FBS). The cells were then
filtered through sterile gauze and centrifuged at 15 × g for three minutes at room
36
temperature. Cells were rinsed by resuspension and centrifugation in collection medium
and the cell pellet was resuspended in collection medium. Cells were counted using a
hemocytometer and cell viability was determined with a trypan blue exclusion test.
Typical hepatocyte viability was 85%.
3.3.5 Biosynthesis studies
Resuspended Leydig cells and hepatocytes were inoculated at a concentration of
approximately 2.0 × 106 cells/ml in Erlenmeyer flasks containing 6 – 12 ml of the culture
mediums described above. Cells were then incubated with radiolabelled [3H]androstenone (20 µM, 17.9 µCi/µmol) dissolved in ethanol for 12 hours at 37ºC under
5% CO2 and 95% air in a shaking water bath with 100 oscillations/minute. Aliquots of 2
– 4 ml were removed in triplicate at 10 min, 1h, 4h, 8h, and 12h, transferred to pre-cooled
test tubes, and centrifuged at 400 × g for five minutes at 4ºC. Supernatants were frozen at
-20º until further use. Medium containing [3H]-androstenone that had not been inoculated
with cells was used as a control.
3.3.6 High-Performance Liquid Chromatography (HPLC)
Thawed supernatants from cell incubations were diluted 1:1 (v/v) with acetonitrile
and centrifuged for 10 minutes at room temperature to precipitate protein. The
supernatant was analyzed for steroids using reversed phase HPLC by injecting a 100µl
aliquot onto a Luna 5µ C18(2) HPLC column (250 × 4.60 mm) (Phenomenex, Torrance,
CA, USA). Elution of radiolabelled steroids was monitored by a β-RAM model 2 isotope
detector (IN/US Systems, Tampa, FL, USA). The HPLC mobile phase, with a flow rate
37
of 1.0 ml/minute, consisted of an isocratic flow with acetonitrile/water (33:67, v/v) for 8
minutes, a linear gradient from 33 – 60% acetonitrile for the next 17 minutes, an isocratic
flow with 100% acetonitrile for the next 5 minutes, and an isocratic flow with
acetonitrile/water (33:67, v/v) for the remaining 10 minutes. In this system, free steroids
eluted between 36 and 38 minutes and conjugated steroids eluted starting at
approximately 20 minutes. The identities of free and conjugated steroids were further
characterized by manually collecting entire peak fractions from the HPLC followed by
further processing as outlined below.
3.3.7 Steroid Conjugate Analyses
Collected fractions were dried using a Savant Speed Vac centrifugal vacuum
concentrator and conjugated steroids present in the fraction were deconjugated as
previously described (Raeside et al., 1999), with modifications. Briefly, dried fractions
were incubated overnight with 5 ml of trifluoroacetic acid/ethyl acetate (1/100, v/v) at
45ºC. This process chemically removes sulfonate groups from steroids, if present. To
remove glucuronide groups, separate fractions were incubated overnight with 1.0 ml of
0.5 M sodium acetate buffer, pH 5.0, containing 2500 units/ml of β-glucuronidase (type
B-1, from bovine liver) at 37ºC. Steroids treated with β-glucuronidase were extracted
twice with 5 ml of diethyl ether and supernatants were pooled. Samples from both
treatments were dried under nitrogen at 45ºC, reconstituted with 1.0 ml of 85%
acetonitrile, filtered with GHP Acrodisc syringe filters (0.2 μm) (Pall Corp., Ville St.
Laurent, Quebec, Canada), and analyzed with HPLC using the same gradient system as
above.
38
3.3.8 Mass Spectrometry
Liquid chromatography-mass spectrometry was performed on entire supernatant
samples from 8 hour cell incubations that contained only unlabelled androstenone and
which were diluted 1:1 (v/v) with acetonitrile, as above. Steroids were analyzed on an
Agilent 1200 HPLC liquid chromatograph interfaced with an Agilent UHD 6530 Q-Tof
mass spectrometer at the Mass Spectrometry Facility of the Advanced Analysis Center at
the University of Guelph. A 50 µl sample volume was injected onto a Poroshell 120 2.7µ
C18 column (150 × 4.6 mm) (Agilent Technologies, Mississauga, ON, Canada). The
HPLC mobile phase system consisted of an isocratic flow of 0.1% formic
acid/acetonitrile with 0.1% formic acid (98:2, v/v) for 1 minute, a linear gradient from 2 –
100% acetonitrile with 0.1% formic acid for the next 19 minutes, an isocratic flow with
100% acetonitrile with 0.1% formic acid for 3.5 minutes, followed by re-equilibration for
the final 10 minutes. The first 2 and last 5 minutes of gradient were sent to waste and not
the spectrometer. The flow rate was maintained at 0.4 ml/minute. The mass spectrometer
electrospray capillary voltage was maintained at 4.0 kV and the drying gas temperature
was 250ºC with a flow rate of 8 L/minute. Nebulizer pressure was 30 psi and the
fragmentor was set to 160. Nitrogen was used as a nebulizing, drying, and collisioninduced gas. The mass-to-charge ratio was scanned across the range of 50 – 1400 m/z in
4GHz positive or negative-ion auto MS/MS mode. Two precursor ions per cycle were
selected for fragmentation. The instrument was externally calibrated with ESI TuneMix
(Agilent Technologies). Chromatograms were analyzed with Agilent Qualitative Analysis
B.06.00 software defining compounds by the Molecular Feature algorithm and generating
possible compound formulas including elements C, H, O, N, S, and Cl.
39
3.3.9 Data Analysis
Statistical analysis was carried out using a Student’s t-test using Microsoft Excel
(2013). The statistical significance level was defined as P < 0.05. All data were expressed
as means ± standard error.
3.4 Results
Characteristic HPLC chromatographs of conjugated and unconjugated peaks in
media from Leydig cells and hepatocytes incubated with [3H]-androstenone are shown in
Figure 3.1. Three major peaks from Leydig cell incubations and two major peaks from
hepatocytes were observed. The peak with a retention time (RT) of approximately 36
minutes, found in incubations from both cell types, was identified as unconjugated
androstenone based on a comparison with authentic steroid standard and confirmed via
LC-MS with an m/z- of 271.2 (data not shown). Authentic steroid standards of 3αandrostenol and 3β-androstenol eluted at approximately the same time as androstenone
when analyzed by HPLC, suggesting that the peak observed at 36 minutes contained a
mixture of unconjugated steroids. The presence of these steroids produced by both
Leydig cells and hepatocytes was also confirmed by LC-MS; however, since these
steroids have an identical mass (m/z- = 273.2), α- and β- linkages were not differentiated.
The steroid peak with an RT of 20.25 minutes found in incubations with Leydig cells, but
not hepatocytes (Figure 3.1A,C), corresponded to the RT of 5α-androstenone following
fraction collection and solvolysis treatment (Figure 3.1B). No change in retention time
was observed with this fraction following treatment with β-glucuronidase. Therefore, this
peak was thought to represent at least one sulfoconjugated steroid. The third major peak
observed (RT 2.7 minutes) was present in Leydig cell and hepatocyte incubations.
40
Figure 3.1 Characteristic HPLC profiles of conjugated and unconjugated steroids
from 8 hour incubations of radiolabelled [3H]-androstenone with cells from the
testis and liver. RT for a reference standard was approximately 36-37 minutes. (A)
Leydig cells; (B) Leydig cells following fraction collection and solvolysis of the peak
with an RT of 20.25 minutes; (C) hepatocytes.
41
However, there was no change in RT following either solvolysis or β-glucuronidase
treatment. LC-MS analysis of this peak showed a number of polymer-related compounds.
None of these compounds had mass-to-charge ratios that corresponded to those expected
for androstenone metabolites. In addition, since this peak was also seen in control
samples, it was thought to be an artifact rather than a product of androstenone
metabolism, although its production over time by Leydig cells was significantly higher (p
< 0.05) when compared to control samples (Figure 3.3A).
Further analysis of conjugated androstenone metabolites from Leydig cells via
LC-MS carried out in negative ionization mode identified an ion at an RT of 16.09
minutes with a mass-to-charge ratio (m/z) of 353.18 (Figure 3.2A). This is identical to the
predicted m/z of androstenol sulfate. The ionized sulfate group can also be seen with m/z96.96. A second ion with an RT of 18.09 minutes and m/z- 367.16 was also identified,
which has an identical m/z to a sulfonated form of androstenone previously reported as 3keto-4-sulfoxy-androstenone (Desnoyer, 2011) (Figure 3.2B). Therefore, it was
concluded that the HPLC peak identified at RT 20.25 minutes contained both androstenol
sulfate and a sulfonated form of androstenone. Steroid glucuronides were not found in
incubations with Leydig cells. LC-MS analysis of hepatocyte incubations was carried out
in positive ionization mode and an ion at RT 12.78 minutes with m/z+ 450.17 was
identified (Figure 3.2C). This m/z corresponds to that of androstenol glucuronide,
although this peak was not observed during HPLC analysis. Steroid sulfonates were not
found in hepatocyte incubations.
Time course data for the production of metabolites by Leydig cell and hepatocyte
incubations with radiolabelled androstenone are shown in Figure 3.3 and Figure 3.4,
42
respectively. Relative peak area, representing relative radioactivity and steroid content,
was significantly different (p < 0.05) in Leydig cell incubations when compared with
control samples for all three major HPLC peaks. The peak representing the artifact
increased from 52.1% ± 3.6 after 10 minutes in Leydig cell incubations to a maximum of
83.5% ± 2.0 after 4 hours. This was similar to data from hepatocyte incubations, which
showed a relative peak area of 85.9% ± 3.5 after four hours; however this was not
significantly different (p > 0.05) when compared to the control sample. When considering
only the sulfoconjugated and unconjugated fractions produced from Leydig cell
incubations, peak area was greatest for unconjugated steroids (98.9% ± 0.8), with a small
amount present in the conjugated form (1.1% ± 0.8) after 10 minutes (Figure 3.5). The
peak area percentage of unconjugated steroids decreased to 79.5% ± 6.9 after 1 hour,
followed by a decrease to 28.9% ± 12.9 after 4 hours, and to 14.3% ± 8.1 after 8 hours.
At 12 hours, the unconjugated fraction increased to 23.7% ± 13.2, however this was not
statistically significant (p > 0.05). Corresponding absolute peak areas, representing
absolute radioactivity and steroid content, for Leydig cell and hepatocyte incubations are
shown in Table 3.1. Peak area was not significantly different (p > 0.05) for peak 1 (RT
2.7 min), identified as an artifact, between Leydig cell and hepatocyte incubations at any
incubation times, with the exception of the 12 hour time point. For peak 3 (RT 36.9),
significantly higher (p < 0.05) peak areas were observed with hepatocyte incubations
when compared to Leydig cells for 0.17, 1, 4, and 8 hour incubation periods. This
difference was not due to the production of sulfoconjugated metabolites by Leydig cells
as the peak areas from hepatocyte incubations were still significantly higher when
absolute areas of peak 2 and 3 were combined for Leydig cells (data not shown).
43
Figure 3.2 Mass spectra of conjugated metabolites of androstenone produced by
the testes and liver. Samples from Leydig cells were analyzed in negative ionization
mode and those from hepatocytes were analyzed in positive ionization mode. (A)
androstenol sulfonate (m/z- = 353.1785) with ionized sulfate group (m/z- = 96.9598)
produced by Leydig cells; (B) 3-keto-sulfoxy androstenone (m/z- = 367.1600)
produced by Leydig cells; (C) androstenol glucuronide (m/z+ = 450.1687) produced by
hepatocytes.
44
A
Relative Peak Area (%)
B
C
Time (hours)
Figure 3.3 Relative peak areas, as determined by HPLC analysis, of
metabolites produced from incubations of radiolabelled [3H]-androstenone
with Leydig cells over time. (A) identified artifact (RT 2.7 min); (B)
sulfoconjugated fraction (RT 20.2 min); (C) unconjugated androstenone and
androstenols (RT 36.9 min); (●) Leydig cell sample; (■) control sample. Values
are plotted as mean ± standard error for five independent experiments performed in
triplicate (n = 5). Significant differences compared to the control (p < 0.05) were
evaluated using a Student’s t-test and are denoted by (*).
45
Relative Peak Area (%)
A
B
Time (hours)
Figure 3.4 Relative peak areas, as determined by HPLC analysis, of metabolites
produced from incubations of radiolabelled [3H]-androstenone with hepatocytes
over time. (A) identified artifact (RT 2.47 min); (B) unconjugated androstenone and
androstenols (RT 37.05 min); (●) hepatocyte sample; (■) control sample. Values are
plotted as mean ± standard error for five independent experiments performed in
triplicate (n = 5). Significant differences compared to the control (p < 0.05) were
evaluated using a Student’s t-test and are denoted by (*).
46
Relative Peak Area (%)
A
B
Time (hours)
Figure 3.5 Relative peak areas, as determined by HPLC analysis, of metabolites
produced from incubations of radiolabelled [3H]-androstenone with Leydig cells
over time when considering only sulfoconjugated and unconjugated steroids. (A)
identified sulfoconjugated fraction (RT 20.2 min); (B) unconjugated androstenone and
androstenols (RT 36.9 min); (●) Leydig cell sample; (■) control sample. Values are
plotted as mean ± standard error for five independent experiments performed in
triplicate (n = 5). Significant differences compared to the control (p < 0.05) were
evaluated using a Student’s t-test and are denoted by (*).
47
48
Leydig Control
76.04 ± 4.032
86.19 ± 5.061
102.1 ± 3.085
110.3 ± 7.892
141.5 ± 7.031
0.2382 ± 0.09832
0.2495 ± 0.1046
0.3734 ± 0.1574
0.6177 ± 0.1954
0.4008 ± 0.1758
391.9 ± 44.68
357.6 ± 30.30
287.4 ± 54.17
226.4 ± 35.43
284.3 ± 50.78
Leydig cells
76.13 ± 7.107
128.7 ± 16.18
229.1 ± 20.44
246.5 ± 7.929
236.0 ± 16.41 *
0.4543 ± 0.1583
11.08 ± 2.662
39.48 ± 6.076
78.64 ± 13.76
51.10 ± 16.44
75.30 ± 4.027 *
51.12 ± 7.909 *
13.12 ± 6.061 *
5.889 ± 2.350 *
13.36 ± 6.553
110.1 ± 6.883 *
78.58 ± 7.643 *
34.46 ± 8.484 *
25.33 ± 5.727 *
11.14 ± 3.920
ND
ND
ND
ND
ND
77.14 ± 4.361
139.9 ± 19.36
228.5 ± 30.95
273.6 ± 22.82
297.8 ± 28.30 *
Hepatocytes
300.7 ± 9.749
247.3 ± 37.40
134.5 ± 53.33
39.47 ± 20.16
21.68 ± 8.503
ND
ND
ND
ND
ND
63.78 ± 0.9153
59.27 ± 4.515
65.95 ± 2.728
76.28 ± 11.20
127.0 ± 26.20
Hepatocyte Control
Note. The data presented in this table represent means ± standard error for five independent experiments performed in
triplicate (n = 5). ND, no detectable metabolite. Significant differences between Leydig cells and hepatocytes (p < 0.05)
were evaluated using a Student’s t-test and are denoted by (*).
Time (hours)
Peak 1 (RT 2.7 min)
0.17
1
4
8
12
Peak 2 (RT 20.2 min)
0.17
1
4
8
12
Peak 3 (RT 36.9 min)
0.17
1
4
8
12
Mean Peak Area (arbitrary units × 104)
Table 3.1 Absolute mean peak areas of metabolites produced by Leydig cells and hepatocytes
3.5 Discussion
In the present study, HPLC and LC-MS were used to investigate the metabolism
of androstenone in porcine Leydig cells and hepatocytes as in vitro models. The results
showed that the Leydig cells produced steroid sulfonates, but not glucuronides, whereas
the hepatocytes produced androstenol glucuronide, but no sulfonates. The production of
16-androstene glucuronides by hepatocytes, but not Leydig cells, has been shown in
previous studies (Sinclair et al., 2005a; Chen et al., 2015) and corresponds to the theory
that glucuronidation serves primarily as a mechanism to facilitate metabolic clearance
(Sinclair et al., 2005b). This is further supported by the high concentrations of androgen,
estrogen, and androstene glucuronides found in the urine and bile, as well as the high
levels of UGT enzymes found in the liver and kidney (Gower and Patterson, 1970;
Sinclair et al., 2005b). The observed androstenol glucuronide in the present study was
only detected by LC-MS analysis and not by HPLC. This is presumably due to
differences in levels of detection between these two techniques and suggests that
glucuronidation activity was relatively low in hepatocytes compared to sulfoconjugation
activity in Leydig cells. This is further supported by the differences observed between
Leydig cells and hepatocytes when comparing absolute peak areas. The combined total
peak area, and thus radioactivity, for metabolites produced from hepatocyte incubations
was significantly higher compared to that of Leydig cells after 0.17, 1, 4, and 8 hour
incubation periods. One explanation is that the Leydig cells may have taken up and
retained more of the radiolabelled substrate compared to hepatocytes which may reflect
the activity/viability of these cells. However, radioactivity was not measured in the cell
pellets.
49
Previous studies have also shown the production of steroid sulfonates by Leydig
cells (Saat et al., 1972, 1974; Ruokonen and Vihko, 1974a,b; Booth, 1975; Sinclair et al.,
2005a); however results from porcine hepatocytes have been conflicting. Sinclair et al.
(2005b) reported that approximately 25% of androstenone was converted into
androstenone, 3α-androstenol, and 3β-androstenol sulfonates in hepatocyte incubations
following deconjugation, derivatization, and GC-MS analysis of steroid metabolites. In
contrast, Chen et al. (2015) did not detect any steroid sulfonates in porcine hepatocytes
incubated with androstenone, which agrees with results from the present study. These
authors suggested that sulfoconjugation may not have occurred because in vitro cell
culture systems have lower sulfate concentrations when compared with in vivo systems.
Alternatively, any number of problems could have occurred during hepatocyte isolation
which may have led to cell contamination or cell lysis. This may also further explain why
androstenol glucuronide was found by LC-MS but not HPLC analysis.
Steroid sulfonates produced by Leydig cells included androstenol sulfonate and a
sulfonated form of androstenone. Time course data for these sulfonates reached a
maximum production level after 8 hours of incubation with approximately 85.7% of
androstenone converted to sulfonated steroid metabolites. This high proportion of
sulfoconjugated steroids is similar to that reported in previous studies using porcine
Leydig cells incubated with androstenone (Sinclair et al., 2005a), as well as 16androstene, androgen, and estrogen steroid sulfonate levels reported in porcine plasma
(Saat et al., 1974; Raeside and Renaud, 1983; Sinclair and Squires, 2005). In the present
study, α- and β- linkages were not differentiated for the androstenol sulfonate identified,
however, previous studies have consistently shown β-androstenol sulfonate as the
50
dominant metabolite in porcine Leydig cells, hepatocytes, and plasma (Saat et al., 1974;
Sinclair et al., 2005a,b; Sinclair and Squires, 2005). Therefore, the metabolite identified
as androstenol sulfonate most likely consists of a relatively high ratio of β-androstenol
sulfonate to α-androstenol sulfonate.
Previously, it was thought that since androstenone did not contain any hydroxyl
groups, it could not undergo sulfonation; however hydroxylated forms of androstenone
have been produced by axillary microflora and porcine hepatocytes (Austin and Ellis,
2003; Sinclair et al., 2005b; Chen et al., 2007b). In addition, a large proportion of
androstenone sulfonate has been found in the plasma of boars, although these results were
obtained indirectly (Sinclair and Squires, 2005). Androstenone is thought to be
sulfoconjugated by prior enolisation of the 3-keto group (Sinclair et al., 2005a,b, 2006)
which involves the removal of a hydrogen atom from a carbon that is replaced on an
oxygen atom to form a hydroxyl group (Toullec, 1982). The exchange of a proton can
occur between either positions 2 and 3 or positions 3 and 4; however, it has generally
been thought that the direction of enolisation of 3-keto-5α-steroids is towards position 2
(Berkoz et al., 1962; Gardi et al., 1962). This newly formed hydroxyl group can then be
acted upon by sulfotransferase enzymes to produce 3-enol androstenone sulfonate (Figure
3.6). However, this metabolite, with an expected mass of 352.5, was not observed in the
present study. Instead, a metabolite with m/z- 367.2 was found which corresponds to a
compound identified as 3-keto-4-sulfoxy androstenone in a similar study (Desnoyer,
2011). In addition, the present study showed that this steroid conjugate possessed a
sulfate group with m/z- 96.96. The m/z of a sulfate ion can vary depending on where it
was attached on the parent compound and this feature has previously been used to
51
Androstenone
Formula: C19H28O
Mass: 272.214
ENO
3-enol
Formula: C19H28O
Mass: 272.214
Hydroxy-androstenone
Formula: C19H28O2
Mass: 288.209
SULT
SULT
3-enol sulfonate
Formula: C19H28O4S
Mass: 352.171
3-keto-sulfoxy androstenone
Formula: C19H28O5S
Mass: 368.166
Figure 3.6 The production of 3-keto-sulfoxy androstenone and 3-enol sulfonate from
androstenone. ENO, enolase; SULT, sulfotransferase (adapted from Sinclair et al.,
2005b; Desnoyer, 2011).
52
identify the structures of sulfated metabolites (Yi et al., 2006). For example, an ion with
m/z [M-H-80]- represents a neutral loss of SO3 and indicates that the metabolite was
produced by the sulfoconjugation of a phenol or an enol with the sulfate attached to an
sp2 carbon. This characteristic ion would have been expected if 3-enol androstenone
sulfonate was observed in the present study. An ion with m/z 97, which was observed in
the current study, represents the formation of a bisulfate anion (HSO4−) and indicates that
the sulfate group was attached to an sp3 carbon and the sulfated product produced by the
sulfoconjugation of an aliphatic alcohol (Yi et al. 2006). Therefore, the sulfated steroid
observed in the current study with m/z- 367.2 would have been the result of the addition
of a hydroxyl group onto androstenone at a position other than C3 followed by
sulfoconjugation of this group to produce 3-keto-sulfoxy androstenone (Figure 3.6).
Hydroxylated forms of androstenone have been shown to be produced by porcine
hepatocytes (Sinclair et al., 2005b). In addition, Austin and Ellis (2003) identified 6hydroxy androstenone as a metabolite produced by axillary microflora. Hydroxylation of
other steroids has also been shown to occur at the C6 position (Zimniak et al., 1991;
Miyata et al., 1994); however, there are multiple sites where this reaction can occur due
to the various enzymes that may be involved (Waxman, 1988; Martucci and Fishman,
1993; Gilep et al., 2011; Robic et al., 2014). Therefore, the position of the added
hydroxyl group, and consequently the sulfate group, remains unknown. Unfortunately,
many steroid conjugates are not yet commercially available as pure standards for
chromatography, making identification of compounds from steroid metabolism difficult;
however, the use of other analytical techniques such as nuclear magnetic resonance
(NMR) spectroscopy has been previously used to determine the position of
53
sulfoconjugation in other compounds (Schaber et al., 2001; Ibrahim et al., 2003; Lerch et
al., 2003) and may be a useful technique for future studies.
Like glucuronidation, sulfoconjugation has typically been regarded as a
mechanism to facilitate metabolic clearance (Sinclair, 2004; Rasmussen et al., 2012b).
However, the presence of SULTs in the testis and other extrahepatic tissues (Roberts and
Lieberman, 1970; Gasparini et al., 1976; Hobkirk, 1985; Falany et al., 1995), as well as
the relatively high proportion of 3-keto-sulfoxy-androstenone produced by the Leydig
cells in the current study, indicates otherwise. Since sulfoconjugated steroids can be
deconjugated via the action of membrane bound sulfatase enzymes (Falany, 1997; Glatt,
2000), sulfoconjugation may regulate the availability of active steroid hormones by
creating a reservoir from which unconjugated steroids could be regenerated by target
tissues (Hobkirk, 1993; Glatt and Meinl, 2004). In addition, sulfoconjugation reactions
generate metabolites that are more water-soluble and thus less likely to accumulate in
adipose tissue (Falany, 1997). Therefore, the testicular production of sulfoconjugated
androstenone has been suggested to impact the level of unconjugated androstenone that is
available to accumulate in fat, which ultimately plays a role in determining whether or
not a boar will develop boar taint (Sinclair et al., 2005a).
It is important to note the techniques used for steroid conjugate analysis in the
present study. Steroids are typically isolated using liquid/solid, liquid/liquid, or column
extraction procedures prior to HPLC and MS analysis (Corpéchot et al., 1981; Guazzo et
al., 1996; Rizea Savu et al., 1996; Bean and Henion, 1997; Chatman et al., 1999; Raeside
et al., 1999). In addition, steroids may be derivatized in order to increase the sensitivity of
detection (Higashi and Shimada, 2004; Penning et al., 2010). These methods are time-
54
consuming and result in indirect and potentially inaccurate data. In the present study,
media samples from Leydig cell and hepatocyte incubations were precipitated with
acetonitrile and entire samples were successfully analyzed using both HPLC and LC-MS.
When steroids were analyzed by LC-MS, sulfoconjugated steroids tended to ionize better
in negative mode, while glucuronidated steroids ionized better in positive mode.
Therefore, a novel method for the analysis of sulfoconjugated and glucuronidated steroids
is proposed in the current study. The ability to analyze an entire steroid profile from one
sample is particularly relevant for studies that have a limited amount of sample, as well as
those that require a rapid turnaround time (Soldin and Soldin, 2009).
In summary, this study used effective HPLC and LC-MS techniques to investigate
the metabolism of androstenone in isolated porcine Leydig cells and hepatocytes. This
study provided new data on pig Leydig cell steroid metabolism. In particular, direct
evidence was provided, for the first time, for the production of a sulfoconjugated form of
androstenone by porcine Leydig cells. This metabolite was suggested to be 3-ketosulfoxy-androstenone, which may be formed via prior hydroxylation of androstenone.
The study also investigated androstenone metabolite production over time and
determined an optimal incubation time of 8 hours for the production of sulfoconjugated
steroids. Future studies should focus on investigating the mechanism responsible for
androstenone sulfoconjugation as well as confirming the structural form of androstenone
sulfonate.
55
CHAPTER 4: THE SULFOCONJUGATION OF ANDROSTENONE AND
DEHYDROEPIANDROSTERONE BY HUMAN AND PORCINE SULT2A1 AND
SULT2B1 ENZYMES
4.1 Abstract
Porcine sulfotransferase 2A1 (pSULT2A1) has previously been identified as a key
enzyme involved in the testicular and hepatic metabolism of androstenone. pSULT2B1
may also be important, although no direct evidence exists for its involvement. The
purpose of this study was to investigate the sulfoconjugation activity of pSULT2B1 and
to further characterize and compare human and porcine sulfotransferase enzymes.
pcDNA 3.1 vectors expressing pSULT2A1, pSULT2B1, human (h) SULT2A1,
hSULT2B1a, and hSULT2B1b enzymes were transfected into human embryonic kidney
(HEK) cells. Transfected cells were then incubated with either androstenone or
dehydroepiandrosterone (DHEA) in both time-course and enzyme kinetics studies. The
production of androstenone sulfonate and DHEA sulfonate increased over time for all
enzymes with the exception of pSULT2B1. Enzyme kinetics analysis showed that
androstenone and DHEA were poor substrates for the human orthologs, hSULT2B1a and
hSULT2B1b. Comparisons between human and porcine SULT2A1 showed substantially
different substrate affinities for androstenone (Km 5.8 ± 0.6 µM and 74.1 ± 15.9 µM,
respectively) and DHEA (Km 9.4 ± 2.5 µM and 3.3 ± 1.9 µM, respectively). However,
these enzymes did show relatively similar sulfonation efficiencies for DHEA (Vmax/Km
50.5 and 72.9 for hSULT2A1 and pSULT2A1, respectively). The findings in this study
provide direct evidence, for the first time, suggesting that pSULT2B1 is not involved
with androstenone metabolism and does not sulfonate DHEA. In addition, these results
56
suggest that data obtained from studies utilizing the pig as a human model be interpreted
with caution, particularly in preclinical toxicological testing of pharmaceuticals where
many sulfotransferase enzymes are required to activate drugs and where species
differences may lead to significantly different responses in certain experiments.
Keywords: boar taint, enzyme kinetics, human, pig, sulfotransferase
4.2 Introduction
Sulfoconjugation, or sulfonation, plays an important role in the biotransformation
of hormones, neurotransmitters, and xenobiotic compounds (Falany, 1997;
Weinshilboum et al., 1997). These reactions generate metabolites that are more watersoluble and thus more easily excreted in the urine and bile (Falany, 1997). In the boar,
this is a particularly relevant metabolic process, as the sulfoconjugation of 16-androstene
steroids will influence the occurrence of boar taint, an unpleasant odour and flavour from
heated pork products. Androstenone, the primary compound responsible for this meat
quality defect, as well its metabolites 3α-androstenol and 3β-androstenol, have been
shown to occur largely in their sulfoconjugated form, reaching levels of up to 72%
relative to their unconjugated form in the peripheral and testicular vein plasma of mature
boars (Sinclair and Squires, 2005).
Sulfonation of steroids involves the transfer of a sulfo group (SO3-) from a donor
molecule to the hydroxyl group of an acceptor molecule to form a sulfonate conjugate
(Falany et al., 1995; Strott, 2002). The donor molecule required for sulfonation is 3’phosphoadenosine 5’-phosphosulfate (PAPS), and all mammalian tissues are able to carry
out its synthesis (Robbins et al., 1956; Falany et al., 1995). Sulfonation reactions are
57
mediated by a family of enzymes termed sulfotransferases (SULTs) which are found in
many tissues including liver, kidney, testis, ovary, intestinal tract, lung, and brain
(Roberts and Lieberman, 1970; Gasparini et al., 1976; Hobkirk, 1985; Falany et al.,
1995). To date, six distinct cytosolic SULT gene families have been identified in
mammals; however only the SULT1 and SULT2 families are capable of sulfonating
steroids (Strott, 2002; Blanchard et al., 2004; Neunzig et al., 2014). Porcine (p)
SULT2A1, in particular, has been identified as a key enzyme involved in the testicular
and hepatic metabolism of androstenone (Sinclair et al., 2005b, 2006). This enzyme is
capable of conjugating a number of different steroids although it is most reactive towards
dehydroepiandrosterone (DHEA), an important precursor steroid in androgen and
estrogen synthesis (Falany et al., 1989, 2006). pSULT2B1 has also been suggested to
play a role in androstenone metabolism as it has been negatively correlated to fat
androstenone concentration and has shown selectivity for 3β-hydroxysteroids (Moe et al.,
2007a,b; Meloche and Falany, 2001). However, direct evidence for the involvement of
this enzyme has yet to be generated. In pigs, only one SULT2B1 enzyme exists; however
in humans this gene encodes two isoforms, SULT2B1a and SULT2B1b, which are
generated by alternate splicing of the first exon (Her et al., 1998; Meloche and Falany,
2001; Kohjitani et al., 2006). These two isoforms have been shown to preferentially
sulfonate pregnenolone and cholesterol, while DHEA was observed to be a poor substrate
(Javitt et al., 2001; Fuda et al., 2002).
Pigs share many similar anatomical and physiological characteristics with humans
making them a valuable experimental model for multiple research applications (Patterson
et al., 2008; Swindle et al., 2012). Consequently, similarities between human and porcine
enzymes may further justify the increased use of pigs in research, particularly in
58
preclinical toxicological testing of pharmaceuticals, as many SULTs are known to
conjugate drugs and in some cases, are required to activate them (Weinshilboum and
Otterness, 1994; Swindle et al., 2012).
The purpose of this study was to further characterize and compare the
sulfoconjugation activity of human and porcine sulfotransferase enzymes towards
androstenone and DHEA.
4.3 Materials and methods
4.3.1 Materials
Human embryonic kidney cells (HEK293) were purchased from ATCC
(Manassas, VA, USA). The pMK-RQ vector containing cDNA encoding porcine
SULT2B1 was custom synthesized by Life Technologies Inc. (Burlington, ON, Canada).
The pKK233-2 vector containing cDNA encoding for human SULT2A1 was kindly
provided by Dr. Charles Falany (University of Alabama at Birmingham, Birmingham,
AL, USA) while pCR3.1 vectors containing cDNA encoding for human SULT2B1a and
human SULT2B1b were generously donated by Dr. Richard Weinshilboum (Mayo
Clinic, Rochester, MN, USA). Radiolabelled [3H]-androstenone (10 Ci/mmol) was
obtained from Moravek Biochemicals (Brea, CA, USA) and radiolabelled [1,2,6,73
H(N)]-DHEA (70.5 Ci/mmol) was purchased from Perkin-Elmer (Boston, MA, USA).
Nonradioactive steroids were purchased from Steraloids Inc. (Newport, RI). Organic
solvents of analytical grade were obtained from Fisher Scientific (Toronto, ON, Canada).
All other chemicals used for the incubation of HEK293 cells were of analytical grade and
were purchased from Sigma–Aldrich Canada, Ltd. (Oakville, ON, Canada) or Fisher
Scientific, unless specified otherwise.
59
4.3.2 Plasmid Constructs
The pSULT2A1 coding sequence was amplified from porcine liver cDNA and the
pSULT2B1, hSULT2B1a, and hSULT2B1b coding sequences were amplified from their
respective vectors by polymerase chain reaction (PCR) using platinum Taq DNA
polymerase and gene-specific primers (Table 4.1). These primers were designed based on
sequences available from the National Center for Biotechnology Information (NCBI).
The PCR procedure consisted of heating at 94°C for 2 minutes, 34 cycles of 94°C for 30
seconds, 60°C for 30 seconds, and 68°C for 1 minute, and finally incubating at 68°C for
10 minutes. Amplified DNA fragments containing appropriate start codon and Kozak
sequence (GCC) were gel-purified using a PureLink quick gel extraction kit (Invitrogen)
and were subcloned into mammalian expression vector pcDNA3.1/V5-His-TOPO using a
TA cloning kit (Invitrogen). Expression vectors for V5-His tagged proteins were
generated in order to detect protein expression by Western blot using anti-V5-horseradish
peroxidase (HRP) antibody (Billen and Squires, 2009). Sequences of all constructs were
confirmed by DNA sequence analysis.
Table 4.1 Gene-specific primers used for PCR of human and porcine SULTs
Gene
Forward Primer (5’→3’)
Reverse Primer (5’→3’)
Accession Number
pSULT2A1
GCCATGACAGAAGAGGAGG
TTGCCATGGGAACAGCTCTT
NM 001037150.1
pSULT2B1
GCCATGGATGGGCCTGCCG
CGGGTGGGGGACCTCG
NM 001243697.1
hSULT2A1
GCCATGTCGGACGATTTCT
TTCCCATGGGAACAGC
NM 003167.3
hSULT2B1a
GCCATGGCGTCTCCCCCAC
TGAGGGTCGTGGGTGC
NM 004605.2
hSULT2B1b
GCCATGGACGGGCCCGCCG
TGAGGGTCGTGGGTGC
NM 177973.1
60
4.3.3 Cell Culture Transfection and Enzyme Expression
HEK293 cells were routinely maintained as monolayer cultures in 75 cm2 cell
culture flasks (Greiner Bio-One North America Inc., Monroe, NC, USA) and grown in
complete growth medium (Dulbecco’s modified eagle’s medium (DMEM) supplemented
with 10% fetal bovine serum, 1% L-glutamine, 1% penicillin/streptomycin, 1% nonessential amino acids, 1% sodium pyruvate, and 1% geneticin) in a humidified
atmosphere at 37ºC and 5% CO2 in air. Cells were plated in triplicate at 7 × 105 cells per
well in 6-well cell culture plates (Greiner Bio-One) with 2 ml/well of complete growth
medium and allowed 24 hours for adherence. Vectors expressing sulfotransferases were
then transfected into cells using Nano Juice core transfection reagent (1:2, w/v of
DNA:reagent) and Nano Juice transfection booster (1:3.2, w/v of DNA:reagent)
(Novagen, Etobicoke, ON, Canada) with 0.08 µg pSULT2A1, 1.25 µg pSULT2B1, 1.25
µg hSULT2A1, 0.045 µg hSULT2B1a, and 0.045 µg hSULT2B1b. Vector amounts were
varied in order to obtain similar levels of protein expression assessed by Western blotting
without compromising activity. Empty pcDNA 3.1 vector was added, when appropriate,
so that the total plasmid DNA level was 1.25 µg/well. Empty pcDNA 3.1 vector was also
used to transfect control samples. After 48 hours, transfected cells were treated with
radiolabelled [3H]-androstenone (20 µM, 31.2 µCi/µmol) and [1,2,6,7-3H(N)]-DHEA (20
µM, 15.5 µCi/µmol) dissolved in ethanol for 4, 8, 16, and 24 hours in a time-course
study. Kinetic parameters were also obtained in three separate experiments by incubating
cells for 24 hours with varying concentrations of androstenone or DHEA prepared by
serial dilution. Media was subsequently removed and diluted 1:1 (v/v) with acetonitrile,
centrifuged for 10 minutes at room temperature to precipitate protein, and analyzed by
HPLC. Cells were harvested and lysed by sonication in 300 µl of radio61
immunoprecipitation assay (RIPA) buffer (0.5% sodium deoxycholate, 1% Nonnidet P40, and 0.1% SDS) with 1% protease inhibitor cocktail (EMD Millipore Corp., San
Diego, CA, USA). Protein concentrations were determined using the Bradford (Bio-Rad)
protein assay (Bradford, 1976).
4.3.4 HPLC
Media samples diluted with acetonitrile were analyzed using reversed phase
HPLC by injecting a 100 µl aliquot onto a Luna 5µ C18(2) HPLC column (250 × 4.60
mm) from Phenomenex (Torrance, CA, USA). Elution of radiolabelled steroids was
monitored by a β-RAM model 2 isotope detector (IN/US Systems, Tampa, FL, USA).
The HPLC mobile phase system for samples incubated with radiolabelled [3H]androstenone consisted of an isocratic flow with acetonitrile/water (33:67, v/v) for 8
minutes, a linear gradient from 33 – 60% acetonitrile for the next 17 minutes, an isocratic
flow with 100% acetonitrile for the next 5 minutes, and an isocratic flow with
acetonitrile/water (33:67, v/v) for the remaining 10 minutes. In this system, free steroids
eluted between 36 and 38 minutes and sulfoconjugated steroids eluted starting at
approximately 20 minutes. Conjugated steroids were further characterized by manually
collecting entire peak fractions from the HPLC system followed by further processing as
outlined below. The HPLC mobile phase gradient for samples incubated with labelled
[1,2,6,7-3H(N)]-DHEA consisted of 50% acetonitrile for 15 minutes, 100% acetonitrile
for the next 5 minutes, and 85% acetonitrile for the last five minutes. In this system,
DHEA eluted at approximately 11 minutes and DHEAS at 2.5 minutes, which was
confirmed using authentic steroid standards. Both gradients were delivered at a flow rate
of 1.0 ml/minute.
62
4.3.5 Steroid Conjugate Analyses
Collected fractions were dried using a Savant Speed Vac centrifugal vacuum
concentrator and any conjugated steroids present in the fraction were deconjugated as
previously described (Raeside et al., 1999), with modifications. Briefly, dried fractions
were incubated overnight with 5 ml of trifluoroacetic acid/ethyl acetate (1/100, v/v) at
45ºC. This process chemically removes sulfonate groups from steroids, if present. To
remove glucuronide groups, separate fractions were incubated overnight with 1.0 ml of
0.5 M sodium acetate buffer, pH 5.0, containing 2500 units/ml of β-glucuronidase (type
B-1, from bovine liver) at 37º. Steroids treated with β-glucuronidase were extracted twice
with 5 ml of diethyl ether and supernatants were pooled. Samples from both treatments
were dried under nitrogen at 45ºC, reconstituted with 1.0 ml of 85% acetonitrile, filtered
with GHP Acrodisc syringe filters (0.2 μm) (Pall Corp., Ville St. Laurent, Quebec,
Canada), and analyzed by HPLC using the same gradient systems as above.
4.3.6 Western Blot Analysis
Protein expression of HEK293 cells transfected with pSULT2A1, pSULT2B1,
hSULT2A1, hSULT2B1a, and hSULT2B1b were analyzed by Western blot as previously
described (Billen and Squires, 2009), with modifications. Briefly, 20 µg protein samples
were separated on 12% sodium dodecyl sulfate – polyacrylamide gel by electrophoresis
(SDS-PAGE) and transferred to polyvinylidene fluoride (PVDF) membranes (EMD
Millipore) by semi-dry transblot transfer. Blots were incubated overnight at 4ºC in 5%
(w/v) dried skim milk in phosphate buffered saline (PBS) with 0.1% Tween 20 and then
incubated with a 1:16000 dilution of primary anti-V5 antibody followed by a 1:8000
63
dilution of secondary goat-anti-mouse-HRP antibody in the 5% skim milk solution, as
above. Protein bands were visualized and quantified by incubating with detection solution
(50 µl of 68 mM p-coumaric acid in dimethyl sulfoxide (DMSO), 5 ml of 1.25 mM
luminol in 0.1 M Tris, pH 8.5, and 15 µl of 3% hydrogen peroxide) for 1 minute and
using a ChemiDoc MP imaging system (Bio-Rad) with Image Lab 5.0 software. Enzyme
activity was normalized to the protein content of expressed sulfotransferase enzymes
based on immunoblot band density.
4.3.7 Data Analysis
Multiple amino acid sequence alignment analysis was carried out using MAFFT
Version 7 software. Statistical analysis was carried out using a Student’s t-test using
Microsoft Excel (2013) and the statistical significance level was defined as p < 0.05.
Enzyme kinetics analysis was carried out using SigmaPlot 12.0 and Microsoft Excel with
the Solver supplement. If no substrate inhibition was observed, data were fitted to the
Michaelis-Menten equation (1) and the Hill equation (2):
𝑣=
𝑉𝑚𝑎𝑥 ⋅ [𝑆]
𝐾𝑚 + [𝑆]
(1)
𝑣=
𝑉𝑚𝑎𝑥 ⋅ [𝑆]𝑛
𝐾𝑚 𝑛 + [𝑆]𝑛
(2)
where v is the rate of substrate sulfoconjugation, Vmax is the apparent maximum rate of
substrate sulfoconjugation, S is the substrate incubation concentration, Km is the substrate
concentration at which the reaction rate is half of Vmax, and n describes binding
cooperativity. The appropriate kinetic model was determined by examining EadieHofstee plots. Where a hook-shaped Eadie-Hofstee plot was observed, indicative of
autoactivation, data were fitted to the Hill equation, and where a linear Eadie-Hofstee
plot was observed, data were fitted to the Michaelis-Menten equation (Houston and
64
Kenworthy, 2000; Hutzler and Tracy, 2002). When substrate inhibition was observed,
kinetic parameters were determined as previously described (Zhang et al., 1998; Geese
and Raftogianis, 2001). Briefly, Vmax and Km were estimated at low concentrations, as
above. The entire data set was then fitted to a partial substrate inhibition equation
(Fromm, 1975):
𝑉 ⋅𝑆
1+𝑉 2 ⋅𝐾
𝑚𝑎𝑥
𝑖
𝑣 = 𝑉𝑚𝑎𝑥 ⋅ (
)
𝐾𝑚 𝑆
1+
+
𝑆
𝐾𝑖
(3)
where v, Vmax, S, and Km are defined as above, V2 is the estimated rate plateau that is
reached when inhibition occurs, and Ki is the substrate concentration at which the Vmax is
decreased by half. Vmax and Km were then determined over substrate concentrations much
less than Ki, where substrate inhibition was negligible. Vmax/Km ratios, representing
enzyme catalytic efficiencies, were also calculated. All data were expressed as mean ±
standard error.
4.4 Results
The amino acid sequence alignments of human and porcine SULTs are shown in
Figure 4.1 and Figure 4.2. Both human and porcine proteins contain the classical PAPS
binding domain that is nearly identical among SULTs (Dong et al., 2012; Falany and
Rohn-Glowacki, 2013). They also contain the highly conserved ‘KTVE’ dimerization
domain. The pSULT2A1 and hSULT2A1 cDNAs both contain open reading frames of
858 base pairs (bp) which encode for 285 amino acids. These proteins share an identity of
66% (Table 4.2) and a similarity of 81%. The pSULT2B1, hSULT2B1a, and
hSULT2B1b cDNAs contain open reading frames of 1017, 1053, and 1098 bp which
65
hSULT2A1 MSDDFLWFEG IAFPTMGFRS ETLRKVRDEF VIRDEDVIIL TYPKSGTNWL
pSULT2A1 -TEEEVR--- -F--KQILSP -M-QE--E-- TFKE---L-- -F-------M
50
50
hSULT2A1 AEILCLMHSK GDAKWIQSVP IWERSPWVES EIGYTALSET ESPRLFSSHL
pSULT2A1 I--I--IL-- --T------- N-D----L-- IS--EN-KNK -G---I----
100
100
hSULT2A1 PIQLFPKSFF SSKAKVIYLM RNPRDVLVSG YFFWKNMKFI KKPKSWEEYF
pSULT2A1 -------A-- K----M--II -----II--- -----STNLV –R-E-L-Q--
150
150
hSULT2A1 EWFCQGTVLY GSWFDHIHGW MPMREEKNFL LLSYEELKQD TGRTIEKICQ
pSULT2A1 ---I--N-P- -------R-- L---DKE-V- I-------R- -RSAV-----
200
200
hSULT2A1 FLGKTLEPEE LNLILKNSSF QSMKENKMSN YSLLSVDYVV DKAQLLRKGV
pSULT2A1 ----K----- -SSVVE---- -V----N--- F---KGLHLG -TGC-----T
250
250
hSULT2A1 SGDWKNHFTV AQAEDFDKLF QEKMADLPRE LFPWE
pSULT2A1 P-----Y--- ----A----- --------Q- ----Q
285
285
Figure 4.1 Amino acid sequence alignment of SULT2A1 enzymes from human and
pig species. The amino acid sequences were aligned using MAFFT with hSULT2A1 as
reference. Only dissimilar residues are displayed. The conserved PAPS binding domain is
doubly underlined, the conserved catalytic residue histidine is highlighted black, loop 3
which makes up the outer surface of both the PAPS and substrate binding pockets is
singly underlined, and the conserved ‘KTVE’ dimerization domain is highlighted gray
(adapted from Falany and Rohn-Glowacki, 2013).
encode for 338, 350, and 365 amino acids, respectively. The pSULT2B1 protein shares
identities of 72 and 76%, with hSULT2B1a and hSULT2B1b, respectively, and is 84%
similar to both enzymes. Both of the human SULT2B1 isoforms contain an extended
identical 50 amino acid carboxy-terminal sequence that is enriched in proline and serine
residues and is not found in other human SULTs (Falany and Rohn-Glowacki, 2013).
pSULT2B1 also has an extended carboxy-terminal sequence that is not found in
pSULT2A1, although it contains 25 amino acid residues and is only 40% identical to
hSULT2B1a and hSULT2B1b. The amino-termini of hSULT2B1a, hSULT2B1b, and
pSULT2B1 are also extended and contain 8, 23, and 23 amino acid residues, respectively.
66
hSULT2B1a MASPPPFH-- ---------- ---SQKLPGE YFRYKGVPFP VGLYSLESIS 35
hSULT2B1b -DG-AEPQIP GLWDTYEDDI SEI------- ---------- ---------- 50
pSULT2B1
-DG-AEPRNQ AEWDPYEKNI SEI--N-S-- ------I--- --V--P---- 50
hSULT2B1a LAENTQDVRD DDIFIITYPK SGTTWMIEII CLILKEGDPS WIRSVPIWER 85
hSULT2B1b ---------- ---------- ---------- ---------- ---------- 100
pSULT2B1
I---AK*-Q- ---------- ---N------ S----D---- --Q-----K- 99
hSULT2B1a APWCETIVGA FSLPDQYSPR LMSSHLPIQI FTKAFFSSKA KVIYMGRNPR 135
hSULT2B1b ---------- ---------- ---------- ---------- ---------- 150
pSULT2B1
S----A-M-- ----N-P--- -----P---L ------N--- ----L----- 149
hSULT2B1a DVVVSLYHYS KIAGQLKDPG TPDQFLRDFL KGEVQFGSWF DHIKGWLRMK 185
hSULT2B1b ---------- ---------- ---------- ---------- ---------- 200
pSULT2B1
--L------- ---------- ------QN-- ---------- ------I--R 199
hSULT2B1a GKDNFLFITY EELQQDLQGS VERICGFLGR PLGKEALGSV VAHSTFSAMK 235
hSULT2B1b ---------- ---------- ---------- ---------- ---------- 250
pSULT2B1
--E------- -------HS- -Q-V-Q---- -------D-- ----A-N--- 249
hSULT2B1a ANTMSNYTLL PPSLLDHRRG AFLRKGVCGD WKNHFTVAQS EAFDRAYRKQ 285
hSULT2B1b ---------- ---------- ---------- ---------- ---------- 300
pSULT2B1
--A---F--- -T----Q--- ------I--- ---------- ----SV--E- 299
hSULT2B1a MRGMPTFPWD EDPEEDGSPD PEPSPEPEPK PSLEPNTSLE REPRPNSSPS 335
hSULT2B1b ---------- ---------- ---------- ---------- ---------- 350
pSULT2B1
---L------ -T-*--AR-- -D---D**** ********** ********-- 326
hSULT2B1a PSPGQASETP HPRPS
hSULT2B1b ---------- ----pSULT2B1
--------V- --***
350
365
338
Figure 4.2 Amino acid sequence alignment of SULT2B1 enzymes from human and
pig species. The amino acid sequences were aligned using MAFFT with hSULT2B1a as
reference. Only dissimilar residues are displayed. The conserved PAPS binding domain is
doubly underlined, the conserved catalytic residue histidine is highlighted black, loop 3
which makes up the outer surface of both the PAPS and substrate binding pockets is
singly underlined, the conserved ‘KTVE’ dimerization domain is highlighted gray, and
gaps are indicated by asterisks (*) (adapted from Falany and Rohn-Glowacki, 2013).
67
Table 4.2 Sequence identities of human and porcine SULT amino acid sequences
pSULT2A1
pSULT2B1
hSULT2A1
pSULT2A1
─
pSULT2B1
45
─
hSULT2A1
66
41
─
hSULT2B1a
41
72
39
hSULT2B1a
─
hSULT2B1b
39
76
37
94
Note. Values are presented as percent identity for each individual pair of SULTs
Interestingly, the Ile20 and Ile23 residues found in the amino-terminal of hSULT2B1b,
which are responsible for the high cholesterol sulfoconjugating activity of this enzyme
(Fuda et al., 2002), are also conserved in the pSULT2B1 sequence.
Vectors containing cDNA encoding SULT proteins were over-expressed in HEK
cells in order to determine relative sulfoconjugation activities over time and to
characterize and compare the kinetic properties between human and porcine SULTs.
Although vector concentrations were varied in order to obtain similar levels of protein
expression, hSULT2B1a and hSULT2B1b protein were still expressed approximately 3
to 9 times more when compared to the other SULTs (Figure 4.3). Therefore, protein
expression was normalized to a standard sample via Western blot analysis.
Characteristic HPLC chromatographs of steroid metabolites produced from
incubations with androstenone and DHEA are shown in Figure 4.4. Unconjugated
androstenone eluted at approximately 36.4 minutes which was confirmed by a
comparison to authentic steroid standard. A sulfoconjugated fraction occurred at
approximately 20.9 minutes which was identified by fraction collection and solvolysis
treatment. Following this treatment, the collected fraction eluted from the HPLC column
at approximately the same time as unconjugated androstenone. A third peak was
68
observed at approximately 2.8 minutes however there was no change in RT following
either solvolysis or β-glucuronidase treatment. Since this peak was also seen in control
samples, and since the production of this peak was not significantly different (p > 0.05)
when compared to controls for all enzymes, it was thought to be an artifact rather than a
product of androstenone metabolism. A fourth peak was also observed at approximately
4.7 minutes, however its appearance was not consistent. In addition, fraction collection
and either solvolysis or β-glucuronidase treatment did not result in an RT change,
suggesting that this peak may have been an artifact as well. DHEA had a retention time
of approximately 11.2 minutes while DHEAS eluted at approximately 2.7 minutes. This
was confirmed by HPLC analysis of authentic steroid standards.
Time course data for all SULTs describing androstenone and DHEA
sulfoconjugation are shown in Figure 4.5. The production of androstenone sulfonate
increased over time for all enzymes with the exception of pSULT2B1. Conversion was
less than 0.3% across all time points for this enzyme and was only significantly different
(p < 0.05) when compared to control samples at 8h. Conversion of androstenone reached
a maximum of 12.3% ± 1.8 after 24 hours by hSULT2A1 activity. Similar results were
found for incubations with DHEA where the production of DHEAS increased over time
for all enzymes, again with the exception of pSULT2B1. Conversion was less than 0.8%
for cells transfected with pSULT2B1 expression vector across all time points but was
significantly different (p < 0.05) when compared to control samples at 4h, 16h, and 24h
incubation periods. The maximum conversion of DHEA was 9.8% ± 0.5 after 24 hours
by hSULT2A1 activity. Since pSULT2B1 activity did not show increased
sulfoconjugation production over time with either substrate, it was not included in
enzyme kinetics analyses. In addition, initial rate enzyme kinetics were carried out using
69
an incubation of 24 hours, as the majority of enzymes had less than 10% conversion of
substrate.
Enzyme kinetics data obtained from incubations with androstenone as substrate
were fit to either the Michaelis-Menten equation or the Hill equation (see Materials and
Methods) (Figure 4.6 and Figure 4.7), while those obtained using DHEA as substrate
showed substrate inhibition and were fit to an equation that describes partial substrate
inhibition (Figure 4.8 and Figure 4.9). Corresponding kinetic parameters are shown in
Table 4.3. Comparisons between human and porcine SULT2A1 showed substantially
different substrate affinities for both androstenone and DHEA. However, these enzymes
did show relatively similar sulfonation efficiencies for DHEA. Interestingly, the
sulfonation efficiency for hSULT2A1 was approximately 5-fold higher than for
pSULT2A1 when using androstenone as substrate. Vmax/Km ratios for hSULT2B1a and
hSULT2B1b were significantly smaller than those of the other enzymes for both
substrates suggesting that androstenone and DHEA are poor substrates for these human
orthologs.
Figure 4.3 Typical Western blot of human and porcine SULT proteins. Lanes
are as follows: 1, protein marker; 2, pSULT2A1; 3, pSULT2B1; 4, hSULT2A1; 5,
hSULT2B1a; 6, hSULT2B1b.
70
Figure 4.4 Characteristic HPLC profiles of conjugated and unconjugated steroids
from HEK cells expressing SULT enzymes and incubated with radiolabelled [3H ]androstenone (A) and [1,2,6,7-3H(N)]-DHEA (B). RTs for reference standards of
androstenone and DHEA were approximately 36-37 and 11-12 minutes.
71
A
B
Figure 4.5 Relative peak areas, as determined by HPLC analysis, of androstenone
sulfonate (A) and DHEAS (B) production over time for human and porcine SULTs.
Values are plotted as mean ± standard error for two independent experiments performed
in triplicate (n = 2). Significant differences compared to the control (p < 0.05) were
evaluated using a Student’s t-test and are denoted by (*).
72
A
B
Figure 4.6 The production of androstenone sulfonate by porcine SULT2A1 (A) and
human SULT2A1 (B) with increasing androstenone concentrations. Values are
plotted as mean ± standard error for three independent experiments performed in
triplicate (n = 3).
73
A
B
Figure 4.7 The production of androstenone sulfonate by human SULT2B1a (A) and
human SULT2B1b (B) with increasing androstenone concentrations. Values are
plotted as mean ± standard error for three independent experiments performed in
triplicate (n = 3).
74
A
B
Figure 4.8 The production of DHEAS by porcine SULT2A1 (A) and human
SULT2A1 (B) with increasing androstenone concentrations. Values are plotted as
mean ± standard error for three independent experiments performed in triplicate (n = 3).
75
A
B
Figure 4.9 The production of DHEAS by human SULT2B1a (A) and human
SULT2B1b (B) with increasing androstenone concentrations. Values are plotted as
mean ± standard error for three independent experiments performed in triplicate (n = 3).
76
Table 4.3 Kinetic constants for porcine and human SULTs
Vmax (pmol/hr/mg protein)
Km (µM)
Vmax/Km
Ki (µM)
pSULT2A1
689.7 ± 92.2
74.1 ± 15.9
9.3
ND
hSULT2A1
276.4 ± 52.5
5.8 ± 0.6
47.7
ND
hSULT2B1a
22.5 ± 5.0
26.8 ± 13.2
0.8
ND
hSULT2B1b
29.3 ± 6.5
46.9 ± 17.9
0.6
ND
pSULT2A1
240.4 ± 8.2
3.3 ± 1.9
72.9
23
hSULT2A1
475.1 ± 15.3
9.4 ± 2.5
50.5
11
hSULT2B1a
18.8 ± 1.1
3.6 ± 3.0
5.2
29
hSULT2B1b
13.7 ± 0.8
2.4 ± 4.6
5.7
16
Androstenone
DHEA
Note. The kinetic constants reported in this table were derived from plots in Figure 4.6 to
Figure 4.9. The data represent means ± standard errors for three independent experiments
performed in triplicate (n = 3). ND, no detectable substrate inhibition.
4.5 Discussion
In the present study, HEK cells were overexpressed with SULT proteins in order
to determine which SULT plays a role in androstenone sulfoconjugation. The
sulfoconjugation activities of porcine and human SULTs were also further characterized
and compared using DHEA as a substrate. Previous studies have identified pSULT2A1 as
a key enzyme involved in the sulfoconjugation of steroids (Sinclair and Squires, 2005;
Sinclair et al., 2006), which is supported by results found in the present study. Porcine
SULT2B1 was also suggested to be important, as hepatic expression was negatively
correlated to androstenone concentration in adipose tissue (Moe et al., 2007a). However,
results from the present study provide direct evidence suggesting that pSULT2B1 is not
involved in the sulfoconjugation of androstenone or DHEA.
Authentic steroid standards of DHEA and DHEAS were commercially available
77
and thus used to confirm and quantify sulfoconjugate production by HPLC. However, an
authentic steroid standard of androstenone sulfonate is not yet commercially available.
Therefore, fraction collection from HPLC and solvolysis treatment were used to identify
sulfoconjugate metabolites. The sulfoconjugated metabolite peak that eluted at
approximately 20.9 minutes in the current study was previously shown to include
androstenol sulfonate and 3-keto-sulfoxy androstenone in incubations with porcine
Leydig cells (Chapter 3). These metabolites were identified by solvolysis treatment as
well as by LC-MS analysis. In the current study, LC-MS was not used for metabolite
analysis so it is unclear whether the sulfoconjugated peak was only androstenol sulfonate,
only 3-keto-sulfoxy androstenone, a mixture of both, or another form of sulfoconjugated
androstenone. Previous studies have shown that expression and activity of 3α- and 3βhydroxysteroid dehydrogenase in HEK cells is minimal (Zhang et al., 2000; Mason et al.,
2004; Udhane et al., 2013), and since androstenone is first metabolized by these enzymes
to produce 3α-androstenol and 3β-androstenol prior to sulfoconjugation, the production
of androstenol sulfonate in the current study is unlikely. However, more studies are
needed to confirm this and to identify the sulfoconjugated compound observed.
Reported calculations of Vmax and Km are usually difficult to compare due to
differing assay conditions, source and purity of the enzyme, and kinetic assumptions
made about substrate inhibition (Liu et al., 2006). However, since sulfonation reactions
for both human and porcine SULTs were carried out under identical assay conditions, a
direct and straightforward comparison can be made here. Both hSULT2B1a and
hSULT2B1b isoforms were not efficient at sulfoconjugating androstenone or DHEA
when compared to pSULT2A1 and hSULT2A1. Previous studies have shown that
hSULT2A1 preferentially sulfonated DHEA but not cholesterol, whereas hSULT2B1b
78
preferentially sulfonated cholesterol, but not DHEA (Javitt et al., 2001; Fuda et al.,
2002). In addition, hSULT2B1b, but not hSULT2A1, was selectively expressed in skin
where cholesterol sulfonation plays an important role in skin barrier development (Javitt
et al., 2001). This suggests that SULTs have selective physiological roles (Strott, 2002).
Since the Ile20 and Ile23 residues present in the amino-terminal of hSULT2B1b were
found to be essential for cholesterol sulfoconjugation (Fuda et al., 2002) and are
conserved in pSULT2B1, and since the full-length cDNA sequence for this latter enzyme
was also generated from skin (Uenishi et al., 2007), it is suggested that pSULT2B1 may
have a specific physiological role and act in a similar manner as hSULT2B1b. However,
the sulfoconjugation activities of pSULT2B1 and hSULT2B1b towards androstenone and
DHEA were noticeably different.
One explanation for the low activity of pSULT2B1 may be related to protein
stability issues. In humans, the native form of hSULT2B1b was shown to be expressed in
E. coli using different expression vectors but was not catalytically active following lysis
of the cells for protein isolation (Meloche and Falany, 2001; Falany and Rohn-Glowacki,
2013). The addition of a histidine (His)-tag allowed for the recovery of the active
enzyme; however removal of the His-tag resulted in an unstable enzyme that was
inactivated by freeze-thawing (Meloche and Falany, 2001; Falany, 2005). Consequently,
it was suggested that the added His-tag stabilized the active form of hSULT2B1b or
allowed for more efficient enzyme folding (Falany et al., 2006; Falany and RohnGlowacki, 2013). Although vectors containing the V5-His-tag were used in the present
study, the pSULT2B1 protein may not be stabilized by this tag due to species differences
in amino acid sequences. Alternatively, the low activity of pSULT2B1 may be normal
79
and this enzyme may simply not sulfoconjugate androstenone or DHEA to any significant
degree. Future studies should therefore examine other possible substrates for this enzyme.
In the present study, hSULT2A1 had a significantly higher binding affinity for
androstenone, but a lower Vmax, and was approximately 5-fold more efficient at
sulfoconjugating this substrate when compared to pSULT2A1. Androstenone has largely
been investigated in pigs in relation to boar taint; however much less is known about its
role in humans. In plasma, androstenone concentrations have been reported to be twice as
high in boars compared with humans (Smals and Weusten, 1991). Androstenone has also
been found in the axillary region in humans (Austin and Ellis, 2003) and has been
suggested to act as a chemosignal that mediates social behaviour and influences mood
and physiology (reviewed by Havlicek et al., 2010). This is similar to the pheromonal
role of androstenone in pigs (Andresen, 1976, 2006). Therefore, results from the current
study warrant further investigation into the significance of androstenone sulfoconjugation
in humans.
Many SULTs are known to conjugate drugs, and in some cases are required to
activate them (Weinshilboum and Otterness, 1994). Since pigs share many similar
anatomical and physiological characteristics with humans, similarities between human
and porcine enzymes may further justify the increased use of pigs in research, particularly
in preclinical toxicological testing of pharmaceuticals (Swindle et al., 2012). However,
care should be taken when interpreting data from these investigations, as orthologous
SULT forms in pigs and humans may have substantially different sulfoconjugation
activities. In the present study, although pSULT2A1 and hSULT2A1 had relatively
similar sulfoconjugation efficiencies for DHEA, pSULT2A1 had an almost 3-fold higher
80
binding affinity for this substrate when compared to hSULT2A1. In addition, previous
studies have shown pronounced differences in substrate specificities toward promutagens
between human and mouse SULT1B1 enzymes as well as between human and rat
SULT2A1 orthologs (reviewed by Glatt, 2000).
Many sulfotransferases are often inhibited by their own substrates (Reed et al.,
2010; Dong et al., 2012), which was demonstrated in the present study with DHEA but
not androstenone. DHEA substrate inhibition has also been shown in previous studies
with hSULT2A1, hSULT2B1a, and hSULT2B1b (Falany, 1997; Geese and Raftogianis,
2001; Chang et al., 2004; Liu et al., 2006). Two major potential mechanisms for substrate
inhibition in SULTs have been proposed (reviewed by Wu, 2011; Dong et al., 2012). In
the first, substrate inhibition may occur due to the presence of both an inhibitory binding
site and an active primary binding site within the enzyme. When substrate concentration
is high, a second DHEA molecule can bind to the inhibitory binding site and affect the
sulfonate transfer efficiency (Figure 4.10) (Gamage et al., 2003; Berger et al., 2011). In
the second mechanism, substrate inhibition may occur via formation of a ternary deadend enzyme complex [SULT•PAP•DHEA] which is non-productive and reaction
inhibitory (Wu, 2011). In many cases, substrate inhibition is thought of as an abnormality
SULT
∙PAPS
Km
SULT∙PAPS
∙DHEA
𝑉𝑚𝑎𝑥
SULT+DHEAS+PAP
Ki
∙PAPS
DHEA∙SULT∙DHEA
V2
SULT+DHEA+DHEAS+PAP
Figure 4.10 Kinetic model for partial substrate inhibition by DHEA
81
that results from using artificially high substrate concentrations in a laboratory setting
(Reed et al., 2010); however several studies have shown that it may be a biologically
relevant regulatory mechanism. The substrate inhibition of DHEA, in particular, has been
suggested to play an important role in steroid homeostasis (Chang et al., 2004).
In summary, this study used a transfected overexpression system in order to
further characterize and compare human and porcine sulfotransferase enzymes. Direct
evidence was provided, for the first time, showing that pSULT2B1 was not involved in
androstenone metabolism. In addition, comparisons between human and porcine
SULT2A1 showed substantially different substrate affinities for androstenone and
DHEA. Future studies should focus on confirming the results observed in the present
study regarding the sulfoconjugation activity of pSULT2B1 as well as on the biological
role of substrate inhibition in both humans and pigs.
82
CHAPTER 5: GENERAL DISCUSSION AND CONCLUSIONS
5.1 Discussion and Future Considerations
The purpose of this project was to investigate the metabolism of androstenone in
the liver and testes, and to further characterize the enzymes involved in androstenone
sulfoconjugation. It was expected that both porcine Leydig cells and hepatocytes would
sulfoconjugate androstenone through the action of SULT2A1 and SULT2B1 enzymes.
Results from primary cell incubation studies showed that the Leydig cells produced
steroid sulfonates, but not glucuronides, whereas the hepatocytes produced androstenol
glucuronide, but not sulfonates. HPLC analysis showed that the steroid sulfonates eluted
as a group at approximately 20 minutes and analysis by LC-MS identified androstenol
sulfonate and a sulfated form of androstenone as the metabolites making up this
conjugated group. In addition, approximately 85.7% of androstenone was metabolized
into these compounds after 8 hours of incubation with Leydig cells. The high proportion
of sulfoconjugates observed in the current study are similar to that found in previous
studies involving 16-androstene steroids, androgens, and estrogens in pigs (Saat et al.,
1974; Raeside and Renaud, 1983; Sinclair and Squires, 2005; Sinclair et al., 2005a), and
are a reflection of the high levels of sulfotransferase enzymes present in Leydig cells
(Hobkirk et al., 1989; Raeside and Renaud, 1983).
Of particular importance was the identification of a sulfoconjugated form of
androstenone. LC-MS analysis in negative ionization mode showed that this metabolite
had an m/z of 367.16 which was comparable to that reported in a similar study
(Desnoyer, 2011). This author suggested a corresponding structure of 3-keto-sulfoxy83
androstneone. It has generally been thought that androstenone is not capable of
undergoing sulfoconjugation as it does not possess any hydroxyl groups, however
androstenone sulfonate has been found in the plasma of boars (Sinclair and Squires,
2005). The production of androstenone sulfonate was suggested to occur through prior
enolisation of the 3-keto group (Sinclair et al., 2005a,b, 2006). This involves the removal
of an alpha hydrogen atom from a carbon that is placed on an oxygen atom to form a
hydroxyl group (Toullec, 1982), which may then be acted on by sulfotransferase
enzymes. The resultant 3-enol sulfonate compound, however, would have an expected
mass of approximately 352.5, which was not observed in the current study. Conversely,
Desnoyer (2011) suggested that, following enolisation, sulfotransferase enzymes may act
on either the C2 or C4 position to produce a 3-keto-sulfoxy androstenone metabolite with
a mass similar to that observed in the current study. However, the sulfotransferase
enzyme responsible for conjugating 16-androstene steroids is a hydroxysteroid SULT and
thus only acts on hydroxyl groups (Strott, 2002; Sinclair and Squires, 2005; Sinclair et
al., 2005b ). Since enolisation of androstenone would result in a hydroxyl group at the C3
position, and not the C2 or C4 positions, the production of 3-keto-4-sulfoxy androstenone
may not be possible through this enolisation mechanism. Instead, the production of the 3keto-sulfoxy androstenone compound observed in the current study is suggested to occur
via the addition of a hydroxyl group at a position other than C3. Hydroxylated forms of
androstenone have been shown to be produced by porcine hepatocytes (Sinclair et al.,
2005b). In addition, Austin and Ellis (2003) identified 6-hydroxy androstenone as a
metabolite produced by axillary microflora. Although hydroxylation of other steroids has
also been shown to occur at the C6 position (Zimniak et al., 1991; Miyata et al., 1994),
84
there are multiple sites where this reaction can occur due to the various enzymes that may
be involved (Waxman, 1988; Martucci and Fishman, 1993; Gilep et al., 2011; Robic et
al., 2014). Previously, identification of conjugated steroids was indirect and potentially
inaccurate, as isolation and derivatization were required prior to analysis (Guazzo et al.,
1996; Higashi and Shimada, 2004; Penning et al., 2010). However, in the present
investigation, entire samples containing both conjugated and unconjugated steroids were
successfully analyzed. Unfortunately, the identification of the sulfoconjugated
metabolites observed in this study could not be confirmed since authentic reference
standards are not yet commercially available. Future research should therefore focus on
identifying the structure of androstenone sulfonate. This could be accomplished by
analyzing samples using various spectroscopy techniques such as nuclear magnetic
resonance (NMR) which determines molecular structures based on the magnetic
properties of atomic nuclei. In addition, future research should also focus on identifying
the enzymes involved in androstenone hydroxylation as this may affect the level of
unconjugated steroid available to accumulate in adipose tissue and will consequently play
a role in the development of boar taint.
In addition to the sulfoconjugated metabolites, androstenol glucuronide was also
produced, although this only occurred in incubations with hepatocytes and not with
Leydig cells. Unfortunately, the amount of metabolite produced could not be quantified,
as it was only observed by LC-MS and not by HPLC. Although the output from LC-MS
provided an abundance value, this could not be compared with those of the sulfonated
steroids since this value is dependent on the ionizability of a compound which changes
with size and chemical nature. Therefore, these values were not reported for any of the
85
metabolites detected. However, the observance of androstenol glucuronide by LC-MS
and not HPLC suggests that the levels of this compound may have been relatively low
and below the limit of detection of the HPLC. This further suggests that the viability of
hepatocytes used in this study may have been compromised, as similar studies have
shown relatively high levels of 3α- and 3β- androstenol glucuronides produced by
porcine hepatocytes (Sinclair et al., 2005b). The production of 16-androstene
glucuronides by hepatocytes, but not Leydig cells, has been shown in previous studies
(Sinclair et al., 2005a,b; Chen et al., 2015) and corresponds to the theory that
glucuronidation serves primarily as a mechanism to facilitate metabolic clearance
(Sinclair et al., 2005b). This is further supported by the high concentrations of androgen,
estrogen, and androstene glucuronides found in the urine and bile, as well as the high
levels of UGT enzymes found in the liver and kidney (Gower and Patterson, 1970;
Sinclair et al., 2005b). Therefore, future research should focus on the metabolism of
androstenone into glucuronide conjugates as well as the enzymes involved in these
processes. Recently, one study investigated gene expression patterns involved in porcine
hepatic androstenone metabolism in high and low androstenone boars from three
different breeds (Sahadevan et al., 2015). Based on co-expression cluster analysis, these
authors showed that UGT2B17, UGT2B18-like, and UGT2B31-like were co-expressed in
one of the clusters from low androstenone boars across all three breeds. The enzyme or
enzymes responsible for glucuronidating the androstenols have not been identified,
although members from the UGT2B family, which are known to conjugate steroids
(Mackenzie et al., 1992; Turgeon et al., 2001), would be the most likely candidates. In
addition, it is not known whether androstenone itself is glucuronidated in the pig although
86
this is possible and would most likely occur through prior enolisation, as described
above. In humans, UGT2B17 was shown to glucuronidate both the 3- and 17-hydroxy
positions of C19 steroids (Beaulieu et al., 1996; Bélanger et al., 2003) while monkey
UGT2B18 was shown to act on the hydroxyl group at the 3α position of C19 steroids
(Beaulieu et al., 1998). If the enzyme responsible for androstenone/androstenol
glucuronidation can be identified, and if enzyme expression and activity can be correlated
to low androstenone concentrations in adipose tissue and plasma, this enzyme could
potentially be used as a genetic marker to select for low boar taint boars.
In the second part of this study, HEK cells were transfected with plasmids
containing porcine and human SULT genes. The objective here was to further
characterize the sulfoconjugation activity of porcine SULTs. Previously, pSULT2A1 was
identified as a key enzyme involved in the metabolism of androstenone, as its activity and
expression were negatively correlated with both fat and plasma androstenone
concentrations in boars (Falany et al., 1989, 1994; Sinclair and Squires, 2005; Sinclair et
al., 2006). Porcine SULT2B1 expression was also shown to be negatively correlated to
fat androstenone concentration, although these results were achieved indirectly since goat
anti-human SULT2B1 antibodies were used to identify the presence of pSULT2B1 via
Western blot (Moe et al., 2007a,b). However, results from the current study suggested
that pSULT2B1 was not directly involved with androstenone metabolism as this enzyme
did not appear to conjugate androstenone to any significant degree. Porcine SULT2B1
did not appear to sulfoconjugate DHEA either. Comparisons with the human orthologs,
hSULT2B1a and hSULT2B1b, showed that although these enzymes were capable of
sulfoconjugating androstenone and DHEA, their efficiencies were relatively low. These
87
human isoforms possess a 50 amino acid carboxy-terminal extension that makes them
structurally unique when compared to other human SULTs (Falany et al., 2006, 2013). In
addition, the amino-termini of hSULT2B1a and hSULT2B1a are also extended and
include 8 and 23 amino acids, respectively (Falany et al., 2013). Fuda et al. (2002)
showed that the hSULT2B1b isoform preferentially sulfonates cholesterol, while
hSULT2B1a sulfonates pregnenolone, but not cholesterol. These authors determined that
isoleucines present at positions 21 and 23 in the extended amino-terminus of
hSULT2B1b were crucial for cholesterol sulfoconjugation. Interestingly, the porcine
SULT2B1 enzyme also contains an extended amino-terminus which is 65% similar to
that of the human ortholog and includes the Ile21 and Ile23 residues. Therefore, like
hSULT2B1b, the pSULT2B1 enzyme may preferentially sulfoconjugate cholesterol.
Further, since cholesterol sulfonate can be metabolized to other sulfoconjugated steroids
via the steroid-sulfonate synthesis pathway (Roberts et al., 1964; Gasparini et al., 1976;
Falany et al., 1995), pSULT2B1 may still play an indirect role in the biosynthesis of
androstenone and, consequently, in the development of boar taint. Future studies should
therefore focus on determining which substrates are capable of being sulfoconjugated by
pSULT2B1. In addition, hSULT2B1 isoforms have shown distinct tissue expression
patterns compared to the hSULT2A1 enzyme (reviewed by Falany and Rohn-Glowacki,
2013). Generally, hSULT2A1 is expressed in tissues that do not express hSULT2B1b and
vice versa. Therefore, studies comparing pSULT2A1 and pSULT2B1 tissue expression
are warranted in order to further understand the function of these enzymes and role that
they play in the development of boar taint. Finally, since pSULT2B1 expression was
negatively correlated to androstenone concentration in adipose tissue (Moe et al.,
88
2007a,b), future studies investigating potential polymorphisms within this enzyme may
be useful for developing genetic markers for boar taint.
5.2 Conclusions
This research has provided direct evidence, for the first time, showing that
androstenone sulfoconjugation does occur in the boar. In addition, porcine SULT2B1 did
not sulfoconjugate androstenone, although it may still be indirectly involved in boar taint.
The results presented in the current investigation add to the further understanding of the
pathways and genes involved in the development of boar taint, and emphasize the
importance of conjugation in steroid metabolism. Although a new method for controlling
boar taint was not discovered here, it is hoped that this research will lead to new lines of
investigation and contribute to the identification of genetic markers, allowing for the
genetic selection of low boar taint boars without negative consequences.
89
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