myocyte androgen receptor modulates body …...myocyte androgen receptor modulates body composition...
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MYOCYTE ANDROGEN RECEPTOR MODULATES BODY COMPOSITION AND
METABOLIC PARAMETERS
by
Shannon M. Fernando
Hon. B.Sc. University of Toronto, 2008
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Institute of Medical Science
University of Toronto
© Shannon M. Fernando, 2010
UNIVERSITY OF TORONTO
July, 2010
All rights reserved. This work may not be
reproduced in whole or in part, by photocopy or other means,
without the permission of the author
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Myocyte androgen receptor modulates body composition and metabolic parameters Shannon M. Fernando, Master of Science, 2010 Institute of Medical Science, University of Toronto
ABSTRACT
Androgens (such as testosterone) have been shown to increase lean body mass and
reduce fat body mass in men through activation of androgen receptors (AR). While this
suggests a potential clinical use for androgens, attempts at utilization of this class of
hormones as a therapeutic are limited by side effects due to indiscriminate AR activation
in various tissues. Thus, a greater understanding of the tissues and cells involved in
promoting these changes would be beneficial. Here we show that selective overexpression
of AR in muscle cells of transgenic (HSA-AR) rodents both increases lean muscle mass
and significantly reduces fat mass in males. Similar effects can be induced in HSA-AR
females treated with testosterone. Metabolic analyses of HSA-AR males show that these
animals demonstrate increased O2 consumption and hypermetabolism. Thus, targeted
activation of AR in muscle regulates body composition and metabolism, suggesting a
novel target for drug development.
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ACKNOWLEDGEMENTS
None of the work contained within these pages could have been accomplished
without the help of many wonderful people. I must begin by thanking my supervisor, Dr.
Ashley Monks, for his incredible guidance and support over the course of my time in his
laboratory. I entered his lab as a very “green” undergraduate, but his knowledge and trust
have allowed me the opportunity to learn the pitfalls and plateaus of science. He took a
chance on me, and for that I am forever grateful. I have thoroughly enjoyed my time
under his tutelage, as well as our many collaborative efforts, and I can only hope that I
have given back as much as he has given to me. I also thank the members of my Program
Advisory Committee, Drs. Angela Lange and Tim Westwood, for their input and
encouragement. I thank the members of the Monks Lab for their consistent help and
support. Most notably, I must acknowledge the efforts of Dr. Lee Niel, who seemingly
helped me in everything I did from the first moment I walked into the lab. I also thank Dr.
Pengcheng Rao, Dr. Melissa Holmes, Dr. Kaiguo Mo, Dr. Diptendu Chatterjee, Marijana
Stagljar and Mutaz Musa for technical support. The learning curve would have been
much steeper if not for you all.
I am very grateful to have the unconditional love and support of Arthur, Jini and
Krystyna Fernando. Many thanks for putting up with my complaints, and for
understanding the importance of the words: “I have work to do.”
Lastly, I must acknowledge the Natural Sciences and Engineering Research
Council of Canada, the Ontario Graduate Scholarship Program, and the University of
Toronto Open Fellowships for the funding of my research.
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TABLE OF CONTENTS
TITLE PAGE..................................................................................................................... I ABSTRACT ...................................................................................................................... IIACKNOWLEDGEMENTS ........................................................................................... IIILIST OF TABLES......................................................................................................... VIILIST OF FIGURES..................................................................................................... VIIICHAPTER 1: INTRODUCTION .................................................................................... 11.1.OVERVIEW..........................................................................................................................11.2.ANINTRODUCTIONTOBODYCOMPOSITION ....................................................................21.2.1.Lean(Muscle)Mass ..................................................................................................................21.2.2.FatMass.........................................................................................................................................61.2.3.Bone.................................................................................................................................................8
1.3.ENERGYBALANCEANDMETABOLISM............................................................................. 101.3.1.WhatisEnergyBalance?..................................................................................................... 101.3.2.EnergyIntake ........................................................................................................................... 121.3.3.EnergyExpenditure ............................................................................................................... 151.3.4.EnergyBalance,BodyCompositionandObesity....................................................... 18
1.4.THEANDROGENSYSTEM ................................................................................................ 211.4.1.AndrogenPharmacology–StructureandProduction........................................... 221.4.2.AndrogenReceptor ................................................................................................................ 25
1.5.ANDROGENS,ARANDMUSCLE....................................................................................... 271.5.1.AnabolicFunctionsofARinMuscle................................................................................ 281.5.2.MuscleARandNeuromuscularDevelopment ............................................................ 311.5.3.SpinalandBulbarMuscularAtrophy ............................................................................ 34
1.6.ANDROGENS,ARANDFAT ............................................................................................. 361.6.1.SexDifferencesinAdiposity ............................................................................................... 361.6.2.AndrogensandAdipocyteDevelopment....................................................................... 40
1.7.WHERETHEACTIONIS:LESSONSLEARNEDFROMARKO ........................................... 421.7.1.WholeBodyAndrogenReceptorKnockout................................................................. 441.7.2.TissueSpecificAndrogenReceptorKnockout............................................................ 46
1.8.THEMETABOLICPOTENTIALOFMUSCLE ...................................................................... 491.8.1.SkeletalMuscleMetabolism............................................................................................... 491.8.2.MuscleMitochondria............................................................................................................. 53
1.9.OBJECTIVESANDHYPOTHESES ....................................................................................... 58CHAPTER 2: MATERIALS AND METHODS ........................................................... 602.1.OVERVIEW....................................................................................................................... 602.2.ANIMALSTRAINS............................................................................................................. 602.2.1.HSAARRats.............................................................................................................................. 602.2.2.TfmRats...................................................................................................................................... 622.2.3.HSAAR/TfmRats ................................................................................................................... 642.2.4.HSAARMice ............................................................................................................................. 65
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2.3.SUBJECTS ......................................................................................................................... 672.3.1.Animals........................................................................................................................................ 672.3.2.Genotyping................................................................................................................................. 67
2.4.EXPERIMENTI:TISSUESPECIFICITYOFTRANSGENEEXPRESSION ................................ 692.4.1.ReverseTranscription(EndPoint)PCR....................................................................... 70
2.5.EXPERIMENTII:BODYCOMPOSITIONANALYSIS ........................................................... 712.5.1.DualEnergyXRayAbsorptiometry............................................................................... 712.5.2.ParametersExaminedandDerived................................................................................ 722.5.3.DissectionsandTissueWeights........................................................................................ 73
2.6.EXPERIMENTIII:TESTOSTERONETREATMENTOFADULTFEMALES ........................... 732.6.1.TCapsuleSurgeries ............................................................................................................... 742.6.2.BodyCompositionAnalysis................................................................................................. 75
2.7.EXPERIMENTIV:ADIPOSEHISTOLOGY.......................................................................... 752.7.1.SamplePreparationandSamplingStrategy.............................................................. 752.7.2.Measurements .......................................................................................................................... 77
2.8.EXPERIMENTV:ENERGYBALANCEANDMETABOLICANALYSES................................... 772.8.1.RestingMetabolismbyIndirectCalorimetry.............................................................. 772.8.2.SpontaneousActivityMeasures........................................................................................ 79
2.9.STATISTICALANALYSES .................................................................................................. 79CHAPTER 3: RESULTS ................................................................................................ 813.1.OVERVIEW....................................................................................................................... 813.2.EXPERIMENTI:TISSUESPECIFICITYOFTRANSGENEEXPRESSION ................................ 813.2.1.TransgenemRNAisExpressedinMuscleTissueofHSAARAnimals .............. 82
3.3.EXPERIMENTII:BODYCOMPOSITIONANALYSIS ........................................................... 823.3.1.HSAARExpressionDoesNotRegulateBodyMassinRats .................................. 833.3.2.IncreasedLeanMuscleMassPercentinHSAARMaleRats ................................ 833.3.3.ReducedFatBodyMassinHSAARMaleRats ........................................................... 853.3.4.NoEffectsonBodyCompositionofHSAARFemaleRats ..................................... 863.3.5.IndividualMuscleandFatPadWeights ....................................................................... 873.3.6.HSAARSimilarlyAffectsBodyCompositioninL78Mice ..................................... 883.3.7.EffectsofHSAARandTfmonBoneContentandDensity .................................... 89
3.4.EXPERIMENTIII:TESTOSTERONETREATMENTOFADULTFEMALERATS ................... 913.4.1.TTreatmentRegulatesBodyCompositionofHSAARFemales......................... 923.4.2.TTreatmentDoesNotAffectBodyCompositionofWildtypeFemales ......... 94
3.5.EXPERIMENTIV:ADIPOSEHISTOLOGY.......................................................................... 943.5.1.HSAARExpressionReducesAdipocyteArea.............................................................. 94
3.6.EXPERIMENTV:ENERGYBALANCEANDMETABOLICANALYSES................................... 953.6.1.HSAARExpressionIncreasesRestingMetabolisminRatsandMice .............. 963.6.2.SpontaneousActivityisNotAffectedbytheTransgene ........................................ 97
3.7.SUMMARY ........................................................................................................................ 97CHAPTER 4: DISCUSSION.......................................................................................... 994.1.OVERVIEW....................................................................................................................... 994.2.MYOCYTEARANDBODYCOMPOSITION......................................................................1004.2.1.LeanMuscleMass .................................................................................................................1014.2.2.AdiposeTissue........................................................................................................................104
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4.2.3.Bone............................................................................................................................................1084.3.THEEFFECTOFTFMONBODYCOMPOSITION..............................................................1114.3.1.ComparisonwithARKOMice...........................................................................................1114.3.2.WhytheDiscrepancy?.........................................................................................................112
4.4.INTERACTIONSWITHESTROGENS.................................................................................1154.4.1.EstrogensandAdiposeTissue .........................................................................................1164.4.2.Estrogens,HSAARandTfm.............................................................................................117
4.5.MYOCYTEARANDOXIDATIVEMETABOLISM ..............................................................1204.5.1.LocalEffectsonSkeletalMuscleMetabolism ...........................................................1204.5.2.MitochondrialBiogenesisandEnzymeActivity ......................................................1214.5.3.EffectsonOtherMetabolicOrgans ...............................................................................124
4.6.COMPARINGHSAARRATSANDMICE........................................................................1264.6.1.PhenotypeComparisons ....................................................................................................1274.6.2.DoesExpressionLevelExaggeratePhenotype?.......................................................128
4.7.ANEWHOPE:SELECTIVEANDROGENRECEPTORMODULATORS ...............................1304.7.1.SelectiveAndrogenReceptorModulators..................................................................1314.7.2.TargetingMuscle ..................................................................................................................132
4.8.FUTUREDIRECTIONS ....................................................................................................1334.8.1.OtherMetabolicParametersinHSAARAnimals...................................................1334.8.2.IdentifyingARandMitochondrialInteractionsinSkeletalMuscle ................1354.8.3.MolecularandBiochemicalAssaysofKeyMetabolicPlayers...........................137
4.9.CONCLUSIONS ................................................................................................................139REFERENCES .............................................................................................................. 141TABLES AND FIGURES ............................................................................................. 161APPENDIX A: STASTICAL VALUES ...................................................................... 174
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List of Tables
Table 1: Summary of animals used in each experiment
Table 2: List of primers used
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List of Figures
Figure 1: Characterization of Transgene Expression
Figure 2: Transgene Expression Regulates Body Composition in Males
Figure 3: No Effect of the Transgene on Body Composition in Female Rats
Figure 4: Excised Fat Pads and Muscles Confirm DXA Findings
Figure 5: Body Composition of HSA-AR Mice
Figure 6: Effects of HSA-AR on Bone Parameters
Figure 7: Body Composition is Altered by T-treatment of HSA-AR Females
Figure 8: Smaller Adipocytes are Found in HSA-AR Males
Figure 9: Increased Oxygen Consumption in HSA-AR Male Rats
Figure 10: Differences in Energy Expenditure in HSA-AR L78 Mice
Figure 11: HSA-AR Expression Does Not Affect Spontaneous Activity Level
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List of Abbreviations
aARKO = adipocyte-specific androgen receptor knockout AICAR = aminoimidazole carboxamide ribonucleotide ALS = amyotrophic lateral sclerosis AMPK = 5’ adenosine monophosphate-activated protein kinase αMSH = α-melanocyte stimulating hormone ANOVA = analysis of variance AR = androgen receptor ARKO = androgen receptor knockout ArKO = aromatase knockout AT = anterior tibialis ATP = adenosine triphosphate AVPV = anteroventral periventricular nucleus BAT = brown adipose tissue BC = bulbocavernosus BMC = bone mineral content BMD = bone mineral density BMR = basal metabolic rate CART = cocaine and amphetamine related transcript CP = creatine phosphate cDNA = complementary DNA C/EBP = CCAAT/enhancer binding protein α CNTF = ciliary neurotrophic factor DBD = DNA-binding domain DEPC = diethyl-pyrocarbonate DHT = dihydrotestosterone DNA = deoxyribonucleic acid DXA = dual-energy X-ray absorptiometry EDL = extensor digitorum longus ETC = electron transport chain FATP = fatty-acid transport protein FBM = fat body mass FSH = follicle-stimulating hormone GAPDH = glyceraldehyde-3-phosphate dehydrogenase GLUT4 = glucose transporter 4 GnRH = gonadotropin-releasing hormone GS = glycogen synthase HD = Huntington’s Disease HKII = hexokinase II HPG = hypothalamic-pituitary-gonadal HSA = human skeletal alpha-actin HSA-AR = human skeletal alpha-actin promoter, androgen receptor transgene IGF-1 = insulin-like growth factor-1 IHC = immunohistochemistry IR = insulin receptor
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kb = kilobase LA = levator ani LBM = lean body mass LH = luteinizing hormone LPL = lipoprotein lipase mARKO = myocyte-specific androgen receptor knockout MC4R = melanocortin 4 receptor MePD = posterodorsal medial amygdala mtDNA = mitochondrial DNA mRNA = messenger ribonucleic acid NPY = neuropeptide Y NTD = amino-terminal transactivation domain O2 = oxygen OCT = optimal cutting temperature embedding medium OD = optical density PAS = periodic acid-schiff PASD = diastase-digested periodic acid-schiff PBF = phosphate buffered formalin PBS = phosphate buffered saline PCOS = polycystic ovarian syndrome PCR = polymerase chain reaction PGC-1α = peroxisome proliferator-activated receptor gamma, coactivator 1 alpha PM = peritubular myoid PND = post-natal day POMC = proopiomelanocortin PPAR = peroxisome proliferator-activated receptor RMR = resting metabolic rate RNA = ribonucleic acid RT-PCR = reverse-transcription polymerase chain reaction SARM = selective androgen receptor modulator SBMA = spinal and bulbar muscular atrophy SCARKO = sertoli cell-specific androgen receptor knockout SD = Sprague-Dawley SDN-POA = sexually dimorphic nucleus of the preoptic area SHBG = sex hormone-binding globulin T = testosterone T2DM = type 2 diabetes mellitus Tfm = Testicular feminization mutation Tg = transgenic Type I = slow-twitch muscle fibers Type IIa = fast-twitch, oxidative muscle fibers Type IIb = fast-twitch, glycolytic muscle fibers UCP = uncoupling protein VEGF = vascular endothelial growth factor WAT = white adipose tissue WT = wild-type
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Chapter 1: Introduction 1.1. Overview
Perhaps the most well known function of androgens is its anabolic role in
regulating skeletal muscle mass. In men, this increase in skeletal muscle mass is largely
believed to be associated with activation of the androgen receptor (AR). However, men
who take exogenous amounts of androgens also experience a decrease in adipose tissue.
Significantly less is known about how androgens are capable of regulating body fat, and
improving body composition. Furthermore, significant controversy exists as to which
tissues possess the AR necessary for achieving these profound effects on body
composition. The principle purpose of this study was to investigate transgenic
(Tg) rats that overexpress AR in muscle fibers, and how this increased expression
contributes to changes in overall body composition and energy balance.
This introductory section provides the reader first with a general review of body
composition and the complex nature of energy homeostasis. This is followed by an
overview of the androgen system. A review of androgen function will be provided,
primarily focused upon how androgens are currently known to regulate muscle mass and
adipose tissue. Finally, discussion will turn toward the current problems in defining
androgen site-of-action in mediating body composition, as well as the metabolic potential
of skeletal muscle and the role it may play in regulating these effects. Finally, objectives
and hypotheses of this study will be described.
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1.2. An Introduction To Body Composition
Most (if not all) mammalian species on the planet share a common set of
components that constitute their collective mass (Corva & Medrano, 2000). These
common components provide a basis for homeostatic maintenance, diverse behaviours,
and development across the lifespan. Scientific study has largely separated these
components into three major types: lean (muscle) mass, fat mass, and bone. All three
serve important roles in the maintenance of mammalian life, and modulation of overall
body composition can have important implications for health and disease. Here we review
these major elements, understanding not only that each component of body composition
differs from others, but also the complexity contained within each.
1.2.1. Lean (Muscle) Mass
While the term “lean mass” is most accurately defined as the collection of non-fat,
non-bone tissue in the mammalian organism, many instead simply define it as overall
muscle mass. Muscle is the major type of contractile tissue contained within animals, and
is derived from the mesoderm layer during development (Orallo, 1996). It represents a
major site of nutrient metabolism, and functions through the movements of individual
contractile filaments that move past each other, subsequently changing the size of the
overall cell. There are three major types of muscle: smooth, cardiac, and skeletal.
The three types of muscle differ in various properties, including structure,
contractile properties, and control mechanisms (Orallo, 1996; Caplice & Deb, 2004;
Allen, Lamb & Westerblad, 2008). Smooth muscle is largely organized into sheet-like
structures and surrounds various hollow organs and tubes, including the stomach,
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intestines, urinary bladder, uterus, blood vessels, and airways in the lungs. It is non-
striated, and contraction of smooth muscle surrounding hollow organs may allow for the
propelling of luminal contents through the organ, or changes in the diameter of the tube.
Smooth muscle myocytes are attached to the hairs of the skin, as well as the iris of the
eye. Control of smooth muscle contraction is involuntary, and dictated by the autonomic
nervous system, endocrine/autocrine/paracrine agents, and other local chemical signals. In
contrast, cardiac muscle is a striated type of muscle that makes up the muscle of the heart.
Contraction of cardiomyocytes propels blood through the circulatory system. Similar to
smooth muscle, its contraction is involuntary, and under the control of similar neuronal
and endocrine signals. Finally, as its name suggests, skeletal muscle is attached to bone,
and is found throughout the body, from the rectus femoris of the leg to the trapezius of the
neck. Contraction of skeletal muscle is responsible for support and movement of the
skeleton. Similar to cardiac muscle, skeletal muscle is striated. However, unlike the other
types of muscle, skeletal muscle contraction occurs under voluntary control, and is
initiated by impulses in the neurons that innervate these muscles.
When studying body composition, skeletal muscle is often the most discussed
muscle-type, due to the fact that it is the most prone to change over the course of the
lifespan. Growth of skeletal muscle is typically associated with resistance exercise, a
prominent method used for achieving change in body composition. Similarly, skeletal
muscle is one of the most metabolically demanding tissues in mammalian bodies, and
thus changes in overall skeletal muscle weight can result in modulation of systemic
metabolism.
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Differences in skeletal muscle fiber types can be elucidated based upon
histochemical staining properties of the myosin adenosine triphosphatase (ATPase)
enzyme. Functional characteristics of each muscle fiber type are based upon its ATPase
enzyme activity. In this way, three major skeletal muscle fiber types have been found:
type I (slow-twitch), type IIa (fast-twitch, oxidative), and type IIb (fast-twitch, glycolytic)
(Staron, 1997). These fiber types can be distinguished on the basis of contraction time,
activity/duration of use, endurance, mitochondrial density, power, and
oxidative/glycolytic capacity. Type I (slow-twitch) fibers are predominantly used for
aerobic activity (Coen et al., 2010). As the name indicates, they have a slow contraction
time, and are characterized by a low activity level of ATPase. They contain large and
numerous mitochondria, and thus, are relatively resistant to fatigue. In contrast, type II
(fast-twitch) fibers demonstrate a significantly higher level of ATPase activity (Klover,
Chen, Zhu & Hennighausen, 2009). They have a much faster contraction time, and are
used during short-term, anaerobic activity. Type II fibers contain much less mitochondria
than type I fibers, and as a result, are far less resistant to fatigue. Type II fibers can also
be distinguished based upon their methods of energy utilization and metabolism. Certain
fast-twitch fibers are similar to slow-twitch fibers in that they strongly rely upon
oxidative metabolism for generation of ATP. These fibers are termed type IIa (fast-twitch,
oxidative). Conversely, type IIb fibers rely primarily upon anaerobic glycolysis for
generation of ATP. This method of energy generation is less efficient than oxidative
metabolism (less ATP per molecule of glucose), but is also faster. These fibers are thus
termed as fast-twitch, glycolytic fibers. Due to their reliance upon oxidative metabolism,
type IIa fibers tend to have relatively slower contraction time than type IIb fibers, but are
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also more resistance to fatigue. Individual muscles typically contain all three fiber types,
although the proportions of certain types are greater in specific muscles. For example, the
gastrocnemius muscle is largely composed of type II fibers, while the soleus muscle
(which together with the gastrocnemius constitutes the calf muscle) is largely composed
of type I fibers (Wang et al., 2004). Thus, even muscle groups in close proximity can be
composed of very different fiber type proportions.
Muscle fiber types can adapt to changing demands by altering size or fiber type
composition (Scott, Stevens & Binder-MacLeod, 2001). This is typically what occurs
when individuals take part in exercise training, whether resistance- or endurance-based.
Both types of training tend to result in muscle fiber hypertrophy, although there is some
disagreement as to which fiber types are preferentially altered by exercise (Adams,
Hather, Baldwin & Dudley, 1993). Both endurance and resistance training result in
similar reductions of MHC coexpression, resulting in a greater number of “pure” fibers.
However, while both types of exercise result in similar trends in fiber type conversion, the
physiological changes induced by each type of exercise are rather different. Endurance
training results in an increase of muscle oxidative capacity (hypertrophy of type I fibers),
while resistance training increases overall muscle fiber strength and size (hypertrophy of
type II fibers).
Muscular contraction is a major energetically demanding process. Contraction of
skeletal muscle mediates all voluntary movement and physical action in mammalian
organisms. The ability of a muscle fiber to generate force and movement depends on the
interaction of the two major contractile proteins: actin and myosin (Kee, Gunning &
Hardeman, 2009). Actin molecules are globular proteins composed of a single
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polypeptide, and polymerize with other actins in order to form intertwined helical chains.
These chains make up the core of a thin filament in the sarcomere. Myosin, conversely, is
composed of two large polypeptide heavy chains, and four smaller light chains. These
polypeptides combine to form a molecule that consists of two globular heads, and a long
tail. The tail of each myosin molecule lies along the axis of the thick filament, and the
two globular heads extend out to the sides, forming the cross-bridges. Each globular head
contains two binding sites, one for actin, and one for ATP. During contraction, ATP is
hydrolyzed (to form ADP and a phosphate group). Using the energy from ATP
hydrolysis, myosin is able to bind to actin, and pull adjacent actin molecules together.
After the end of a contraction sequence, the binding of a new ATP molecule is required in
order for the link between actin and myosin to be broken. Thus, one can see that muscle
contraction requires high amounts of energy in the form of ATP.
1.2.2. Fat Mass
Triglyceride (fat) consists of three fatty acids, which are linked to a glycerol
group. Fat accounts for approximately 80 percent of the energy that is stored in the body
(Friedman, 2009). Under typical resting conditions, approximately half of the energy
utilized by muscle, liver and kidney is derived from catabolism of fatty acids. In
mammalian systems, although most cells do store some small amounts of fat, the
predominant localization of these molecules is in adipose tissue (Rosen & Spiegelman,
2000). This is a collection of loose connective tissue composed of individual fat cells
called adipocytes. Adipose tissue is largely localized to specific depots within the body. It
either exists in various deposits underlying the skin (termed ‘subcutaneous fat’) or in
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larger aggregates in body cavities such as the abdomen (termed ‘visceral fat’). The
function of individual adipocytes is to synthesize and store triglycerides during periods of
food uptake and then, when food is not being absorbed from the intestinal tract, to release
fatty acids and glycerol into the blood so that they may be taken-up and used as substrates
for the harvest of energy (in the form of ATP). Here, this review will very briefly focus
on the two most important pathways involving fatty acids: fat synthesis (the formation of
triglycerides and incorporation into adipocytes) and fat catabolism (breakdown of fat
molecules in order to derive energy).
Fat synthesis (the production of triglyceride molecules) and integration into
adipocytes are now understood to be complex biochemical processes, with various
cellular and molecular players (Lefterova & Lazar, 2009). That being said, the pathway
through which fat synthesis occurs remains the same. The enzymes that mediate fat
synthesis are largely localized to the cytoplasm, whereas the breakdown of fat (as will be
discussed shortly) largely occurs because of enzymes in the mitochondria. Fatty acid
synthesis begins with cytoplasmic acetyl coenzyme A, which transfers its acetyl group to
another molecule of acetyl coenzyme A. This forms a four-carbon chain. Repetition of
this process builds up long-chain fatty acids (two carbons at a time). These newly
synthesized fatty acids are then transported across the membrane of adipocytes, where
they are combined with glycerol in an esterification reaction to form triglycerides, which
are then incorporated into these cells. The mechanism by which this occurs remains
unclear, with various proteins in the adipocyte plasma membrane being implicated in
fatty acid transport (Pohl, Ring, Korkmaz, Ehehalt & Stremmel, 2005).
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The utilization of fatty acids is a major source of energy in mammalian
physiology (Floyd, Medes & Weinhouse, 1947). Release of fatty acids from adipocytes is
the first major step in fat metabolism. Once the fatty acids have been taken-up by a
metabolic tissue (such as skeletal muscle or liver), they are transported into the
mitochondria, where ATP harvest occurs. The breakdown of a fatty acid is initiated by
linking a molecule of coenzyme A to the carboxyl end of the fatty acid. The coenzyme A
derivative of the fatty acid is subsequently taken through a series of reactions that are
collectively known as β-oxidation. In this process, a molecule of acetyl coenzyme A is
removed from the end of the fatty acid, which includes the transfer of two pairs of
hydrogen atoms. The hydrogen atoms from the coenzymes then enter the oxidative
phosphorylation pathway to form ATP. When an acetyl coenzyme A is split from the end
of a fatty acid, another coenzyme A is added, and the sequence is repeated. In this way,
the initial fatty acid is shortened by two carbons each time, until all the carbon atoms
have been transferred to coenzyme molecules. These molecules subsequently produce
carbon dioxide and ATP via the Krebs cycle and oxidative phosphorylation. The result is
a relatively large yield of ATP from a single fatty acid molecule, making β-oxidation of
fatty acids one of the most efficient methods of energy generation.
1.2.3. Bone
The final major component of body composition is bone, which makes up the
mammalian skeleton. Bone is a tissue that consists of a protein (collagen) matrix upon
which calcium salts (notably calcium phosphates) are deposited. For descriptive purposes,
a growing long bone is divided into the ends (epiphyses) and the remainder (the shaft).
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Bone remodeling is a complex process that mediates growth and loss of bone tissue
(Luisetto & Camozzi, 2009). Osteoblasts, the bone-forming cells at the shaft edge of the
epiphyseal growth plate, function by converting the cartilaginous tissues at the edge into
bone. Meanwhile, specialized cells called chondrocytes lay down new cartilage in the
interior of the plate. In this way, the epiphyseal growth plate remains intact, and is
gradually pushed away from the center of the bony shaft as lengthening of the shaft
occurs. Conversely, specialized cells called osteoclasts mediate a process called
resorption (Hattner, Epker & Frost, 1965). Here, osteoclasts function by removing the
mineralized matrix and breaking up organic bone. The minerals released during bone
resorption are subsequently transported to the blood. Bone growth and decline are
important indicators of overall health. Work in endocrinology has shown that bone
density and strength are strongly influenced by a variety of different hormonal
contributors (Olney, 2009). In this way, bone structure is a commonly used indicator of
endocrine status and possible pathology.
In summary, therefore, it can be seen that body composition is characterized by
different components, each with different properties. In the context of energy
homeostasis, most work is focused on delineating differences in skeletal muscle and
adipose tissue. White adipose tissue (WAT) serves as one of the most important stores of
energy. Skeletal muscle performs similar storage functions (although not to the degree of
adipose or liver), but is also capable of energy catabolism, and is one of the most
energetically demanding tissues in the body. In this way, constant interplay between
skeletal muscle and adipose has become a focus of metabolic research.
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1.3. Energy Balance and Metabolism
Animal physiology is predicated on the idea of homeostasis: the need for the body
to maintain a particular state, with perturbation resulting in deviation back towards the
mean or “steady state”. A delicate balance also maintains the state of energy within an
organism. The execution of various processes (both voluntary and involuntary) requires
energy (typically defined as ATP, but also in other forms). While the body expends
energy in order to conduct these processes, energy must also be taken in, in order to
replace that which is spent. This homeostatic mechanism is termed “energy balance”, and
maintenance of energy balance is under strict regulatory control within the organism.
Inability to maintain this balance can have disastrous consequences for overall health of
the organism (Spiegelman & Flier, 2001).
1.3.1. What is Energy Balance?
Animals intake energy sources (food) for the purposes of deriving various organic
molecules for use as fuel. The breakdown of these organic molecules (metabolism)
liberates the energy locked in their molecular bonds. This released energy is what cells
use in order to perform the various forms of biological work – including muscle
contraction, active transport within a cell, and molecular synthesis. Energy derivation and
expenditure within an organism is governed by the laws of thermodynamics, which
account for all energy in the universe. The first law of thermodynamics states that energy
can neither be created nor destroyed, but can only be converted from one form to another.
Therefore, the internal energy liberated during breakdown of an organic molecule can
either dissipate as heat, or be used to perform work (Spiegelman & Flier, 2001).
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The majority of energy derived from the breakdown of organic molecules is
utilized to perform work. The energy to be used for work must first be incorporated into
molecules of ATP. The subsequent breakdown of ATP serves as the immediate energy
source for the work. Biological work can generally be described as either “external” work
(ex. movement of limbs by contracting skeletal muscles); or “internal” work (cellular
activity, such as molecular transport and synthesis).
Thus, we can think of energy balance as a constant struggle between energy intake
and energy expenditure (Spiegelman & Flier, 2001). Energy intake is characterized by our
feeding habits, the nutrient sources that makeup (or are absent from) our diets.
Conversely, energy expenditure is the utilization of these nutrient sources for resting
metabolic processes, physical activity, and adaptive thermogenesis. “Tipping the scales”
too far in either direction can be thought of as maladaptive. If energy intake surpasses
energy expenditure, energy stores become larger and larger. In mammals, this can
manifest itself in metabolic syndrome, characterized by insulin resistance (and
subsequently diabetes) as well as obesity (significant fat accumulation). Similarly, if
expenditure of energy surpasses levels of intake, energy stores become depleted
(starvation). Without enough energy to maintain itself, a cell will eventually die.
Subsequently, if exacerbated, this would ultimately lead to coma and death of the
organism. Therefore, like all physiological systems under homeostatic control, energy
balance must be tightly regulated by complex mechanisms.
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1.3.2. Energy Intake
While human beings tend to think of their feeding behaviour as largely being
under conscious and voluntary control, the truth is that there are significant molecular
regulators of this process. Understanding these processes provides researchers with
significant insight about how the body manages to control the intake of food, and how
this can be affected by an individual’s energy expenditure. The molecular study of
feeding behaviour is made complex by the fact that this process is not only influenced by
unconscious, homeostatic direction, but also by psychological factors. In addition to food
availability, feeding is affected by metabolic, neuronal and hormonal factors; and is also
modified by powerful sensory, emotional and cognitive inputs. It is the interplay of all of
these factors (in higher-level organisms) that contributes to energy intake (Schwartz et al.,
2000).
In understanding the idea of physiological homeostasis, possibly no brain area has
received as much attention as the hypothalamus. This region of the brain is critical for
homeostatic processes such as feeding, thermoregulation, and reproduction (Elmquist et
al., 1999). The hypothalamus receives inputs from neural, endocrine and metabolic
signals. It then integrates this information and triggers various response pathways. The
role of the hypothalamus in regulating feeding behaviour is well established. Lesion
studies demonstrate that lesions in the ventromedial hypothalamus result in obesity, while
lesions of the lateral hypothalamus cause leanness (Elmquist et al., 1999). We now
understand that the hypothalamus is able to regulate feeding through secretion of various
neuropeptides. These neuropeptides are essentially neurotransmitters, which function
through complex metabolic signaling pathways. They are classified as either being
13
orexigenic (appetite-stimulating) or anorexigenic (appetite-inhibiting). The most
prominent hypothalamic orexigenic neuropeptide is neuropeptide Y (NPY). When
administered into cerebral hemispheres or specific hypothalamic nuclei, NPY robustly
and rapidly increases feeding and suppresses energy expenditure, thereby promoting
obesity (Stanley et al., 1986; Billington et al., 1994). Conversely, the major anorexigenic
neuropeptides produced by the hypothalamus are cocaine and amphetamine related
transcript (CART), and α-melanocyte stimulating hormone (α-MSH), which are both
derived from proopiomelanocortin (POMC). These neuropeptides function by activation
of melanocortin 4 receptor (MC4R). Activation of MC4R by α-MSH reduces food intake,
while suppression of MC4R by pharmacological antagonists increases feeding (Fan et al.,
1997). Similarly, genetic deletion of MC4R results in obesity in mice (Huszar et al.,
1997), while this same phenotype also occurs due to mutation in the POMC gene,
disrupting synthesis of CART/α-MSH (Krude et al., 1998). Thus, the hypothalamus is
capable of regulating feeding through secretion of neuropeptides, which then have
downstream effects upon metabolic signaling pathways.
But how does the hypothalamus know whether to secrete orexigenic or
anorexigenic neuropeptides? It is this question that is critical to understanding the
homeostatic control of energy balance. The hypothalamus secretes neuropeptides in
response to external molecular signals. The most prominent of these signals are endocrine
in nature. It was the discovery of the adipose-derived hormone leptin that provided the
best evidence for endocrine control of feeding (Friedman & Halaas, 1998). Leptin is
primarily manufactured in the adipocytes of WAT, and therefore the level of circulating
leptin is directly proportional to the total amount of fat. With regards to feeding
14
behaviour, leptin serves as a satiety signal, inhibiting appetite and further food intake.
Mice with homozygous mutations in the Ob gene, which codes for the leptin protein,
develop obesity characterized by hyperphagia (Zhang et al., 1994). Treatment of these
mutant mice with the missing hormone attenuates weight gain and appetite (Halaas et al.,
1995). Expression of leptin is found in other tissues, such as skeletal muscle, suggesting
that other metabolic organs may act to inhibit appetite (Wang et al., 1998). The
mechanism through which leptin suppresses appetite is not entirely clear. However, what
is known is that leptin binds to receptors in the hypothalamus, and regulates secretion of
hypothalamic neuropeptides. For example, leptin administration has been shown to
reduce expression of orexigenic neuropeptides, such as NPY (Elias et al., 1998).
Conversely, leptin action on the hypothalamus increases expression of anorexigenic
peptides, such as CART and α-MSH (Elmquist et al., 1999). Through these simultaneous
effects, leptin is able to reduce food intake through appetite suppression.
In contrast to leptin is the hormone ghrelin. This peptide hormone is expressed
primarily in stomach and brain (with the stomach being its major site of synthesis), but
the control of its expression is not completely understood (Lutter et al., 2008). Ghrelin
action opposes that of leptin, acting on the hypothalamus to induce hyperphagia and
energy intake (Nakazato et al., 2001). Daily administration of ghrelin causes increased
food intake, reduced fat utilization, and eventual obesity in mice and rats (Tshcop, Smiley
& Heiman, 2000). Similar to leptin, however, the mechanism of ghrelin action has not
been completely elucidated. There is existing evidence to suggest that ghrelin acts upon
the hypothalamus to increase NPY release, and stimulation of hunger (Tschop et al.,
15
2000). Thus, it can be concluded that energy intake is largely controlled by neuronal and
endocrine signals, with the overall purpose being the maintenance of energy homeostasis.
1.3.3. Energy Expenditure
In order for the organism to maintain energy homeostasis, the energy generated
through intake (i.e. feeding) must be split in order to drive various processes. Energy
expenditure is largely accomplished by three major factors: basal metabolism, physical
activity, and adaptive thermogenesis. Here we will briefly review each of these factors,
and how they can contribute to overall energy balance. When quantifying the energy of
metabolism, scientists will often use a particular unit of measurement, referred to as the
calorie. Energetically speaking, one calorie is the amount of heat required to raise the
temperature of one liter of water by one degree Celsius. The energy stores in food are
quite high relative to a calorie, so in the context of energy balance, the term kilocalorie
(Calorie) will be used. Total energy expenditure is often described in Calories, with
expenditure per unit of time referred to as the metabolic rate.
Many factors influence overall metabolic rate, including age, sex, height, and
overall health (Spiegelman & Flier, 2001). Thus the most common methodology for
evaluating it specifies certain standardized conditions and measures what is known as the
basal metabolic rate (BMR). This is often termed the ‘metabolic cost of living’ and refers
to the energy necessary to sustain autonomic cellular processes in major energetically
demanding tissues such as heart, liver, kidney and brain. When in the basal condition, the
subject is at mental and physical rest in a room at a comfortable temperature, and is often
in the fasted state (i.e. having not eaten for at least 12 hours). When animals are digesting
16
food, but not expending energy through other means (such as physical activity), energy
expenditure is defined as ‘resting metabolic rate’ (RMR). Several factors are known to
influence BMR (Spiegelman & Flier, 2001). The thyroid hormones are the single most
important determinant of BMR, regardless of an organism’s size, age, or sex.
Hyperthyroidism (pathological secretion of thyroid hormone) increases oxygen
consumption and heat production, with the overall result of net cellular catabolism and
loss of body weight (Kahaly, 2010). Another major regulator of BMR is the
hormone/neurotransmitter epinephrine. Release of epinephrine occurs during sympathetic
stimulation of the autonomic nervous system, resulting in increased heart rate, contraction
of blood vessels, and dilation of air passages. Epinephrine stimulates glycogen and
triglyceride catabolism, since ATP splitting and energy liberation occur during
breakdown and re-synthesis of these metabolic substrates. Thus, we can see that BMR is
determined through the interaction of several influencing factors, and can control an
organism’s basic level of energy expenditure.
‘Non-resting’ conditions are those that occur during periods of activity. The factor
that can most increase metabolic rate (energy expenditure) is altered skeletal muscle
activity. Even minimal increases in muscle contraction can significantly increase
metabolic rate, and strenuous exercise might raise energy expenditure more than
fifteenfold (Jones & Killian, 2000). While we have already described metabolic
differences in fiber-type, and will later discuss the metabolic potential of muscle, it is
important to know how physical activity (particularly exercise) can influence energy
homeostasis, and the role of other important organs in mediating these effects. During
exercise, large quantities of fuels must be mobilized to provide required energy for
17
muscle contraction. These fuels include circulating plasma glucose, fatty acids, and
glycogen (stored glucose polymers). A major role is played by the liver in regulating this
process (Jones & Killian, 2000). The plasma glucose used during exercise is largely
provided by the liver, both by breakdown of its glycogen stores, and by gluconeogenesis
(synthesis of new glucose molecules). Lipolysis of triglycerides creates a surplus of
glycerol for the liver, and fatty acids for metabolism by muscle. Changes in plasma
glucose during exercise are small in the short-term, but long-term exercise does lead to
transient decreases in blood glucose, due to an inability of hepatic gluconeogenesis to
keep up with glucose utilization. The metabolic profile seen in an exercising person
(increased hepatic gluconeogenesis, triglyceride breakdown, fatty acid utilization, etc) is
similar to that seen in a fasting person. This relates back to the main equation of energy
balance. An energy deficit (and its resultant metabolic profile) can be achieved through
increased energy expenditure, or decreased energy intake.
In earlier discussion on thermodynamics, it was mentioned that liberated energy
could be either used for work or heat. It has been demonstrated that liberated energy from
the breakdown of organic molecules can be used to create ATP (which are split to
perform work). However, as thermodynamics tells us, energy can also be lost in the form
of heat. In the context of metabolism, this heat loss leads to discussion of the third and
final component of energy expenditure: adaptive thermogenesis. Physiologically,
thermogenesis is primarily controlled by brown adipose tissue (BAT). The evolutionary
importance of BAT and adaptive thermogenesis is clear: it functions to generate body
heat, and is particularly abundant in newborns and hibernating animals, which are
incapable of shivering (Enerback, 2010). Mitochondrial density is significantly greater in
18
BAT than WAT. It is through mitochondria that BAT is able to generate heat.
Mitochondria are the major energy generators in the cell, and function to produce ATP
through glycolysis and fatty acid oxidation. This is accomplished by movement of
protons across a gradient and into the mitochondrial matrix. However, protons may “leak”
back across the inner mitochondrial membrane in a manner not linked to ATP production.
This uncouples energy storage from oxygen consumption, a process referred to as
‘uncoupled respiration’. Inherently, a certain degree of uncoupled respiration is expected
to occur in mitochondria of all cells. However, this process is greatly accelerated in BAT,
due to high expression of uncoupling proteins (UCP), which function as specialized
protein channels that provide a route for protons to return across the mitochondrial
membrane. The end result is energy expenditure, but largely in the form of heat. Mice that
lack BAT are prone to obesity, suggesting that this mechanism is an important avenue of
energy expenditure (Lowell & Flier, 1997). Evidence now exists for several different
types of UCP. UCP1 is predominantly found in BAT, while UCP2 is expressed in a
variety of different tissues (Enerback et al., 1997). UCP3 is the major form found in
skeletal muscle, and muscle-specific overexpression of UCP3 results in leanness and
hyperphagia (Clapham et al., 2000). Thus, it seems as though thermogenesis occurs in
tissues other than BAT, and elucidating these processes will provide scientists with a
greater understanding of whole-body energy expenditure.
1.3.4. Energy Balance, Body Composition and Obesity
Understanding the key molecular players governing whole-body energy intake
and energy expenditure are key for translational study of body composition. Deregulation
19
of energy balance results in manifestation of changes in body composition, and
subsequently impacts overall health (Friedman, 2009). As mentioned, when energy
expenditure exceeds energy intake, the result is cell death due to insufficient energy. If
persistent, death of the organism will occur. However, the more pressing problem (at least
in Western society) is a deregulation in energy balance caused by an increase in energy
intake, and subsequently limited energy expenditure. The result is increased adiposity (fat
accumulation), higher levels of circulating glucose, and increased glycogen storage. Over
time, this results in obesity, type 2 diabetes mellitus (T2DM), and metabolic dysfunction.
Obesity (excess fat accumulation) is a major health concern. Those who suffer
from obesity are at significantly greater risk to develop heart disease, cancer, and T2DM.
The prevalence of obesity has increased steadily, particularly in developed, Western
countries (Friedman, 2009). While the etiology of obesity is complex (linking both
genetic susceptibility and environmental factors), at its root, all instances of obesity are
characterized by a dysfunction in energy balance. A chronic imbalance between energy
intake and energy expenditure can lead to an increase in both fat cell size and fat cell
number. Therefore, a successful obesity therapy must impact energy intake, energy
expenditure, or both.
The use of drugs that affect energy intake have been met with limited success.
After the isolation of leptin came the promise that exogenous administration of this
hormone might be capable of inhibiting appetite and inducing weight loss (Friedman &
Halaas, 1998). However, long-term treatment of subjects with leptin results in reduced
efficacy, due to the development of leptin resistance (Mantzoros & Flier, 2000). Scientists
believe that leptin resistance (due to higher levels of the circulating hormone from
20
increased adipose tissue) may exacerbate energy intake in obese individuals. Thus,
scientists have instead focused upon the discovery of leptin receptor agonists. This
avenue has yielded some surprising insights, as some cytokines have been shown to have
leptin-like effects, suppressing appetite and promoting weight loss. Most notable among
these is ciliary neurotrophic factor (CNTF), a neurocytokine that has been shown to
reverse obesity in leptin-resistant mice (Gloaguen et al., 1997), although the mechanism
of action of these cytokines is unknown. With that in mind, some scientists have
suggested that a more appropriate method might be to simply target hypothalamic
neuropeptides themselves, as the secretion of these neuropeptides is what is controlled by
the endocrine signals. To that end, there has been development of small-molecule
antagonists for NPY, the major orexigenic neuropeptide; as well as agonists for the
MC4R receptor, which has anorexigenic effects on appetite (Gehlert, 1999). Only time
will tell whether success in regulation of energy balance can be efficiently achieved
through modulation of feeding behaviour.
On the other hand, some believe that treatments geared toward increasing
metabolic rate (and energy expenditure) may be a more promising avenue for the
treatment of metabolic disease. Feeding behaviour is still influenced by social and
environmental factors, but this is less applicable for regulation of metabolic rate. As body
weight declines, metabolic rate subsequently decreases as well, and therefore a strong
treatment may be used to either prevent this drop, or increase overall metabolic rate.
Although thyroid hormone is the main regulator of BMR, use of exogenous (i.e.
supraphysiological) doses of thyroid hormone can have negative side-effects, including
goiter, loss of lean muscle mass, and blindness (Kahaly, 2010). Conversely, physicians
21
often recommend that patients who suffer from hypometabolism take part in resistance
exercise training regimens, with the purpose of building lean muscle (Harrison &
Leinwand, 2008). This is because skeletal muscle is a major metabolically demanding
tissue, and increased muscle mass is sufficient to induce significant changes in resting
metabolism. Later discussion in this review will focus on how skeletal muscle can
influence systemic metabolism, but for now it is worth mentioning that scientists have
begun to focus on manipulation of this tissue as a possible method of regulating energy
expenditure and overall energy homeostasis. Factors that influence skeletal muscle
growth (such as androgens) may be looked upon as treatment measures. By targeting the
energy balance equation, scientists are attempting to find novel methodologies for treating
metabolic disease.
1.4. The Androgen System
The term ‘androgens’ is used to collectively refer to the class of male sex
hormones. Most prominent of the androgens are the hormones testosterone (T) and
dihydrotestosterone (DHT), which are known to have diverse functions in mammalian
physiology. These can be contrasted with the estrogens, the class of female sex hormones
(including 17β-estradiol and estrone). Much is known about androgens, and a large
amount of work has elucidated details regarding their pharmacology, synthesis and
mechanism of action.
22
1.4.1. Androgen Pharmacology – Structure and Production
Androgens fall into a class of hormones termed ‘steroid hormones’. Included in
this class are not only the male sex hormones, but also the female sex hormones,
glucocorticoids and mineralocorticoids. All hormones within this class have certain
characteristics with regards to their structure and stability (McEwen, 1992). Steroids are
lipid molecules, and all of these hormones are composed of four interconnected rings of
carbon atoms. A few polar hydroxyl groups may be attached to these rings, however the
overall number of hydroxyl groups are not numerous enough to make steroids into polar
molecules. For this reason, sex hormones are not soluble in the blood, but rather need to
be transported by binding to an albumin (water-soluble protein), known as sex hormone-
binding globulin (SHBG). However, the lipid, non-polar nature of steroid hormones
allows them to diffuse easily across the cell membrane, unlike peptide hormones (a
separate class that includes insulin and glucagon).
Steroidogenesis describes the biochemical process by which steroids are generated
from cholesterol, and subsequently transformed into other steroids (Aizawa et al., 2008).
There is no specific gene that codes for androgen, but rather androgens are derived by
enzymatic reactions from lipid precursors. In the case of de novo androgen synthesis (in
males), this process takes place in the leydig cells of the testes and (to a significantly
lesser degree) in the adrenal cortex. Cholesterol precursors are initially cleaved by
CYP11A, a mitochondrial cytochrome P450 oxidase. The loss of 6 carbon atoms from the
cholesterol molecule results in the prohormone pregnenolone. The cleavage of two more
carbons from this precursor by the CYP17A enzyme yields a variety of 19-carbon
steroids. Subsequent oxidation of the 3-hydroxyl group by the enzyme 3-β-
23
hydroxysteroid dehydrogenase produces androstenedione. Finally, in the rate-limiting
step, this intermediate is reduced by 17-β-hydroxysteroid dehydrogenase to yield T. Thus,
T is not produced directly by protein synthesis from a gene, but rather it is the expression
of these important enzymes that regulate its biosynthesis.
What is important to remember is that steroid hormone biosynthesis does not end
with T production. T itself, despite having its own functions, also acts as a precursor to
other sex hormones. In fact, T serves as the major precursor for the synthesis of estrogens.
This reaction is mediated by the enzyme aromatase, which oxidizes the T molecule and
removes a methyl group, resulting in the production of estradiol. Female aromatase-null
(ArKO) mice demonstrate significantly elevated T levels, significantly reduced estrogen
levels, and severe defects in sexual differentiation, such as underdeveloped external
genetalia and uteri (Fisher, Graves, Parlow & Simpson, 1998). Expression of aromatase is
highest in the gonads, but it is also found in brain, adipose tissue, placenta and
endometrium.
Another major metabolite of T is the androgen dihydrotestosterone (DHT), which
is largely expressed in prostate gland, testes, hair follicles, and adrenal glands (Zitzmann,
2009). Conversion of T to DHT occurs via the enzyme 5-α-reductase, which reduces the
4,5 double-bond in the T molecule through the addition of two hydrogen atoms. DHT
functions in a similar fashion as other androgens, but differs from T in that it is
considered to be more potent. This is for two primary reasons. First, DHT has stronger
affinity for androgen receptor (AR), which is the nuclear receptor that mediates androgen
action; as well as for SHBG, which is the major transporter of androgens through the
bloodstream. Secondly, unlike T, DHT cannot be aromatized to estrogens, and thus does
24
not serve as a precursor, which would limit its yield. DHT’s increased potency has made
it a target for treatment of disorders associated with hyperandrogenicity, such as benign
prostatic hyperplasia (prostate growth) and androgenic alopecia (male-pattern baldness).
This is typically accomplished through the use of a 5-α-reductase inhibitor, which inhibits
the enzyme, and subsequently reduces conversion of T to DHT (Walsh, 2010).
Secretion of androgens from the testes is under strict homeostatic control in
normal males (Tobet, Bless & Schwarting, 2001). Similar to release of glucocorticoids
from the adrenal glands, androgen secretion is controlled by hormonal signals from the
brain (specifically the hypothalamus and the pituitary gland). Together, these hormones
constitute a feedback loop that allows them to regulate one another. Gonadotropin-
releasing hormone (GnRH) is released from hypothalamic nuclei, which then travels to
the anterior pituitary gland. Stimulation of the anterior pituitary by GnRH results in the
release of the gonadotropins: luteinizing hormone (LH) and follicle-stimulating hormone
(FSH). As the name suggests, the gonadotropins stimulate the gonads. In males, LH acts
upon the leydig cells of the testes, and contributes to the production of T and other
androgens. In contrast, FSH stimulates maturation of the seminiferous tubules of the
testes in males, and also acts upon sertoli cells, functioning to promote spermatogenesis.
LH also indirectly plays a role in spermatogenesis, as it stimulates T production, and
androgens are involved in regulating this process. Also, T acts upon hypothalamic AR in
order to reduce secretion of GnRH. This negative feedback maintains concentrations of
these hormones in homeostatic balance, thus allowing for fulfillment of normal
androgenic function.
25
1.4.2. Androgen Receptor
As mentioned, androgens exert their functions by binding to the androgen receptor
(AR). Like other sex steroid hormone receptors, AR is a ligand-inducible transcription
factor belonging to the nuclear receptor subfamily (Chang, Kokontis & Liao, 1988). The
gene encoding AR is located at position q11-12 of the X chromosome, and is comprised
of eight exons. It encodes a protein composed of 919 amino acids, with a molecular mass
of approximately 110 kDa (Lubahn et al., 1988).
The structure of the AR protein itself is not too dissimilar from that of other
nuclear receptors (Chawla, Repa, Evans & Mangelsdorf, 2001). It consists of four
functional domains: an amino-terminal transactivation domain (NTD, encoded by exon
1), containing stretches of glutamine, proline and glycine; a highly conserved central
DNA-binding domain (DBD, encoded by exons 2 and 3); a hinge region (encoded by the
5’ part of exon 4); and a carboxy-terminal ligand-binding domain (LBD, encoded by the
3’ part of exon 4, as well as exons 5-8), to which the androgen binds.
Androgens (such as T and DHT) are lipid-based molecules, and as such, can
easily diffuse across the lipid-bilayer membrane of the cell (without the aid of a transport
protein). Upon binding of the androgen to the LBD of AR, an important sequence of
events is initiated (Lee & Chang, 2003). When not bound to its androgen ligand, AR is
maintained in a stable conformation in the cytoplasm through association with chaperone
molecules. The specific chaperone molecule involved is usually dependent upon the cell
type (for example, heat shock protein 70 maintains AR conformation in prostate cells –
see He et al., 2004). Upon binding of the androgen ligand, the conformation of AR
undergoes change and the chaperone molecule disassociates (Chang et al., 1995). AR
26
then forms a homodimer, which translocates to the nucleus. This homodimer then binds
to specific DNA motifs, termed androgen response elements (AREs), which then results
in recruitment of various coactivators to form the transcription complex (Roche, Hoare &
Parker, 1992). These AREs are situated in or near various target genes, and largely fall
into two classes. The ‘classical’ AREs are also recognized by other steroid receptors
(Claessens et al., 2001), while ‘selective’ AREs are only responsive to activation by AR
(Shaffer et al., 2004).
Expression of AR is found in many tissues of the body, albeit to different degrees
(Ting & Chang, 2008). The highest expression of AR is believed to be found in the
adrenal gland, followed by epididymis, prostate, skeletal muscle, kidney, liver, and heart.
With respect to skeletal muscle, higher expression of AR is found in androgen-dependent
muscles (such as the levator ani) as compared to non-androgen-dependent muscles, such
as the extensor digitorum longus (EDL) (Monks, Kopachik, Breedlove & Jordan, 2006).
AR expression is also controlled by androgens themselves. Androgens have been shown
to regulate both AR mRNA as well as protein. However, it seems as though the effects
that androgens have on AR expression (i.e. whether they increase it or decrease it) are
tissue-dependent. Treating human prostate cancer cells with T reduces AR mRNA almost
two-fold (Trapman et al., 1990). Conversely, androgen action increases AR expression in
human hepatocellular carcinoma cells (Wiren et al., 1997), as well as in rat hippocampus
(Kerr, Allore, Beck & Handa, 1995). In skeletal muscle, it appears that both T and DHT
increase AR mRNA expression (Lee & Chang, 2003).
The binding of the homodimer androgen-AR complex with AREs (and subsequent
modulation of gene expression) characterizes the most well-understood mechanism of AR
27
function. However, recent evidence has uncovered non-transcription related (termed
‘non-genomic’) functions of AR (Baron et al., 2004). These non-genomic mechanisms
typically involve interactions of AR with other cell-signaling pathways. This includes
increases in intracellular calcium, and activation of various kinase enzymes, such as
protein kinase A, protein kinase C, and MAP kinase; leading to diverse cellular effects,
such as smooth muscle relaxation, neuromuscular and junctional signal transmission, and
neural plasticity (Heinlein & Chang, 2002). These T-dependent effects on second
messenger signals are unaffected by inhibitors of transcription and translation, and occur
within seconds to minutes, which are time courses that are believed to be too rapid to
represent changes in gene expression (Fix, Jordan, Cano & Walker, 2004). Thus it
appears that AR does not simply exert itself through modulation of gene expression
(although this does represent its major function), but also plays a role in regulation of cell
signaling.
1.5. Androgens, AR and Muscle
As mentioned previously, androgens have notable effects on body composition.
Perhaps most well-studied of these effects is the anabolic actions of androgens on muscle.
Muscle has long been known to be a prominent androgen target. AR expression has been
demonstrated in all three types of muscle: skeletal, cardiac and smooth (Lee & Chang,
2003). The functions of AR in these types of muscle, and the subcellular mechanisms
involved, are amazingly diverse.
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1.5.1. Anabolic Functions of AR in Muscle
The classical role of muscle AR is its anabolic function. It has largely been
demonstrated that androgen/AR action is responsible for development and maintenance of
muscle fibers (Bhasin et al., 1997; MacLean & Handelsman, 2009). There is considerable
evidence suggesting that androgens promote not only myogenesis by acting upon
progenitor satellite cells, but also mediate hypertrophy of mature individual muscle fibers.
While muscle growth during mammalian development is largely due to myogenesis and
the creation of new muscle fibers, growth in adulthood is largely believed to be due to
hypertrophy of existing fibers (Sinha-Hikim et al., 2002). Various molecular and cellular
mechanisms have been implicated in mediating each of these effects, in addition to
overlapping mechanisms that control both myogenesis and fiber hypertrophy.
Singh et al. (2003) showed that T has the capability to promote differentiation of
mesenchymal pluripotent stem cells toward the myogenic lineage, thereby driving
myogenesis. Further research has shown that similar effects can be found with regards to
development of satellite cells, when treated with T. Satellite cells are found between the
basal lamina and the plasma membrane of muscle fibers (Mauro, 1961). These specialized
progenitor cells have the ability to differentiate into muscle fibers, or fuse and augment
existing fibers. In this way, they can be likened to adult stem cells, and represent the
major source for the addition of new myonuclei into the growing muscle fiber. Satellite
cells are maintained in a quiescent state, until activated by external signals, which then
promote myogenesis (Tedesco, Dellavalle, Diaz-Manera, Messina & Cossu, 2010).
There are several lines of evidence that have been used to show that androgens
can act as an external signal to stimulate the proliferation of satellite cells in muscle. First,
29
AR expression has been readily demonstrated in satellite cell cultures, suggesting that
these precursor cells do serve as androgen targets (Doumit, Cook & Merkel, 1996). Work
with cultured human satellite cells has shown that AR protein expression increases when
cells are exposed to T (Sinha-Hikim, Taylor, Gonzalez-Cadavid, Zheng & Bhasin, 2004).
Powers and Florini (1975) were the first to successfully demonstrate that T, but not
estradiol, was capable of stimulating the mitotic activity of satellite cells in established
myoblast culture systems. Similarly, Sinha-Hikim and colleagues (2003) showed that, in
men, T supplementation led to a significant increase in the total number of satellite cells
found. Thus, effects of T on myogenesis (and proliferation of myogenic factors) have
been well established.
Whether androgens are capable of regulating hypertrophy of adult myofibers is
less clear. Most of the work in this context has been performed on human populations,
and have involved exogenous T-treatment, making it difficult to tease out effects on
myogenesis from those on muscle hypertrophy. Some evidence indicates that T induces
the hypertrophy of both type I (oxidative) and type II (glycolytic) fibers. When comparing
muscle biopsies from athletes who abuse anabolic steroids against those who have not
used such supplementation, the greatest disparities are seen in type II muscle fibers (Kadi,
2008). However, this belief is rather controversial, and there is some evidence suggesting
that androgens may have their primary effects on type I fibers. In comparison of biopsies
taken from trapezius muscle of androgen users and controls, the largest disparity is seen
between type I fibers, although there is also a significant difference in size of type II
fibers (Kadi, Eriksson, Holmner & Thornell, 1999). Similar differences are also seen in
the vastus lateralis muscle (Eriksson, Kadi, Malm & Thornell, 2005). It is believed that
30
this more robust effect on type I fibers is due to the fact that these fibers are more
sensitive to the effects of T treatment (Hartgens & Kuipers, 2004). For example, Sinha-
Hikim and colleagues (2002) found that type II fibers required twice as much exogenous
T as type I fibers in order to demonstrate hypertrophy. However, the above study also
found no differences in fiber type proportion, a finding that is disputed by related studies
(Kadi et al., 1999). Thus, specific effects of androgens upon muscle fiber type (both in
terms of proportion of total fibers and hypertrophy of individual fibers) remain unclear.
Further evidence indicates that activation of AR is necessary for promotion of
androgen’s anabolic functions upon skeletal muscle. Blockade of AR through the use of
the AR-antagonist oxendolone suppresses muscle hypertrophy of rat muscle fibers (Inoue,
Yamasaki, Fushiki, Okada & Sugimoto, 1994). Also, knockout of AR (either
ubiquitously, or selectively in myocytes) results in muscular atrophy and reduced skeletal
muscle strength (MacLean et al., 2008; Ophoff et al., 2009). Similar to precursor satellite
cells, AR expression can be modulated by T administration, as mentioned previously.
AR-containing myonuclei in human skeletal muscle become more numerous after the use
of anabolic steroids (Kadi, Bonnerud, Eriksson & Thornell, 2000). These results were
consistent among biopsies taken from the neck (trapezius muscle), as well as the limb
(vastus lateralis).
These anabolic effects are seemingly not limited to skeletal muscle, and androgens
have also been shown to be important in promoting hypertrophy of cardiac muscle. In
vitro work performed on cultured human cardiomyocytes found that T administration
resulted in increased incorporation of [3H]phenylalanine into cardiomyocytes, as well as
atrial natriurectic peptide secretion – both of which are significant indicators of
31
cardiomyocyte hypertrophy (Marsh et al., 1998). These T-mediated effects were
subsequently abolished when the cultured cells were first treated with the anti-androgen
cyproterone acetate. These findings were then confirmed by in vivo work performed on
ARKO mice. Whole-body ARKO mice demonstrate impaired cardiac growth, and smaller
overall heart mass (Ikeda et al., 2005). In addition, various cardiac impairments are found
in these ARKO mice, including cardiac fibrosis and impaired ventricular function. Taken
together, these results indicate that androgen-AR signaling is necessary for normal
cardiomyocyte growth and heart function. However, it also suggests that
supraphysiological doses of T may result in pathological hypertrophy of cardiac muscle,
providing a potential explanation for the increased incidence of sudden cardiac death in
athletes that abuse anabolic steroids (Luczak & Leinwand, 2009). Relatively little
research has been conducted in this area, and any links between androgens and
hypertrophic cardiomyopathy (and associated heart disease) are surrounded by a large
degree of controversy.
1.5.2. Muscle AR and Neuromuscular Development
Androgens and AR are important regulators of sexual differentiation, and strongly
contribute to the development of sexually dimorphic muscles, as well as the motoneurons
that innervate them. Most well-studied of these muscle systems is the spinal nucleus of
the bulbocavernosus (SNB) neuromuscular system (Sengelaub and Forger, 2008). These
motoneurons project from the lumbar spinal cord into the striated muscles of the
perineum: the bulbocavernosus (BC) and levator ani (LA). The muscles themselves are
attached to the base of the penis, and their contraction mediates male sexual behaviour.
32
SNB neurons are larger and more numerous in males than females (Breedlove & Arnold,
1980). In rats, during gestation, both sexes possess the SNB motoneurons and target
muscles, but prenatal T secretion allows for the neuromuscular system to be maintained in
males, whereas females undergo apoptosis (Nordeen, Nordeen, Sengelaub & Arnold,
1985). Not surprisingly, the development of the motoneurons that innervate the sexually
dimorphic muscles (BC/LA) are also under the control of circulating steroid hormones
(Breedlove & Arnold, 1980). These motoneurons innervate the striated BC and LA
muscles, and mediate copulatory behaviour in males. Conversely, in females, the SNB
motoneurons develop prenatally, but undergo significant postnatal cell death with
significantly fewer neurons seen in adult females, as compared to age-matched males
(Nordeen et al., 1985). Thus, in adulthood, SNB motoneurons are larger in number and
size in males (Breedlove & Arnold, 1980; McKenna & Nadelhaft, 1986).
It is now well-accepted that the survival of SNB motoneurons and their target
musculature occurs in males due to actions of circulating androgens in early postnatal
development (Sengelaub & Forger, 2008). Activation of AR by androgens is necessary
for masculinization of the SNB system to occur, and evidence for this is demonstrated
through the use of androgen-insensitive Testicular feminization mutation (Tfm) male rats.
The use of Tfm male rats has supported the necessary role of androgens/AR in
development and masculinization of the nervous system (Zuloaga, Puts, Jordan &
Breedlove, 2008). Tfm males, which have non-functional AR, demonstrate a feminized
SNB system, further highlighting the fact that this effect is AR-mediated (Breedlove &
Arnold, 1981). Similarly, administering the potent AR-antagonist flutamide to neonatal
male rats also suppresses SNB development (Breedlove & Arnold, 1983). However,
33
despite the fact that activation of AR is a requisite of SNB development, it is not yet clear
which tissues or cells are necessary site(s) of androgen action.
Several lines of evidence indicate that AR in the perineal muscles themselves
(BC/LA) are the necessary site of androgen action. AR expression is significantly higher
in sexually dimorphic muscles (such as the LA), as compared to non-sexually dimorphic
muscles (such as the EDL), with enrichment at the neuromuscular junction (Monks,
O’Bryant & Jordan, 2004). Furthermore, AR immunoreactivity is found in perineal
muscles during the critical postnatal period for SNB development (Jordan, Padgett,
Hershey, Prins & Arnold, 1997). Removal of the BC/LA muscles on the first day of birth
results in a dramatic loss of SNB motoneurons by PND10, an effect that could not be
rescued by subsequent T treatment (Kurz, Cover & Sengelaub, 1992). Finally, injection of
flutamide (an AR antagonist) directly to the perineum of females results in fewer SNB
motoneurons than when flutamide is delivered systemically (Fishman & Breedlove,
1992). Taken together, these results suggest that AR in muscle fibers plays a critical role
in mediating development of the SNB neuromuscular system. However, in contrast to this
theory, Niel and colleagues (2009) found that transgenic expression of AR only in muscle
fibers of Tfm males was not capable of rescuing the SNB system in these mutants. Thus,
while significant evidence suggests the involvement of muscle AR in SNB development,
it appears that it is not sufficient, and AR in other tissue(s) may also be responsible for
this effect.
34
1.5.3. Spinal and Bulbar Muscular Atrophy
Up to this point, we have shown that androgens/AR have significant effects upon
normal muscular development in males. Genetic (XY) males that harbor mutations in the
AR gene (or who are unable to synthesize androgens) demonstrate severe defects in these
contexts. However, androgens are also explored in the field of neuromuscular pathology,
as they have become linked with a disease known as Spinal and Bulbar Muscular Atrophy
(SBMA), also known as ‘Kennedy’s Disease’.
SBMA is a progressive neurodegenerative disease, largely affecting males, which
is characterized by progressive weakness and motoneuron death (Kennedy et al., 1968).
Prevalence of SBMA is very rare, with manifestation of the disease in only 1/400,000
(Fischbeck, 1997). Genetic analysis of SBMA first found that the disease was X-linked,
through localization of the defective allele using polymorphism linkage analysis
(Fischbeck et al., 1986).
It is now understood that the development of SBMA is related to the number of
glutamine (CAG) repeats in the AR gene (Monks et al., 2008). Polyglutamine repeats
exist in the previously discussed NTD region of the AR gene. The existence of this repeat
region is thought to contribute to differences between individuals in androgen sensitivity.
Shorter AR repeats increase receptor transactivation, with longer repeats decreasing
transactivation. After the determination of the X-linked inheritance pattern of SBMA,
scientists eventually discovered that the disease is associated with an increased number of
CAG repeats in exon 1 of the AR gene (La Spada et al., 1991). Not only that, but the
number of glutamine repeats in the gene was predictive of age of onset, as well as
severity of the disease, with an increasing number of CAG repeats indicative of earlier
35
age of onset and increased severity (Atsuta et al., 2006; La Spada et al., 1992). Mouse
models that attempt to recapitulate this disease, through induction of glutamine repeats in
the AR gene, do show some of the symptoms of SBMA (Sopher et al., 2004).
The mechanism of how polyglutamine expansion in the AR gene subsequently
results in neurodegeneration and cellular death remains controversial. The prevailing idea
within the field is that the disease is neurogenic (Monks et al., 2008). That is, the
pathology manifests itself primarily in motoneurons, with the resultant toxicity causing
motoneuron death and subsequent muscular atrophy as a secondary result of this
motoneuron death (Monks et al., 2008). This is supported by in vitro work demonstrating
that polyglutamine expanded AR are sufficient to cause motoneuron death in cultured
motoneurons (Merry et al., 1998). However, recent work has cast doubt on this initial
premise. Most notably, polyglutamine expansion of the AR gene only in motoneurons of
transgenic mice is not capable of inducing symptoms of SBMA (Abel et al., 2001).
It now appears that SBMA may be myogenic in nature (i.e. it manifests itself
initially in muscle, with muscular atrophy resulting in motoneuron death). It has been
shown experimentally in an SBMA transgenic mouse model that muscular atrophy does
in fact precede motoneuron death (Yu et al., 2006). Furthermore, overexpression of AR
(without polyglutamine expansion) in muscle fibers creates a phenotype consistent with
most symptoms of SBMA (Monks et al., 2007). The resultant myopathy in this model is
associated with deregulation of gene expression in muscle fibers, and mitochondrial
abnormalities (Musa et al., In Prep). Thus, although SBMA appears to be AR-mediated,
the site-specific etiology of this disease remains unclear. Further characterization of
36
disease models, as well as patient populations, will be necessary in order for scientists to
gain a better understanding of this neuropathology, and subsequently design treatments.
In summary therefore, androgen action on skeletal muscle is important for
mediating a variety of processes. This includes increased muscle mass, which occurs
through androgen-mediated myogenesis and muscle hypertrophy. Furthermore, muscle
AR plays an important role in regulating development of innervating motoneurons, as is
exemplified by the SNB neuromuscular system. Finally, recent evidence seems to link
AR in myocytes to SBMA incidence, suggesting a potential myogenic cause for disease
onset.
1.6. Androgens, AR and Fat
The anabolic effects of androgens in mediating muscle growth have mostly been
well elucidated and understood. Less well studied, however, is how androgens regulate
body fat. In males, androgens are largely thought to reduce overall body fat (Bhasin et al.,
1996). In females, the effects of T and other androgens on body fat are clouded by the
influence of estrogens. Teasing apart the effects of these two hormone classes is
paramount to understanding exactly how androgens are capable of modulating whole
body fat.
1.6.1. Sex Differences in Adiposity
Although much of the work in the field of mammalian sexual differentiation has
focused upon dimorphisms in reproductive organs and the nervous system, scientists are
now finding that there are significant differences between males and females in amount
37
and distribution of body fat, as well as in the incidence of various metabolic disorders,
including obesity and T2DM. Women under the age of 50 have significantly reduced
likelihood of developing such diseases, but their prevalence increases markedly after the
onset of menopause (Ford, Giles & Dietz, 2002). Among adolescents, incidence of
metabolic disease is significantly higher among males than females (Ford et al., 2002).
Overall, the epidemiological data seems to suggest that ovarian hormones (ex: estrogens)
appear to be protective against metabolic disease because, prior to menopause, the
prevalence of these diseases is higher among males than females; however, after
menopause, females are more likely to develop these diseases (Shi, Seeley & Clegg,
2009). In addition, although older men are less likely to develop these diseases than older
females, older men are significantly more likely to do so than younger men. This suggests
a similar protective effect of androgens, since these hormones decline significantly with
age.
The increased risks due to obesity vary depending upon the location of adipose
accumulation (Bjorntorp, 1997). More specifically, concentration of adipose tissue
distributed in the abdominal (visceral) region carries a much greater risk for metabolic
disease than adipose accumulation in subcutaneous regions (Bjorntorp, 1997). This is
because different adipose depots have different properties. There are distinct sex
differences in adipose accumulation. Overall, females tend to have more subcutaneous fat
than males do (Lonnqvist, Thorn, Large & Arner, 1997), whereas accumulation of
adipose in visceral regions tends to occur to a greater degree in males (Mujica et al.,
2008). These effects also tend to change over the lifespan. As women age (particularly
after menopause, when estrogen levels decline), they tend to gain visceral adipose tissue.
38
As men age, they also tend to gain visceral adipose tissue, although this gain has been
shown to be correlated with a reduced level of androgens in the bloodstream (Shi et al.,
2009). Thus, it appears that sex hormones do play a profound role in mediating adiposity
in both males and females, although this regulation is drastically different.
In males, androgens largely promote decreases in adiposity (i.e. reduction in
adipose tissue accumulation). Lines of evidence for this effect come from both treatment
of adult males with exogenous T, as well as natural reductions in T that occur during
aging. T supplementation decreases fat mass in both hypogonadal and eugonadal men
(Wilson, 1988; Bhasin et al., 2001). Furthermore, these effects of T on fat mass are
strongly correlated with the dose of T provided, as well as circulating levels in the
bloodstream (Bhasin et al., 1997; Snyder et al., 1999). These effects also appear to be
depot-specific, as T supplementation in older men preferentially reduces fat mass in
visceral depots (Marin et al., 1996). Studies of hypogonadal men seem to indicate similar
associations. A Finnish cohort study demonstrated that men who developed obesity in
late-adulthood had a 2.6-fold increased risk of having hypogonadism (Laaksonen et al.,
2005). Among men suffering from Klinefelter Syndrome (who have hypergonadotropic
hypogonadism), the prevalence of obesity and T2DM is raised by a factor of four, as
compared to age-matched controls (Bojesen et al., 2006; Ishikawa et al., 2008). Thus,
studies in humans seem to strongly suggest that androgens largely function to reduce
adiposity, primarily from visceral depots. Many therefore believe that the increase in
adiposity seen in aging men is directly related to age-related decline in androgen
signaling.
39
The picture of sex hormones and adiposity is made significantly more complex
when one considers the role of estrogens (the female sex hormones). In order to more
fully contextualize the effects of androgens on adiposity, one must first understand how
estrogens contribute to this phenomenon. Similar to androgens, estrogens act upon their
own nuclear receptor (estrogen receptor – ER), which also functions as a transcription
factor. As mentioned previously, estrogens are derived from T, in a reaction mediated by
the enzyme aromatase. Significant evidence suggests that estrogens also function to
reduce adipose accumulation in females. In women, adipose accumulation increases
significantly after the onset of menopause, when circulation of estrogens declines
(Gambacciani et al., 1997), and these increases can be reversed by treatment of females
with exogenous estrogen (Haarbo, Marslew, Gotfredsen & Christiansen, 1991). This
evidence is supplemented by clinical investigation of women suffering from Polycystic
Ovarian Syndrome (PCOS). In PCOS, the ovary of the woman secretes abnormally high
levels of androgens, resulting in negative feedback on the hypothalamus and significant
suppression of estrogen secretion. 50% of women with PCOS are obese, demonstrating an
increase in visceral adipose tissue (Dunaif et al., 1987). While this was originally taken to
be evidence that androgens actually increase adipose tissue, it is now better understood
that the increase in adiposity is a consequence of reduced estrogen signaling in PCOS
patients. This postulation is supported by evidence from rodents. Ovariectomy of female
rats results in increased visceral fat, while administration of exogenous 17-β estradiol
subsequently reduces this adipose accumulation (Clegg, Brown, Woods & Benoit, 2006).
Furthermore, ablation of ERα results in increased adiposity and body weight in female
mice (Heine et al., 2000). Interestingly, knockout of ERα also affected adiposity in males,
40
as male mice demonstrate increased fat accumulation in late adulthood. However, more
recent evidence suggests that the site-of-action for ER-mediated reduction in adiposity
appears to be the ventromedial hypothalamus, an area previously discussed to be a major
regulator of feeding behaviour. Selective knockdown of ERα in the hypothalamus
induces obesity and metabolic disease through hyperphagia (Musatov et al., 2007). Thus,
taken together, these studies demonstrate that action of estrogens have similar effects
upon adipose tissue as androgens. Furthermore, these effects do seem to be important in
both sexes, although to a larger degree in females. Lastly, it appears that estrogens
mediate their effects by regulating feeding behaviour, likely through interaction with
leptin (Asarian & Geary, 2002; Asarian & Geary, 2006).
1.6.2. Androgens and Adipocyte Development
While it appears that estrogens regulate adiposity by acting upon ERα in the brain
and modulating feeding, less is understood about how androgens exert their effects upon
adipose tissue. Generally speaking, reductions in adiposity are largely achieved through
two major methods (Shi et al., 2009). The first is through inhibition of adipocyte
differentiation and development. Adipocytes are derived from precursor cells (termed
‘preadipocytes’), and inhibition of their differentiation from these precursor cells would
subsequently reduce their accumulation. The second avenue by which to achieve a
reduction of adiposity is the more well-known mechanism of fat metabolism. Here
mature, developed adult adipocytes release triglyerides, which are lysed for the purposes
of deriving fatty acids. These released fatty acids subsequently undergo β-oxidation in
mitochondria of various tissues in order to harvest energy in the form of ATP.
41
When it comes to androgens, it is largely believed that these hormones exert their
effects on adipose tissue through the first pathway. That is, androgens are largely believed
to inhibit the development of adult adipocytes from their precursor cells (Gupta et al.,
2008). Indirect evidence from animal models has suggested that high androgen levels
modulate the proliferation and differentiation of preadipocytes differentially in specific
fat depots (Garcia et al., 1999). The difficulty associated with studying precursor cells in
an in vivo setting provides a rationale for why this possibility has largely been probed
from an in vitro perspective. Nevertheless, several lines of evidence have supported the
idea that androgens are capable of inhibiting differentiation and development of
preadipocytes into mature adult cells. Treatment of epididymal (reproductive)
preadipocyte cells with either T or DHT inhibits the activity of an adipose-specific
isoform of the enzyme glyceraldehyde-3-phosphate-dehydrogenase (GAPDH)
(Dieudonne et al., 2000). Some work in this context on mature adipocytes shows that T-
treatment actually suppresses lipoprotein lipase (LPL) activity and lipid uptake, thus
inhibiting adipocyte growth. This provides a third potential mechanism by which
androgens might inhibit increased adiposity. Singh and colleagues (2003) found treatment
of mouse C3H 10T1/2 pluripotent cells (which are capable of differentiation into muscle,
fat, cartilage, and bone cells) with T reduced the number of mature adipocytes found. Pre-
treatment of cells with bicalutamide (a potent AR-antagonist) blocked the T-induced
inhibition of adipogenesis. Therefore, the preceding studies suggest that androgens are
capable of reducing adipose tissue by acting directly upon AR in preadipocytes and
inhibiting their development into mature cells.
42
AR expression is well-documented in adipocytes and pre-adipocytes of both
humans (Dieudonne et al., 1998) and rodents (Dieudonne et al., 2000). It is believed that
AR’s function as a transcription factor may be harnessed in adipocytes and preadipocytes,
through downregulation of adipogenic genes (Matsumoto, Takeyama, Sato & Kato,
2003). Mature adipocytes treated with T demonstrate reduced expression of LPL and
GAPDH, which both promote adipocyte growth. Treatment of mouse C3H 10T1/2
pluripotent cells with T (Singh et al., 2003) reduced expression PPARγ2 mRNA and
protein, as well CCAAT/enhancer binding protein α (C/EBP). Both proteins are key
transcription factors necessary for adipogenic differentiation (Rosen & Spiegelman, 2000;
MacDougal & Mandrup, 2002). In a separate study, Singh et al. (2006) treated mouse
pre-adipocyte 3T3-L1 cells with T, and measured gene expression of various adipogenic
genes. Here, the scientists were able to recapitulate the downregulation of both PPARγ2,
as well as C/EBP. They also examined the Wnt gene family, which have been implicated
largely in the control of preadipocyte differentiation and adipogenesis (Cadigan & Nusse,
1997). It was found that activation of AR affected expression of various Wnt proteins in
the preadipocytes, suggesting a possible mechanism by which AR might modulate
adipogenesis. Once again, these effects were abolished by pretreatment of the cells with
bicalutamide. Overall, the evidence seems to suggest that androgen-mediated reduction in
adiposity occurs due to decreased expression of adipogenic factors by AR.
1.7. Where The Action Is: Lessons Learned from ARKO
Previous in vitro work concerning the role of AR in modulation of body
composition has demonstrated rather direct effects of AR action. As described previously,
43
androgen-activation of AR has been shown to induce commitment of pluripotent cells to
the myogenic lineage (Singh et al., 2003). In addition, androgens have been shown to act
upon pre-cursor myoblast cells, promoting their development into mature myocytes
(Singh et al., 2009). Also, as mentioned, androgens have been shown to have opposite
effects on adipose, inhibiting differentiation of pluripotent stem cells into the adipogenic
lineage (Singh et al, 2006), and largely preventing development of pre-adipocytes into
mature cells (Gupta et al, 2008). Thus, these avenues of research have largely supported
the notion that androgens have direct effects upon their target tissues, and play a major
role in either promoting or inhibiting cellular differentiation and development.
While the previously mentioned studies have been fruitful in demonstrating in
vitro effects of androgens on various tissues (across various species), they are by
definition limited by the fact that they are conducted in isolation of other physiological
systems and cells. The physiology of mammalian organisms is inherently complex, with
tremendous interaction between various systems. For that reason, scientists ideally wish
to study physiological systems in vivo (i.e. in living organisms), in order to observe how
manipulations within one system can affect whole-body function. The advent of gene
targeting made such investigation possible. Traditional gene targeting uses homologous
recombination in order to alter a gene. Changes made include whole-gene deletion
(knockout), removal of portions of the coding region of a gene, addition of a gene
(transgenic technology), and induced point mutations. These genetic alterations can be
ubiquitous (i.e. occurring in all cells throughout the organism), or tissue-specific. With
regards to AR, gene targeting provides a powerful tool for understanding of sites of
androgen action. As discussed, AR expression is abundant in various tissues (Lee &
44
Chang, 2003). By altering AR expression within a particular tissue (via either loss- or
gain-of-function), scientists can study the role of that tissue-specific AR, and its
contribution to various phenotypes (Kerkhofs, Denayer, Haelens, & Claessens, 2009).
1.7.1. Whole-Body Androgen Receptor Knockout
Using gene-targeting technology, the first androgen receptor knockout (ARKO)
mouse was produced in 2002 (Yeh et al., 2002). For decades prior to this advancement,
Testicular feminization mutation (Tfm) rats and mice had served as the classical animal
models in understanding loss of AR function, albeit primarily in males (Yarbrough et al.,
1990; Lyon and Hawkes, 1970). Since the AR gene is located on the X chromosome, and
is vital for male fertility, the scientists were unable to use traditional knockout
technology, and instead created a conditional knockout model. This was done through the
use of a cre-lox strategy. Such a system uses the expression of the P1 phage cre
recombinase (Cre) enzyme, which catalyzes the excision of DNA located between
flanking loxP sites (Holt and May, 1993). By using this methodology, researchers could
generate male founder mice carrying the floxed AR gene, who could then be mated with
female mice ubiquitously expressing Cre (through the use of the CMV promoter).
Resultant offspring would express the ARKO mutation in all tissues.
Upon examination of the initial ARKO mice, several striking phenotypes were
noted (Yeh et al., 2002). Similar to Tfm mice (Lyon and Hawkes, 1970), the external
genitalia of male ARKO mice show an ambiguous or feminized appearance. Agenesis of
the vas deferens, epididymis, seminal vesicles and prostate were reported, along with
significant decrease in the size of testes.
45
Interesting findings were reported with regard to body composition in ARKO
mice. Yeh and colleagues (2002) found that cancellous bone volumes are lower in the
ARKO mice than in both female and male WT littermates. Furthermore, osteoclast (cells
involved in bone resorption) levels were higher in the ARKO mice, and they
demonstrated osteopenia. Similar effects of AR on bone resorption were reported by a
separate research group (Kawano et al., 2003). Here it was shown that whole-body
ablation of AR in males resulted in reduced trabecular and cortical bone mass, but no
change in bone shape. These findings were not found in ARKO females. Taken together,
these results provide strong evidence that androgen/AR action is important for
suppression of bone resorption.
Examination of skeletal muscle physiology in ARKO mice seemed to confirm
previously postulated ideas regarding the role of AR in muscle. As stated previously, it is
largely believed that androgens have anabolic effects on muscle mass, resulting in muscle
fiber hypertrophy (Ting and Chang, 2008). Others have suggested that, in addition to
mediating hypertrophy of individual muscle fibers, androgens act to promote myogenesis
and development/differentiation of mature myocytes from precursor cells (Singh et al.,
2009). MacLean and colleagues (2008) found marked change in skeletal muscle of
ARKO mice. Muscle mass is largely decreased in ARKO males, with no such differences
seen in ARKO females. Furthermore, muscle function appears to be impaired in ARKO
males. Isolated type II (EDL) muscles were found to demonstrate reduced maximum
tetanic force. This study largely demonstrates that peak skeletal muscle mass and function
in ARKO males is achieved through androgen/AR action.
46
Perhaps most interesting of the phenotypes observed in ARKO mice relates to
adipose tissue and metabolism. The original study of ARKO mice found that overall body
weight in ARKO males closely resembled that of WT and ARKO females (Yeh et al.,
2002), with overall body fat and adipocyte size actually lower in the 8-week old mutant
males, as compared to WT brothers. These findings led the authors to postulate that AR
may actually be necessary for adipogenesis. However, these same scientists later recanted
this possibility once they had followed the male ARKO mice into late adulthood (Lin et
al., 2005). Aging ARKO males were found to demonstrate accelerated weight gain, even
overtaking WT males after 20 weeks of age. This weight gain resulted in excess adiposity
and obesity in later adulthood. Onset of obesity was also co-morbid with hyperglycemia,
progressively reduced insulin sensitivity and impaired glucose tolerance, suggesting that
adult ARKO males also develop T2DM. This late-onset obesity in ARKO males was
confirmed by a separate study (Fan et al., 2005). Here it was reported that ARKO males
also had lower levels of basal metabolism (as demonstrated through reduced oxygen
consumption), in addition to reduced rates of spontaneous activity. Taken together, the
above results all seem to suggest a model by which androgen activity is important for
reduction of adiposity, at least in late adulthood. T levels in males naturally decrease as
they age (Bhasin et al., 1996), which can partially account for increases in adiposity due
to aging (Shi et al., 2009).
1.7.2. Tissue-Specific Androgen Receptor Knockout
While the previously discussed research using ARKO mice has been fruitful, it
has largely just confirmed what has been believed about the role of androgens and body
47
composition in males. That is, T is associated with increases in lean muscle mass, and
decreases in overall fat mass (Bhasin et al., 1996; Forbes et al., 1992). These studies still
do not indicate which tissues and cells are important for the effects of androgens on body
composition. An innovative way to tackle this problem is through the use of tissue-
specific AR knockout. One advantage behind the nature of ARKO mice (i.e. the floxed
AR gene) is that they can be mated with animals having tissue-specific Cre-recombinase
expression (Yeh et al., 2002). Such a mating would produce offspring that have null-AR
in the Cre-expressing tissue. This provides a very powerful tool for the study of
androgen-AR function in specific tissues. To date, these tissue-specific ARKO mice have
provided great insight into the distinctive roles of AR in mammary glands (Yeh et al.,
2003), reproductive tissue (Hu et al., 2004; Zhang et al., 2006; Wu et al., 2007) and the
immune system (Chuang et al., 2009; Lai et al., 2009).
While many questions remain regarding site-of-action for androgenic effects on
body composition, tissue-specific ARKO have provided significant insight in this realm.
Scientists first aimed to decipher the androgenic site for reduction of adipose tissue. As
discussed previously, it is largely believed that androgens are capable of reducing WAT
by acting upon AR in adipocytes themselves, inhibiting the differentiation of precursor
pre-adipocytes into mature cells, and down-regulating expression of adipogenic factors
(Singh et al, 2006; Gupta et al, 2008). Such a mechanism may account for the late-onset
obesity seen in male ARKO mice. To test this principle, ARKO mice were developed
with selective knockout of AR in adipocytes (aARKO; Yu et al., 2008). Interestingly,
body weight and adiposity of male aARKO mice did not differ from age-matched WT
brothers, and these animals did not develop obesity in later adulthood. This study
48
revealed that the characteristic obesity seen in adult ARKO mice was not due only to loss
of AR in adipocytes. This issue was further probed by knockout of AR in hepatocytes
(hARKO; Lin et al., 2008). The liver is a major physiological site of fatty-acid oxidation,
and impaired hepatocyte function is associated not only with localized fatty-acid
accumulation (hepatic steatosis), but also obesity (Kammoun et al., 2009). However,
selective knockout of AR in hepatocytes also failed to recapitulate the obesity phenotype
found in whole-body ARKO mice. When the mice were put on a high-fat diet, hARKO
male mice developed hepatic steatosis and insulin resistance (likely as a consequence of
the steatosis), which was not found in WT brothers. These findings suggest that liver AR
may be important for local oxidization of fatty acids, but (similar to adipocyte AR) are
not sufficient in mediating androgenic reduction of adiposity.
To identify the role of AR in muscle, Ophoff and colleagues (2009), developed
ARKO mice lacking AR in myocytes (mARKO mice). Male mARKO mice demonstrate
reduced body mass and lean body mass (LBM). While there was a marked reduction in
size of the androgen-dependent levator ani (LA) muscle in male mARKO mice,
differences in other individual muscles were either very small or non-existent. Scientists
also found no differences in muscle contractile function, although they noted that there
was a significant conversion of muscle fiber type from fast to slow-twitch in male
mARKO mice. This study provided strong evidence that myocyte AR is important for
regulation and maintenance of skeletal muscle mass and fiber-type, but not function or
strength. Thus, taken together, tissue-specific ARKO has revealed that myocyte AR is
important for maintenance of mature skeletal muscle, but has been unable to identify a
site-of-action for androgenic reduction in adiposity.
49
1.8. The Metabolic Potential of Muscle
Discussion of mammalian metabolism largely focuses upon organ systems as its
major players, including the brain, the liver, and the pancreas. For this reason, skeletal
muscle has been largely ignored as a prominent contributor to systemic metabolism.
However, in the last several decades, scientists have become acutely aware that changes
in skeletal muscle mass (and even atrophy/hypertrophy of individual fibers) can
drastically alter whole-body metabolism and energy homeostasis (Harrison & Leinwand,
2008). Muscle is a major storage site for glycogen, and a prominent target for insulin and
serum glucose uptake. Furthermore, as mentioned, catabolism of energy substrates by
skeletal muscle accounts for a significant proportion of systemic metabolism. We also
find that this metabolic capability is largely mediated by skeletal muscle mitochondria,
the oxidative energy builders within the cell.
1.8.1. Skeletal Muscle Metabolism
As briefly discussed previously, energy utilization in skeletal muscle is largely
dependent upon the muscle fiber-type in question. Type I (slow-twitch) muscle fibers are
characterized by a high mitochondrial content, while type II (fast-twitch) muscle have
less mitochondrial content, relying moreso upon glycolysis in order to meet energy
demands (Scott et al., 2001). In mammals, the type II fibers can be further subdivided into
the more oxidative type IIa fibers, and the more glycolytic type IIb fibers.
Muscle fibers require high levels of ATP in order to maintain their cycle of
contraction and relaxation (Hultman & Spriet, 1986). Skeletal muscle has three major
50
methods by which it can form ATP. A majority of the ATP synthesized in muscle is
derived from muscle glycogen stores. First, glycogen can be directly broken down to
form ATP and lactic acid through the process of glycolysis. This method is relatively
inefficient, but can be performed quickly and in the absence of oxygen. It is the most
common method of energy utilization for type II fibers. Alternatively, ATP can be
derived through oxidative phosphorylation in muscle mitochondria. This process is
aerobic (requiring oxygen), but can derive far more ATP molecules per glucose molecule
than glycolysis. Additionally, it is capable of using fatty acids and amino acids as
substrates, while glycolysis is only capable of employing glucose. This type of energy
utilization is most common among type I fibers, although type IIa fibers use this method
to a larger degree than type IIb fibers. Finally, ATP can be synthesized in skeletal muscle
through phosphorylation of ADP by the molecule creatine phosphate (CP) and the
enzyme creatine kinase (CK) (Klivenyi et al., 1999). CP is created through
phosphorylation of the molecule creatine, which is derived from dietary amino acids, and
produced in the kidneys and liver. After production, it is subsequently transported into the
blood. Creatine is used almost exclusively by skeletal muscle, although a small
percentage of serum creatine is also transported to the brain and heart. After creatine is
taken up by skeletal muscle, it is phosphorylated to form energy stores in the form of CP.
When CP is broken, the released energy catalyzes the phosporylation of ADP to ATP.
While this process is relatively quick, it is inherently limited by the initial concentration
of CP in the cell. For this reason, weight lifters commonly use creatine supplements for
the purposes of increasing muscle energy, allowing them to lift more weight (Kadi et al.,
2008).
51
The method of ATP synthesis used is largely dependent upon the intensity and
duration of muscle contraction. At moderate levels of skeletal muscle activity (and for
longer duration of exercise), most of the ATP used for muscle contraction is derived from
oxidative phosphorylation, and is driven by the breakdown of muscle glycogen stores into
glucose. As exercise progresses, and glycogen stores deplete, the muscles will turn to
fatty acid as substrate for energy utilization. As the intensity of muscle activity increases,
a greater fraction of total ATP production is formed by anaerobic glycolysis. Once muscle
activity is complete, CP and glycogen levels have largely decreased, and the muscle must
look to build up energy again by replenishing these stores.
The major source of energy supply in skeletal muscle comes from glycogen,
which are highly branched polysaccharide stores, composed of glucose subunits. The
uptake of individual glucose molecules by skeletal muscle is a highly specialized process,
which is mediated by the metabolic hormone insulin (Bouzakri et al., 2006). Synthesized
by the β cells of the pancreas, insulin is the major endocrine regulator of glucose
homeostasis, allowing for its uptake from the blood by peripheral organs, and facilitating
its storage as glycogen. Insulin functions by binding to the insulin receptor (IR), which
when activated results in phosphorylation of insulin receptor substrates (IRSs), eventually
facilitating transport of glucose across its transporter and into the cell. Expression of IR
and the major glucose transporter 4 (GLUT4) are readily found in skeletal muscle
(Holloszy, 2008). Once glucose enters the cell, it is subsequently packaged by the enzyme
glycogen synthase (GS) into glycogen. The activity of GS seems to be readily regulated
by insulin action, as IR knockout mice demonstrate reduced GS enzyme activity in both
skeletal muscle and brain (Araki et al., 1994). Once packaged into glycogen stores, they
52
are ready to be utilized for ATP synthesis during periods of muscle contraction. Also, as
previously mentioned, fatty acids can serve as an important substrate for skeletal muscle
ATP derivation through oxidative phosphorylation. Fat is stored in the form of
triglycerides within specialized adipocyte cells. Triglycerides are broken down by the
enzyme LPL into glycerol and fatty acids, which are packaged with proteins into
aggregates called lipoproteins. This packaging allows for fatty acids to be transported
through the blood. Fatty acids enter muscle tissue through the fatty acid transport protein
(FATP; which also imports fatty acids into the mitochondria), and are then capable of
being used as a substrate for ATP production.
Muscle energy homeostasis may play an important role in the pathophysiology of
type 2 diabetes mellitus (T2DM), a major metabolic disease that is characterized by high
levels of serum glucose, and an inability for target tissues to uptake this glucose. In the
past, much research into T2DM has focused upon the β cell of the pancreas (where
insulin is produced), with β cell death being primarily linked to the disease. However,
scientists now understand that insulin resistance in target tissues may play as large of a
role in the development of T2DM as β cell failure (Biddinger & Kahn, 2006). In other
words, although the β cell is capable of producing significant levels of insulin, this
hormone may not be able to bind to the IR, or if it does, is incapable of generating the
signal cascade necessary for glucose transport. This leads to heightened levels of glucose
in the blood (characteristic of T2DM), as well as increased levels of insulin (a condition
known as hyperinsulinemia). Much of the research in this field has focused upon insulin
resistance in target tissues such as the brain and liver, but it is now clear that skeletal
muscle insulin resistance alone can result in T2DM. Insulin resistance is skeletal muscle
53
largely occurs due to higher levels of fatty acids within the cell (Coen et al., 2010). Thus,
while fatty acids are an important substrate for the generation of ATP in skeletal muscle,
higher levels of fatty acids are capable of causing significant disruption of glucose
transport and subsequent glycogen synthesis (Turner et al., 2007).
Exactly how lipids accumulate within skeletal muscle myocytes is still a matter of
debate, and there are several potential mechanisms proposed. Largely, these mechanisms
can be grouped into two premises: (1) Increased fatty acid transport into the cell from the
blood or (2) Reduced fatty acid oxidation by muscle mitochondria for the generation of
ATP (or some combination of the two). It has been shown in humans (Bonen et al., 2004)
and rodents (Turcotte et al, 2001; Hegarty, Cooney, Kraegen, Furler, 2002) that
disruption of insulin signaling is induced by increased uptake of fatty acids into muscle.
This suggests that high levels of circulating triglycerides and lipoproteins may be
sufficient to induce insulin resistance, regardless of muscle oxidative capacity. A separate
body of evidence seems to suggest that defects in oxidative phosphorylation might
account for the increased fatty acid content of skeletal muscle myocytes (Hancock et al.,
2008).
1.8.2. Muscle Mitochondria
While nutrient transporters and membrane-bound receptors (and their subsequent
signaling targets) are known to play important roles in muscle metabolism, most research
is directed to the primary energy generator of the cell: the mitochondria. Even though
transport proteins and enzymes are important for packaging energy stores, the subsequent
breakdown of these stores and the derivation of ATP is largely accomplished by the
54
mitochondria. This is especially true for type I (slow-twitch) muscle fibers, which rely
almost solely upon aerobic cellular respiration for energy for contraction.
Muscle mitochondria are a direct indicator of oxidative capacity in skeletal muscle
(Holloszy, 2008). As discussed previously, muscle fibers that rely upon oxidative
metabolism for ATP generation tend to be rich in mitochondria. Modulation of oxidative
capacity in adult skeletal muscle myocytes is largely achieved by two primary methods:
(1) mitochondrial biogenesis, and (2) mitochondrial enzyme activity (most notably the
enzymes of the electron transport chain and ATP synthase). Increasing either of these two
parameters will augment oxidative metabolism (create more ATP, while reducing the
amount of nutrient substrate available), while decreasing them will reduce oxidative
metabolism.
Increasing muscle mitochondrial content (mitochondrial biogenesis) occurs
naturally over the course of myocyte development. In mature adult cells, mitochondrial
content can be augmented through physical activity and exercise. There are several
consequences that arise from increased mitochondrial biogenesis (Holloszy, 2008). Not
only is the cell capable of generating greater amounts of ATP, there is also a significant
decrease in disturbance of energy homeostasis during times of profound muscle
contraction (such as seen during exercise). What this means is that there is a smaller
decrease in ATP and creatine phosphate consumed, as well as a smaller increase in ADP,
AMP and lactate produced (Constable et al., 1987). Furthermore, muscle fatigue is greatly
reduced (Dudley, Tullson & Terjung, 1987). Finally, and perhaps most interestingly,
proliferation of mitochondria is often linked with a change in substrate utilization toward
more oxidation of fatty acid, as opposed to breakdown of glucose, reducing fat stores
55
throughout the body (Coggan et al., 1990; Hurley et al., 1986). Thus, increased
mitochondrial biogenesis not only alters methods of energy utilization within myocytes
themselves, but also improves whole-body systemic metabolism.
Various molecular players have been implicated in mediating muscle-specific
mitochondrial biogenesis and enzyme activity. The first major factor is linked to the
Peroxisome Proliferator-activated Receptor (PPAR) family. The PPAR nuclear receptors
(α, β, γ, δ) are activated by polyunsaturated fatty acids, and play an important role in fatty
acid homeostasis (Chawla et al., 2001). A specific co-activator of PPARγ, termed
peroxisome proliferator-activated receptor γ coactivator-1α (PGC-1α), has become well
known as one of the significant modulators of mitochondrial biogenesis. Modifying PGC-
1α expression not only alters mitochondrial content, but also has significant effects on
whole-body metabolism and body composition. Muscle-specific overexpression of PGC-
1α results in large increases in functional mitochondria (Wu et al., 1998; Lehman et al.,
2000, Lin et al., 2002) and GLUT4 expression (Wende et al., 2007). PGC-1α mediates its
effects on mitochondrial biogenesis by docking on and activating the transcription factors
that regulate nuclear genes encoding mitochondrial proteins and that induce expression of
mitochondrial transcription factor A, which regulates mitochondrial DNA transcription
(Lin, Handschin & Spiegelman, 2005).
Another major cellular modulator of mitochondrial fatty-acid oxidation is the
enzyme 5’ AMP-activated protein kinase (AMPK), which is activated during muscle
contraction (Chen et al., 2003), as well as by external endocrine signals (Tamauchi et al.,
2002). The effects of AMPK activation are diverse in nature. In skeletal muscle, AMPK
functions by enhancing fatty acid uptake (Steinberg et al., 2004), and increasing fatty acid
56
transport into mitochondria for β-oxidation (Munday, Carling & Hardie, 1988). It also has
pronounced effects upon glucose homeostasis in skeletal muscle, increasing GLUT4
content in the sarcolemma (Merrill, Kurth, Hardie & Winder, 1997). In these ways,
AMPK is capable of enhancing substrate utilization for ATP in skeletal muscle, reducing
serum glucose and fatty acids, and increasing systemic metabolism. AMPK has also been
shown to alter energy homeostasis through its effects on gene expression (McGee &
Hargreaves, 2010). Sufficient research shows that AMPK is capable of altering the
expression of several key genes in skeletal muscle metabolism. This includes genes
involved in angiogenesis, such as vascular endothelial growth factor (VEGF); and
glycolysis, such as hexokinase II (HKII), which phosphorylates glucose molecules in
preparation for energy harvest (Stoppani et al., 2002). AMPK also increases expression of
FATP, which increases transport of fatty acids into mitochondria (Barnes et al., 2005).
Substantial evidence also shows that AMPK is capable of increasing expression of
mitochondrial enzymes, particularly those involved in the aforementioned ETC
(Jorgensen et al., 2007; Garcia-Roves, Osler, Holmstrom & Zierath, 2008). Thus, taken
together, this work shows that AMPK plays an important role in mitochondrial biogenesis
and respiration in skeletal muscle.
Induction of regulators of skeletal muscle mitochondrial biogenesis tends to
increase metabolism in mammalian organisms. Overexpression of PGC-1α in skeletal
muscle fibers has several beneficial effects in aging mice (Wenz, Rossi, Rotundo,
Spiegelman & Moraes, 2009; Calvo et al., 2008). As compared to WT littermates at 20
months of age, these mice demonstrate reduced body fat, increased lean muscle mass, and
higher levels of basal metabolism. With regards to glucose metabolism, MCK-PGC-1α
57
transgenic mice show increased insulin sensitivity and improved insulin signaling.
Similar findings have been demonstrated in the context of AMPK. In a key study, Ronald
Evans’ group showed that treatment of sedentary mice with the AMPK agonist AICAR
induced increases in oxidative capacity of skeletal muscle, fiber-type changes, and
induction of various metabolic genes (Narkar et al., 2008). They also found that these
mice show improved muscle-endurance when engaged in exercise, and show a synergistic
effect (in terms of muscle physiology and gene expression) as compared to mice that
exercised without the AMPK-agonist. These studies provide evidence of metabolic
benefits achieved through direct activation of mitochondrial biogenesis in skeletal
muscle.
While activation of these molecular players has been shown to have beneficial
effects on health, disruption of skeletal muscle mitochondria have been shown to result in
various pathologies. Mitochondrial myopathy (muscular disease characterized by
mitochondrial dysfunction) causes major disruption of metabolism and can result in
severe impairment or death (Johannsen & Ravussin, 2009). Several diseases (both
myogenic and neurogenic in etiology) that were originally thought to be due to unknown
mechanisms surrounding cellular toxicity are now thought to be related to mitochondrial
disease. For example, amyotrophic lateral sclerosis (ALS) is a neurodegenerative disease,
characterized by motoneuron loss and muscular atrophy. Recent evidence has found that
mitochondria and impaired energy homeostasis may play a role in ALS (Menzies, Ince &
Shaw, 2002), with many ALS mouse models demonstrating skeletal muscle
hypermetabolism. Treating ALS mouse models with creatine (which, as described, is used
as an energy source in skeletal muscle) results in amelioration of ALS symptoms, and
58
increased longevity (Klivenyi et al., 1999; Dupuis et al., 2004). Similar findings have
been shown in the context of Huntington’s Disease (HD), a neurodegenerative genetic
disease believed to be caused by neurotoxicity. Treatment of HD model mice with
creatine also has neuroprotective effects in aiding to slow progression and onset of the
disease (Ferrante et al., 2000). Interestingly, muscle biopsies taken from HD patients
show reduced expression of PGC-1α and lower levels of oxidative metabolism in isolated
myocytes (Chaturvedi et al., 2009).
PGC-1α-null mice demonstrate myopathy, coupled with reduced insulin-
sensitivity, T2DM and increased mortality (Lin et al., 2004), with similar deficits seen in
humans that lack the protein (Petersen, Dufour, Befroy, Garcia & Shulman, 2004).
Finally, myopathic symptoms can be attenuated by induction of transgenic PGC-1α
expression and subsequent mitochondrial biogenesis in skeletal muscle (Wenz, Diaz,
Spiegelman & Moraes, 2008). Skeletal muscle mitochondrial dysfunction has also been
implicated in pathogenesis of spinal and bulbar muscular atrophy (SBMA) (Ranganathan
et al, 2009; Musa, et al., In Prep). Thus, not only can augmentation of skeletal muscle
mitochondria improve metabolic parameters, but dysfunction of these organelles can also
result in manifestation of a wide variety of pathologies.
1.9. Objectives and Hypotheses
Identifying the site of action by which T mediates body composition has
significant implications for human health and medicine. The preceding discussion has
highlighted the fact that skeletal muscle is an important regulator of whole-body
metabolism, and changes within this tissue (whether gain or atrophy) have far-reaching
59
effects on systemic metabolism. With that in mind, this study aimed to investigate
transgenic rats that overexpress AR in muscle fibers. Body composition was probed, with
special attention being paid to adipose tissue. Whole fat pads were dissected and analyzed
from animals, with specific differences in individual adipocytes noted. The necessity of T
for the effects seen was also investigated in transgenic and wild-type females. Finally, the
study aimed to discover whether differences in body composition could be pinpointed to
underlying differences in resting metabolism.
Since a loss-of-function knockout of AR in skeletal muscle fibers was sufficient to
induce muscular atrophy (Ophoff et al., 2009), it was hypothesized that overexpression of
AR in skeletal muscle fibers would increase skeletal muscle mass, likely by mediating
fiber hypertrophy. The importance of skeletal muscle to whole-body metabolism
suggested that an increase in muscle mass might be sufficient to increase resting
metabolism and reduce adiposity (Harrison & Leinwand, 2008). Thus, it was expected
that these transgenic rats would subsequently demonstrate reduced fat body mass, and
enhanced resting metabolism. Finally, it was unclear as to whether these effects could be
seen in transgenic females, since despite overexpression of AR in skeletal muscle, T
levels would be too low for any measurable differences to be noted. Therefore, it was
hypothesized that the differences found between transgenic and wild-type males could be
manifest in females through acute T-treatment.
60
Chapter 2: Materials and Methods
2.1. Overview
The described study consisted of five separate experiments aimed at identifying
the role of muscle AR in regulation of body composition and metabolism. All 5
experiments used the rat species (Rattus norvegicus), while Experiments II and V
supplemented the work on rats by also employing the mouse species (Mus musculus). The
specific strains employed are discussed in further detail below. Since androgens (namely
T) have more prominent functions in males, most of the experiments (i.e. Experiments I,
II, IV and V) use this sex predominantly, if not exclusively. Female animals are also
investigated in Experiment II, while only Experiment III is performed exclusively with
females. A summary of all of the animals used is provided in Table 1.
Most of these studies involve investigation of skeletal muscle. For this purpose,
the extensor digitorum longus (EDL) muscle was typically examined. The EDL is often
used as a representative skeletal muscle since it is easily extracted, and its use has been
well characterized in studies of androgens and non-sexually dimorphic skeletal muscles
(Monks et al., 2006). Finally, subject genotyping as well as Experiment I involved the use
of various primer sequences, which are summarized in Table 2.
2.2. Animal Strains
2.2.1. HSA-AR Rats
HSA-AR transgenic rats were created and generously provided by Ligand
Pharmaceuticals Inc. (San Diego, CA). In these animals, overexpression of human AR is
61
driven specifically in muscle fibers through the use of the Human Skeletal α-Actin (HSA)
promoter sequence. Expression of the α-skeletal actin (ACTA1) gene is largely restricted
to the striated muscles (more in skeletal muscle than cardiac) in all vertebrate species
studied, and is not found in major androgen target tissues such as the prostate or preputial
gland (Brennan & Hardeman, 1993). Similarly, androgens are capable of activating the
ACTA1 gene and promoter in isolated muscle cells, but not in liver, prostate or breast
cancer cells (Hong et al., 2008). Therefore, the HSA promoter is commonly used to drive
transgene expression specifically in skeletal muscle myocytes (Ex: Clapham et al., 2000;
Rao & Monks, 2009).
Generation of HSA-AR Tg rats was performed as described previously (Niel et
al., 2009). Human AR cDNA was obtained from a clone (as provided by Dr. S. T. Liao,
University of Chicago). A 3400-bp of the clone was ligated into a HSA expression
cassette, using BglII and ClaI restriction sites. The HSA promoter used was a fragment of
that found in the ACTA1 gene. Thus, the promoter used for development of this transgenic
strain was a truncated version of the full HSA promoter. The sequence and orientation of
obtained constructs was confirmed by sequencing. This was followed by pronuclear
microinjection of transgene DNA into Sprague-Dawley (SD) strain zygotes. Identification
of animals carrying the HSA-AR transgene was accomplished through PCR
amplification. Founders were subsequently backcrossed onto WT SD rats for at least 7
generations.
Initial characterization of HSA-AR rats was performed in order to determine
location and level of transgene expression (Niel et al., 2009). Reverse transcription
polymerase chain reaction (RT-PCR) using transgene-specific primers demonstrates
62
expression of the transgene in limb muscle and BC/LA (sexually dimorphic skeletal
muscles) of neonates. No expression is found in muscle from WT neonates. Transgene
expression is also found in LA muscle of adult HSA-AR rats. RT-PCR was not performed
on non-sexually dimorphic muscle, nor was it performed on other tissue types. These
differences in mRNA expression of the transgene were also confirmed for protein
expression through western blot. AR was probed using a non-specific AR antibody
(probes for WT and Tg AR). In tissue taken from HSA-AR and WT adult rats, it was
demonstrated that the amount of AR protein was roughly 1.6 fold higher in LA muscle,
and 3.8 fold higher in EDL muscle of HSA-AR rats. Thus, taken together, it has been
shown that HSA-AR rats demonstrate higher levels of AR expression in skeletal muscle.
However, it remains unclear as to whether transgene expression is found in other tissue
types (including other muscle types) in HSA-AR animals. Functionality of the transgene
was assessed through gene expression of myoglobin, which is a typical marker of local T
action in skeletal muscle (Manttari, Anttila & Jarvilehto, 2008). Myoglobin mRNA levels
were found to be higher in EDL muscle of HSA-AR adult male rats, as compared to WT
male adults (Niel et al., 2009). This provides evidence of functionality of transgene-
derived AR.
2.2.2. Tfm Rats
For the last several decades, Testicular feminization mutation (Tfm) rodents have
provided the dominant model for understanding androgen insensitivity in mammalian
systems (Zuloaga et al., 2008). The Tfm phenotype is the product of a single-base pair
mutation in the AR gene (Yarbrough, et al., 1990). More specifically, it is the mutation of
63
guanine to adenine within exon E, resulting in an amino acid change of arginine 734 to
glutamine within the ligand-binding domain of the AR gene. Thus, even though the AR
protein is successfully translated, due to the mutation in the ligand-binding domain, this
AR protein is not capable of binding to androgens, nor translocating to the nucleus to alter
expression of androgen-responsive genes. However, due to the simple nature of the
mutation (single amino acid alteration), it has been argued that there is some residual
sensitivity to androgens in Tfm rats, although it is believed to be largely negligible
(Zuloaga et al., 2008).
The translation of non-functional AR in Tfm rats results in a phenotype that is
analogous to that seen in human androgen-insensitivity syndrome (AIS) (Zuloaga et al.,
2008). Homozygous Tfm rats that are genetically male (i.e. possessing the XY
chromosome configuration) appear phenotypically female. They possess nipples typical
of female rats. Furthermore, their external genitalia demonstrate an ambiguous (intersex)
phenotype, with no discernible phallus. They are also infertile, due to defects in
spermatogenesis. Due to the X-linked nature of the mutation, only genetically male (XY)
carriers can demonstrate androgen insensitivity. Since Tfm males are infertile, obtaining
Tfm animals is therefore achieved by crossing a WT male with a heterozygous Tfm-carrier
(XTfmX) female.
Finally, it is important to discuss endocrine parameters in Tfm male rats. Most
notably, T levels appear to be extremely high in males, with serum concentrations
comparable to the high male range (Naess et al., 1976). This is presumably due to
uncoupling of HPG negative feedback. The mutation of AR in the hypothalamus means
that T will not have action on this region of the brain, and therefore will not be able to
64
suppress secretion of LHRH or gonadotropins (FSH and LH). Concomitant with this
explanation, FSH and LH levels are also elevated in Tfm male rats (Naess et al., 1976).
This would account for the increased levels of T seen in these animals. Lastly, there have
been some differences noted in activity of the enzyme aromatase in Tfm rats. Recall that
aromatase is the enzyme that mediates the conversion of T into 17β-estradiol. Since
ArKO mice develop obesity (Fisher et al., 1998), in this context it is important to
understand any enzymatic differences between strains. In Tfm males, studies have found
impairment of aromatase activity in several parts of the brain, thus limiting conversion to
estrogens in these animals (Roselli, Salisbury & Resko, 1987; Rosenfeld, Daley, Ohno &
YoungLai, 1977). To date, differences in aromatase activity in other tissues are unclear.
Therefore, scientists must keep in mind the possibility that estrogen signaling may also be
impaired in males of this mutant strain.
2.2.3. HSA-AR/Tfm Rats
Obtainment of HSA-AR/Tfm animals is typically accomplished by breeding an
HSA-AR male with a heterozygous Tfm-carrier female. It could also be done through the
mating of a WT male with an HSA-AR/Tfm-carrier female. Resultant male offspring that
are homozygous for the Tfm mutation, but also express the HSA-AR transgene,
demonstrate a very interesting AR-expression phenotype. These animals possess non-
functional AR in all tissues (due to Tfm), except for muscle fibers, where AR is expressed
due to the transgene (Niel et al., 2009). As a result, these animals serve as a powerful
genetic model for investigation of AR function only in myocytes.
65
Preliminary analysis of HSA-AR/Tfm males demonstrates increased expression of
AR in limb muscle fibers of neonatal rats, as compared to Tfm males (through
immunohistochemistry and confocal microscopy) (Niel et al., 2009). Phenotypically,
rescue of AR in skeletal muscle fibers of these rats does not promote development of the
sexually dimorphic SNB neuromuscular system in Tfm males. Post-natal maintenance of
perineal muscles (BC/LA) or innervating motoneurons is not found in this animal model,
suggesting that AR in skeletal muscle is not sufficient for development of this
neuromuscular system.
In this study, HSA-AR/Tfm male rats will be used to understand how AR within
muscle fibers alone can alter body composition and metabolism. For most purposes, they
will be compared and contrasted with Tfm littermates.
2.2.4. HSA-AR Mice
Transgenic mice overexpressing AR in skeletal muscle were developed by Monks
et al. (2007). Validation of the HSA promoter used for development of these mice was
performed through the creation of a LacZ (β-galactosidase) reporter strain, and it was
shown that the HSA promoter drives transcription only in skeletal muscle myocytes.
Transgenic HSA-AR mice were created using this promoter, driving expression of rat WT
AR cDNA. Obtained HSA-AR mice were then crossed with mice possessing the Tfm
mutation. Unlike in rats, Tfm mutation in mice is a single base deletion, resulting in a stop
codon (Zuloaga et al., 2008). For that reason, homozygous Tfm mice do not translate AR
protein. Subsequent HSA-AR/Tfm mice had tissues processed and were probed for AR
protein expression. Immunoblot assays confirm the specificity of the HSA promoter, with
66
AR protein expression found in skeletal muscle only, and not in other tissues (such as
spinal cord, testes or heart).
Two major founding lines expressing the HSA-AR transgene were predominantly
used: L78 and L141 (Monks et al., 2007). There were significant differences in transgenic
AR expression found between these two lines. L141 animals have a significantly higher
transgene copy number (approximately 100 copies/pg RNA in L78, contrasted with
nearly 1000 copies in L141). Furthermore, L141 mice demonstrate significantly higher
AR expression in skeletal muscles, with differences noted in both AR mRNA and protein
level.
In terms of phenotype, marked differences also exist between the L78 and L141
lines. L141 mice demonstrate significant muscular atrophy, with male viability at birth
significantly reduced as compared to WT and L78 males. L141 male mice (or females
acutely treated with T) demonstrate reduced motor ability, body weight, and muscular
strength. Since the study described in this paper exclusively employs the L78 line, focus
will be paid towards the phenotype of this founding line. In contrast to L141 mice, L78
males are largely asymptomatic. Similarly, treating female L78 mice with exogenous T
does not result in muscular atrophy, as seen in L141 females. Body weight is reduced in
adult L78 males (as compared to WT brothers from the same line), but no defects in
motor functioning were noted. Some muscular pathology was noted in histological assays
of muscle fibers, with a reduced number of total EDL fibers recorded (as compared to
WT brothers from the same line). However, the degree of this pathology is significantly
reduced when compared to that seen in L141 mice.
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2.3. Subjects
2.3.1. Animals
All experiments described herein were accomplished using transgenic HSA-AR
and/or Tfm Sprague Dawley rats. HSA-AR and Tfm rats were also crossed in order to
produce the HSA-AR/Tfm strain, which expresses AR exclusively in myocytes.
Experiments II and V also employed transgenic L78 HSA-AR mice on a C57BL/6J
background. Please see Table I for breakdown of the number of animals used in each
experiment, and the genotypes of those animals. All animals were bred locally at the
University of Toronto at Mississauga. All procedures and experiments using live animals
were approved by the Office of Research Ethics at the University of Toronto, and adhered
to federal and National Institutes of Health guidelines. All animals were housed at
approximately 23oC, on 12h:12h light-dark cycles. Food and water were available ad
libitum.
2.3.2. Genotyping
Determination of animal genotype (including transgene expression or mutation)
was accomplished using polymerase chain reaction (PCR). At postnatal day (PND) 21,
animals were weaned from their mothers and housed in separate cages with food and
water. At this time, ear samples were taken by tissue puncture, placed in microcentrifuge
tubes on ice, and subsequently stored at -80oC for next-day lysis. The following day,
100µL of lysis buffer and 2µL of proteinase K were added to each microcentrifuge tube
containing a tissue sample. Samples were heated at 55oC for two hours, with brief
vortexing of samples after 30 minutes (to mix contents). Samples were then heated at
68
95oC for 10 minutes, and spun in a centrifuge (10,000 RPM for 2 minutes) in order to
separate lysate from un-lysed tissue. This concentrated DNA was then diluted (1:10) in
distilled water for use in PCR.
PCR was performed using specific primers for each genotype. For HSA-AR rat
genotyping, DNA samples were analyzed using transgene-specific HSA-AR primers (F:
GGACAGGGCACTACCGAG; R: GGCTGAATCTTCCACCTAC), as well as primers
for the housekeeping gene glyceraldehyde 3-phosphate dehydrogenase (GAPDH), used as
a control (F: ATGGGAAGCTGGTCATCAAC; R: GGATGCAGGGATGATGTTCT).
All PCR reactions include a known positive control, a known negative control, and a null
control. The reaction was run for a total of 31 cycles. Subsequent PCR product was run
on a 1.2% agarose gel for 30-35 minutes at 100 V. The gel was then removed and
visualized using ethidium bromide. HSA-AR is a 192 bp product, while GAPDH is a 440
bp product, creating two easily distinguishable bands. For genotyping of HSA-AR mice,
DNA samples were analyzed using transgene-specific HSA-AR primers (F:
AGTAGCCAACAGGGAAGGGT; R: GAGGCAGCCGCTCTCAGGGTG), with
primers for the housekeeping gene thyroid-stimulating hormone (TSH), used as a control:
(F: AACGGAGAGTGGGTCATCAC; R: CATTGGGTTAAGCACACAGG). The
reaction was run for 35 cycles. Subsequent PCR product was run on a 1.2% agarose gel
for 30-35 minutes at 100 V. The gel is then removed and visualized using ethidium
brominde. HSA-AR is a 650 bp product, while TSH is a 383 bp product, creating two
easily distinguishable bands.
Genotyping for the Tfm mutation was performed using PCR with primers for the
rat AR gene (F: GCAACTTGCATGTGGATGA; R: TGAAAACCAGGTCAGGTGC).
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Since this gene should exist in all rats, a housekeeping gene is not necessary. PCR
reaction ran for 35 cycles. Since the Tfm mutation is the result of a single base-pair
mutation, a portion of PCR product from each sample was digested using the restriction
enzyme Sau96I, which recognizes the 5’ GGNCC sequence. Tfm AR, due to the single
base pair mutation, cannot be cleaved by Sau96I, whereas WT AR should be cleaved into
two fragments. For each sample, restriction enzyme-digested PCR product was run next
to a non-digested PCR product (as a control). Gel electrophoresis was performed on a 3%
agarose gel for 30-35 minutes at 100 V. The gel was then removed and visualized using
ethidium bromide. All non-digested PCR product should demonstrate a single band,
signifying the AR gene. The presence of a single band in digested PCR product is
indicative of Tfm AR, while the presence of two bands indicates WT AR. Since
generation of homozygous Tfm females is not possible, the presence of only the non-
cleaved band in the digested-product is indicative of a Tfm male. Heterozygous Tfm-
carrier females can be identified from digested PCR product, through the presence of both
the single (non-cleaved) Tfm AR band, and the double WT AR band.
2.4. Experiment I: Tissue Specificity of Transgene Expression
Previous work on HSA-AR rats has demonstrated transgene expression (and AR
overexpression) in skeletal muscle (Niel et al., 2009). However, the fact that these
animals were produced using a truncated version of the HSA promoter implies the
possibility that transgene expression may be “leaky”, and found in other tissue types. For
this purpose, tissue specificity of transgene expression was analyzed.
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2.4.1. Reverse-Transcription (End-Point) PCR
At 16 weeks of age, WT and HSA-AR male rats were euthanized with an
overdose of sodium pentobarbital and dissected. EDL (skeletal muscle), heart (cardiac
muscle), urinary bladder (smooth muscle), kidney and white adipose tissue (WAT) were
removed. It is important to mention that separate tools were used for dissection of each
tissue, and were thoroughly cleaned after each animal, in order to limit any RNA
contamination. Tissues were removed, frozen in liquid nitrogen, and subsequently stored
at -80oC. Isolation of RNA was performed using TRIzol reagent. Tissues were slightly
degraded before being homogenized in the presence of 1 mL of TRIzol. Residual tissue
that could not be homogenized was discarded. Homogenized samples were allowed to
incubate for 5 minutes on ice before 200 µL of chloroform was added. Samples were
vortexed and components separated using centrifugation (14,000 RPM for 15 min). The
RNA exists exclusively in the colourless upper aqueous phase, from where it was
isolated. The remaining organic phase was discarded. Precipitation of RNA pellets was
accomplished by addition of isopropyl alcohol, manual shaking, and incubation at room
temperature for 2 hours. Following incubation, component separation through
centrifugation (12,600 RPM for 10 min) was carried out, resulting in the formation of an
RNA pellet. Next was an RNA wash phase, where the supernatant was removed, and the
pellet was washed with 1 mL of 75% ethanol. Centrifugation was carried out once again
(7,600 RPM for 5 min), after which point ethanol was removed. Finally, RNA was re-
dissolved. The pellet was allowed to briefly air-dry, before being washed in
diethylpyrocarbonate (DEPC) water (approximately 25 µL, potentially more if the pellet
could not be dissolved). Following this, the isolated RNA was stored at -80oC.
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After successfully isolation, RNA samples were analyzed for stability (non-
degradation) through glyoxylation and electrophoresis. Stable RNA was next treated with
DNase in order to remove any residual DNA. Next, the RNA was converted to cDNA
through a reverse transcriptase-mediated reaction. Newly acquired cDNA was then used
in a RT-PCR reaction (35 cycles) with transgene-specific primers for human AR (F:
AGGAAGCAGTATCCGAAGGCA; R: GGACACCGACACTGCCTTACA). Expression
of the housekeeping gene GAPDH was analyzed in a separate reaction (using previously
described primers), in order to act as a control. PCR products were electrophoresed on a
1.5% agarose gel. Presence of an electrophoresed band would be indicative of gene
expression. Analysis was performed for both WT (n = 3) and HSA-AR (n = 3) male rats.
2.5. Experiment II: Body Composition Analysis
Since androgens have been shown to have significant effects on body composition
(namely increasing lean muscle mass and decreasing fat mass), this possibility was
investigated in rat strains (WT, HSA-AR, Tfm, HSA-AR/Tfm), as well as comparative
mouse strains (WT, L78 HSA-AR).
2.5.1. Dual-Energy X-Ray Absorptiometry
Body composition was analyzed using dual-energy X-ray absorptiometry (DXA)
scanning. DXA scanning is a common method of measuring body composition, and is
employed in basic science animal studies, as well as in clinical contexts with patients
(Malina, 1969). These scans emit X-rays, which are directed at the subject, and
subsequently absorbed by the tissues of the body. Different tissue-types (namely lean
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mass, fat mass, and bone) absorb X-ray emissions to different degrees. Using these
differences in absorption, the machine is able to derive estimates of total body mass, as
well as the mass of each individual component of body composition.
Animals were scanned using a standard DXA machine (Hologic QDR4500;
Hologic Inc., Waltham, MA). For each DXA session, the machine was first calibrated
through the use of standardized phantom models. Measured parameters of these phantom
scans were maintained across sessions, in order to ensure the comparative validity of
scans performed on different days. Before scanning, animals were anaesthetized using
isoflurane, and maintained under anesthesia for the duration of the experiment. DXA
small animal scans take approximately 2 minutes each.
For rats, analysis was performed on WT males (n = 11), WT females (n = 8),
HSA-AR males (n = 11), HSA-AR females (n = 9), Tfm males (n = 7), Tfm-carrier
females (n = 7), HSA-AR/Tfm males (n = 9) and HSA-AR/Tfm-carrier females (n =7).
Rats were group-housed, with 3-5 animals per cage. Males and females were caged
separately. Rats were scanned bi-weekly, beginning at 4 weeks of age, and ending at 10
weeks of age. For mice, analysis was performed on the L78 line males: WT (n = 5) and
HSA-AR (n = 6). Mice were measured once by DXA at 24 weeks of age.
2.5.2. Parameters Examined and Derived
DXA analysis provides the operator with estimates of various parameters of body
composition. The includes total body mass, bone mineral content (BMC; the total amount
of bone mass), bone mineral density (BMD; the amount of bone per unit of bone area),
total fat body mass (FBM; the amount of adipose mass on the body), lean mass + bone
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mineral content (the total amount of non-FBM on the body), and FBM% (the percentage
of total mass accounted for by fat). Given these parameters, lean body mass (LBM) could
be derived. LBM was derived by taking the Lean+BMC parameter and subtracting the
known BMC value. LBM% was then determined using this total LBM and the known
body mass. For all scans, animal body weight was measured using a weighing scale, and
compared with the provided total body mass from the DXA scan, in order to validate
DXA accuracy. It was found that the measured weight by DXA never differed from the
actual mass by more than 2%, suggesting that the DXA machine was highly accurate in
its measures.
2.5.3. Dissections and Tissue Weights
In order to determine if the findings from DXA analysis could be confirmed in
individual tissues, dissections were carried out to measure weights of individual muscles
and fat pads of WT, HSA-AR, Tfm and HSA-AR/Tfm male rats. At 16 weeks of age, rats
were euthanized through overdose with sodium pentobarbital. Individual EDL and AT
muscles were dissected and weighed. WAT weight was measured by removal of
perigonadal (reproductive) fat pads from rats. Dissections were carried out for WT (n =
10), HSA-AR (n = 12), Tfm (n = 7) and HSA-AR/Tfm (n = 10) males.
2.6. Experiment III: Testosterone Treatment of Adult Females
This experiment attempted to recapitulate effects of the transgene seen in HSA-
AR male rats through acute treatment of adult HSA-AR female rats with exogenous T.
This was a within-group study, with all animals having body composition measured
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during treatment with T, as well as when T capsules were removed (and replaced with
vehicle capsules). Analysis was carried out on WT (n = 8) and HSA-AR (n = 9) female
rats, aged 24-30 weeks.
2.6.1. T-Capsule Surgeries
Treatment of females with exogenous T was done through implantation of silastic
capsules containing the hormone. This methodology has been used before in many
studies, and is a standard method of treating animals with exogenous hormone (Monks,
Vanston & Watson, 1999; Monks & Watson, 2001; Monks et al., 2001a; Monks et al.,
2001b). T-capsules containing exogenous T are SILASTIC brand (1.57mm inner diameter
X 3.18mm outer diameter, Dow Corning, Midland, MI), containing T (20 mm in length,
Steraloids, Newport, RI). After baseline DXA scanning, all animals were anaesthetized
with isoflurane. A small incision was made on the dorsal side of the rat, and two T
capsules were inserted subcutaneously (just beneath the skin). Incisions were
subsequently stitched and sealed, and animals were treated with an analgesic (Anafen) to
relieve any pain. Animals were also single-housed for three days after surgery, in order to
ensure proper closure of the wound. Following 4 weeks of exogenous T-treatment, all
animals were returned to surgery and anaesthetized again with isoflurane. The dorsal
incision was re-opened, and the T capsules were removed. Capsules were checked to
ensure that excess T had not been released. After T capsules had been removed, they were
replaced by two vehicle capsules (Silastic capsules that were empty), and the incision was
then sealed and stitched as described above.
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2.6.2. Body Composition Analysis
Female rats were subjected to DXA scanning before T treatment, in order to
establish baseline measures of body composition. Following T treatment, animals were
scanned weekly for 4 consecutive weeks. Immediately following the final scan of this
regimen, T capsules were removed, and vehicle capsules replaced, as described above.
Females were then scanned by DXA weekly for 4 consecutive weeks, in order to
determine if and how subsequent T withdrawal would alter body composition in HSA-AR
females.
2.7. Experiment IV: Adipose Histology
In order to determine effects of muscle AR on WAT at a cellular level, histology
of adipose tissue was conducted to examine the size of individual adipocytes.
2.7.1. Sample Preparation and Sampling Strategy
At 16 weeks of age, animals were euthanized through overdose with sodium
pentobarbital and dissected. Reproductive perigonadal fat pads were dissected in the same
way from all animals. Following dissection, removed fat pads were placed in 10%
phosphate-buffered formalin (PBF) for fixation for at least 5 days. Following fixation, a
sample of the whole fat pad was used to perform histology. This sample was taken from
the same part of the fat pad in all animals. Perigonadal fat pads are directly attached to the
testes in males, and so the sample was removed from the part of fat pad furthest from the
testes. After fixation, adipose samples were processed to prepare for paraffin embedding.
Samples were stored in individual plastic cassettes and placed in an automated basket,
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which takes the cassettes through each solution. Tissue was incubated in various solutions
(distilled water, 70% ethanol, 70% ethanol, 95% ethanol, 95% ethanol, 100% ethanol,
100% ethanol, xylene, xylene, paraffin, paraffin) for 1 hour per solution. Following
processing, tissue samples were properly oriented and embedded into paraffin blocks
using molds. The tissue was allowed to set overnight before sectioning.
Paraffin blocks were sliced on a microtome at room temperature, and at a
thickness of 6µm. Sections were mounted on slides using gelatin solution. Slides were left
to dry overnight on a slide heater. The following day, the slides were stained. Slides were
first cleared in xylene for 13 minutes (to remove paraffin), followed by dehydration in
graded alcohols (100%, 100%, 95%, 70%) for 3 minutes each. Slides were washed with
distilled water for 3 minutes, before being incubated in hematoxylin (Sigma-Aldrich,
Oakville, Ontario, Canada) for 5 minutes. Following hematoxylin staining, slides were
washed in distilled water for 1 minute, before being processed with eosin and
subsequently coverslipped.
Resultant sections needed to be analyzed systematically in order to maintain
consistency across subjects. Images were acquired using an Olympus bright-field
microscope (model BX51; Olympus, Tokyo, Japan), a 4X objective, and a colour
videocamera (Cool Snap Pro Color; Roper Scientific, Duluth Georgia) with Image Pro
Plus Software (Media Cybernetics Inc., Silver Spring, MD). Images were taken at 40X
magnification, at a resolution of 680 X 512 pixels. For each adipose cross-section, three
photomicrographs were taken for analysis. All photomicrographs were taken in the same
area of the cross-section for all subjects, in order to maintain consistency.
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2.7.2. Measurements
In order to quantify adipocyte size, photomicrographs were imported into ImageJ
software (National Institutes of Health, Bethesda, MD), which was used to trace cell size.
Calibration of the software was performed prior to measurements of size, so as to provide
accurate area measures in µm2. Due to the nature of the hematoxylin and eosin stain, not
all individual adipocytes were clearly demarked. For that reason, only cells whose
complete membrane border could be seen were subsequently traced and analyzed. For
each animal, a minimum of 150 measures were taken, which were then averaged in order
to determine the mean adipocyte size. Analysis was conducted on WT (n = 6), HSA-AR
(n = 6), Tfm (n = 6), and HSA-AR/Tfm (n = 4) males.
2.8. Experiment V: Energy Balance and Metabolic Analyses
We investigated parameters of energy expenditure in order to determine if any
changes in whole body composition might be associated with alterations in these
parameters.
2.8.1. Resting Metabolism by Indirect Calorimetry
Evaluation of resting metabolism is typically accomplished through the use of
indirect calorimetry, which uses indicators of metabolism in order to derive an estimate of
RMR. The most common method of indirect calorimetry is oxygen (O2) consumption.
Consumption of oxygen is closely related to metabolism, as O2 molecules are required by
cells undergoing oxidative phosphorylation. O2 is reduced by the transfer of two electrons
from cytochrome C oxidase in order to produce two water (H2O) molecules and is a
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necessary step in the production of ATP. For that reason, higher levels of O2 consumption
are indicative of higher rates of metabolism (Holloszy, 2008).
At 12 weeks of age, metabolic measures were taken from male rats through
analysis of O2 consumption. Animals were placed inside a 700 mL (8.5 cm diameter, 13.0
cm length) cylindrical gas-exchange chamber (model G114; Qubit Systems, Kingston,
Ontario, Canada). Calibration of the system was first achieved by passing nitrogen gas
through the system (which provides a “0” O2 concentration baseline level). Room air was
then pumped through the chamber at a flow rate of 400 mL/min, and outflow O2
concentration was measured by a flow-through oxygen analyzer (model S102; Qubit
Systems). Concentration of outflow O2 was analyzed and displayed by gas-exchange
software (Logger Pro version 3; Vernier Software, Beaverton, OR). Animals were kept in
the chamber until outflow O2 concentration levels were stable, which was defined by no
deviation in concentration of more than 0.02% over a period of 30 minutes. The
difference between final outflow O2 concentration and inflow concentration (i.e.
concentration of O2 in room air) was used to determine oxygen uptake (in µL/min).
Animals were matched for time of day at which the measures were recorded. In order to
derive RMR in (J/g min), values of O2 uptake were corrected for body mass. Indirect
calorimetry was measured for WT (n = 10), HSA-AR (n = 12), Tfm (n = 7), and HSA-
AR/Tfm (n = 10) male rats. Indirect calorimetry was also measured for WT (n = 5) and
HSA-AR (n = 6) L78 male mice, aged 22-23 weeks.
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2.8.2. Spontaneous Activity Measures
Animals were measured bi-weekly, starting at the age of 4 weeks until the age of
10 weeks. These ages were chosen as they correspond to the periods at which body
composition was measured in male rats. Locomotor activity was assessed in clear, acrylic
glass boxes (L 43 X W 22 X H 25 cm), with clear, ventilated acrylic glass lids. The boxes
were separated by opaque screens, which prevented the rats from seeing one another
during testing, and thus eliminated visual conspecific cues. Each activity box was
equipped with two arrays of 16 X 16 infrared photo-beams, spaced 2.5 cm apart
(constructed by the Centre for Addiction and Mental Health, University of Toronto,
Toronto, Ontario). The bottom array, positioned 3 cm above the floor of the chamber,
recorded horizontal movement. The top array, positioned 15 cm above the floor of the
chamber, recorded rearing behaviour. These arrays were connected to a computer via an
interface that detected interruptions in the photo-beam (i.e. beam breaks induced by
activity). Each activity session lasted for 60 minutes, and animals were counterbalanced
for the time of day at which they were measured. Activity measures were calculated for
WT (n = 4), HSA-AR (n = 6), Tfm (n = 3) and HSA-AR/Tfm (n = 5) male rats.
2.9. Statistical Analyses
Experiments II, IV and V (between-group studies involving HSA-AR and Tfm
rats) were analyzed using two-way multivariate ANOVA, with HSA-AR and Tfm as
between-subject factors (for body composition analysis, an ANOVA was calculated for
each week that animals were tested, and all between-group comparisons were made at
each age). Differences between groups were analyzed using independent sample student
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t-tests. Parts of experiments II and V that employed L78 mice were analyzed using
independent sample student t-tests to compare WT and HSA-AR groups.
Experiment III, which involved T-treatment of WT and HSA-AR adult female
rats, was a within-group study, in which both groups were exposed to both treatments (T
and vehicle). Data was examined using a repeated-measures ANOVA with HSA-AR as a
between-subject factor and total body mass, LBM%, FBM%, raw LBM or raw FBM as
the within-subject factor. Statistical analyses were run separately for the first 5 weeks
(baseline measures and 4 weeks of T-treatment) and for weeks 4-8 (final week of T-
treatment, and 4 weeks following removal of T). Pairwise comparisons were performed
using unprotected paired t-tests if an interaction was found between the within-subject
and between-subject factors. Otherwise, a Dunnett correction for family-wise error was
applied. Pairwise comparisons were made with each week either being compared to
baseline (applies for weeks 1-4) or week 4 (applies for weeks 5-8). The use of these two
epochs would allow for evaluation of effects of T-treatment (for the former weeks), and
subsequent T withdrawal (for the latter weeks). For all studies, α was set at P < or = 0.05.
Complete statistical analyses are included in Appendix A.
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Chapter 3: Results
3.1. Overview
HSA-AR animals were characterized to determine tissue specificity of transgene
expression. Subsequently, male and female rats (HSA-AR and WT) were analyzed for
changes in body composition. HSA-AR animals were generally compared against WT
littermates on the same Tfm background (i.e. comparisons were largely WT vs. HSA-AR,
and Tfm vs. HSA-AR/Tfm). This is due to the fact that Tfm male rats demonstrate
significantly reduced body mass, which strongly confounds any analyses of body
composition made when comparing these rats to WT males. After discovering differences
in body composition of male HSA-AR rats, but not females, these effects were induced
by acute T-treatment of HSA-AR females. Finally, differences in overall energy balance
were investigated, in order to gain some understanding of the physiological mechanism
behind the differences seen.
Where available, studies in rats were supplemented with similar experiments in
HSA-AR mice. This would allow for investigation of the generality of the effects seen
(i.e. whether they are found in other species).
3.2. Experiment I: Tissue Specificity of Transgene Expression
Due to the fact that HSA-AR rats were generated using a truncated version of the
full HSA promoter, it was necessary to qualitatively analyze various tissues of these rats
in order to determine if they expressed the transgene.
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3.2.1. Transgene mRNA is Expressed in Muscle Tissue of HSA-AR Animals
Dissected tissues from adult (16-week old) WT and HSA-AR male rats were
analyzed using RT-PCR and transgene-specific primers (Fig 1). EDL (skeletal muscle),
urinary bladder (smooth muscle). heart (cardiac muscle), WAT and kidney were all
examined for both genotypes. RT-PCR found no expression of the transgene in any
tissues taken from WT animals. Conversely, transgene mRNA expression was found in
skeletal muscle, smooth muscle, and cardiac muscle of HSA-AR males. No transgene
expression was demonstrated in adipose tissue or kidney of HSA-AR males. Thus, HSA-
AR mRNA expression seems to be found in myocytes of HSA-AR animals only.
3.3. Experiment II: Body Composition Analysis
Bi-weekly DXA scanning was performed on WT, HSA-AR, Tfm and HSA-
AR/Tfm rats (from 4 weeks of age until 10 weeks), in order to determine whether AR in
muscle is capable of modulating body composition. Since T secretion increases markedly
at the onset of puberty (roughly 6 weeks in rodents, see Lee & Chang, 2003), examining
these time points would allow for some delineation of T-dependency. If the action of
post-natal T on the transgene did indeed affect body composition, such differences likely
should not arise prior to the onset of puberty. Various parameters were examined,
including whole body mass, lean body mass (LBM), LBM%, fat body mass (FBM),
FBM%, bone mineral content (BMC) and bone mineral density (BMD). Representative
images of male rats (scanned at 10 weeks of age), including WT (Fig 2a), HSA-AR (Fig
2b), Tfm (Fig 2c) and HSA-AR/Tfm (Fig 2d) are shown. Due to the large amount of
statistical analyses carried out in examination of body composition, only pertinent
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statistics will be presented in text. This largely relates to main effects as determined by
two-factor (HSA-AR and Tfm) ANOVA. For complete statistical analyses (including
results of all ANOVA and between group t-tests), please refer to Appendix A.
3.3.1. HSA-AR Expression Does Not Regulate Body Mass in Rats
We first used DXA scanning to determine if the HSA-AR transgene could alter
whole body mass in male rats (Fig 2e). Interestingly, no differences in total body mass
were found due to HSA-AR. This was the case for males at 4 weeks (F3,25 = 0.359, P =
0.553), 6 weeks (F3,40 = 0.408, P = 0.528), 8 weeks (F3,34 = 1.181, P = 0.286) and 10
weeks (F3,34 = 3.339, P = 0.077). As expected, a main effect of Tfm was found to
influence whole body mass. No effect was found at 4 weeks (F3,25 = 0.817, P = 0.373),
however Tfm males were found to have lower body mass than non-Tfm males at 6 weeks
(F3,40 = 4.517, P < 0.05), 8 weeks (F3,34 = 20.641, P < 0.001) and 10 weeks (F3,34 =
22.116, P < 0.001). This was not surprising, as Tfm males develop an intersex phenotype,
and are found to have lower overall body weight (Zuloaga et al., 2008). No interaction
was found between HSA-AR and Tfm for any age.
3.3.2. Increased Lean Muscle Mass Percent in HSA-AR Male Rats
Selective knockout of AR in myocytes alone has been shown to be sufficient to
induce muscular atrophy in mice (Ophoff et al., 2009). With this in mind, we investigated
lean muscle mass of our HSA-AR rats using DXA scanning. We hypothesized that
overexpression of AR in myocytes should increase lean muscle mass. We first compared
LBM%, which is the total amount of lean mass per unit of total body mass (Fig 2f). It was
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found that HSA-AR is capable of increasing LBM%. At 4 weeks, no effect of HSA-AR is
found (F3,25 = 1.219, P = 0.278). However, effects become significant after the onset of
puberty at 6 weeks (F3,40 = 17.748, P < 0.001), as well as 8 weeks (F3,34 = 10.630, P <
0.01) and 10 weeks (F3,34 = 6.510, P < 0.05). Unexpectedly, we found a similar effect of
Tfm in increasing LBM%. Similar to the main effect of HSA-AR on this parameter, no
effect of Tfm is seen at 4 weeks of age (F3,25 = 0.205, P = 0.654), but mutation increases
LBM% at 6 weeks (F3,40 = 4.396, P < 0.05), 8 weeks (F3,34 = 27.393, P < 0.001) and 10
weeks (F3,34 = 11.549, P < 0.01). This finding was surprising as it was contrary to the
previously cited literature on mARKO mice (Ophoff et al., 2009), as well as the muscular
atrophy noted in whole-body ARKO (MacLean et al., 2008). Here it seems that lack of
functional AR may potentially result in increasing LBM%. No interaction between HSA-
AR and Tfm was found for any age.
Interestingly, the effects due to HSA-AR and Tfm do not directly carry over when
examining raw muscle mass. Here, raw LBM values are provided by DXA, but not
corrected for overall body weight (Fig 2h). No significant effect was found for HSA-AR
on LBM at any age measured. Pair-wise comparisons between groups on the same genetic
background (i.e. WT vs. HSA-AR and Tfm vs. HSA-AR/Tfm) also demonstrate no
difference. As expected, there is a main effect of Tfm, with Tfm males having lower levels
of raw LBM at 8 weeks (F3,34 = 44.909, P < 0.001) and 10 weeks (F3,34 = 11.560, P <
0.01). This was not surprising as Tfm males have lower overall body mass, when
compared to WT littermates, and thus would be expected to have lower levels of raw
LBM. Once again, no interaction was found between HSA-AR and Tfm.
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In summary, these results suggest a disparity between HSA-AR and WT animals
in LBM%, with HSA-AR animals having higher levels. However, this difference in
proportion of LBM is not accounted for by simply increased LBM, suggesting that other
components of body composition may additionally be altered. We also demonstrate an
interesting effect of Tfm in increasing LBM%, which was contrary to expectations.
3.3.3. Reduced Fat Body Mass in HSA-AR Male Rats
With the existing disparity between WT and HSA-AR rats demonstrated in
LBM%, we investigated body fat to determine if this component of body composition
might be altered due to transgene expression. Whole-body ARKO male mice develop
increased adipose tissue and eventual obesity (Lin et al., 2005; Fan et al., 2005),
suggesting that androgens and AR can mediate this major energy storage site. This effect
cannot be recapitulated by knockout of AR in adipocytes (Yu et al., 2008) and
hepatocytes (Lin et al., 2008). Unexpectedly, knockout of AR in myocytes was found to
decrease fat mass (Ophoff et al., 2009), but the authors were unable to provide a
satisfactory explanation for this effect. Thus, whether myocyte AR is capable of
modulating fat body mass (FBM) is currently unknown.
Investigation of FBM% (total amount of fat mass per unit of whole body mass) in
male rats shows a main effect of the transgene in reducing this parameter (Fig 2g).
Similar to effects on LBM%, no differences are seen at 4 weeks of age (F3,25 = 1.200, P =
0.282). However, decreased FBM% is noted in HSA-AR rats at 6 weeks (F3,40 = 16.275, P
< 0.001), 8 weeks (F3,34 = 10.004, P < 0.01) and 10 weeks (F3,34 = 6.360, P < 0.05). Once
again, Tfm is found to act similarly to HSA-AR in improving body composition by
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reducing FBM% in male rats. Effects emerge at 6 weeks (F3,40 = 4.718, P < 0.05), and are
also seen at 8 weeks (F3,34 = 11.640, P < 0.01) and 10 weeks (F3,34 = 12.837, P < 0.001).
Unlike the differences seen in LBM%, the effects on FBM% carry over when one
examines raw FBM (not corrected for weight; Fig 2i). That is, HSA-AR male rats
actually demonstrate reduced levels of FBM as compared WT littermates. Similar to other
effects, no differences are seen at 4 weeks (F3,25 = 2.216, P = 0.147), but are present at 6
weeks (F3,40 = 10.541, P < 0.01), 8 weeks (F3,34 = 7.769, P < 0.01) and 10 weeks (F3,34 =
6.662, P < 0.05). Tfm also reduced raw FBM, as adiposity is lower in male rats expressing
this mutation at 6 weeks (F3,40 = 4.718, P < 0.05), 8 weeks (F3,34 = 11.640, P < 0.01) and
10 weeks (F3,34 = 12.837, P = 0.001). This is to be expected, due to lower levels of whole
body mass found in Tfm males, as compared to WT littermates.
These results tell us that transgene expression reduces FBM in male rats (both raw
fat, and when corrected for body weight). Similar main effects are seen due to Tfm, with
mutant males demonstrating reduced FBM% (as well as lower raw FBM value, which
was expected due to lower overall body mass in the mutants). This finding was surprising,
as it identifies a novel role for muscle AR in reducing FBM.
3.3.4. No Effects on Body Composition of HSA-AR Female Rats
We also examined body composition of WT, HSA-AR, Tfm and HSA-AR/Tfm
female rats. It was not expected that the transgene would have pronounced effects on
females, due to significantly reduced levels of circulating T in this sex.
Analysis of whole body mass showed no differences between groups at any time
point (Fig 3a). Females were also investigated for effects on LBM% (Fig 3b), and FBM%
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(Fig 3c). No significant differences were found in body composition between females at 4
weeks, 8 weeks or 10 weeks (see Appendix A for complete statistics). Interestingly, a
transient effect of HSA-AR was found at 6 weeks of age. Analyses of body composition
found that HSA-AR increases LBM% (F3,44 = 13.249, P < 0.01) and decreases FBM%
(F3,44 = 14.335, P < 0.001). These effects occur in the same direction as those seen due to
HSA-AR in males. No effects due to Tfm were found, although females can only be
heterozygous for the mutation. As mentioned, these effects of the transgene were
transient, and disappeared when females were again measured at 8 weeks and 10 weeks.
Generally speaking, therefore, differences in body composition due to HSA-AR
expression are not seen in females, likely due to low levels of circulating T.
3.3.5. Individual Muscle and Fat Pad Weights
In order to confirm the differences in body composition seen in HSA-AR males,
adult male rats were dissected (16 weeks of age), and individual skeletal muscles and fat
pads were weighed. Analysis of EDL muscle mass (Fig 4a) shows no effect of HSA-AR
on total mass (F3,45 = 0.650, P = 0.847). Tfm males did have lower EDL mass (F3,45 =
6.083, P < 0.05), although this was expected due to lower overall body weight found in
male mutants. Similarly, mass of individual AT muscles (Fig 4b) also was not affected by
HSA-AR expression (F3,45 = 3.70, P = 0.546). Reduction in AT mass due to Tfm was not
quite statistically significant, although it strongly approached the set α value (F3,45 =
3.874, P = 0.055). Thus, the mass of individually dissected skeletal muscles does not
differ between HSA-AR and WT animals. This confirms the analysis derived from DXA
scanning, which found no differences in raw LBM due to HSA-AR expression.
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To confirm differences found in FBM through DXA, visceral WAT fat pads were
dissected from perigonadal depots of male rats and weighed. Analysis of fat pad mass
(Fig 4c) found that these pads are significantly reduced in mass in HSA-AR rats (F3,45 =
16.370, P < 0.001). Thus, transgene expression seems to decrease raw FBM. No effect of
Tfm was found (F3,45 = 2.200, P = 0.147). No interactions were found. Pairwise
comparisons of animals on the same genotypic background showed that HSA-AR reduces
fat pad mass. HSA-AR males have lighter fat pads than WT males (t29 = 1.877, P < 0.05),
and HSA-AR/Tfm males have lighter fat pads than Tfm males (t16 = 3.557, P < 0.05).
Therefore, analysis of individual fat pads confirms the analysis derived from DXA,
suggesting that HSA-AR (or, more generally, myocyte AR) is capable of reducing FBM.
3.3.6. HSA-AR Similarly Affects Body Composition in L78 Mice
We conducted body composition analysis on HSA-AR L78 mice (Monks et al.,
2007) in order to generalize the effects of the transgene in other species, as well as rule
out the possibility of regulation of these effects by ectopic expression of the transgene in
other muscle tissues (such as smooth and cardiac muscles). The HSA-AR mouse lines
have been shown to express the transgene only in skeletal muscle (Monks et al., 2007),
whereas our previously discussed RT-PCR data reveals transgene expression in heart and
smooth muscle of HSA-AR rats. DXA analysis was conducted on WT and L78 HSA-AR
male mice at 24 weeks of age, with representative DXA X-ray images shown (Fig 5a).
Measures of whole body mass (Fig 5b) reveal that L78 mice have lower levels of body
mass than WT littermates (t9 = -5.594, P < 0.001), which is consistent with previous
measures of body mass in this HSA-AR line (Monks et al., 2007). Measures of body
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composition indicate striking similarities to the effects seen in HSA-AR rats. L78 males
demonstrate increased LBM% (Fig 5c; t9 = 2.940, P < 0.05) and reduced FBM% (Fig 5d;
t9 = -3.860, P < 0.01). Analysis of raw values indicates that L78 mice have reduced LBM
as compared to WT littermates (Fig 5e; t9 = -2.994, P < 0.05), which is characteristic of
mild muscular atrophy seen in this line (Monks et al., 2007). L78 mice also exhibit
drastically reduced raw FBM (Fig 5f; t9 = -3.860, P < 0.01).
Taken together therefore, analysis of body composition in L78 HSA-AR male
mice reveals effects that largely parallel those seen in HSA-AR rats, including increased
LBM%, and decreased FBM (both raw and when corrected for body mass). This strongly
implicates AR in skeletal muscle myocytes as a major regulator of rodent body
composition.
3.3.7. Effects of HSA-AR and Tfm on Bone Content and Density
Finally, we finished our analysis of body composition in HSA-AR rats by
examining bone mineral content (BMC) and density (BMD). Bone resorption is impaired
in ARKO mice (Callawaert et al., 2009; Kawano et al., 2003), suggesting that AR is
important for maintenance of bone strength. We examined parameters of bone strength in
male and female rats, as well as male L78 mice. BMC refers to the total amount of bone
mass in the body. BMD is measured by correcting BMC for total body volume. However,
because the DXA machine is incapable of determining total body volume, it instead uses
scanned bone area in order to determine BMD values. For that reason, differences in total
body/bone area can strongly skew any differences in BMD.
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Unexpectedly, we found that HSA-AR significantly decreases BMC in male rats
(Fig 6a). No differences were found at 4 weeks, but a main effect of HSA-AR is evident
at 6 weeks (F3,42 = 5.659, P < 0.05), 8 weeks (F3,34 = 6.096, P < 0.05), and 10 weeks (F3,34
= 9.542, P < 0.01). Tfm was also found to decrease BMC at these age points as well,
which was to be expected due to lower body mass in the mutant males. When BMD
values were derived for males, however, these effects due to HSA-AR were abolished
(Fig 6c). No differences in BMD were found due to HSA-AR between male rats at 6
weeks (F3,45 = 0.410, P = 0.526), 8 weeks (F3,34 = 0.042, P = 0.839) and 10 weeks (F3,34 =
0.650, P = 0.431). Interestingly, a main effect of Tfm was found in reducing BMD in
males at 4 weeks of age (F3,25 = 5.655, P < 0.05). While this effect is similar to that seen
in male ARKO mice (Kawano et al., 2003), it is transient, and does not persist post-
puberty.
Examination of bone density in females seems to support the results derived from
males. For the most part, no differences are found in BMC in females (Fig 6b). However,
a transient effect of HSA-AR is found at 6 weeks of age, with transgene expression
reducing BMC in females (F3,44 = 10.951, P < 0.01). The directionality of this effect is
consistent with that seen in males, and is also seen transiently at 6 weeks (which is
consistent with other changes in body composition seen in HSA-AR females). The lack of
differences at other age points likely reflects lower circulating T levels in females. No
effects of Tfm are found with regards to BMC. Analysis of BMD in females (Fig 6d)
finds no significant effect of either HSA-AR or Tfm.
Finally, we examined parameters of bone strength and density in HSA-AR L78
male mice. Similar to the effects seen in HSA-AR rats, L78 male mice also have lower
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levels of total BMC, as compared to WT littermates (Fig 6e; t9 = 2.298, P < 0.05).
However, as previously mentioned, there is a significant difference in total body mass
between WT and L78 males. Analysis of BMD from these mice reveals no significant
differences between groups (Fig 6f; t9 = 0.249, P = 0.809).
Therefore, in summary, these results indicate that transgene expression results in
reduced BMC in HSA-AR male rats and mice, but not female rats (although this effect
exists transiently at 6 weeks of age). The fact that this effect persists in male mice
suggests that AR in skeletal muscle may possibly be mediating bone development.
Whether this difference in BMC is related to direct action of myocyte AR, or due to
indirect influences (such as differences in FBM), remains to be seen.
3.4. Experiment III: Testosterone Treatment of Adult Female Rats
We found that transgene expression had significant effects on body composition
of male HSA-AR rats and mice, but not in female rats. Thus, we attempted to induce
these previously described effects in adult female rats through acute treatment with
exogenous T. Females had body composition measures completed at baseline. Following
baseline measures (Week 0), they were treated with T capsules for a period of 4 weeks
(Weeks 1-4), during which time they had their body composition analyzed weekly. After
this (at Week 4), T capsules were removed, replaced with vehicle (control) capsules, and
body composition was measured weekly again for 4 weeks (Weeks 5-8). We were mainly
interested in seeing how exogenous T treatment could regulate body composition in HSA-
AR females. Separate ANOVAs were carried out for each phase of the treatment regimen:
the ‘T-Treatment’ phase (Weeks 1-4, using Week 0 as a baseline reference) and the ‘T-
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Withdrawal’ phase (Weeks 5-8, using Week 4 as a baseline). ANOVAs examined within-
subject effects on body composition parameters over the treatment, between-subject
effects of HSA-AR, and an interaction between the two. When an interaction was found,
a pair-wise t-test was conducted between the week being measured, and the relative
baseline week (for comprehensive statistical values, see Appendix A). Representative
DXA images of a WT female and her HSA-AR littermate over the course of the treatment
regimen are depicted (Fig 7a).
3.4.1. T-Treatment Regulates Body Composition of HSA-AR Females
We first examined HSA-AR females, in order to determine if their body
composition was significantly affected by treatment with exogenous T.
During the ‘T-Treatment’ phase, LBM% significantly increased in HSA-AR
females, with a main effect of HSA-AR found between-subjects over this treatment
regimen (Fig 7b). HSA-AR females experienced a significant increase in LBM% at 3
weeks (t8 = -3.433, P < 0.01) and 4 weeks (t8 = -2.614, P < 0.05), as compared to baseline
values. HSA-AR females also experienced a significant reduction in FBM% over this
phase of treatment (Fig 7c). A main effect of reduced FBM% was found within-subjects.
Pairwise comparisons show that HSA-AR females had lower FBM% after 3 weeks (t8 =
3.406, P < 0.01) and 4 weeks (t8 = 2.687, P < 0.05) of the ‘T-treatment’ phase, as
compared to baseline values. These differences due to treatment carried over when HSA-
AR animals were examined for changes in raw mass. Unlike HSA-AR males measured
previously, T-treated HSA-AR females experience increases in raw LBM (Fig 7d).
Compared to baseline, HSA-AR females show increases in raw LBM beginning as early
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as week 1 (t8 = -3.036, P < 0.05), with consistently elevated LBM at weeks 2 (t8 = -3.972,
P < 0.01), 3 (t8 = -5.617, P < 0.001) and 4 (t8 = -5.064, P < 0.001). Similar changes are
seen in raw FBM of HSA-AR females over the course of T-treatment. Overall, during this
phase, HSA-AR females experience a decrease in raw FBM (Fig 7e). While differences
in LBM are seen rather quickly, reduction in raw FBM is not seen until 3 weeks
following T-treatment (t8 = 2.470, P < 0.05). Thus, it appears that T-treatment of adult
HSA-AR females improves body composition, but that effects in lean muscle tend to
precede those occurring in WAT.
After 4 weeks, T-capsules were removed and replaced with empty vehicle
capsules, followed by weekly body composition measures. During this ‘T-Withdrawal’
phase, HSA-AR females found their body composition parameters returning toward their
original baseline levels. LBM% was significantly lower than week 4 (last week of T-
treatment) by week 6 (t8 = 4.604, P < 0.01), and persisted at week 7 (t8 = 2.337, P < 0.05)
(Fig 7b). Similarly, FBM% was increased in HSA-AR females following T-withdrawal,
with FBM% being significantly higher than week 4 by week 6 (Fig 7c; t8 = -4.724, P <
0.001). Raw values of body composition also seemed to regress back toward original
(week 0) baseline measures following removal of T. The impressive increases in LBM
were followed by a reduction in this parameter by week 6 (Fig 7d; t8 = 5.675, P < 0.001).
FBM, which had been reduced over the course of T-treatment, also increased following
T-withdrawal (Fig 7e). As compared to week 4, FBM values had significantly increased
by week 6 (t8 = -3.720, P < 0.01). This data supports the postulation that the effects of the
transgene are dependent upon circulating T, as subsequent removal of exogenous T
results in abolishment of these effects.
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3.4.2. T-Treatment Does Not Affect Body Composition of Wild-type Females
Next, body composition of WT females was examined over the course of the
treatment regimen. Unlike HSA-AR females, it was found that T treatment did not
significantly alter body composition parameters in WT females. No significant effects of
HSA-AR were found between subjects across treatment, nor were any within-subject
differences found for any of the parameters, including LBM% (Fig 7b), FBM% (Fig 7c),
raw LBM (Fig 7f) and raw FBM (Fig 7g). This suggests that the increase in exogenous T
given to WT females had no effect on overall body composition. Thus, this data indicates
that in HSA-AR females, the induction of changes in body composition are due to
transgene expression in myocytes, and not simply due to increased levels of circulating T.
3.5. Experiment IV: Adipose Histology
Previously discussed results indicate that muscle AR is sufficient for the reduction
of adiposity that is seen in HSA-AR rats. This finding was unexpected, and we chose to
examine it further on a cellular level through analysis of individual adipocytes. Thus, at
16 weeks of age, adult male rats were overdosed with sodium pentobarbital and
perigonadal fat pads were dissected for adipose histology.
3.5.1. HSA-AR Expression Reduces Adipocyte Area
Representative images of adipocytes from WT, HSA-AR, Tfm, and HSA-AR/Tfm
adult male rats are shown (Fig 8a). We measured adipocyte area in order to determine if
the size of these cells was altered by transgene expression. Interestingly, it was found that
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HSA-AR expression reduces overall adipocyte area (Fig 8b). A main effect of HSA-AR
was found, with male HSA-AR animals demonstrating reduced adipocyte area, as
compared to WT littermates (F3,18 = 16.370, P < 0.001). No main effect of Tfm was found,
although Tfm males (without the transgene) were found to have the largest adipocyte area,
when compared to WT animals, which had the next highest area (t10 = -2.645, P < 0.05).
No interaction was found between HSA-AR and Tfm.
Frequency distributions were then calculated, in order to visually depict
differences in proportions of cell size (Fig 8c). As can be seen, male rats expressing the
transgene experience a shift toward the left (smaller cells), as compared to WT littermates
on the same genetic background. This indicates that HSA-AR males have an increased
proportion of smaller adipocytes than WT males. Taken together, therefore, these results
indicate that differences in adiposity between WT and HSA-AR males can be
demonstrated at a cellular level, with reduced size of individual adipocytes noted in HSA-
AR animals.
3.6. Experiment V: Energy Balance and Metabolic Analyses
We hypothesized that the differences seen in body composition of HSA-AR males
(particularly the reduction of FBM) were mediated by changes in overall energy balance
and metabolism. Reduction in overall FBM may occur due to increases in energy
expenditure. Thus, we examined energy expenditure in HSA-AR animals by examining
basal metabolism (using the indirect measure of O2 consumption) and spontaneous
activity level.
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3.6.1. HSA-AR Expression Increases Resting Metabolism in Rats and Mice
Analysis of resting metabolic rate (RMR) was accomplished through the use of
indirect calorimetry. In this case, O2 consumption was measured, with higher levels of
this parameter associated with increased RMR (Holloszy, 2008). At 12 weeks of age,
male rats were placed in a gas-exchange chamber, and O2 consumption was measured
through the use of an O2 sensor (Fig 9a). Interestingly, a main effect of HSA-AR was
found, with transgene expression being associated with higher levels of O2 uptake (F3,35 =
10.817, P < 0.01). No main effects for Tfm, or an HSA-AR X Tfm interaction, were
found. However, when RMR was derived after correction for body weight (Fig 9b), a
main effect of HSA-AR was not noted, although it did approach significance (F3,35 =
7.574, P = 0.1). Here Tfm actually demonstrated a significant effect (F3,35 = 14.348, P <
0.001), which was to be expected due to the lower body mass of Tfm males. An
interaction between HSA-AR and Tfm was found (F3,35 = 1.40, P < 0.01), with HSA-
AR/Tfm males demonstrating the highest RMR levels.
We also examined O2 consumption and RMR in HSA-AR L78 mice, in order to
test the generality of our effects (Fig 10a). WT and HSA-AR male mice were measured in
adulthood (approximately 24-30 weeks of age), using a similar paradigm as that which
was used for rats. Although there was a trend toward higher O2 consumption in HSA-AR
L78 male mice (as compared to WT littermates), this effect was not significant (t9 = -
0.828, P = 0.429). However, derivation of RMR (which accounts for disparities in overall
body mass), indicates that RMR levels are increased in HSA-AR L78 males (Fig 10b; t9 =
-2.286, P < 0.05). Therefore, this evidence seems to indicate that HSA-AR male rodents
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may be hypermetabolic, resulting in increased rate of energy expenditure, and
subsequently providing a mechanism for the reduction in adiposity seen in these animals.
3.6.2. Spontaneous Activity is Not Affected by the Transgene
Another major component of energy expenditure is physical activity, and thus we
examined this parameter in HSA-AR male rats. ARKO mice have been shown to
demonstrate reduced voluntary activity, suggesting that AR-activation may be important
in mediating this behaviour (Ophoff, Callewaert et al., 2009). Animals were investigated
at the same ages at which body composition was analyzed (4, 6, 8 and 10 weeks of age).
No differences in spontaneous activity were found between groups at any age of
examination (Fig 11). No main effects of HSA-AR or Tfm (or interaction between the
two) were found for male rats at any time-point in question. Thus, it remains unlikely that
AR in myocytes is capable of modulating physical activity in these animals.
3.7. Summary
Characterization of HSA-AR expression in transgenic animals shows that the
transgene is expressed only in myocytes. Expression of the transgene is associated with
profound changes in body composition of HSA-AR male rats and mice, which includes:
Increased LBM%, decreased FBM%, decreased raw FBM, and reduced BMC. These
differences are confirmed by dissection and weighing of individual muscles and fat pads.
All of these findings are also demonstrated in HSA-AR/Tfm rats, suggesting that AR in
myocytes alone is sufficient to stimulate these effects. While these changes in body
composition are not found in HSA-AR female rats, they can be induced through acute T
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treatment, showing that these effects are likely T-dependent, and do not occur as a
consequence of transgene expression alone. Finally, it has been shown that HSA-AR
expression modulates metabolic parameters, inducing energy expenditure by elevating
RMR, but not affecting activity level. Please refer to Appendix A for full statistical
analyses.
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Chapter 4: Discussion 4.1. Overview
Taken together, our results present an in vivo model whereby increased androgen
signaling in myocytes alone is sufficient to induce robust changes in body composition
(increasing LBM% and reducing FBM%) and systemic metabolism. These differences
were also found in HSA-AR L78 male mice, indicating that they are a general result of
overexpression of AR in skeletal muscle myocytes. In addition, several of our results
suggest that these effects are dependent upon T. First, in males, the effects were not
evident until 6 weeks of age, when these animals reach puberty and circulating T levels
increase drastically. Secondly, in female rats, these effects could be induced through
acute treatment with exogenous T, indicating that transgene expression alone was not
sufficient (although one cannot rule out the potential roles of female sex chromosomes
and ovarian hormones). Finally, in these T-treated female rats, removal of T abolished
these effects, and females returned to their baseline levels of body composition within 1-2
weeks. Therefore, it is clear that the differences induced in body composition by myocyte
AR are dependent upon activation of the nuclear receptor by T in this cellular population.
It seems likely that these differences in body composition result from regulation of
systemic metabolism, evidenced by increased oxygen consumption and RMR in HSA-AR
male rats and mice. Finally, no differences were found in spontaneous activity level of
rats, indicating that the augmented RMR level of these animals is likely the driving force
behind their increased energy expenditure and changes in body composition.
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4.2. Myocyte AR and Body Composition
This set of experiments provides insight into how androgens affect body
composition. Notably, they provide evidence for androgenic action on muscle to reduce
fat via increases in metabolic rate. Despite widespread belief that androgen
supplementation alters body composition, experimental support for this idea has only
recently been obtained (Bhasin et al., 1997). Furthermore, T-treatment of older men has
been successful in limiting the negative effects of aging on body composition, with some
scientists citing the reduced T levels that men experience as they age as a prominent
contributing factor to late-onset metabolic disease (Bhasin et al., 2006). These effects of
androgens on body composition have been verified by basic science, gene-targeting
research on ARKO mice. Ablation of AR throughout the body results in atrophy of
skeletal muscle (MacLean et al., 2008), adult-onset obesity (Lin et al., 2005; Fan et al.,
2005) and hypometabolism (Fan et al., 2005). While these effects are interesting, and
provide further credence to the idea that androgens are capable of modulating body
composition, they do not identify the tissue-specific AR that are necessary for mediating
them. Identifying these targets is a major goal of research in androgen therapy, for the
purposes of exploiting them as treatments for obesity and metabolic disease. Our study
employed a gain-of-function transgene (HSA-AR) that results in overexpression of AR
protein in myocytes. Furthermore, by crossing HSA-AR rats with those possessing the
Tfm mutation, we studied the resultant HSA-AR/Tfm progeny, which express AR only in
myocytes. Using these powerful genetic tools, we have gained significant insight into
how AR in myocytes contributes to changes in body composition.
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4.2.1. Lean Muscle Mass
In popular culture, the association between androgens and skeletal muscle is a
common one (Kadi, 2008). The use of anabolic-androgenic steroids (which are
pharmacological agents analogous in structure to endogenous androgens, and that also
bind to AR) has been linked with performance-enhancement in various sports – with the
belief that use of these drugs leads to increased lean muscle mass, and thus increased
strength and power. More specifically, human studies in adult men have found that T
supplementation has widely been associated with hypertrophy of skeletal muscle fibers
(Sinha-Hikim et al., 2002). T-dependent hypertrophy has been found in young men, as
well as through T-treatment of elderly males (Bhasin et al., 2001). Androgens have found
limited clinical use, but have been shown to have beneficial effects in increasing muscle
mass of patients suffering from HIV wasting (Bhasin et al., 2000), renal disease
(MacDonald et al., 2007) and burns (Wolf et al., 2006). However, despite these
associative clinical studies, the idea that T and other androgens directly increase lean
muscle mass is not completely accepted, due to limited evidence of sufficient functional
benefits in T-treated humans (MacLean & Handelsman, 2009).
For this reason, basic science research using cell cultures and animal models have
attempted to bridge this gap of knowledge by identifying cellular and molecular
mechanisms by which androgens/AR can influence lean muscle mass. Developmental
work has shown direct function for T in progenitor cells. Treatment of pluripotent stem
cells (or more mature satellite cells) with T enhances commitment of these cells to the
myogenic lineage, proliferation of satellite cells, and myoblast/myofiber protein accretion
(Singh et al., 2003; Joubert & Tobin, 1995; Chen, Lee, Zajac & MacLean, 2008).
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Treatment of these cells with an AR antagonist (such as flutamide) abolishes these
effects. Therefore, it is largely well-accepted that androgens have developmental effects
in promoting myogenesis. However, identifying mechanisms by which androgens can
contribute to hypertrophy of mature skeletal muscle remains elusive.
Work using animal models has provided some promising evidence of AR effects
on adult muscle. Abolishment of AR in ARKO mice results in reduced muscular strength
and muscular atrophy (MacLean et al., 2008). However, the nature of this knockout
means that AR has been deleted from all cells, including important myocyte precursor
cells. Thus, teasing out the effects of AR in myogenesis from those in adult muscle using
whole-body ARKO is not feasible. To get around this problem, Ophoff and colleagues
(2009) used the Cre/Lox approach to generate myocyte-specific ARKO mice (mARKO).
This was done using the myocyte-creatine kinase (MCK) promoter to drive Cre-
expression, so that AR is deleted only in mature myocytes that express this promoter, but
not precursor cells (such as satellite or embryonic stem cells). The muscle phenotype in
mARKO mice is very distinct, with KO mice demonstrating reduced limb muscle mass
and strength. These mARKO mice also exhibit a change in fiber type of soleus muscle,
with an increase in type I fibers. It is also worth noting that the atrophy found in skeletal
muscle of mARKO mice is not as severe as that seen in whole-body ARKO mice,
implying the importance of AR in progenitor cells, as well as the possibility of
involvement of AR in other tissues.
In contrast to the loss-of-function paradigm employed by Ophoff et al., we used a
gain-of-function transgenic model to overexpress AR in skeletal muscle fibers. This was
done through the use of the HSA promoter. We find that overexpression of AR in adult
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muscle fibers increases LBM%, but that these effects do not carry over when
investigating raw values in males. That is, adult HSA-AR males (6 to 10 weeks) do not
demonstrate increases in raw LBM. This lack of difference was confirmed through
dissection of individual muscles, as EDL and AT muscle weight did not differ between
WT and HSA-AR littermates. However, T-treatment of adult HSA-AR female rats
resulted in significant increases in LBM, effects that were not seen through T-treatment of
WT females. Furthermore, removal of T caused a significant decline in LBM. Thus, it is
possible that myocyte AR have acute, local functions in increasing skeletal muscle mass.
Histological analysis of skeletal muscle of HSA-AR male rats by our lab also
reveals interesting insights (Fernando et al., 2010). Dissected EDL muscles from intact
HSA-AR and WT rats were obtained at 10 weeks of age. Analysis of number of
myofibers revealed no differences between WT and HSA-AR animals. Furthermore,
fiber-type analysis shows no shift in the fiber-type proportion. Interestingly, however,
selective hypertrophy of type IIb (fast-twitch, glycolytic) muscle fibers was noted. These
muscle fibers showed larger area, while other fiber-types were not significantly different
between groups. Ophoff et al. (2009) found that myocyte-specific ARKO also had no
effect on fiber-type proportion in EDL, although they did not report any differences in
fiber hypertrophy either.
On the face of it, our findings seem largely incongruent to those reported by
Ophoff et al. in their mARKO mouse model (2009). While these scientists report that
knockout of AR in myocytes reduces muscle mass, we found no effect on raw muscle
mass through overexpression of AR in myocytes (particularly in the EDL). However,
very recent evidence from Chambon and colleagues (2010) seems to support our findings.
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Here, scientists developed another mARKO model through the use of the HSA promoter.
In this study, it was found that (in contrast to Ophoff et al.) selective ablation of AR in
myocytes did not affect limb muscle mass (including the EDL), but rather had significant
effects on the strength of these muscles. Furthermore, the authors also show reduced
cross-sectional area of type II muscle fibers in the EDL, which nicely compliments the
selective hypertrophy of these fibers seen in our transgenic rats. Thus, our HSA-AR
model seems to indicate that androgens may have effects on mature myocytes. However,
the nature of these effects remains unclear. Whether myocyte AR plays a significant role
in increasing muscle mass in non-sexually dimorphic muscles will continue to be a hotly
debated topic.
4.2.2. Adipose Tissue
While research into androgen effects on skeletal muscle mass have resulted in
controversial results, very little work has been undertaken in order to determine the nature
of how androgens affect adipose tissue. Clinical studies on patient populations show that
exogenous T treatment can have beneficial effects in reducing adiposity in young and
older men (Bhasin et al., 1996; Bhasin et al., 2001). In addition, epidemiological surveys
reveal that serum T levels are inversely related to whole-body and regional fat mass in
men (Siedell et al., 1990; Derby et al., 2006). However, identification of the cellular and
molecular mechanisms involved in this androgenic reduction of fat has not been well
elucidated.
In the previous section, it was mentioned that androgens are believed to have dual
effects on skeletal muscle, mediating the development of mature myocytes from
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progenitor cells, as well as hypertrophy of existing, mature cells. However, in study of
adipose tissue, androgens (such as T) have been implicated largely in inhibiting the
development of mature adipocytes from precursor cells (termed ‘pre-adipocytes’). This
process by which progenitor precursor cells develop into mature adipocytes is known as
adipogenesis. While originally believed to be a process that occurred only during the
early stages of life, it is now known that adipogenesis occurs across the lifespan (Rosen &
Spiegelman, 2000). For this reason, reduction of adiposity (lower fat accumulation) in
adulthood can either be the result of increased fatty acid β-oxidation (metabolism of fat)
or inhibition of adipogenesis. As mentioned, androgens have traditionally been associated
with the latter process. Significant in vitro work using human and rat pre-adipocytes has
shown that differentiation of pre-adipocytes into mature adipocytes is impaired when
cells are treated with T (Singh et al., 2003; Singh et al., 2006; Gupta et al., 2008). Gene
expression assays of these cells reveal significant down-regulation of major adipogenic
factors, such as PPARγ and C/EBPα. Furthermore, this androgenic regulation of
adipogenesis is believed to be mediated by activation of AR, as pre-treatment of cells
with an AR antagonist abolishes these effects. Thus, it has largely been believed that
androgens exercise their effects on adipose tissue by binding to AR in adipocytes, and
subsequently down-regulating major adipogenic genes.
Manipulation of AR using in vivo animal models has been largely consistent with
the idea of androgen-mediated reduction in adipose tissue. Whole-body ablation of AR
results in adult-onset obesity, with significantly increased adipocyte size, and higher
frequency of larger adipocytes in ARKO mice (Lin et al., 2005; Fan et al., 2005).
Interestingly, ARKO mice were shown to have reduced expression in adipose tissue of
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adipogenic genes (including PPARγ and C/EBPα), but also decreased expression of
major genes in liver and skeletal muscle that are involved in β-oxidation (such as
PPARα). Thus, it was unclear as to whether the increased adiposity seen in late-adulthood
of ARKO mice was due to reduced adipogenesis or reduced fatty acid metabolism (or
both). Also, although these findings were interesting, they did not shed any light on which
tissue-specific AR were responsible for the increase in WAT seen in ARKO mice. With
this in mind, scientists generated mice with selective knockout of AR in adipocytes
(aARKO; Yu et al., 2008), since (as mentioned) it was largely believed that AR in these
cells mediate T’s effects on WAT by inhibiting adipogenesis. Unexpectedly, these
aARKO mice did not demonstrate the adult-onset obesity seen in whole-body ARKO.
These same scientists then examined selective knockout of AR in liver hepatocytes
(hARKO), a major organ involved in fat catabolism (Lin et al., 2008). While hARKO
mice demonstrate reduced hepatic PPARα expression, and hepatic steatosis (local
accumulation of fat in the liver), they also do not show adult-onset obesity typical of
ARKO mice. Thus, it appears that neither AR in adipocytes nor hepatocytes are sufficient
for regulation of the androgenic reduction in adiposity.
We examined FBM in our HSA-AR rats and mice, which overexpress AR in
skeletal muscle myocytes. Skeletal muscle is an attractive but often overlooked tissue in
the study of regulation of FBM. Myocytes are energetically-demanding cell populations,
and disruption of fatty acid transport into myocytes is often associated with metabolic
diseases (Zitzmann, 2009). We found that FBM is drastically reduced in HSA-AR rats
and mice, a finding that is confirmed through dissection and weighing of individual fat
pads. In rats, these effects are seen shortly after the onset of T secretion during puberty,
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suggesting that they are T-dependent. Examination of L78 mice in late adulthood shows
that this effect persists. Furthermore, T-treatment of HSA-AR female rats reduces
adiposity significantly – an effect which is abolished upon withdrawal of T. Examination
of adipocyte size in HSA-AR males reveals that they have smaller cell area than WT
littermates, and an increased proportion of these smaller cells.
These findings, that HSA-AR expression reduces fat mass and adipocyte size,
were largely surprising because AR overexpression is limited to myocytes (in L78 mice,
only in skeletal muscle myocytes). Furthermore, they were also seen in HSA-AR/Tfm
rats, which express AR only in myocytes, as provided by the transgene. This suggests that
myocyte AR is sufficient to reduce FBM. These effects are in contrast to the more
traditional model of androgens reducing WAT through activation of AR in adipocytes,
and subsequent inhibition of adipogenesis. While our model was not designed to test the
function of AR in adipocytes, it can be said with some certainty that action of AR in
WAT is not responsible for the results that we see. This is for several reasons. First, HSA-
AR mRNA expression was not found in WAT of rats through RT-PCR. Examination of
β-galactosidase expression in L78 mice also found no expression of AR in this cellular
population (Monks et al., 2007). Furthermore, these effects were also seen in HSA-
AR/Tfm male rats, which do not express AR in any non-muscle tissue, including WAT
tissue. Finally, adipogenesis is a process that is largely mediated by actions in WAT
tissue itself (Rosen & Spiegelman, 2000). It is therefore highly unlikely that local actions
of AR in muscle could be capable of inhibiting adipogenesis. Given this unlikely
scenario, logic therefore suggests that myocyte AR reduces adiposity by increasing fatty-
acid metabolism (i.e. β-oxidation). This would explain why knockout of AR only in
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adipocytes (Yu et al., 2008) or hepatocytes (Lin et al., 2008) is incapable of recapitulating
the adult-onset obesity that is seen in whole-body ARKO mice (Lin et al., 2005). It would
also explain the hypometabolism that is found in whole-body ARKO mice (Fan et al.,
2005), a finding that will be elaborated upon in a later section.
4.2.3. Bone
Sex hormones are well-known to be mediators of bone development and
remodeling during adulthood (Bland, 2000; Compston, 2001). The most prominent
effects of sex hormones in this context are often attributed to estrogens, which directly act
upon ER in bone tissue in order to increase bone strength and remodeling (Simpson &
Davis, 2000; Couse & Korach, 1999). This explains the significant increase in incidence
of osteoporosis after menopause in women, when secretion of estrogens declines
significantly – an effect that can be reduced with treatment of exogenous estrogens
(Riggs, Khosla & Melton, 2002).
The role of androgens in bone development and remodeling, however, is less
understood. AR is expressed in bone tissues, including osteoblasts and osteoclasts
(Compston, 2001). Furthermore, androgens have been shown to modulate expression of
various growth factors and cytokines involved in bone remodeling (Compston, 2001). For
example, T treatment of an osteoblastic cell line increases expression of Insulin-like
growth factor 1 (IGF-1), which is a prominent growth factor that promotes bone
development (Gori, Hofbauer, Conover & Khosla, 1999). Taken as a whole, most
associative research seems to indicate that androgens have protective functions on bone,
resulting in increased bone mass and density in males and females (Bilezikian, 2002;
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Hofbauer & Khosla, 1999). Investigation of bone parameters in whole-body ARKO mice
seems to support these findings (Kawano et al., 2003). 8-week old male ARKO mice
demonstrate high bone turnover with increased bone resorption. Overall bone mineral
content (BMC) values are reduced in these ARKO mice, and analysis of primary
osteoblast/osteoclast cultures from ARKO mice reveals that reduced androgen signaling
removes inhibition of osteoclastogenesis, resulting in breakdown and resorption of bone.
Therefore, androgens are believed to be major regulators of bone development and
remodeling, and act by binding to AR in bone and directly inhibiting osteoclastogenesis.
We investigated both BMC and bone mineral density (BMD) in HSA-AR male
rats and mice. Surprisingly, we found that BMC was significantly reduced in male HSA-
AR animals, both in rats and mice. This effect was also not present at 4 weeks of age, but
emerged after the onset of puberty, suggesting that it is T-dependent. Furthermore, this
effect was seen transiently at 6 weeks in HSA-AR female rats, a time period where other
changes in body composition were found (LBM% and FBM%). Reduced BMC is also
seen in HSA-AR/Tfm males, as compared to Tfm males, suggesting that AR in myocytes
alone is sufficient for this reduction in bone mass. No effects of HSA-AR were found on
BMD, however this measure may be confounded, as it is incorrectly calculated by the
DXA machine using bone area and not total volume. Interestingly, no effects of Tfm were
found with regards to BMD, although at 4 weeks, male rats possessing this mutation
exhibited lower BMD as compared to WT controls.
Our findings lie in stark contrast to the effects found in ARKO mice (Kawano et
al., 2003), and the typically beneficial effects on bone attributed to androgens (Bilezikian,
2002). While these studies suggest that androgens are important in maintaining and
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increasing bone mass, we find a reduction of BMC in animals overexpressing AR in
myocytes, and those expressing AR only in myocytes. These results suggest two possible
explanations. First, it is possible that action of AR directly in myocytes is contributing to
reduction of BMC. Although we did not examine expression of various bone growth
factors and cytokines, it is unlikely that myocyte AR down-regulates these factors. In
actuality, existing lines of evidence seem to indicate the opposite: that AR activation in
muscle actually promotes increases in expression of IGF-1 signaling molecules, although
whether these molecules are then capable of circulating throughout the body is unknown
(Svensson et al., 2010). A more likely explanation for the reduction in BMC seen in
HSA-AR animals is that it is due to indirect effects of the transgene – namely the
reduction in adipose tissue. Increases in WAT are paradoxically known to be associated
with decreased serum levels of adiponectin, an adipocyte-derived cytokine, which is
sometimes classified as a hormone (Arita et al., 1999). Very recent work examining
adiponectin-knockout mice reveals that these animals have higher levels of BMC, and
lower bone fragility (Williams et al., 2009). While we did not examine adiponectin
concentrations in our animals, it is reasonable to assume that WT animals would have
lower levels of the hormone, due to increased FBM. Lower levels of adiponectin would
thus explain the increase in BMC seen in WT animals, similar to those seen in
adiponectin-knockout mice. Nevertheless, more research is necessary in order to
determine mechanisms of muscle-bone and WAT-bone crosstalk.
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4.3. The Effect of Tfm on Body Composition
Our work also allows for the understanding of how whole-body ablation of AR
can contribute to changes in body composition, through the use of male Tfm rats. To that
end, we find several interesting differences in body composition between WT and Tfm
male rats that are worth discussing.
4.3.1. Comparison with ARKO Mice
As mentioned previously, several distinct phenotypes are found with regards to
body composition in ARKO mice (as compared to WT littermates). In the context of
skeletal muscle, ARKO males demonstrate muscular atrophy as well as reduced muscle
mass, coupled with impaired force production (MacLean et al., 2008). These findings also
extend to cardiac muscle, with reduced heart mass and atrophy of cardiomyocytes in
ARKO males (Ikeda et al., 2005). FBM also appears to differ in ARKO mice, as male
ARKO demonstrate adult-onset obesity (development of excess adiposity after 30 weeks
of age), coupled with increased size of individual adipocytes and heavier fat pads (Lin et
al., 2005; Fan et al., 2005). Lastly, differences in bone were also found in ARKO mice
(Kawano et al., 2003). Male ARKO mice show reduced bone mass, increased resorption
of bone (due to higher rates of osteoclastogenesis) and osteopenia. Taken together, we
can identify a body composition phenotype of ARKO mice characterized by reduced
LBM and BMC, but increased FBM. These findings were only found in males of this line,
and not females.
Our examination of body composition in Tfm males reveals several striking
differences between these mutants and their WT littermates. Examination of LBM per
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unit of body mass reveals higher levels of LBM% in Tfm males. With regards to WAT,
we also find that Tfm males demonstrate reduced FBM%, as compared to WT littermates.
However, examination of adipocytes of Tfm males in late adulthood reveals that cells
from these animals have the largest area. Finally, BMD was examined in Tfm males. It
was found that the mutation resulted in reduced BMD at 4 weeks of age, but this effect
was transient, and not found in examination of rats at later ages. In summary, therefore,
we find that body composition of Tfm males is marked by increased LBM%, reduced
FBM% and no effects on BMD, as compared to WT littermates. These differences are
largely contradictory to those previously described to have been found in ARKO male
mice.
4.3.2. Why the Discrepancy?
While it is tempting to directly compare Tfm rats with ARKO mice, there are
several important phenotypic discrepancies between the males of these strains (aside from
the obvious species difference) that should be noted. First, the ARKO is derived using
Cre/Lox technology, and subsequent deletion of the AR gene (Yeh et al., 2002). Thus, in
theory, no AR protein can be translated in any cells where this gene has been deleted. In
practice however, the efficacy of Cre-induced recombination in ARKO mice has been
called into question, as the degree of Cre-mediated excision is unknown (MacLean &
Handelsman, 2009). For example, protein expression assays of moderate sensitivity
demonstrate some AR expression in muscle of mARKO mice, although the levels are
significantly lower than those seen in WT controls (Ophoff et al., 2009). Thus, the
potential for residual AR activation remains. In contrast to the Cre/Lox technology used
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to generate ARKO, Tfm results from a single base-pair mutation, resulting in the
alteration of a single amino acid in the AR polypeptide (Yarbrough et al., 1990). The
amino acid alteration occurs in the ligand-binding domain of the AR protein, and though
the protein is translated, it is incapable of binding to AR, and thus being activated.
Nevertheless, Tfm AR protein can be identified using immunohistochemistry (IHC), and
some residual AR activation is found in tissues of Tfm males (Zuloaga et al., 2008).
Therefore, there are significant differences between ARKO mice and Tfm rats in terms of
AR protein translation and function.
Another major difference between ARKO mice and Tfm rats is exemplified by
circulating hormone levels. In ARKO mice, serum T and DHT levels are drastically
reduced, as compared to WT males (Yeh et al., 2002; Kawano et al., 2003). This is
believed to be due to atrophy of testes in these animals. Similar findings are found in Tfm
mice (Jones et al., 2003), who demonstrate decreased circulating T levels, due to atrophy
of testes. In contrast, Tfm male rats have been found to have significantly higher levels of
circulating T, reaching the highest limits seen in WT rats (Naess et al., 1976). This occurs
despite testicular atrophy in these animals, and is believed to be due to lack of AR in the
hypothalamus, and thus uncoupling of negative feedback in these animals. This is further
evidenced by increases in gonadotropin (FSH and LH) levels in these rats. Therefore, not
only do ARKO mice and Tfm rats differ in translation of AR protein, but their hormonal
profiles are also rather distinct.
Our findings regarding body composition in Tfm rats are largely contradictory to
the existing literature regarding androgens and body composition (Zitzmann, 2009). First,
it is important to note that these findings in Tfm rats do not necessarily cast doubt on our
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findings regarding the HSA-AR transgene and myocyte AR. While Tfm does improve
body composition, rescue of AR in myocytes of Tfm males shifts them further in the same
direction. Therefore, it is unlikely that loss of AR in myocytes of Tfm males causes the
improvement in body composition. Nevertheless, it is puzzling that inactive AR in any
tissue would be capable of improving these parameters.
Various explanations may account for these counterintuitive effects of Tfm. First,
it is important to note the differences in developmental periods at which ARKO mice
were measured, as compared to our experiments in Tfm rats. For example, in the original
study employing ARKO mice, differences in FBM were investigated at 8 weeks of age
(Yeh et al., 2002). Here, the scientists found reduced adiposity and smaller adipocytes in
ARKO mice (similar to our findings in Tfm rats), and initially concluded that AR might
be necessary for adipogenesis. It was not until these animals were subsequently followed
into late-adulthood (30 weeks of age) that increased adiposity was found (Lin et al.,
2005). We investigated Tfm males from only 4 weeks of age until 10 weeks, so it is
possible that these mutants may develop obesity in later adulthood.
The higher levels of circulating T also complicate the picture of Tfm males.
Aromatization of T results in synthesis of estrogens, which have diverse functions (the
possible role of estrogens in mediating differences in LBM and FBM that we see will be
discussed in the next section). Although activity of aromatase enzymes is shown to be
reduced in brain regions of Tfm males (Roselli, Salisbury & Resko, 1987), higher serum
concentrations of estrogens have also been reported in these mutant males, presumably
due to increased levels of T substrate (Vanderschueren, Boonen & Bouillon, 1998). While
we did not measure levels of serum estrogens in our animals, it is possible that elevated
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levels would contribute to major differences between Tfm and WT brothers, as estrogens
have major regulatory effects on body composition (namely through reduction of adipose
tissue).
Study of bone structure in Tfm males shows that these animals are able to obtain
similar trabecular bone volumes as WT male littermates (Vanderschueren et al., 1993).
Furthermore, gonadectomy of Tfm males significantly reduces BMD in these animals
(Vanderschueren et al., 1994). This is strong evidence that this effect in Tfm males is a
result of T levels. However, since AR is non-functional in these animals, a metabolite of
T is likely mediating this effect. The authors hypothesize that the metabolite in question
may be estrogen. We have previously discussed that estrogens have beneficial effects in
improving BMC and BMD (Simpson & Davis, 2000). Treatment of male rats with an
aromatase inhibitor (which prevents the conversion of T to estrogens) during
development impairs BMD, highlighting the importance of estrogens in mediating BMD
in both males and females (Vanderschueren et al., 1997). Thus, our experiments show
that Tfm male rats do not display the impaired BMD that is characteristic of ARKO male
mice. However, this protection against osteopenia in Tfm rats (as evidenced by our
results) may potentially be a result of increased estrogen action in these rats; a
consequence of high T levels.
4.4. Interactions with Estrogens
The influence of estrogens is an important aspect that must be discussed, as
estrogens are known to have profound effects upon body composition in both males and
females. These effects have significance not only for the previously mentioned Tfm male
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rats (who demonstrate increased estrogen levels), but also for our T-treated adult females,
whose estrogen signaling was likely impaired due to treatment with exogenous T. The
withdrawal of T from these rats originally results in regression toward baseline values
(that is, increased FBM% and decreased LBM%). However, toward the end of
monitoring, both groups display non-significant trends in body composition that mimic
those that occur due to transgene expression (decreased FBM% and increased LBM%).
These latter changes may reflect an increase in estrogenic signaling in females of both
genotypes, due to removal of T, and subsequent increase in circulating gonadotropins.
Thus, understanding of any estrogenic influences on effects of the transgene are important
for fully elucidating the sub-cellular mechanisms that lead to the phenotypes seen.
4.4.1. Estrogens and Adipose Tissue
It has been long understood that estrogen signaling is important in reducing fat
mass in females (Shi et al., 2009). Estrogen receptor null (ERKO) mice demonstrate
increased adiposity as early as 6 weeks of age (Heine et al., 2000). Furthermore, these
effects are apparent in male mice as well as female mice, a finding that is not replicated in
ARKO mice, whose effects are only seen in males (Lin et al., 2005). Thus, estrogens
appear to be important in mediating adiposity in both sexes, although the effects are more
pronounced in females (Heine et al., 2000). In fact, many have made the argument that
androgens exert their effects on body composition in males only indirectly – through
aromatization to estrogens. Aromatase-knockout (ArKO) male mice exhibit obesity in
adulthood, despite elevated serum T levels, and this effect was abolished through
treatment with exogenous 17β-estradiol (Jones et al., 2000). Furthermore, treatment of
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gonadectomized male mice in adulthood with DHT (a non-aromatizable androgen) results
in increased fat mass, as compared to a control group treated with 17β-estradiol
(Moverare-Skrtic et al., 2006). Strangely, however, the DHT treated group did not differ
from sham-operated animals. Therefore, it is clear that estrogens have significant effects
on WAT, and many have argued that these effects are more prominent than (or are the
consequence of) effects of androgens.
4.4.2. Estrogens, HSA-AR and Tfm
Given these robust effects of estrogens, it is important to dissect any influence
they may have on the effects that we see. There are two separate venues from our
experiments where estrogens may be especially important. First, as mentioned, Tfm male
rats have been shown to have higher levels of estrogens (Vanderschueren, Boonen &
Bouillon, 1998), which may explain their surprising differences in body composition.
Secondly, we treated WT and HSA-AR adult female rats with exogenous T, in order to
determine the necessity of androgens in mediating this effect. In doing this, estrogen
signaling was likely attenuated significantly, complicating the hormonal state of these
animals.
Before considering the role of estrogens in each of these separate contexts, one
must first confront the mechanism of estrogenic effects on WAT. Interestingly, ERKO
mice do not exhibit significant differences in energy intake, suggesting that this parameter
does not account for the increased adiposity seen in these animals (Heine et al., 2000).
While feeding behaviour was not different between genotypes, energy expenditure (in the
form of O2 consumption) was found to be reduced in male and female ERKO mice.
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However, the authors did not disclose whether these differences were due to alterations in
BMR, adaptive thermogenesis or activity level (or some combination of these factors).
Furthermore, it was also unclear as to which tissues were involved in mediating this
estrogenic reduction in adiposity. An answer to both of these questions was provided by
Musatov et al. (2007), who delivered an ER-specific siRNA directly to the ventromedial
hypothalamus, reduced ER expression in this brain area, and found that it resulted in
obesity due to reduced activity levels in treated animals. This work was supported by
research on aromatase-null mice, which also exhibit normal RMR and feeding, but
reduced spontaneous activity (Jones et al., 2000). Despite the evidence cited in these
papers, relatively new work indicates that estrogen signaling in the hypothalamus does
seem to influence feeding behaviour (Shi et al., 2009), and is believed to function by
heightening anorexigenic signaling molecules of the leptin pathway (Gao et al., 2007).
Thus, it appears that estrogens influence body composition (more specifically WAT)
through direct actions on the hypothalamus, and by modulating both feeding behaviour
and activity level.
With this information in mind, one can postulate as to whether estrogen plays a
role in mediating effects that we see due to HSA-AR. First, it is highly unlikely that
estrogen plays a major role in regulating the differences seen in WAT in HSA-AR males.
No differences were found in spontaneous activity of HSA-AR males. Earlier work
monitoring feeding behaviour also shows no significant effect of transgene expression on
food intake (Rao & Monks, unpublished data). Thus, there is little reason to believe that
the effects seen in HSA-AR males can be attributed to influence of estrogen.
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Conversely, with regards to the T-treated females, estrogenic effects become a
real possibility, since we did not ovariectomize these females. Thus, one might argue that
the effects seen due to T-treatment might actually be secondary effects caused by
suppression of ovarian hormones. While we did not measure energy intake or expenditure
in these females, this possibility is also unlikely. First, as has been discussed, estrogen
signaling is important for reduction of WAT, but we see a similar result from suppression
of estrogen signaling, suggesting minimal effect of ovarian hormones. Secondly, these
effects manifest themselves only in HSA-AR females, indicating that the dosage of
exogenous T itself (and subsequent suppression of estrogens) was not responsible for
these effects, but rather binding of this T to the transgenic AR. Thus, while estrogens are
important in mediating WAT in females, it is unlikely that they are involved in regulating
the effects that we see in our T-treated females.
Finally, the influence of estrogens in regulating WAT in Tfm males is possibly a
very real one. It has been shown that estrogen levels are significantly elevated in Tfm
males (Vanderschueren, Boonen & Bouillon, 1998). We did not measure energy intake in
Tfm males, and thus it is not known whether this parameter differs from normal WT
males. Examination of activity level found no differences between Tfm and WT males.
Aside from measuring these parameters, one might also treat Tfm males with an
aromatase inhibitor, in order to limit estrogen levels. Finally, gene targeting may also be
employed. Ablation of AR in the brain could result in increased brain estrogen levels.
While a model of AR ablation only in the nervous system was generated very recently
(Juntti et al., 2010), energy balance parameters have not been measured. Thus, we
propose that the unexpected shift in body composition seen in Tfm male rats (increased
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LBM% and decreased FBM%) may be due to higher levels of estrogens, which is a
consequence of higher serum T levels. Similar effects are not seen in other rodent models
of androgen-insensitivity (i.e. Tfm mice or ARKO mice), as these strains exhibit
significantly lower levels of circulating T. Such a potentiality represents the major
interplay between androgens, estrogens and body composition.
4.5. Myocyte AR and Oxidative Metabolism
Overall, our results suggest a model whereby increased (or sole) expression of AR
in myocytes is sufficient to induce hypermetabolism, and subsequent improvement of
body composition through reduction of FBM. This is a novel finding, and thus,
elucidation of the mechanism by which myocyte AR is able to alter systemic metabolism
remains an important step.
4.5.1. Local Effects on Skeletal Muscle Metabolism
Our investigation of basal metabolism shows that RMR is heightened in HSA-AR
male rats and mice. This is evidenced by increased O2 consumption in these animals – a
major indicator of elevated systemic metabolism. We also found no differences between
male rats in physical activity, nor was feeding behaviour altered between groups (Rao &
Monks, unpublished data), suggesting that this disparity in RMR was likely the major
driving force behind the changes we see in body composition.
It is likely that these differences might be due to local action of androgens in
skeletal muscle. It is now well understood that alterations in skeletal muscle can strongly
contribute to changes in systemic metabolism (Harrison & Leinwand, 2008). Hypertrophy
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of skeletal muscle fibers is associated with increased uptake of glucose and fat, and the
subsequent catabolism of these substrates for the purpose of energy harvest. Therefore, it
is possible that myocyte AR may have local effects on muscle glycogen content. This
possibility was examined in mARKO mice (Ophoff et al., 2009). Here, scientists
examined skeletal muscle glycogen content using biochemical enzymatic fluorometric
assay, and found no significant differences between groups, although a trend toward
lower glycogen content in mARKO males was seen. In addition, treatment of adult rats
with exogenous T has been shown to increase glycogen accumulation in soleus muscle
(Cunha et al., 2005). Furthermore, glycogen content is increased in HSA-AR male mice
(both L78 and L141; Musa et al., In Prep). Here, electron microscopy of skeletal muscle
sections from EDL of HSA-AR animals revealed increased aggregation of glycogen,
while Periodic acid-Schiff (PAS) staining was heightened in HSA-AR EDL sections.
Thus, it appears possible that myocyte AR may have local effects on muscle glycogen
accumulation.
4.5.2. Mitochondrial Biogenesis and Enzyme Activity
Differences in skeletal muscle metabolism are most often linked with muscle
mitochondria, which are the major sources of energy generation in the cell, and are
responsible for catabolism of glucose and fatty acids (Holloszy, 2008). Therefore,
alterations in muscle mitochondrial biogenesis or enzyme activity will likely have major
implications for systemic metabolism and body composition. Recent work from our lab
has shown that muscle mitochondria are significantly altered in HSA-AR male rats and
mice. In HSA-AR male rats, dissected EDL muscles were homogenized and mitochondria
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were isolated. Examination of electron transport chain (ETC) enzymes reveals higher
activity of these proteins in HSA-AR males (Fernando et al., 2010). Activity of all four
ETC complexes was heightened in HSA-AR males, and interestingly activity of these
enzymes is also reduced in muscles of Tfm males (as compared to WT males). HSA-
AR/Tfm male rats demonstrate equivalent ETC activity to that seen in WT males,
presumably due to contrasting action of HSA-AR and Tfm on skeletal muscle
mitochondria in these males.
Similar findings are reported in HSA-AR mouse lines (Musa et al., In Prep). ETC
activity is heightened in L78 males, as well as T-treated L78 and L141 females. This
occurs despite significant muscular atrophy in T-treated L141 females. Electron
microscopy of EDL from these animals shows that HSA-AR males (and HSA-AR
females treated with T) demonstrate proliferation of mitochondria. These organelles are
larger in L78 males than WT littermates, and more numerous in L141 males than WT
littermates. Similar results are found in T-treated females of both lines. Although
symptomatic T-treated females and L78 males demonstrate a loss of oxidative fibers,
mitochondrial content appears to be augmented in L78 males. Thus, it seems that
mitochondrial biogenesis and enzyme activity are both increased in HSA-AR mice.
These alterations in myofiber mitochondria provide some insight of a mechanism
by which increased myocyte AR contributes to heightened systemic metabolism and
improved body composition (through increased RMR). Increased myocyte mitochondrial
biogenesis has been associated with increased cellular respiration and β-oxidation of fatty
acids (Cha et al., 2006; Holloszy, 2008). Similarly, increased activity of ETC enzymes in
skeletal muscle has been shown to have similar results (Birch-Machin & Turnbull, 2001).
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Reduction in content of skeletal muscle mitochondria is found in patients suffering from
obesity and T2DM (Ritov et al., 2005). Finally, deficits in muscle ETC enzyme activity
are also found in these patient populations (Ritov et al., 2009). Thus, regulation of
skeletal muscle mitochondria has important implications for whole-body physiology and
human health.
The question therefore surrounds exactly how androgens/AR are capable of
modulating mitochondria in skeletal muscle. The most likely potential pathway involves
AR-mediated expression of major factors involved in mitochondrial biogenesis and β-
oxidation (ex: PGC-1α, PPAR family proteins). Unfortunately, very little evidence of
such effects actually exist. Analysis of gene expression in skeletal muscle of ARKO mice
reveals down-regulation of PPARα (Lin et al., 2005). Agonists of this nuclear receptor
have been shown to stimulate mitochondrial fatty acid β-oxidation in skeletal muscle
(Minnich, Tian, Byan & Bilder, 2001). Therefore, it is possible that AR may mediate
expression of skeletal muscle PPARα, thus contributing to normal fatty acid utilization in
myocytes. In addition, it is possible that androgens may be affecting expression of
mitochondrial DNA. Hormone response elements – DNA sequences to which nuclear
receptor/ligand complexes bind in mediating gene expression – have been found in the
mitochondrial genome, suggesting that steroid hormones can act directly upon
mitochondria (Psarra, Solakidi & Sekeris, 2006). More specifically in skeletal muscle,
Weber et al. (2002) showed that treatment of rats with glucocorticoids (another steroid
hormone) resulted in mitochondrial biogenesis. Here these scientists demonstrated that
glucocorticoid receptor was found in skeletal muscle mitochondria, further evidence that
these hormones act directly upon mitochondria. However, to date, similar findings have
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not been investigated in the context of androgens, AR and skeletal muscle. In summary,
research surrounding skeletal muscle mitochondria shows that these organelles can be
regulated by muscle AR, although the mechanisms by which this regulation occurs is
largely unknown.
4.5.3. Effects on Other Metabolic Organs
While it is most likely that the differences in body composition and metabolism
between HSA-AR and WT animals are due to direct local actions of androgens in skeletal
muscle, it is important to not forget potential indirect effects on other metabolic organs.
Aside from skeletal muscle, the two most commonly studied tissues known for utilization
of energy substrate (i.e. glucose and fatty acids) are the brain and the liver. Both of these
organs are also known to express AR abundantly (Ruizeveld de Winter et al., 1991).
Whether myocyte AR can have indirect effects on these tissues has not been extensively
studied.
The liver is a major metabolic organ (Kammoun et al., 2009). It regulates glucose
homeostasis through uptake and storage as glycogen, and also helps provide glucose to
major organs that require it (such as the brain and skeletal muscle), through
gluconeogenesis. It is also a major site of fatty acid uptake, lipid storage and β-oxidation.
Recent research reveals that local changes in skeletal muscle can have major implications
for the liver. In a groundbreaking study, Izumiya et al. (2008) report a skeletal muscle
manipulation in transgenic mice that results in a phenotype that is strikingly similar to
that seen due to HSA-AR. Here, scientists use a skeletal muscle-specific, Dox-inducible
transgenic mouse model that overexpresses a constitutively active form of the major
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signaling molecule Akt1. Induction of the transgene in adulthood results in selective
hypertrophy of type IIb (fast-twitch, glycolytic) muscle fibers, reduced fat mass, and
increased O2 consumption (and subsequently increased energy expenditure). As
mentioned, these findings largely mimic the effects seen in HSA-AR male rats.
Interestingly, here the scientists found that major oxidative genes (such as PPARγ and
PGC-1α) were down-regulated in skeletal muscle of HSA-AR animals, while major
glycolytic genes were up-regulated. Puzzled by the contrasting increase in systemic
oxidative metabolism, but decreased oxidative gene expression in skeletal muscle, the
authors investigated liver of these animals and found that fatty acid metabolism in this
organ was significantly increased. They hypothesized that this influence on liver
metabolism may occur through systemic action of muscle-secreted factors, termed
‘myokines’. These myokines have been shown to have beneficial effects on body
composition, although their mechanism of action remains unclear (Pederson et al., 2007).
With regards to our results, it is possible that HSA-AR could be operating under a similar
mechanism. Although we did not investigate the liver of HSA-AR males, it may have
played a potential role in β-oxidation of fatty acids, resulting in the reduced adiposity we
see.
The other major target organ for substrate uptake and utilization is the brain.
Whether the brain is involved in mediating the effects of HSA-AR is less likely than the
potential influence of the liver. First, the brain largely does not use fatty acids as an
energy source. Thus, it is highly improbable that it plays a direct role in reducing the
adiposity that we see in HSA-AR. Secondly, relatively little evidence for cross-talk
between skeletal muscle and the brain exists. The little work that has been done in this
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field has focused upon efferent influences of the brain on skeletal muscle through the
sympathetic nervous system. For example, the hypothalamus is able to induce
mitochondrial biogenesis in skeletal muscle in response to increased levels of fatty acid
substrate (Cha et al., 2006). That being said, there is some evidence that shows that
skeletal muscle is capable of secreting myokine signals that can act upon the brain and
influence its metabolism (Pederson & Febbraio, 2005). Several muscle-secreted factors
(such as interleukin-6) have been shown to cross the blood-brain barrier (Banks, Castin &
Broadwell, 1995). While interleukin-6 has been shown to have effects on brain
metabolism (Penkowa et al., 2003), these influences have not been well-established, and
thus the influence of muscle on brain metabolism remains hypothetical at best.
In summary, while differences in systemic metabolism and body composition of
HSA-AR male animals may result from local actions of AR in skeletal muscle, it is also
important to remember the roles of other metabolic tissues, and how their influence may
potentially be involved in the effects that we see.
4.6. Comparing HSA-AR Rats and Mice
In order to generalize the role of AR in muscle, we studied two separate lines of
transgenic animals overexpressing AR in this tissue: HSA-AR rats, and HSA-AR L78
mice. Both strains utilize the HSA promoter to drive transgene expression (although a
truncated version is employed in the rats). Comparing the phenotypes of these two species
(elucidated through the work of this study, as well as from previous work) will allow for
greater understanding of what AR does in skeletal muscle.
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4.6.1. Phenotype Comparisons
Several similarities between HSA-AR rats and mice exist, but these species are
also marked by major differences. With regards to body composition, similar findings are
seen in both HSA-AR male rats and L78 HSA-AR male mice (as compared to WT
counterparts). Both display increased LBM%, decreased FBM% (including raw FBM)
and reduced BMC. Thus, the existence of these effects in both species suggests that
myocyte AR is capable of improving body composition, primarily by reducing FBM.
Furthermore, study of oxidative metabolism between these two strains also demonstrates
similarities. Both groups show evidence of heightened RMR, indicating that increased
expression of AR in skeletal muscle is capable of generating higher rates of systemic
metabolism. Local examination of skeletal muscle mitochondrial ETC enzymes also
reveals parallel results between strains (Fernando et al., 2010; Musa et al., In Prep). HSA-
AR male rats, L78 male mice, and T-treated L141 female mice all show increased activity
of ETC enzymes in EDL muscle. Thus, taken together, it can be hypothesized that AR
functions in myocytes to aid in regulation of mitochondria, thus resulting in increased
systemic metabolism, and reduction of FBM.
However, there are differences between the two strains that may cloud the details
of this mechanism. Namely, it is demonstrated through HSA-AR mice that
overexpression of AR in muscle can lead to acute muscular pathology and death (Monks
et al., 2007). In contrast, HSA-AR rats show no deleterious effects of transgene
expression. These rats are healthy, and at birth, litters yield the expected 1:1 proportion of
HSA-AR and WT males. Conversely, HSA-AR mice experience robust muscular atrophy,
a phenotype that is more predominant in the L141 line (Monks et al., 2007). The great
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majority of males of this line die perinatally. The few that do survive gestation typically
die by PND 10. Although this phenotype is not as severe in L78 male mice,
histopathological analyses of these animals do display some muscle wasting, while we
also found reduced levels of raw LBM in this strain, consistent with mild atrophy.
Ultrastructural analyses reveal that mitochondria in HSA-AR mice are more numerous
and larger, and that myofibril width is narrowed (Musa et al., In Prep). These deleterious
effects in HSA-AR mice complicate the role of AR in muscle, since we find that this
manipulation has largely beneficial effects in our transgenic rats.
4.6.2. Does Expression Level Exaggerate Phenotype?
Despite differences in tissue specificity of expression, the major disparity between
HSA-AR rats and mice exists in the level of AR expression. Western blot analysis
indicates AR expression in EDL muscle of HSA-AR male rats is roughly 2-3 times the
level seen in WT male littermates (Niel et al., 2009). Therefore, the overexpression of AR
in HSA-AR rats is relatively modest. In contrast, the level of overexpression in HSA-AR
mice is significantly higher (Monks et al., 2007). While protein levels were not
quantitatively analyzed in these mice, transgene constructs reveal orders of magnitude
difference in AR copy number, with L78 mice having roughly 100 copies of AR/pg RNA,
and close to 1000 copies of AR/pg RNA in L141 mice. Therefore, the differences we see
between these transgenic strains and species may be related to the level of AR expression
found.
A modest degree of AR overexpression in muscle of rats appears mostly
beneficial, reducing FBM and elevating RMR. A significantly higher level of
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overexpression in L78 males seems to induce similar effects; however some
abnormalities in skeletal muscle fibers are now noted, with reduced LBM in these animals
and histopathological indications of muscular atrophy. Finally, boosting overexpression
of AR by another order of magnitude (as is seen in L141 males) results in significant
muscular atrophy, proliferation of mitochondria, aggregation of glycogen and resultant
death.
With these differences in expression level and resultant phenotype in mind,
hypothetical explanations can be put forward in order to account for the compounding
effects of AR in muscle. AR’s most well understood function is as a transcription factor,
regulating the expression of various genes. Previous work in whole-body ARKO mice
reveals that mRNA expression of PPARα (a protein involved in mitochondrial
biogenesis) in skeletal muscle of these animals is significantly reduced (Lin et al., 2005).
Therefore, it is possible that AR in skeletal muscle may be responsible for regulating
expression of proteins involved in mitochondrial biogenesis. Higher and higher amounts
of AR protein would exaggerate this response, thus accounting for the higher levels of
ETC activity and mitochondrial proliferation seen in HSA-AR mice (Musa et al., In
Prep). The subsequent increase in systemic metabolism of these animals would be robust.
When energy expenditure exceeds energy intake, an energy deficit occurs, with
exacerbation of this deficit eventually leading to cell death. This is seen in various models
of neurodegenerative and neuromuscular disease (such as ALS and HD), which are now
being regarded as having significant metabolic components (Chaturvedi et al., 2009). In
fact, scientists are now able to alleviate pathological symptoms in mouse models of these
diseases using creatine supplementation, a muscle-specific energy supplement (Ferrante
130
et al., 2000; Klivenyi et al., 1999). Thus, while hypermetabolism is often seen as
beneficial, pushing this system to an extreme could easily result in cell death. Similarly,
the modest degree of AR overexpression in HSA-AR rats has largely positive effects in
increasing metabolism and reducing LBM. This effect appears to be exaggerated in HSA-
AR mice, likely due to significantly higher AR expression, and subsequently resulting in
cell death.
4.7. A New Hope: Selective Androgen Receptor Modulators
Our findings may have significant translational potential for application in the
treatment of human disease. Physicians have often prescribed androgen therapy for
patients suffering from sexual dysfunction, as well as other pathologies. However,
previous investigation into androgen therapy shows that the efficacy of these hormones in
a clinical setting is largely limited by their deleterious side effects (Bhasin et al., 2001;
Basaria et al., 2010). These side effects include increased incidence of prostate cancer in
men, masculinization of secondary sexual characteristics in females, and increased
incidence of heart disease in both sexes. This occurs because use of exogenous androgens
causes indiscriminant activation of AR in all tissues. For that reason, a large degree of
investigation in this field has gone into identification of tissue-specific AR that give rise
to the beneficial effects of androgens. By targeting only these AR (and avoiding AR in
prostate and heart), scientists can devise pharmacological agents aimed at treating a
whole host of pathologies, with minimal side effects. Such technology is now on the rise
with the advent of Selective Androgen Receptor Modulators (SARMs), which are a class
of isolated endogenous agents that bind only to AR in specific tissue (Bhasin et al., 2006).
131
Our research suggests that a SARM targeted toward skeletal muscle might have beneficial
effects in increasing LBM while decreasing FBM, allowing for a powerful treatment
against muscular atrophy and obesity.
4.7.1. Selective Androgen Receptor Modulators
SARMs are not synthetically bioengineered molecules, but are rather endogenous
ligands that are found to bind to AR in specific tissues. For example, 27-
hydroxycholesterol has been found to impair cardiac function not only by forming
plaques in arteries, but also by binding to ER only in cardiovascular tissue and blocking
the beneficial effects of estrogens on vascular function (Umetani et al., 2007). SARMs are
typically not steroid-based, and thus differ from T and steroidal androgens in several
ways. Unlike T, which is converted to active metabolites (namely DHT and estrogen),
nonsteroidal SARMs do not undergo aromatization or 5-α-reduction (Bhasin et al., 2006).
Some specific nonsteroidal SARMs have actually been shown to have more favourable
pharmokinetics, greater AR binding capability, and increased amenability to structural
modifications than their androgen counterparts. SARM technology was first harnessed to
mimic the anabolic functions of androgens in bone, through isolated extraction of
tetrahydroquinoline (Hanada et al., 2003). While treatment of rats with this extract led to
significant improvement in bone density, there were some effects noted in reproductive
tissues. To date, scientists have been able to successfully alter bone composition with
little to no effect on other tissues (Gao & Dalton, 2007).
The potential mechanism of SARM tissue selectivity is still not well understood
(Bhasin et al., 2006). Various mechanisms have been proposed, but none have been
132
explicitly tested. The most commonly cited mechanism focuses upon alterations in AR
conformation induced by SARMs, and subsequent ability to recruit co-activators (Bhasin
et al., 2006). Binding of T to AR induces conformational change in the ligand-binding
domain of AR. These conformational changes could modulate surface topology and
protein-protein interactions between AR and specific coactivators. Since the androgen
response elements differ between tissues (and the subsequent genes that are regulated also
differ), recruitment of coactivators is specific to particular tissues (Ting & Chang, 2008).
Thus, one can hypothesize that a SARM binds to AR in all tissues, however the
conformational change induced in the AR protein due to SARM binding only facilitates
recruitment of certain tissue-specific coregulators and coactivators. This would lead to
up-regulation (or down-regulation) of androgen responsive genes in specific tissues, and
thus provide a rationale for the tissue-specificity noted through the use of SARMs.
4.7.2. Targeting Muscle
Our research suggests that SARMs created to target myocyte AR may have
significant effects in maintaining (possibly increasing) LBM and decreasing FBM. The
use of SARM technology targeted toward skeletal muscle has already been researched,
for the purposes of harnessing the anabolic effects of androgens on skeletal muscle (Yin
et al., 2003). Gao and colleagues (2005) utilized such a SARM on gonadectomized adult
male rats. They found that treatment of these rats with the SARM increased muscle mass
and bone density. These levels were comparable with those seen in animals treated with
DHT. Interestingly, SARM treatment did not decrease FBM, although DHT did. Similar
findings were reported by Schmidt et al. (2009). Here the authors used a newly isolated
133
SARM and found that it had anabolic functions on muscle and bone. Microarray analysis
of skeletal muscle showed that treatment with the SARM induced changes in gene
expression that were largely consistent with those induced by DHT. Thus, these findings
suggest that the use of SARMs may have beneficial effects on skeletal muscle and body
composition, but further insight must be gained regarding the mechanism of action by
which these endogenous ligands function.
4.8. Future Directions
Although the findings from our experiments provide us with significant insight
regarding the effects of the HSA-AR transgene and the role of myocyte AR in mediating
whole-body physiology, there are still a tremendous degree of questions that remain
unanswered. Here we will briefly focus on potential avenues for future research, which
would help to further elucidate the function of AR in muscle.
4.8.1. Other Metabolic Parameters in HSA-AR Animals
Our investigation of metabolism in HSA-AR rats and mice largely focused on the
effects of FBM. Once it was demonstrated that HSA-AR animals had significantly
reduced FBM and heightened systemic metabolism, our attention was immediately turned
to local effects of AR on skeletal muscle metabolism. It is now known that AR in skeletal
muscle has significant effects on muscle mitochondria (Fernando et al., 2010; Musa et al,
In Prep.). However, little is known regarding other major metabolic processes in these
animals.
134
A major contributor to metabolic status is glucose homeostasis. Glucose is the
major fuel of the body, and its catabolism allows for the generation of ATP. Furthermore,
skeletal muscle is known to be one of the major sites of glucose utilization, as it is stored
in this tissue in the form of glycogen, and broken down in muscle mitochondria (Harrison
& Leinwand, 2008). Whole-body ablation of AR in mice results in hyperglycemia and
reduced insulin sensitivity (Lin et al., 2005). With the knowledge that mitochondrial ETC
enzyme activity is significantly enhanced in skeletal muscle of HSA-AR animals, and
since glucose is a major substrate for oxidative phosphorylation, it is reasonable to
assume that glucose parameters may differ between HSA-AR and WT animals. First and
foremost, serum glucose levels can be measured in order to determine if there are any
baseline differences between groups. Animals can also be tested on a glucose tolerance
test, where glucose is administered (either orally or via injection), and blood glucose rates
are then measured. Faster rates of glucose clearance (marked by larger decreases in blood
glucose after glucose administration) suggest increased insulin sensitivity and higher
glucose uptake by target tissues (Chen et al., 2003). Furthermore, expression of major
glucose regulatory proteins (such as GLUT4) could be measured. It is possible that AR in
muscle might be involved in increasing expression of GLUT4 (or FATP) and thus
increasing clearance of glucose and fatty acids from the blood. Such a finding would help
in explaining the higher aggregation of glycogen found in HSA-AR L78 and L141 mice
(Musa et al., In Prep). It is now understood that T2DM and obesity are inextricably linked
(Zitzmann, 2009), and thus treatment of one disease is typically helpful in treating the
other.
135
Examination of liver in HSA-AR animals will also be important. Accumulation of
fatty acids in the liver (hepatic steatosis) is a major cause of liver disease, and its
incidence is significantly increased in individuals with higher levels of FBM (Kammoun
et al., 2009). Furthermore, Izumiya et al. (2008) reported a manipulation in skeletal
muscle of HSA-AR mice that did not have local effects on skeletal muscle oxidative
capacity, but rather increased β-oxidation by the liver. The findings reported in that study
(selective hypertrophy of fast-twitch, glycolytic muscle fibers with concomitant decrease
in fat mass and improved metabolic parameters) strongly parallel those seen in HSA-AR
rats (Fernando et al., 2010). While we do find local effects in metabolic function of
skeletal muscle, investigation of gene expression and oxidative capacity in liver could
provide an alternative route by which fatty acid β-oxidation may be occurring.
Furthermore, histological analysis of fatty acids in hepatocytes may indicate whether
myocyte AR can reduce the incidence (or magnitude) of hepatic steatosis when animals
are placed on a high-fat diet. A similar effect has already been shown in mice, where
knockout of AR only in hepatocytes results in hepatic steatosis (Lin et al., 2008). This
finding highlights the role of AR in fatty acid metabolism.
4.8.2. Identifying AR and Mitochondrial Interactions in Skeletal Muscle
As noted, work from our lab has demonstrated that expression of HSA-AR in
skeletal muscle has significant local effects on muscle mitochondria. This includes
increase in ETC enzyme activity in HSA-AR rats, coupled with reduced activity in Tfm
rats (Fernando et al., 2010) and similar effects on enzyme activity with increased
mitochondrial proliferation in HSA-AR mice (Musa et al., In Prep). These findings are
136
interesting, particularly because it has been shown that there are sex differences in
expression of mitochondrial proteins in skeletal muscle of adult humans, with higher
expression levels found in men (Welle, Tawil & Thornton, 2008). Whether these
differences are entirely due to androgens is unclear. Nevertheless, identification of the
mechanisms by which AR is able modulate mitochondria represent an important step in
elucidation of AR’s role in skeletal muscle. However, to date it is not know whether AR
even binds and colocalizes to mitochondria in skeletal muscle myocytes. Solving this
unknown will provide information as to whether AR acts directly upon muscle
mitochondria.
Previous studies have also linked AR to mitochondrial function, with
demonstration of AR-mitochondria colocalization in non-muscle tissues. Ranganathan
and colleagues (2009) found that AR possessing a polyglutamine expansion mutation (as
seen in models of SBMA) results in altered expression of various mitochondrial proteins,
as well as mitochondrial permeability. This study also found that AR associates with
mitochondria in cultured PG12 cells. Transmission electron microscopy showed that AR
localized within mitochondria as well as along the membrane. However, it is unclear as to
whether this modulatory effect is due to indirect effects on the transcription of
mitochondrial genes encoded in the nucleus, or direct effects of the mutant AR protein on
mitochondria (or both). Further to this, AR has been detected in mitochondria of LNCaP
cells (androgen-sensitive human prostate adenocarcinoma cells) as well as the midpiece
region of human sperm cells (Solakidi et al., 2005). It remains unclear as to what function
AR may perform in the mitochondria of these cells. It has been hypothesized that these
receptors may act by upregulating the transcription of various mitochondrial genes, as
137
well as genes involved in oxidative phosphorylation (Psarra & Sekeris, 2008). This has
been demonstrated in skeletal muscle, where other nuclear receptors (namely
glucocorticoid receptor) have been shown to be involved in mitochondrial biogenesis
(Weber et al., 2002). Glucocorticoid receptor expression has also been found in
mitochondria (Psarra et al., 2006).
There are several methods that can be used to determine if AR colocalizes with
skeletal muscle mitochondria. This can be done mostly simply through isolation of
skeletal muscle mitochondria and then subsequently probing for AR in the mitochondria
using an immunoblot. A positive signal would suggest the existence of an AR-
mitochondria complex. This could alternatively be accomplished through immunogold
labeling for AR, followed by electron microscopy, as employed by Ranganathan et al.
(2009). The benefits of this method are the increased level of resolution obtained, and the
fact that quantification can be achieved through counting of AR-positive mitochondria.
Determining whether AR colocalizes with mitochondria is an important step in
elucidating the mechanisms by which myocyte AR can influence this organelle.
4.8.3. Molecular and Biochemical Assays of Key Metabolic Players
AR’s role as a transcription factor suggests that this nuclear receptor might
influence body composition by altering expression of various metabolic genes. Thus,
gene expression assays will be especially useful for potentially determining what muscle-
specific factors are involved in regulating the effects that we see on body composition of
HSA-AR animals. The most well understood regulator of skeletal muscle metabolism is
the PPARγ coactivator PGC-1α, with increased expression of skeletal muscle PGC-1α
138
being associated with mitochondrial biogenesis, increased systemic metabolism, and
reduction of FBM (Calvo et al., 2008; Wenz et al., 2009). Similar effects are known to
occur for PPARα, a gene whose expression is reduced in skeletal muscle of ARKO mice
(Lin et al., 2005). Thus, examination of expression of these and other important metabolic
genes in skeletal muscle of HSA-AR animals could provide significant insight into the
mechanisms by which mitochondrial biogenesis and reduced FBM occur. Analysis of
gene expression could be accomplished using traditional methods of examining mRNA,
such as Quantitative PCR and Northern blot analyses, along with quantification of protein
expression through Western blot.
Alternatively, it is possible that AR may exert its action through nongenomic
actions. In a nongenomic mechanism, binding of AR by its ligand results in formation of
secondary messengers and influence of the secondary messengers on cell signaling. For
example, glial cell culture experiments show that activation of AR by DHT results in
increased phosphorylation of both ERK and Akt, key effectors in the MAPK and PI3-
kinase signaling pathways, and that phosphorylation of these molecules in inhibited by
administration of an AR antagonist (Gatson, Kaur & Singh, 2006). Such effects can be
contrasted with the classical role of AR in regulation of gene expression. Several lines of
evidence indicate that AR functions using such nongenomic mechanisms in adipose tissue
(Mayes & Watson, 2004). Exhibition of such mechanisms in skeletal muscle by AR have
not been found. However, it is nevertheless possible that AR may exert such effects on
key metabolic signaling molecules in skeletal muscle, such as AMPK. Activation of
AMPK and phosphorylation of its targets has shown to be an important process capable
of regulating mitochondrial biogenesis and fiber type switching in skeletal muscle, as
139
well as increasing metabolic parameters (de Lange et al., 2007; Narkar et al., 2008). We
can evaluate activation of AMPK through the use of biochemical assays, and detection of
phosphorylation of AMPK targets. Administration of AR antagonists will indicate
whether any such effects are T-dependent.
4.9. Conclusions
This study revealed the changes in body composition and metabolism associated
with overexpression of AR in muscle fibers. More specifically, we find that
overexpression of AR in muscle increased LBM%, and decreased FBM, FBM% and
BMC. These findings are seen in both rat and mouse HSA-AR strains. Furthermore, all of
these findings are seen in male rats only expressing AR in muscle fibers, indicating that
androgen activity in this cellular population is sufficient for induction of the effects that
we see. Treatment of female rats with exogenous T recapitulates our findings in males,
suggesting that they are likely dependent upon the androgen ligand, and are not simply
the result of transgene expression only.
Perhaps most interesting, we show that AR in myocytes is sufficient for
androgenic reduction of FBM, which is confirmed through dissection of individual fat
pads, and analysis of adipocyte size. This is a novel finding, as androgens are typically
thought to reduce FBM by acting upon AR in adipocytes, and inhibiting adipogenesis
during development (Singh et al., 2003; Singh et al., 2006). However, our findings
explain why ablation of AR only in adipocytes (Yu et al., 2008) is incapable of
recapitulating the adult-onset obesity found in whole body ARKO (Lin et al., 2005; Fan et
al., 2005). Investigation of metabolic parameters shows that our HSA-AR males exhibit
140
heightened resting metabolism, which is complimentary to the reduced RMR values
found in ARKO mice (Fan et al., 2005). Future research should focus upon elucidation of
the mechanisms involved in heightened metabolism. Recent work from our lab shows that
AR is capable of modulating skeletal muscle mitochondria (Fernando et al., 2010; Musa
et al., In Prep). Therefore, a more comprehensive analysis of how AR is capable of
regulating mitochondria in muscle will provide greater insight into how androgens and
AR influence body composition and energy homeostasis.
141
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Tables and Figures
TABLE 1: Summary of Animals Used
Experiment WT M
Rats
Tg M
Rats
Tfm M
Rats
Tfm /Tg M
Rats
WT F
Rats
Tg F
Rats
Tfm F
Rats
Tfm /Tg F
Rats
WT M
Mice
L78 M
Mice
Exp. I
RT-PCR 3 3
Exp. 2
Body Composition 11 11 7 9 8 9 7 7 5 6
Dissections 10 12 7 10
Exp. 3
Testosterone Treatment 8 9
Exp. 4
Adipose Histology 6 6 6 4
Exp. 5
RMR 10 12 7 10 5 6
Activity Measures 4 6 3 5
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TABLE 2: Primer Sequences
Application Gene Name (Species) Sequenece PCR Human AR (Rat) F: GGACAGGGCACTACCGAG
R: GGCTGAATCTTCCACCTAC PCR Rat AR (Rat) F: GCAACTTGCATGTGGATGA
R: TGAAAACCAGGTCAGGTGC PCR GAPDH (Rat) F: ATGGGAAGCTGGTCATCAAC
R: GGATGCAGGGATGATGTTCT PCR Rat AR (Mouse) F: AGTAGCCAACAGGGAAGGGT
R: GAGGCAGCCGCTCTCAGGGTG PCR TSH (Mouse) F: AACGGAGAGTGGGTCATCAC
R: CATTGGGTTAAGCACACAGG Endpoint PCR Human AR (Rat) F: AGGAAGCAGTATCCGAAGGCA
R: GGACACCGACACTGCCTTACA
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Figure 1: Characterization of Transgene Expression. RT-PCR reveals expression of the transgene in urinary bladder, heart and skeletal muscle of Tg animals only.
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Figure 2: Transgene Expression Regulates Body Composition in Males. Representative DXA images from WT (A), HSA-AR (B), Tfm (C), and HSA-AR/Tfm (D) male rats are shown. Note the reduced mass in the abdominal region of Tg animals. (E) HSA-AR has no effect on body mass, but this parameter is reduced in Tfm males. HSA-AR expression increases LBM% (F) and decreases FBM% (G). Evaluation of raw values shows that HSA-AR expression has no effect on raw LBM (H), but decreases raw FBM (I). (#, Significant main effect of Tfm; *, Significant main effect of HSA-AR).
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Figure 3: No Effect of the Transgene on Body Composition in Female Rats. Transgene expression did not affect body mass (A), LBM% (B) and FBM% (C) in female rats. However, a transient effect of HSA-AR is found at 6 weeks. (*, Significant main effect of HSA-AR).
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Figure 4: Excised Fat Pads and Muscles Confirm DXA Findings. No effect of HSA-AR was found on mass of dissected individual anterior tibialis (AT – A) and extensor digitorum longus (EDL – B) muscles. However, HSA-AR male rats were found to have lighter perigonadal fat pads (C). (*, Significant difference between groups).
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Figure 5: Body Composition of HSA-AR Mice. Representative DXA images of WT and Tg L78 mice (A). Note the reduced abdominal mass in the Tg animal. L78 male mice demonstrate reduced total body mass (B), with increased LBM% (C) and decreased FBM% (D). Evaluation of raw body composition reveals decreased LBM in L78 mice (E), as well as reduced FBM (F). (*, Significant difference between groups).
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Figure 6: Effects of HSA-AR on Bone Parameters. HSA-AR expression was found to decrease BMC in Tg males (A), but not females (B). Derivation of BMD however found no difference between groups for males (C) or females (D). Similar findings were seen in HSA-AR L78 mice. Here also, transgene expression reduced BMC in Tg males (E), but had no effects on BMD (F). (*, Significant main effect of HSA-AR; #, Significant main effect of Tfm).
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Figure 7: Body Composition is Altered by T-treatment of HSA-AR Females. WT and Tg females were treated with T for 4 weeks, at which point T was removed. Body composition was measured weekly. Representative DXA images of a WT female (top) and her Tg sister (bottom) are shown over the course of treatment (A). Note the reduced abdominal mass in the Tg female at 4 weeks, and concomitant increase to baseline at 8 weeks. T treatment increases LBM% (B) and decreases FBM% (C) in Tg females only. Tg females show a T-dependent increase in raw LBM (D) and decrease in FBM (E) over the course of treatment. WT females show no differences in raw LBM (F) and FBM (G). (*Significant within-group difference from baseline for Weeks 0-4 OR Significant within-group difference from Week 4 for Weeks 5-8).
170
Figure 8: Smaller Adipocytes are Found in HSA-AR Males. Representative adipocytes from WT, HSA-AR, Tfm and HSA-AR/Tfm adult males (A). Transgene expression results in reduced adipocyte size (B), with Tfm males showing the largest cells. Distribution of cell size shows that Tg animals have a larger proportion of smaller cells (C). (*, Significant difference between groups).
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Figure 9: Increased oxygen consumption in HSA-AR Male Rats. Tg adult males show increased oxygen consumption, as compared to WT controls (A). Although significant differences in RMR were not found due to HSA-AR (B), a trend toward higher levels of RMR in Tg animals was seen. (*, Significant difference between groups).
172
Figure 10: Differences in Energy Expenditure in HSA-AR L78 Mice. While Tg L78 adult male mice were not found to significantly differ in oxygen consumption (A), correction for body mass reveals that L78 males have significantly higher RMR than WT brothers. (*, Significant difference between groups).
173
Figure 11: HSA-AR Expression Does Not Affect Spontaneous Activity Level. Male rats were measured for activity using a laser-grid activity box, where the number of laser beam breaks was measured over a period of 1 hour. Rats were analyzed at the same ages at which differences in body composition were found. No main effect of HSA-AR or Tfm were found at any time point.
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APPENDIX A: STASTICAL VALUES
Experiment II: Body Composition Analysis MALE RATS Analysis of Variance (ANOVA) of HSA-AR Genotype (Between subjects, 2 levels) X Tfm Genotype (between subjects, 2 levels) * denotes statistical significance Main effect of HSA-AR Week 4 Body weight F=0.359 p=0.553 Lean body mass % F=1.219 p=0.278 Fat body mass % F=1.200 p=0.282 Uncorrected Lean body mass F=0.181 p=0.673 Uncorrected Fat body mass F=2.216 p=0.147 Week 6 Body weight F=0.408 p=0.528 Lean body mass % F=17.748 p<0.001* Fat body mass % F=16.275 p<0.001* Uncorrected Lean body mass F=0.154 p=0.698 Uncorrected Fat body mass F=10.541 p=0.003* Week 8 Body weight F=1.181 p=0.286 Lean body mass % F=10.630 p=0.003* Fat body mass % F=10.004 p=0.003* Uncorrected Lean body mass F=0.305 p=0.585 Uncorrected Fat body mass F=7.769 p=0.009* Week 10 Body weight F=3.339 p=0.077 Lean body mass % F=6.510 p=0.016* Fat body mass % F=6.360 p=0.017* Uncorrected Lean body mass F=0.923 p=0.344 Uncorrected Fat body mass F=6.662 p=0.015* Main effect of Tfm Week 4 Body weight F=0.817 p=0.373 Lean body mass % F=0.205 p=0.654 Fat body mass % F=0.123 p=0.728 Uncorrected Lean body mass F=0.556 p=0.461
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Uncorrected Fat body mass F=1.173 p=0.287 Week 6 Body weight F=4.517 p=0.042* Lean body mass % F=4.396 p=0.044* Fat body mass % F=4.718 p=0.038* Uncorrected Lean body mass F=3.568 p=0.068 Uncorrected Fat body mass F=7.706 p=0.009* Week 8 Body weight F=20.641 p<0.001* Lean body mass % F=27.393 p<0.001* Fat body mass % F=11.640 p=0.002* Uncorrected Lean body mass F=44.909 p<0.001* Uncorrected Fat body mass F=44.909 p<0.001* Week 10 Body weight F=22.116 p<0.001* Lean body mass % F=11.549 p=0.002* Fat body mass % F=12.837 p=0.001* Uncorrected Lean body mass F=11.560 p=0.002* Uncorrected Fat body mass F=20.914 p<0.001* Interaction between HSA-AR and Tfm Week 4 Body weight F=0.241 p=0.627 Lean body mass % F=0.207 p=0.652 Fat body mass % F=0.272 p=0.606 Uncorrected Lean body mass F=0.136 p=0.715 Uncorrected Fat body mass F=0.968 p=0.333 Week 6 Body weight F=0.448 p=0.508 Lean body mass % F=0.071 p=0.792 Fat body mass % F=0.019 p=0.891 Uncorrected Lean body mass F=0.432 p=0.516 Uncorrected Fat body mass F=0.216 p=0.645 Week 8 Body weight F=1.183 p=0.285 Lean body mass % F=0.540 p=0.468 Fat body mass % F=0.401 p=0.531 Uncorrected Lean body mass F=0.831 p=0.369 Uncorrected Fat body mass F=0.796 p=0.379 Week 10
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Body weight F=0.847 p=0.365 Lean body mass % F=0.463 p=0.501 Fat body mass % F=0.354 p=0.556 Uncorrected Lean body mass F=0.504 p=0.483 Uncorrected Fat body mass F=0.538 p=0.469 Student’s t test: Body Weight WT/WT vs WT/Tfm Wk4 p=0.994 Wk6 p=0.521 Wk8 p=0.013* Wk10 p=0.006* WT/WT vs HSA-AR/WT Wk4 p=1.000 Wk6 p=0.997 Wk8 p=0.961 Wk10 p=0.977 HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.666 Wk6 p=0.440 Wk8 p<0.001* Wk10 p<0.001* WT/Tfm vs HSA-AR/Tfm Wk4 p=0.919 Wk6 p=0.886 Wk8 p=0.765 Wk10 p=0.727 LBM% WT/WT vs WT/Tfm Wk4 p=1.000 Wk6 p=0.834 Wk8 p=0.003* Wk10 p=0.045* WT/WT vs HSA-AR/WT Wk4 p=0.992 Wk6 p=0.002* Wk8 p=0.022*
177
Wk10 p=0.108 HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.850 Wk6 p=0.010* Wk8 p<0.001* Wk10 p=0.001* WT/Tfm vs HSA-AR/Tfm Wk4 p=0.799 Wk6 p<0.001* Wk8 p=0.004* Wk10 p=0.071 FBM% WT/WT vs WT/Tfm Wk4 p=1.000 Wk6 p=0.790 Wk8 p=0.001* Wk10 p=0.027* WT/WT vs HSA-AR/WT Wk4 p=0.944 Wk6 p=0.002* Wk8 p=0.024* Wk10 p=0.103 HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.865 Wk6 p=0.013* Wk8 p<0.001* Wk10 p=0.001* WT/Tfm vs HSA-AR/Tfm Wk4 p=0.781 Wk6 p<0.001* Wk8 p=0.004* Wk10 p=0.088 LBM WT/WT vs WT/Tfm Wk4 p=0.996 Wk6 p=0.775 Wk8 p=0.091 Wk10 p=0.004*
178
WT/WT vs HSA-AR/WT Wk4 p=1.000 Wk6 p=0.866 Wk8 p=0.777 Wk10 p=0.164 HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.755 Wk6 p=0.115 Wk8 p=0.001* Wk10 p<0.001* HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.966 Wk6 p=0.986 Wk8 p=0.992 Wk10 p=0.108 FBM WT/WT vs WT/Tfm Wk4 p=1.000 Wk6 p=0.577 Wk8 p<0.001* Wk10 p=0.004* WT/WT vs HSA-AR/WT Wk4 p=0.961 Wk6 p=0.057 Wk8 p=0.153 Wk10 p=0.164 HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.275 Wk6 p=0.001* Wk8 p<0.001* Wk10 p<0.001* HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.482 Wk6 p=0.002* Wk8 p=0.011* Wk10 p=0.108
179
FEMALE RATS Analysis of Variance (ANOVA) of HSA-AR Genotype (Between subjects, 2 levels) X Tfm Genotype (between subjects, 2 levels) * denotes statistical significance Main effect of HSA-AR Week 4 Body weight F=2.737 p=0.112 Lean body mass % F=0.485 p=0.494 Fat body mass % F=0.529 p=0.475 Week 6 Body weight F=13.992 p=0.001* Lean body mass % F=13.249 p<0.001* Fat body mass % F=14.335 p<0.001* Week 8 Body weight F=5.622 p=0.025* Lean body mass % F=.002 p= 0.968 Fat body mass % F=0.107 p=0.746 Week 10 Body weight F=2.845 p=0.163 Lean body mass % F=3.650 p=0.067 Fat body mass % F=3.044 p=0.092 Student’s t test: * denotes statistical difference LBM% WT/WT vs WT/Tfm Wk4 p= 0.025* Wk6 p=0.658 Wk8 p=0.379 Wk10 p=0.259 WT/WT vs HSA-AR/WT Wk4 p= 0.025* Wk6 p=0.004* Wk8 p=0.226 Wk10 p=.040* HSA-AR/WT vs HSA-AR/Tfm
180
Wk4 p=0.141 Wk6 p=0.026* Wk8 p=0.924 Wk10 p=0.253 WT/Tfm vs HSA-AR/Tfm Wk4 p=0.653 Wk6 p=0.053 Wk8 p=0.338 Wk10 p=0.927 FBM% WT/WT vs WT/Tfm Wk4 p= 0.025* Wk6 p=0.517 Wk8 p=0.477 Wk10 p=0.203 WT/WT vs HSA-AR/WT Wk4 p=0.023* Wk6 p=0.003* Wk8 p=0.224 Wk10 p=0.039* HSA-AR/WT vs HSA-AR/Tfm Wk4 p=0.124 Wk6 p=0.0260 Wk8 p=0.9970 Wk10 p=0.238 WT/Tfm vs HSA-AR/Tfm Wk4 p=0.652 Wk6 p=0.039* Wk8 p=0.490 Wk10 p=0.821 MALE MICE Student’s t test Total body mass t=-5.594 p<0.001* LBM% t=2.94 p=0.016* FBM% t=-3.860 p=0.004* Raw LBM t=-2.944 p=0.016* Raw FBM t=-3.860 p=0.004*
181
Experiment III: T-Dependence Analysis Analysis of Varience (ANOVA) of HSA-AR Genotype (Between subjects, 2 levels) X Time (within subjects, 5 levels) * denotes statistical significance ANOVAs were run separately for the 4 weeks of testosterone treatment, and the 4 weeks following cessation of treatment. For both ANOVAs, a fifth baseline week is included as a reference (Week 0 for the first and W4 for the second ANOVA). Unprotected t-tests were used only when an interaction between the within subjects factor and HSA-AR was observed. Otherwise, Dunnett’s correction was applied to alpha as indicated. ANOVA I: Weeks 0-4 LBM Within subjects LBM F=3.864 p=0.068 Between subjects HSA-AR F=7.472 p=0.015* Interaction LBM X HSA-AR F=8.863 p=0.009* Pairwise comparisons: HSA-AR W1 t=0.138 p=0.829 HSA-AR W2 t=-1.691 p=0.129 HSA-AR W3 t=-3.433 p=0.009* HSA-AR W4 t=-2.614 p=0.031* WT W1 t=-1.41 p=0.292 WT W2 t=1.251 p=0.251 WT W3 t=-0.129 p=0.901 WT W4 t=0.532 p=0.611 FBM Within subjects FBM F=991.293 p<0.001* Between subjects HSA-AR F=3.871 p=0.068 Interaction FBM X HSA-AR F=6.846 p=0.019* Pairwise comparisons: HSA-AR W1 t8=-0.133 p=0.897 HSA-AR W2 t8=1.725 p=0.123 HSA-AR W3 t8=3.406 p=0.009* HSA-AR W4 t8=2.687 p=0.028* WT W1 t7=1.242 p=0.254 WT W2 t7=-1.142 p=0.291 WT W3 t7=0.167 p=0.872 WT W4 t7=-0.455 p=0.663 Raw LBM
182
Within subjects Raw LBM F=19.883 p<0.001 Between subjects HSA-AR F=0.112 p=0.742 Interaction Raw LBM X HSA-AR F=20.955 p<0.001 Pairwise comparisons: HSA-AR W1 t=-3.036 p=0.016* HSA-AR W2 t=-3.972 p=0.004* HSA-AR W3 t=-5.617 p=0.001* HSA-AR W4 t=-5.064 p=0.001* WT W1 t=-1.580 p=0.158 WT W2 t=1.126 p=0.297 WT W3 t=-0.035 p=0.973 WT W4 t=-0.534 p=0.610 Raw FBM Within subjects Raw FBM F=0.993 p=0.335 Between subjects HSA-AR F=6.977 p=0.019* Interaction Raw FBM X HSA-AR F=4.600 p=0.019* Pairwise comparisons: HSA-AR W1 t=-1.105 p=0.301 HSA-AR W2 t=0.988 p=0.352 HSA-AR W3 t=2.470 p=0.039* HSA-AR W4 t=1.625 p=0.143 WT W1 t=1.205 p=0.267 WT W2 t=-1.088 p=0.313 WT W3 t=-0.007 p=0.994 WT W4 t=-0.545 p=0.602 ANOVA II: weeks 4-8 LBM Within subjects LBM F=12.774 p=0.003* Between subjects HSA-AR F=8.826 p=0.10 Interaction LBM X HSA-AR F=0.599 p=0.451 Pairwise comparisons (Dunnett correction applied, alpha set at p<0.012): HSA-AR W5 t=0.502 p=0.629 HSA-AR W6 t=4.604 p=0.002* HSA-AR W7 t=2.337 p=0.048 HSA-AR W8 t=2.057 p=0.074 WT W5 t=-0.600 p=0.567 WT W6 t=1.369 p=0.213
183
WT W7 t=1.914 p=0.097 WT W8 t=0.710 p=0.501 FBM Within subjects FBM F=13.841 p=0.002* Between subjects HSA-AR F=8.651 p=0.010* Interaction FBM X HSA-AR F=0.625 p=0.441 Pairwise comparisons (Dunnett correction applied, alpha set at p<0.012): HSA-AR W5 t=-0.484 p=0.641 HSA-AR W6 t=-4.724 p=0.001* HSA-AR W7 t=-2.350 p=0.047 HSA-AR W8 t=-2.181 p=0.061 WT W5 t=0.651 p=0.536 WT W6 t=-1.370 p=0.213 WT W7 t=1.923 p=0.096 WT W8 t=-0.740 p=0.484 Raw LBM Within subjects Raw LBM F=0.048 p=0.829 Between subjects HSA-AR F=0.002 p=0.969 Interaction Raw LBM X HSA-AR F=2.768 p=0.117 Pairwise comparisons (Dunnett correction applied, alpha set at p<0.012): HSA-AR W5 t=2.706 p=0.027 HSA-AR W6 t=5.675 p<0.001* HSA-AR W7 t=2.957 p=0.018 HSA-AR W8 t=1.670 p=0.133 WT W5 t=-0.087 p=0.933 WT W6 t=0.524 p=0.617 WT W7 t=-0.032 p=0.976 WT W8 t=-0.961 p=0.369 Raw FBM Within subjects Raw FBM F=16.980 p=0.001* Between subjects HSA-AR F=7.452 p=0.016* Interaction Raw FBM X HSA-AR F=0.50 p=0.826 Pairwise comparisons (Dunnett correction applied, alpha set at p<0.012): HSA-AR W5 t=0.143 p=0.890 HSA-AR W6 t=-3.720 p=0.006* HSA-AR W7 t=-1.872 p=0.098 HSA-AR W8 t=-1.802 p=0.109
184
WT W5 t=0.324 p=0.755 WT W6 t=-1.298 p=0.236 WT W7 t=-2.036 p=0.081 WT W8 t=-1.721 p=0.129 Experiment IV: Adipose Histology Average adipocyte size Analysis of Variance (ANOVA) of HSA-AR Genotype (Between subjects, 2 levels) X Tfm Genotype (between subjects, 2 levels) * denotes statistical significance HSA-AR: F = 27.187 p<0.001* Tfm: F = 1.760 p=0.201 HSA-AR X Tfm: F = 1.282 p=0.272 Student’s t test * denotes statistical significance WT/WT vs WT/Tfm p=0.025* WT/WT vs HSA-AR/WT p=0.026* HSA-AR/WT vs HSA-AR/Tfm p=0.921 WT/Tfm vs HSA-AR/Tfm p < 0.001* Experiment V: Energy Balance and Metabolic Analyses Oxygen Consumption – Male Rats Student’s t test * denotes statistical significance WT/WT vs WT/Tfm t = 0.242; p=0.812 WT/WT vs HSA-AR/WT t = -2.025 ;p=0.056* HSA-AR/WT vs HSA-AR/Tfm t = -0.243; p=0.811 WT/Tfm vs HSA-AR/Tfm t = -2.551;p < 0.022* RMR – Male Mice Student’s t test * denotes statistical significance WT vs L78 t =-2.286; p=0.048*
185
Activity Box – Male Rats Analysis of Variance (ANOVA) of HSA-AR Genotype (Between subjects, 2 levels) X Tfm Genotype (between subjects, 2 levels) * denotes statistical significance 4 Weeks HSA-AR: F = 0.197; P = 0.664 Tfm: F = 0.600; P = 0.451 HSA-AR X Tfm: F = 4.277; p = 0.058 6 Weeks HSA-AR: F = 1.054; P = 0.322 Tfm: F = 2.108; P = 0.169 HSA-AR X Tfm: F = 0.575; p = 0.461 8 Weeks HSA-AR: F = 0.245; P = 0.628 Tfm: F = 2.889; P = 0.111 HSA-AR X Tfm: F = 4.114; p = 0.062 10 Weeks HSA-AR: F = 0.972; P = 0.341 Tfm: F = 1.506; P = 0.240 HSA-AR X Tfm: F = 1.603; p = 0.226