FATTY ACID SYNTHASE MODULATES MAMMARY GLAND LACTATION AND

PROSTATE CANCER PROGRESSION IN VIVO

By

JANEL KATHRYN SUBURU

A Dissertation Submitted to the Graduate Faculty of

WAKE FOREST UNIVERSITY GRADUATE SCHOOL OF ARTS AND SCIENCES

in Partial Fulfillment of the Requirements for the Degree of

DOCTOR OF PHILOSOPHY

Cancer Biology

May 2014

Winston-Salem, NC

Approved by:

Yong Q. Chen, Ph.D., Advisor

John Parks, Ph.D., Chair

Timothy Pardee, M.D., Ph.D.

Guangchao Sui, Ph.D.

Steven J. Kridel, Ph.D.

ii

DEDICATION

I dedicate this work to my unwaveringly supportive and loving parents. Your passionate

love and support have carried me through life – from my first steps to my first day of school, and

from kindergarten through 22nd

grade. You have supported me and my endeavors at every

graduation, every soccer game, every race, every day… Your incredible commitment to my well-

being in every imaginable aspect has enabled me to achieve the goals you have inspired me to

set. I am truly blessed and forever thankful to have been raised by such awesome parents, whose

whole-hearted dedication to their children has taught me the amazingness of what it means to be

a part of a supportive and loving family.

iii

ACKNOWLEDGEMENTS

I have many people to thank that have contributed to my graduate education, as well as

my growth as a scientist and person.

First and foremost, this work could not have been done without the guidance and support

of my advisor and mentor, Dr. Yong Q. Chen. He has supported my education with a challenging

project, as well as my growth as a young professional in allowing me to partake in internship and

teaching opportunities. Additionally, I was extremely fortunate to have been welcomed by an

entire lab full of mentors and scientists to guide me through my graduate studies. Many thanks to

the Chen lab members – Dr. Lihong Shi, Dr. Jiansheng Wu, Dr. Shihua Wang, Dr. Zhennan Gu,

Dr. Isabelle Berquin, and Jaiozhong Cai – whom have all contributed to my learning experience

and growth as a scientist.

I would like to thank my committee members, Dr. Timothy Pardee, Dr. John Parks, Dr.

Guangchao Sui, and Dr. Steven Kridel for their support, advice, constructive criticisms, and

motivations to make me a better scientist. A special thank you is due to Dr. Steven Kridel for his

advice and guidance as my occasional, surrogate advisor. Also, thank you Dr. Steven Kridel and

Dr. Isabelle Berquin, for allowing me to rotate through their lab as a first year graduate student.

Thank you Dr. Paul Cao, Dr. Caroline Schnegg, and Dr. Tam Nguyen-Cao for reminding

me that there is light at the end. I am fortunate to have studied among exceptionally bright and

talented Wake Forest students, whom played their part well as studious, ambitious role models

for life during and after graduate school.

A huge thanks and group hug is due to the amazing friendships I have developed during

my graduate studies at Wake. Coach Katherine Skarbek, Mrs. Alexis Ward, Jeremy Ward, and

iv

my entire United FC and Shades of Red soccer teams – you are all amazing people on, and

especially, off the field. Thank goodness for year-round soccer at the Plex and the leagues and

pick-up games that we rock on such a regular basis. Mai Tran, Rachel Haggin, Oakley Cline, Dr.

Laura Painter, Dr. Brad Yentzer, and Dr. Matthew Belford, you are all focus points to my life

and hobbies here in Winston. Life in graduate school would have not have been the same without

all my awesome friends. You brighten my life and make all the work 110 times worth the play!

I would also like to acknowledge several people who have supported me from afar during

this ambitious journey. Thank you to my brothers, Ryan Suburu and Eric Suburu, who have been

patient and supportive of my studies, but who have continued to remind me that there’s no place

like home. Thank you also to my entire extended family on the West Coast for their incredible

support and understanding of my endeavors. I have missed several family gatherings being the

only person on the East Coast, but I have never been forgotten. I have a very special place in my

heart for my grandmothers that will not get to see me finish graduate school, as I know they

would have loved to be at my graduation. I am so blessed with such a loving family; home is

truly where the heart lies. A shout out to my UC San Diego amigos Marcus Kouma and David

Ko who helped champion the 3-way FaceTime chat on Apple devices. And finally, a HUGE hug

and happy dance with my best and longest known friend, Edgar Viveros. Edgar, no matter how

far, no matter how much time has passed, it’s never too late just to say hello. I can always count

on Edgar to have my best interests in mind. I miss your face!

v

LIST OF ILLUSTRATIONS

CHAPTER I

Figure 1. Schematic Representation of Palmitate Synthesis by Fatty

Acid Synthase. 39

Figure 2. Organization of the Enzymatic Domains of Fatty Acid Synthase. 40

Figure 3. Cancer Cell Metabolism Promotes Fatty Acid Synthesis. 41

CHAPTER II

Figure 1. Epithelial cell-specific deletion of Fasn in the mammary gland. 69

Figure 2. Deletion of FASN hinders growth and survival of nursing pups. 71

Figure 3. Pre-weanling death occurs as a whole litter. 72

Figure 4. KO mammary glands are sparse in alveolar structures. 73

Figure 5. FASN deletion induces early involution and cell death. 74

Figure 6. Deletion of FASN hinders alveolar development, but not

secretory activation during lactation. 76

Figure 7. FASN deletion changes the fatty acid profile of mammary

gland milk. 78

Figure 8. Phenotypic rescue of offspring by cross-fostering mothers

at birth. 79

vi

Table 1. Litter statistics for growth and survival curves. 80

Table 2. Litter statistics for swapped litter growth and survival curves. 80

CHAPTER III

Figure 1. Genotypic outcomes from animal model breeding. 105

Figure 2. Heterozygous loss of FASN reduces tumor burden. 106

Figure 3. Heterozygous loss of FASN hinders prostate cancer

progression. 107

Figure 4. Heterozygous deletion of Fasn decreases FASN and

cytoplasmic pAKT473

. 108

Figure 5. FASN facilitates prostate cancer cell proliferation. 109

Figure 6. Hemizygous deletion of FASN has no effect on cell death

in Pten knockout mice. 110

Figure 7. Total prostate weights of inducible Pten/Fasn knockout

mouse model. 112

Figure 8. Histology of inducible Pten/Fasn knockout mouse model. 113

Figure 9. Immunohistochemical staining for pAKT473

and FASN in

Pb-CreERT mice. 114

vii

LIST OF ABBREVIATIONS

SAFA saturated fatty acid

MUFA monounsaturated fatty acid

PUFA polyunsaturated fatty acid

FADS1 Δ6 desaturase enzyme

FADS2 Δ5 desaturase enzyme

SCD1 stearoyl-CoA desaturase 1 or Δ9 desaturase enzyme

ELOVL1-7 elongase of very long chain fatty acids 1-7

ATP adenosine triphosphate

ACLY ATP-citrate lyase

ACACA acetyl-CoA-carboxylase 1 (cytosolic)

ACC1 acetyl-CoA-carboxylase 1 (cytosolic)

Fasn fatty acid synthase

FASN fatty acid synthase protein

FASN human fatty acid synthase gene

Fasn mouse fatty acid synthase gene

AMP adenosine monophosphate

viii

AMPK AMP-activated kinase

ACC2 acetyl-CoA-carboxylase 2 (mitochondrial)

SREBP sterol response element binding protein 1

kDa kilodaltons

MAT malonyl acetyl transferases

ACP acyl carrier protein

KS ketoacyl synthase

NADPH nicotinamide adenine dinucleotide phosphate

KR β-ketoacyl reductase

DH β-hydroxyacyl dehydratase

ER enoyl reductase

TE thioesterase

ΨME pseudo-methyltransferase

ΨKR pseudo-ketoreductase

Sp1 stimulatory protein 1

NF-Y nuclear factor Y

USF1 upstream stimulatory factor 1

ix

USF2 upstream stimulatory factor 2

FASKOL liver-specific deletion of FASN

PPARα peroxisome proliferator activating receptor alpha

mTORC1 mammalian target of rapamycin complex 1

FASKO hypothalamus and β-islet cell-specific deletion of FASN

FASTie endothelial cell-specific deletion of FASN

FAS-villin intestine-specific deletion of FASN

eNOS endothelial nitric oxide synthase

Muc2 mucin 2

FASKard myocardiocyte-specific deletion of FASN

Ca++

calcium

CaMKII Ca++

/calmodulin dependent protein kinase II

shRNA short hairpin ribonucleic acid

FASKOS skeletal muscle-specific deletion of FASN

CaMKβ Ca++

/calmodulin dependent protein kinase β

SERCA1 sarco/endoplasmic reticulum calcium ATPase 1

SERCA2 sarco/endoplasmic reticulum calcium ATPase 2

x

GLUT4 glucose transporter 4

FASKOF adipose-specific deletion of FASN

PPARγ peroxisome proliferator activating receptor gamma

PexRAP peroxisomal reductase activating PPARγ

FASKOM macrophage-specific deletion of FASN

LXRα liver X receptor alpha

LDL low density lipoprotein

CD36 cluster of differentiation 36

TCA tricarboxycylic acid

G6P glucose-6-phosphate

HK1-4 hexokinase 1-4

PPP pentose phosphate pathway

PKM2 pyruvate kinase M2

PKM1 pyruvate kinase M1

ACO1 cytosolic aconitase

IDH1 cytosolic isodehydrogenase

ME1 malic enzyme 1

xi

HIF1α hypoxia-inducible factor 1 alpha

EGF epidermal growth factor

USP2a ubiquitin-specific protease 2a

PTEN phosphatase and tensin homolog

PI3K phosphatidylinositol-3 kinase

LKB1 liver kinase B1

HER2 human epidermal growth factor receptor 2

mTOR mammalian target of rapamycin

ERK1/2 extracellular signal regulated kinases 1/2

HIF-1α hypoxia inducible factor 1 alpha

NOX5 NADPH oxidase 5

ROS reactive oxygen species

AR androgen receptor

β2-M beta-2 microglobulin

MAPK mitogen activated protein kinase

AR8 androgen receptor splice variant 8

CRPC castration resistant prostate cancer

xii

EGFR epidermal growth factor receptor

SCAP SREBP cleavage activating protein

KO knockout

WAP whey acidic protein

WT wild-type

L[#] lactation day [#]

IP intraperitoneal

IHC immunohistochemistry

HRP horseradish peroxidase

IF immunofluorescence

TUNEL terminal deoxynucleotidyl transferase dUTP nick end labeling

rTdT recombinant terminal deoxynucleotidyl transferase

FAME fatty acid methyl ester

ErbB2 mouse epidermal growth factor receptor 2

L[X]P[Z] lactation day [X] of pregnancy [Z]

s.e.m. standard error of the mean

Pb-Cre4 Cre recombinase controlled by the probasin promoter

xiii

Pb-CreER tamoxifen-inducible Cre recombinase controlled by the probasin promoter

4-OHT 4-hydroxytamoxifen

ER estrogen receptor

H&E hematoxylin and eosin

EGCG epigallocatechin-3-gallate

xiv

Table of Contents

DEDICATION ii

ACKNOWLEDGEMENTS iii

LIST OF ILLUSTRATIONS v

LIST OF ABBREVIATIONS vii

ABSTRACT xvi

CHAPTER I

GENERAL INTRODUCTION

(Sections from a publication in: Prostaglandins Other Lipid Mediat. Lipids and prostate cancer. 2012

May;98(1-2):1-10.)

I.A Abstract 2

I.B Fatty Acid Metabolism 3

Types of fatty acid: their sources and roles

Dietary fatty acid metabolism

I.C de novo Fatty Acid Synthesis 6

The pathway enzymes and regulation

Fasn: the enzyme

Fasn: regulation

Fasn: expression

I.D de novo Lipogenesis in Cancer 17

Cancer versus normal metabolism

Cancer metabolism favors fatty acid synthesis

Fatty acid synthesis and cancer

Utility of de novo fatty acids in cancer

Oncogenic signaling pathways promote fatty acid synthesis

I.E Overview 25

I.F References 27

I.G Figures and Figure Legends 39

xv

CHAPTER II

FATTY ACID SYNTHASE IS REQUIRED FOR MAMMARY GLAND

DEVELOPMENT AND MILK PRODUCTION DURING LACTATION

(Published online as an Article in Press in the American Journal of Physiology – Endocrinology and

Metabolism on March 25, 2014)

II.A Abstract 44

II.B Introduction 45

II.C Experimental Procedures 48

II.D Results 52

II.E Discussion 58

II.F References 65

II.G Acknowledgements 68

II.H Figures and Figure Legends 69

II.I Tables 80

CHAPTER III

FATTY ACID SYNTHASE REGULATES DISEASE PROGRESSION IN THE

PTEN KNOCKOUT PROSTATE CANCER MOUSE MODEL

III.A Abstract 82

III.B Introduction 83

III.C Experimental Procedures 85

III.D Results 87

III.E Discussion 94

III.F References 100

III.G Acknowledgements 104

III.H Figures and Figure Legends 105

xvi

CHAPTER IV

GENERAL DISCUSSION

(Sections from a published book chapter in: Polyunsaturated Fatty Acids: Sources, Antioxidant

Properties and Health Benefits, A. Catalá, Editor. 2013. New York: Nova Science Publishers.)

IV.A Biological Implications of de novo Fatty Acid Synthesis 115

IV.B Inhibiting De Novo Fatty Acid Synthesis 117

Effects in normal biology

Effects in cancer biology

Inhibiting FASN

Other mechanisms of inhibiting the fatty acid synthesis pathway

Inhibitors of fatty acid synthesis

IV.C Dietary and Metabolic Considerations during FASN Inhibition 124

Dietary fat and cancer

Obesity and cancer

Adipose tissue and cancer

Cancer associated adipocytes

Inhibiting FASN in cancer and obesity

IV.D Concluding Remarks 127

IV.E References 129

V. CURRICULUM VITAE 135

xvii

ABSTRACT

Fatty acid synthesis is the process of producing de novo fatty acids from carbohydrate

and protein carbon sources. The enzyme responsible for this task is appropriately termed Fatty

Acid Synthase (FASN). FASN is ubiquitously expressed throughout the body, but because fatty

acid synthesis is a very energy-demanding process, its stimulation in cells is heavily regulated to

maintain metabolic homeostasis under normal physiological conditions. In contrast, cancer cells

undergo numerous advantageous, metabolic alterations to promote the activation of the fatty acid

synthesis pathway, and increase the expression of FASN. Through the development of two

different Fasn knockout models, the data in this dissertation outlines the requirement of FASN to

fulfill tissue-specific functions in both a normal and pathological setting. Chapter II shows that

deletion of FASN in the lactating mammary gland results in lost functionality and premature

involution of the gland to a near, pre-pregnancy state. Although, secretory activation was not

disturbed, milk fatty acids were significantly decreased and nursing pups experienced growth

retardation and perished prior to weaning, which was rescued by cross-fostering with a wild-type

mother. Chapter III demonstrates the requirement of FASN for the pathological progression of

prostate cancer. Prostates from Pten knockout mice showed overall decreased total prostate

weight, histopathological progression, proliferation, and activation of AKT. These studies

emphasize the requirement of FASN to sustain tissue-specific functions in both normal and

cancer cells in vivo. Importantly, the data validates FASN as an attractive therapeutic target for

the development of new cancer drugs.

1

CHAPTER I:

GENERAL INTRODUCTION

The following introduction contains sections that were published by Janel Suburu and Yong Q.

Chen [1].

The manuscript was prepared by Janel Suburu and edited by Dr. Yong Q. Chen, whom also acted

as an advisory.

2

I.A ABSTRACT

Fatty acids are essential components of life, and there two ways a cell can acquire them –

dietary fat consumption or de novo lipid synthesis. The metabolism and availability of dietary

fatty acids in peripheral tissues is encumbered by numerous factors regulating absorption,

transport, and cellular uptake. In contrast, de novo fatty acid synthesis is a mechanism by which

cells can generate fatty acids from carbohydrate and protein taken up by the cell as primary

energy sources. The de novo fatty synthesis pathway is an energy-expensive pathway, and

therefore, is heavily regulated by various hormones and signaling molecules to guarantee cellular

metabolic efficiency. Fatty acid synthase (FASN) is the primary enzyme of the fatty acid

synthesis process that performs the function of sequentially adding carbons in a cyclic fashion to

produce the saturated 16-carbon fatty acid, palmitate. Although the expression of FASN is fairly

ubiquitous, few tissues demonstrate high expression of FASN. In contrast, strong expression of

FASN has been demonstrated in the vast majority of cancers. Evidence suggests the activation of

fatty acid synthesis in cancer is a unique characteristic of cancer cells. Numerous metabolic

alterations and a shift from catabolic to anabolic metabolism feed the process of fatty acid

synthesis in cancer cell, which may use the fatty acids for membrane biogenesis, protein

modification, or lipid ligands. Finally multiple cancer signaling pathways have been shown to

promote the activation of fatty acid synthesis, suggesting the fatty acid synthesis pathway is an

oncogenic addiction that may be exploited as a target for therapeutic intervention.

3

I.B FATTY ACID METABOLISM

Types of fatty acid: their sources and roles

Fatty acids are essential components of cellular biology. They are a unique class of

organic molecules that vary in their structure, source, and use within the body. In general, fatty

acids are composed of a carbon chain that extends from 4 to 36 carbons; however, only chains of

24 carbons or less are found in humans. In addition to the variation in their carbon chain length,

fatty acids also differ in their degree of saturation, or their number of double bonds. These

structural biochemistry variations among fatty acids dictate their use and their influence in a

biological system [2].

A fully saturated fatty acid (SAFA) contains zero double bonds; the most common types

of SAFA are palmitate (16:0) and stearate (18:0). SAFA are primary constituents of biological

membrane and are found within the various species of glycerolipids, phospholipids, and

sphingolipids. SAFA may also serve as post-translational protein modifiers, which can regulate

protein activation and localization within a cell. Dietary sources enriched in SAFA include

animal fat, milk, coconut oil, and palm oil [3]; however, SAFA may also be produced de novo

from carbohydrate and protein sources through the fatty acid synthesis pathway .

Monounsaturated fatty acids (MUFA) contain one double bond. Oleate (18:1) is the most

common type of MUFA; its double bond is observed at the omega-9 carbon position of the acyl

chain. Similar to oleate, most MUFA contain their double bond at the omega-9 position, but for

some MUFA it resides at the omega-7 position. MUFA are also major constituents of biological

membrane and found in complex lipid molecules such as glycerolipids, phospholipids, and

sphingolipids. Olive oil, almond oil, and palm oil are examples of highly enriched sources of

4

dietary MUFA, but like SAFA, MUFA may also be produced de novo from carbohydrate and

protein carbon sources, as well as dietary SAFA [3].

Polyunsaturated fatty acids (PUFA) contain more than one and up to six double bonds,

and are quite unique fatty acids relative to SAFA and MUFA. In their natural form, the double

bonds of fatty acids are situated in cis conformation, making the shape of highly unsaturated

PUFA (>2 double bonds) exceptionally bulky compared to SAFA and MUFA. As such,

incorporation of PUFA into membranes can significantly increase the membrane fluidity, as well

as disrupt cellular signaling platforms in the plasma membrane [4-6]. Their high degree of

unsaturation also makes them particularly susceptible to peroxidation by intracellular free

radicals, making PUFA enrichment in the cell a potential source for increased reactive oxygen

species [7]. Finally, PUFA also serve as precursors to a large variety of important signaling

mediators for the body’s immune and cardiovascular systems. Most importantly, PUFA cannot

be synthesized de novo in mammals due to the absence of the Δ12 desaturase enzyme that is

required to produce the limiting substrate for PUFA synthesis; therefore, PUFA must be acquired

from the diet and are thereby considered essential fatty acids. Soybean, sunflower, safflower,

corn, and fish oils are all PUFA-enriched dietary fats [3].

Dietary fatty acid metabolism

The use of dietary fat to fulfill the biological roles of fatty acids is encumbered by a

lengthy physiological process that requires many enzymes, transporters, and chaperones to

facilitate their absorption, distribution, and uptake. First, dietary fatty acids are typically

consumed in the form of triglycerides and must be broken down in order to be efficiently

absorbed in the gut. Pancreatic lipase cleaves two fatty acids from the glycerol backbone and the

resultant free fatty acids and monoacylglycerol are emulsified by bile salt to facilitate their

5

transport across the intestinal lining. The fatty acids are then reincorporated into triglycerides,

loaded onto lipoprotein particles, and enter the lymphatic system. At the thoracic duct, the

lipoprotein particles enter the blood circulatory system where they can bind to their respective

receptors for cellular uptake or be released as free fatty acids by endothelial enzymes. The

release of fatty acids from lipoprotein particles requires the enzymatic action of lipoprotein

lipase; thus, the use of dietary fatty acids in peripheral tissue is largely dependent on the

expression and activity of various lipase enzymes [2, 8].

Following their release from lipoprotein particles, dietary fatty acids may be modified by

desaturation or elongation. While mammals lack the Δ12 desaturase enzyme to produce long

chain PUFA, other desaturase enzymes have been characterized: FADS1 (Δ6 desaturase

enzyme), FADS2 (Δ5 desaturase enzyme), and stearoyl-CoA desaturase (SCD1, also known as

the Δ9 desaturase enzyme). FADS1 and FADS2 specifically act on PUFA , whereas SCD1 is

specific for the conversion of SAFA to MUFA [9]. SCD1 introduces a double bond between the

9th

and 10th

carbons of most long chain SAFA with stearate as the preferred substrate, which is

converted to oleate (18:1) [10]. Fatty acids can also be elongated by one of seven known

mammalian elongase enzymes (ELOVL1-7) through the addition of malonyl-CoA, some of

which seem to have selective fatty acid substrate specificity [9]. Ultimately, the resultant fatty

acids are utilized for energy, protein modification, or incorporation into complex lipid structures

for cellular signaling and membrane integrity.

6

I.C DE NOVO FATTY ACID SYNTHESIS

The pathway enzymes and regulation

As an alternative to acquiring fatty acids from the diet, cells have the potential to produce

fatty acids through the fatty acid synthesis pathway. Fatty acid synthesis occurs in the cytoplasm

and begins with the production of acetyl-CoA from citrate by ATP citrate lyase (ACLY). Acetyl-

CoA is then converted to malonyl-CoA by acetyl-CoA carboxylase (ACACA, commonly

referred as ACC1); this is the rate-limiting step of fatty acid synthesis. Fatty acid synthase

(FASN), the central enzyme to the process, then uses one acetyl-CoA and seven malonyl-CoA

molecules through a series of catalytic domains to produce palmitate, a saturated 16-carbon fatty

acid (16:0). Palmitate is the primary product of FASN, representing about 80-90% of total fatty

acids produced; however, FASN also produces myristate (14:0) and stearate (18:0) [11, 12].

FASN will be the major topic in later sections.

Fatty acid synthesis is a very energy demanding process. The following equation

generalizes the major substrates and products of fatty acid synthesis:

8 Acetyl-CoA + 7 ATP + 14 NADPH → 1 Palmitic acid + 7 CO2 + 14 NADP+

Because of its large energy expenditure, the pathway is very tightly regulated, at both

transcriptional and post-transcriptional levels. Activation of fatty acid synthesis normally occurs

in response to excessive caloric consumption of a high-carbohydrate diet, especially during re-

feeding following a period of fasting; hence, the pathway is controlled both rapidly and slowly

by multiple hormones and nutritional signals, including insulin and glucagon [13, 14].

Acute regulation of the pathway occurs through ACC1 [13]. Expression of ACC1 is

transcriptionally induced by activation of sterol response element binding protein 1 (SREBP-1)

7

[15]; however, it is largely controlled at the post-transcriptional level to facilitate acute

regulation of fatty acid synthesis. ACC1 is allosterically activated by citrate and allosterically

inhibited by fatty acids [13]. Additionally, an inhibiting phosphorylation of ACC1 is

accomplished by the activation of AMP-activated protein kinase (AMPK), the cell’s central

energy sensor [16]. Accordingly, a high intracellular AMP to ATP ratio, indicating low cellular

energy levels, activates AMPK, and intuitively, inhibits the energy-expensive fatty acid synthesis

pathway via phosphorylation of ACC1 [17].

Although ACC1, located in the cytoplasm, is primarily implicated in fatty acid synthesis,

a second mitochondrial isoform, ACC2, also contributes to metabolic homeostasis in the liver,

heart, skeletal muscle, and lung, by producing mitochondrial malonyl-CoA to inhibit fatty acid

oxidation during active fatty acid synthesis [18, 19]. Emphasizing the importance of fatty acid

synthesis in growth and development, deletion of ACC1 in mice results in an embryonic lethal

phenotype [20], whereas deletion of ACC2 in mice promotes constitutive fatty acid oxidation

and protection from diet-induced obesity, fatty-liver, and diabetes [21-24]. The two enzymes are

believed to provide distinct roles in regulating fatty acid synthesis and fatty acid oxidation, since

most reports have not indicated any compensatory mechanisms [20, 25, 26]; however, one report

does demonstrate that the liver can increase transcription of ACC2 as a means of compensating

for ACC1 inhibition in mice [27].

Fasn: the enzyme

There are two types of fatty acid synthases: type I FAS (commonly referred to as FASN)

and type II FAS. In bacteria and plants, fatty acid synthesis (via type II FAS) occurs through a

step-wise enzymatic process performed by individual peptides, whereas animal fatty acid

synthesis (type I FAS) is performed by is a multifunctional, single polypeptide that is transcribed

8

from one gene. Despite these differences, the functional domains of type I and type II FAS are

highly conserved across all organisms, suggesting de novo fatty acid synthesis has been essential

throughout evolution and is a critical component to sustaining life [28, 29].

The FASN protein is a highly sophisticated homodimer with multi-enzyme activity.

Translation of the mRNA transcript results in a 2,504 amino acid sequence protein of about

260kDa that contains 7 functional domains. The malonyl/acetyl transacylase (MAT) domain

loads its substrates to the acyl carrier protein (ACP), which acts as a coenzyme as it binds

reaction intermediates via a 4’-phophopantetheine group and shuttles the acyl chain across the

various enzymatic domains. The β-ketoacyl synthase (KS) domain condenses the substrates to

lengthen the acyl chain. The following reduction reaction is then performed in a three step,

NADPH-dependent process by the β-ketoacyl reductase (KR), β-hydroxyacyl dehydratase (DH),

and enoyl reductase (ER) domains. This cycle is then repeated for further elongation or

terminated by the thioesterase (TE) domain, which cleaves the fatty acid from the ACP [13] (Fig.

1). Two orientations of the FASN dimer have been described: head-to-tail and head-to-head. The

classical head-to-tail orientation was proposed from early evidence that FASN requires

dimerization of the two polypeptides for full functionality, whereby the KS domain of one

subunit is only active when juxtaposed with the ACP of the other subunit [30]. The head-to-tail

model was widely accepted for well over 20 years until the early 2000s when work from Stuart

Smith’s group began to challenge the model, showing evidence of a new head-to-head

orientation [31, 32]. In 2008, the head-to-head model of FASN was confirmed from a high, 3.3 Å

resolution crystal structure of porcine FASN complexed with and without NADP+ [33]. FASN is

structurally organized into an “X” shaped figure, containing an underside, condensing function

from the MAT and KS domains and an upper side with β-carbon-modifying function from the

9

KR, DH, and ER domains; unfortunately, the ACP and TE domains remained unresolved, but

were most likely situated between the bottom and upper segments (Fig. 2). The dimer is held

together by interactions occurring in the KS, ER, and DH domains. Because the dimeric enzyme

can operate with only one functioning KS or MAT domain [31], the authors hypothesized a

rotational, or twisting, mechanism between the upper and lower segments of the “X”.

Interestingly, analysis of the crystal structure also identified two new non-enzymatic domains

linearly located between the DH and ER domains, formerly known as the interdomain, and

structurally located as a protrusion of the enzyme’s upper segment. The new, non-enzymatic

domains had homology to the methyltransferase and ketoreductase enzyme families, and thus,

were coined the “pseudo-methyltransferase” (ΨME) and “pseudo-ketoreductase” (ΨKR)

domains, respectively. While their true functions remain unclear, the authors hypothesized that

these non-enzymatic domains may act as roadblocks to restrict the movement of the ACP, since

the ACP of FASN was previously reported to not sequester its covalently bonded acyl chain, a

distinct mechanism from type II FAS ACP [34]. Due to the dynamic movement of the ACP, its

structural resolution has been very difficult to achieve. However, a recent publication described a

trapping mechanism that allowed high resolution structural analysis of the type II FAS ACP in

two different conformations, providing action shots of ACP movement [35]. While the type I and

type II FAS differ in their respective ACP mechanisms [34], this may provide insight guiding

future biochemical studies of the FASN structure.

Fasn: regulation

Located on chromosome 17q25 of the human genome, the FASN gene has 42 relatively

small introns, 2 promoter regions, and a coding sequence of approximately 7512 base pairs [12,

36]. FASN cDNA from goose, chicken, rat, mouse, pig, and human have been characterized, and

10

sequence conservation is high at both the nucleotide and amino acid level [28, 29]. Activation of

FASN is heavily regulated by a multitude of nutritional and hormonal indicators to ensure

metabolic homeostasis. Increased expression of FASN occurs in response to insulin, thyroid

hormone, and corticosteroids, whereas inhibition of FASN is induced by glucagon, leptin, growth

hormone, essential amino acid deprivation, and PUFA [29, 37-41].

Early evidence demonstrated that FASN expression is mainly regulated by transcriptional

activation [42]. Several response elements including ones for insulin, carbohydrate, and sterol

are all present in the FASN promoter region, as well as an E-box motif. Transcriptional activation

at the promoter region is largely regulated by Stimulatory Protein 1 (Sp1), Nuclear Factor Y

(NF-Y), Upstream Stimulatory Factor 1 (USF1), Upstream Stimulatory Factor 2 (USF2), and

SREBP-1 transcription factor binding, but promiscuous activation at the E-box motifs by other

transcription factors, such as MYC, is also possible [29, 43-46]. Interestingly, the second

promoter region in the first intron was shown to largely exist as an inhibitor of the first promoter

region, which hints at tissue-specific regulation of FASN expression [36, 47].

Although, transcriptional regulation is long believed to be the primary mechanism of

induced FASN expression, post-transcriptional modulation has also been reported. In particular,

increased mRNA stability by insulin and glucocorticoids and FASN protein stabilization are

known to positively regulate Fasn [40, 41, 48-50]. In contrast, phosphorylation of FASN has

been reported to either activate or inhibit FASN, depending on the cell type. The phosphorylation

sites for FASN activation remain unknown; however, phosphorylation at threonine 1029 and

1033, which flank the catalytic residue of the dehydratase domain, inhibit FASN, and

consequently, PPARα activation, in response to feeding or insulin [51-53]. Finally, hepatic

FASN was also reported to be acetylated. Although, the regulatory mechanism of FASN

11

acetylation was not defined, protein lysine acetylation appears to affect most metabolic enzymes,

modulating both enzyme activity and stability. Importantly, enzyme acetylation was regulated by

nutrient status, suggesting it is an acute regulatory mechanism of metabolism [54].

Fasn: expression and biology

Expression of FASN is relatively weak for many differentiated, adult tissues; however,

expression is ubiquitous and certain tissue types show particularly high expression of FASN [12,

55]. Its expression is observed in both proliferating and resting cells, but particularly in hormone-

sensitive and high lipid-metabolizing tissues, as well as tissues responsible for producing lipid

barriers [56]. As such, FASN appears have several tissue-specific functions. In the liver, FASN

produces fatty acids for storage as triglycerides in response to excess food consumption [13].

The lactating mammary gland uses FASN to produce fatty acids for milk triglyceride production

[13, 56]. In the lung, type II alveolar cells express FASN to produce lipid-enriched lung

surfactant [56, 57]. The brain was found have relatively higher FASN expression [12], and it was

shown to regulate energy homeostasis [58]. The cycling endometrium shows biphasic expression

of FASN that correlates with the estrogen and progesterone phases of menstruation and

development of the decidua [56, 59]. Finally, the sebaceous gland cells in the skin were shown to

have very high expression of FASN, where it likely acts to provide fatty acids for the

maintenance of the skin’s lipid barrier [56]. Other tissue types, including pancreas, heart, and

kidney, demonstrated a low, basal-like level of expression of FASN [12, 55]; however, it is

believed that nearly all tissues are capable of inducing a lipogenic phenotype through the

transcriptional activation of FASN [60].

In contrast to its selectively high expression in adult tissues, FASN is strongly expressed

in nearly all fetal cell types, especially actively proliferating cells [56]. Highlighting its

12

importance during development, genetic deletion of Fasn in mice leads to an embryonic lethal

phenotype. Fasn-null embryos perish prior to even uterine implantation (before embryonic day

3.5), and only 30-40% of heterozygotes survive to parturition, suggesting a degree of haploid

insufficiency. Heterozygous females that do survive to adulthood rapidly lose fecundity after

only 3 months of breeding and progressively produce even fewer Fasn+/-

mice, most likely due to

impaired production of Fasn- gametes. These findings strongly implicate the significance of

FASN for growth and development [61]. Despite the lethal phenotype of whole body Fasn

knockout, several animal models of conditional, tissue-specific Fasn deletion have been reported,

highlighting unique roles for fatty acid synthesis in the liver, hypothalamus, myocardium,

endothelium, and intestine [58, 62-65]. Additionally, FASN in skeletal muscle, adipose, and

macrophage appears to play a significant role in diet-induced pathogenesis [66-68].

The first tissue-specific deletion of Fasn was reported in the liver of mice. Mice

harboring the liver-restricted deletion of Fasn (FASKOL) showed no major phenotype until they

were fed a zero-fat diet that forced mobilization of peripheral fat stores. Upon dietary restriction,

FASKOL mice experienced severe hypoglycemia and hepatosteatosis, similar to the

characteristics of a PPARα knockout mouse [69]. Restoration of the zero-fat diet-induced

phenotype was achieved by administration of a PPARα agonist, demonstrating the importance of

liver FASN-mediated activation of PPARα in the liver [62]. Not long after, a FASN-derived

phosphatidylcholine species was identified as the major signaling mediator of the FASN-

mediated activation of PPARα [70], and a new report [53] recently resolved the seemingly

paradoxical interface between these two energy-storing (fatty acid synthesis) and energy-burning

(PPARα activation) pathways. The liver is now known to possess two distinct subcellular pools

of FASN – one localized to the cytoplasm and the other membrane-associated. During fasting,

13

the cytoplasmic FASN is responsible for producing PPARα ligands to stimulate β-oxidation,

whereas insulin stimulation from feeding induces mTORC1-mediated phosphorylation and

inhibition of cytoplasmic FASN, while simultaneously activating the membrane-associated

FASN for triglyceride production [53].

Conditional deletion of Fasn in the mouse brain (FASKO) demonstrated a significant

contribution of hypothalamic fatty acid synthesis to energy homeostasis and feeding behavior.

FASKO mice showed hypophasia mediated by impaired PPARα activation in the hypothalamus

that was rescued by intracranial injection of a PPARα agonist. Because FASKO mice were

generated with a Cre transgene under the rat insulin 2 promoter, deletion also occurred in

pancreatic β-islet cells [58]. Regulation of food intake and pancreatic β-cell function are tightly

interrelated [71]. Therefore, it was hypothesized that deletion of Fasn in β-islet cells may also

regulate feeding; however, β-cell-specific deletion of Fasn had no effect on islet function, despite

significantly decreased FASN activity and increased malonyl-CoA [58]. Although unable to

rescue the hypophagic phenotype, FASKO mice fed a high-fat diet experienced significant

protection from diet-induced obesity and inflammation. Compared to wild-type mice, FASKO

mice showed decreased weight gain and adiposity, improved metabolic performance, resistance

to fatty liver, reduced oxidative stress, and decreased endotoxin-induced cytokine release.

Together these studies highlight a complex role for hypothalamic FASN in regulating

bioenergetics and systemic inflammation during both normal and high-fat feeding [72].

Continued evidence of normal physiological roles of de novo fatty acid synthesis came

from mouse models with a conditional deletion of Fasn in vascular endothelial cells (FASTie)

and intestinal epithelial cells (FAS-villin), which demonstrated its requirement for protein

palmitoylation and regulation of systemic inflammation [64, 65]. Endothelial cells lacking FASN

14

from Cre expression under the Tie2 promoter showed decreased palmitoylation of endothelial-

nitric oxide synthase (eNOS), which prohibited its membrane localization. Although expressed

protein levels of eNOS remained unchanged, its decreased bioavailability at the plasma

membrane led to increased vascular permeability that reflected a heightened inflammatory

response and decreased angiogenesis [64]. Intestinal deletion of Fasn from inducible-Cre

expression under the Villin promoter significantly decreased FASN expression in the distal small

intestine and the colon, which caused major disruption of the intestinal barrier function.

Decreased palmitoylation and, consequently, secretion of mucin 2 (Muc2), a major protein

implicated in the intestinal mucosal lining [73], strongly increased the intestinal permeability to

gut microbiota, causing systemic inflammation and sometimes death to FAS-villin mice within 2

weeks of inducing Cre expression [65]. These studies demonstrate the importance of FASN-

mediated protein acylation in two very different physiological functions.

Mice bearing a conditional knockout of Fasn in the myocardium (FASKard) showed that

de novo lipogenesis in the heart is essential for regulating cardiac function during stress. No

differences in fatty acid and glucose oxidation, nor cardiac output, were observed between

FASKard and control mice at rest. However, FASKard mice with surgically-induced left

ventricular hypertrophy experienced severe arrhythmia and cardiac failure that lead to death

within 1 hour. Unlike previous models of Fasn deletion [58, 62], an analysis of PPARα and its

target genes revealed it was not involved in the observed phenotype. Instead, hyperactivation of

L-type calcium channels by increased autophosphorylation of Ca++

/calmodulin protein kinase II

(CaMKII) was shown to, at least in part, mediate the cardiac dysfunction. Administration of a

CaMKII inhibitor to FASKard mice significantly ameliorated the effects of Fasn deletion, but

administration of palmitate had no effect on cultured myocytes treated with Fasn shRNA.

15

Additionally, FASKard mice had a significantly decreased lifespan compared to control mice, as

nearly 50% of mice perished within one year of age. Signs of progressive heart failure in

FASKard mice as early as 6 months suggested cardiac FASN is critical for general heart health

[63].

Mice with a conditional deletion of Fasn in skeletal muscle (FASKOS) showed aberrant

calcium signaling upon feeding a high-fat diet, which led to significantly increased insulin

sensitivity and enhanced glucose tolerance. When fed a high-fat diet, control mice and FASKOS

mice showed no difference in adiposity or lean mass, nor serum glucose, free fatty acids,

triglycerides, cholesterol, insulin, or leptin; they also had similar skeletal muscle metabolites.

However, the deletion of Fasn resulted in increased activation of AMPK and increased cytosolic

calcium in skeletal muscle cells. Although deletion of PPARα in skeletal muscle is known to

activate AMPK [74], the FASN-PPARα signaling axis was not the mediator in this scenario, but

rather, occurred in a CaMKβ-dependent manner. Although a direct mechanism remains unclear,

FASN was found to interact with sarco/endoplasmic reticulum calcium ATPase 1 and 2

(SERCA1 and SERCA2) and significantly changed the phospholipid profile of the sarcoplasmic

reticulum by increasing the phosphatidylcholine to phosphatidylethanolamine ratio. This

evidence suggested FASN activates CaMKβ and AMPK by regulating intracellular calcium via

SERCA1 and SERCA2 [66]. The activation of AMPK in FASKOS mice fed a high-fat diet

increased translocation of the glucose 4 transporter (GLUT4) to the plasma membrane and

increased glucose uptake [66], even though it has been previously demonstrated that a high-fat

diet inhibits skeletal muscle AMPK [75]. Although the increased cytosolic Ca++

benefited

FASKOS mice in preventing diet-induced insulin resistance, it came at the expense of decreased

muscle strength [66]. This study, in combination with evidence from FASKard mice, suggests

16

FASN is not required for resting muscle physiology, but has implications for muscle function

under certain physiological stresses [63, 66].

Deletion of Fasn in adipose tissue of mice (FASKOF) showed no effect on adiposity or

glucose and insulin tolerance compared to control mice, unless they were fed a high-fat diet.

Upon high-fat feeding, FASKOF mice showed reduced adiposity by adipocyte size, but not

adipocyte number, increased lean weight, energy expenditure, and glucose and insulin tolerance

[67]. Because a high-fat diet is actually known to decrease FASN expression [76], the reduction

in adiposity and improved metabolic profile in FASKOF mice most likely occurred in response

to reduced PPARγ activation, which controls both white and brown fat adipogenesis [77, 78],

and consequentially, increased PPARα activation and thermogenic reprogramming [67]. Indeed,

FASKOF mice on a high-fat diet showed increased expression of uncoupling protein 1 (UCP1)

and fatty acid oxidation in inguinal, but not epididymal, fat, and switching to a zero-fat diet

exacerbated UCP1 expression and other thermoregulatory genes in white adipose tissue.

Knockdown of peroxisomal reductase activating PPARγ (PexRAP) in C57Bl/6 mice with high-

fat feeding showed a very similar phenotype to FASKOF mice, which was reversed by treatment

with rosiglitazone, a PPARγ agonist [67]. This data, along with evidence from FASKOS mice,

strongly implicate FASN in adipose and skeletal muscle as contributors to diet-induced obesity

and diabetes [66, 67].

Finally, the inhibition of Fasn by its conditional knockout in peritoneal macrophage of

apoE-null mice (FASKOM) showed a 20-40% decrease in atherosclerotic lesions in multiple

areas of the aorta. Atherosclerosis in apoE-null mice was shown to be regulated by a FASN-

mediated inhibition of liver X receptor α (LXRα). The release of LXRα inhibition led to

increased cholesterol efflux by ABCA1 and decreased uptake of oxidized low density lipoprotein

17

(LDL) by CD36. Although macrophage accumulation at aortic lesions was unaffected by FASN

knockout, together these effects resulted in significantly decreased cholesterol levels in both

macrophage and atherosclerotic lesions. Overall, this study implicates an important role for

FASN in diet-induced atherosclerosis [68].

The many conditional, tissue-specific knockout mouse models of Fasn have demonstrated

its functional role in maintaining physiological homeostasis across a wide variety of biological

systems in the body. In several cases it relates to metabolic regulation, whereas others cases

pertain to activation of tissue-specific proteins. Under both normal feeding and in a setting of

high-fat diet feeding, FASN appears to regulate various cellular and physiological processes that

had otherwise not been implicated. Continued development of other tissue-specific Fasn

knockout mouse models will further identify unique roles for FASN and de novo fatty acids in

modulating physiological processes.

I.D DE NOVO LIPOGENESIS IN CANCER

Cancer versus normal metabolism

Unlike normal cells, which are highly regulated when it comes to balancing energy

consumption and expenditure, cancer cells are characterized by a large number of metabolic

alterations. One major change from their normal counterpart is a shift from catabolic to anabolic

metabolism. Otto Warburg first described this metabolic shift nearly a century ago [79]. The

proclaimed Warburg Effect describes the unique phenomenon of aerobic glycolysis in a cancer

cell, whereby cancer cells consume a large quantity of glucose, metabolize it via glycolysis, and

release the majority as lactic acid into the extracellular space – most peculiarly, this occurs under

normal oxygen conditions. Normally, each molecule of glucose consumed by a cell is

18

metabolized through glycolysis to two molecules of pyruvate. Pyruvate is then converted to

acetyl-CoA in the mitochondria where it enters the tricarboxylic acid (TCA) cycle to produce

redox substrates for oxidative phosphorylation, the cell’s major energy-producing, catabolic

pathway. Interestingly, cancer cells seem to consume an excessive quantity of glucose, and

utilize it both catabolically, as just described, and anabolically, whereby the carbons are used as a

source to synthesize and fulfill the macromolecular demand of their proliferative phenotype.

Although the production of amino acids and nucleic acids is certainly an integral characteristic of

the cancer anabolic phenotype, cancer cells seem to require de novo fatty acids and the specific

use of alternative enzymes, metabolic pathways, and common oncogenic pathways to promote

the synthesis of fatty acids.

Cancer metabolism favors fatty acid synthesis

Cancer cells have adapted the use of alternative metabolic pathways and enzymes to

provide the necessary substrates for lipogenesis and prevent its inhibition. These alternative

pathways include isoform switching, truncation of the TCA cycle and the use of TCA cycle

intermediates for fatty acid synthesis, as well as increased use of other non-canonical metabolic

enzymes, such as the pentose phosphate pathway and cytosolic TCA cycle enzymes (Fig. 3).

Metabolism for fatty acid synthesis ultimately begins with cellular uptake of glucose.

Cancer cells typically consume a large quantity of glucose as their major energy source. Upon

entering a cell, glucose is phosphorylated by hexokinase (HK) to glucose-6-phosphate (G6P).

Humans possess four isozymes of HK (HK1-4) and most cell types express HK1 as the

housekeeping enzyme for normal catabolic metabolism of glucose [80]. However, cancer cells

seem to preferentially express HK2, which is thought to support anabolic metabolism [81].

Prostate cancer cells strongly upregulate HK2 and lipid synthesis in response to androgen [82].

19

G6P, produced by HK2, may take one of two fates: metabolism by the pentose phosphate

pathway (PPP) or glycolysis. Cancer cells can shunt G6P through the PPP to produce NADPH,

which is a critical reductant for FASN. PPP intermediates can also return to glycolysis,

producing pyruvate, which may be used later for fatty acid synthesis.

Tumor cells preferentially express the embryonic isoform of pyruvate kinase (PKM2), the

terminal enzyme of glycolysis. Compared to PKM1, the splice variant found in most adult

tissues, PKM2 is characterized by a weaker substrate affinity and decreased activity [83, 84].

Insights to the advantages of which PKM2 imposes on cancer cells have surfaced only recently.

Studies by Cantley and colleagues have shown PKM2 promotes tumor growth and survival by

maintaining a high glycolytic rate, low oxidative phosphorylation, and redox homeostasis [85,

86]. Expression of PKM2 in prostate cancer was also shown to be associated with an alternative

mechanism of producing pyruvate in the absence of ATP production [87]. This suggests

expression of PKM2 in prostate cancer cells promotes anabolic metabolism, such as fatty acid

synthesis, by allowing high glucose consumption as a carbon source for fatty acids, but without

inducing ATP-mediated inhibition of glycolysis [87].

Typically, glucose-derived citrate is processed through the mitochondrial TCA cycle to

produce intermediates for oxidative phosphorylation. In contrast, cancer cells shunt citrate from

the TCA cycle to the cytoplasm and use alternative methods to metabolize citrate that promote

fatty acid synthesis. Cytosolic citrate can be converted to isocitrate by cytosolic aconitase

(ACO1), and subsequently to α-ketoglutarate by cytosolic isocitrate dehydrogenase (IDH1); the

latter reaction produces additional NADPH for fatty acid synthesis. Overexpression of IDH1 in

male mice leads to significantly increased epididymal fat weight and adipogenesis, coinciding

with hyperlipidemia and obesity [88]. Interestingly, oncogenic mutations in IDH have been

20

characterized in several cancer types [89, 90]. These findings suggest IDH1 plays a significant

role promoting fatty acid synthesis and lipid metabolism in cancer.

Cytosolic citrate, in addition to being metabolized outside of the TCA cycle to produce

NADPH, can also be converted to acetyl-CoA by ACLY, and can be used for both fatty acid and

sterol synthesis. A byproduct of this reaction is oxaloacetate, which is converted to malate by

malate dehydrogenase. Cytosolic malate may be converted back to pyruvate by cytosolic malic

enzyme (ME1) to produce yet more NADPH, or it can be recycled back to the mitochondria by

the citrate transport protein in the mitochondrial membrane, which coordinates the influx of

cytosolic malate with the efflux of mitochondrial citrate [91], completing a loop for driving

mitochondrial carbon sources toward fatty acid synthesis. Prostate tissue is unique in its transport

system of citrate in that normal prostate cells produce large amounts of intracellular citrate and

release it into the prostatic fluid [92], where it can act as an energy source for sperm [93].

Prostate cancer cells, however, have significantly decreased intracellular citrate levels. This is

accompanied by increased ACO1 expression and activity [94] and increased ACLY [95].

Collectively, these findings suggest that prostate cancer cells rapidly utilize citrate for

lipogenesis, either to produce NADPH or shunt as a carbon source for fatty acid production.

Although de novo synthesized fatty acids are typically derived from glucose, some cancer

cells rely on glutamine metabolism. Because glucose carbons are shunted from the mitochondria

for fatty acid synthesis, prostate cancer cells convert glutamine to α-ketogluterate and shuttle it

into the mitochondria to replenish TCA cycle intermediates for anaplerotic reactions [96, 97].

Additionally, cancer cells experiencing selective pressure by hypoxia, or a defective electron

transport chain, use glutamine, rather than glucose, as the major lipid precursor. In such a

scenario, stabilized hypoxia-inducible factor 1α (HIF-1α) inhibits the conversion of pyruvate to

21

acetyl-CoA, and IDH1 and ACO1 work in reverse to convert α-ketoglutarate to citrate, which is

then metabolized by ACLY for lipid synthesis [98, 99] (Fig. 3).

Fatty acid synthesis and cancer

It was recognized over 50 years ago that cancer cells exhibit increased production of de

novo fatty acids [100]. In line with an altered metabolism, cancer cells shunt glucose-derived

carbons from the mitochondrial TCA cycle to the cytosol to feed fatty acid synthesis. This

increased efflux of carbon toward fatty acid synthesis is accompanied by upregulation of the

enzymes in the fatty acid synthesis pathway. The major transcriptional regulator of enzymes in

this pathway is the sterol response element binding protein-1c (SREBP-1c). SREBP-1c is a

lipogenic transcription factor that regulates the expression of glycolytic enzymes and fatty acid

synthesis enzymes, including ACLY, ACACA, FASN, and SCD-1 [101]. Expression of SREBP-

1c can be stimulated by androgens and epidermal growth factor (EGF) in prostate cancer cells

[102, 103], and in turn, expression of the androgen receptor is regulated by SREBP-1c [104],

creating a positive feedback loop for the expression of lipogenic enzymes. Overexpression of

SREBP-1 is sufficient to increase tumorigenicity and invasion of prostate cancer cells [104]. Not

surprisingly, increased expression of FASN and the lipogenic phenotype has been documented in

various cancer types, including prostate [105], breast [106], ovarian [107], colorectal [108], and

many others [60, 109, 110]. Pharmacological or small-interfering RNA-mediated inhibition of

enzymes in the de novo fatty acid synthesis pathway, namely ACLY, FASN, or SCD-1 inhibits

prostate tumor growth [95, 111, 112]. Additionally, inhibition of ACACA, FASN, or SCD-1

induces cancer cell death [95, 111-116], demonstrating the requirement for de novo fatty acids

for prostate cancer survival.

22

Upregulation of fatty acid synthase in cancer can arise as a consequence of a variety of

events. Increased gene copy number of FASN in prostate adenocarcinoma coordinates with

overexpression of FASN protein [117]. Increased transcriptional activation of FASN has been

well described and is very common in prostate cancer [102, 103, 105, 113]. Stabilization of

FASN protein by ubiquitin-specific protease-2a (USP2a) has been shown to increase fatty acid

synthesis in prostate cancer [49]. Loss of tumor suppressor proteins, such as PTEN and LKB1,

results in prostate neoplasia and activation of AKT [118, 119], a well-described activator of

FASN expression [120]. Finally, FASN itself has been postulated as an oncogene, as its

overexpression in mouse prostate leads to prostatic intraepithelial neoplasia [121]. Collectively,

these cellular perturbations demonstrate that fatty acid synthesis is a unique characteristic of

cancer and, hence, an attractive therapeutic target for the treatment of prostate cancer.

Utility of de novo fatty acids in cancer

Unlike their normal counterparts, cancer cells require fatty acid synthesis for growth and

survival; hence, they likely utilize these de novo fatty acids to fulfill specific needs. Because

cancer cells divide rapidly, one major purpose for increased fatty acid synthesis is to supply the

heavy demand for new membrane biogenesis. In fact, Swinnen et al. [122] showed that de novo

fatty acids used for membrane biogenesis also preferentially localize in the membrane’s

signaling raft domains. Membrane composition has been shown to affect the conformation and

activation of membrane-bound proteins [123]; thus, de novo fatty acids may play a significant

role in propagating oncogenic signals from the plasma membrane. Indeed, FASN regulates

HER2 expression and activation of AKT [124]. Moreover, enrichment of saturated fatty acids by

FASN in tumor cell membranes protects cells from lipid peroxidation and oxidative stress-

induced cell death [125]. Overexpression of FASN in prostate cancer supports palmitoylation

23

and activation of oncogenic proteins [126]. Additionally, lipid signaling ligands can be derived

specifically from the de novo fatty acid pool [70], suggesting an absolute requirement of fatty

acid synthesis to activate specific signaling pathways. Although lacking evidence of a direct role

for fatty acids, FASN and ACACA are also required for cell cycle progression [116, 127].

Overall, prostate cancer cells may utilize de novo fatty acids for a variety of uses, all of which

facilitate proliferation and survival of cancer cells.

Oncogenic signaling pathways promote fatty acid synthesis

Further demonstrating the importance of fatty acid synthesis in prostate cancer, some of

the major disease-driving signaling pathways regulate the expression of lipogenic enzymes. As a

lipid phosphatase, PTEN opposes the action of PI3K by removing a phosphate group from

phosphatidylinositol-3,4,5-triphosphate, an essential activator of AKT [128]. Loss of PTEN, is

one of the most common events observed in prostate cancer, occurring in approximately 40% of

primary cancers and over 70% of castration resistant and metastatic prostate cancers [129-132].

PTEN deletion leads to invasive prostatic adenocarcinoma in mice [118], and induces expression

of FASN [133]. Treatment with a PI3K inhibitor or reintroduction of PTEN in Pten-null prostate

cancer cells reduces FASN expression [133]. Conversely, inhibition of FASN reduces expression

and activation of AKT [134], which was recently shown to occur by increased degradation of

AKT and its downstream effectors [135]. Additionally, cell growth inhibition by inhibiting

FASN can be overcome by hyperactivation of AKT [135]. Collectively, these studies strongly

suggest coordinate feedback between lipogenesis and PTEN/PI3K/AKT oncogenic signaling to

promote cancer growth and tumor progression.

HER2 is a well-recognized oncogene and diagnostic marker in multiple cancers [136].

Overexpression of HER2 in the absence of amplification has been identified in approximately

24

20% of prostate cancers, and significantly associates with high Gleason grade, proliferation, and

tumor recurrence [137]. It activates fatty acid synthesis via activation of the PI3K/AKT signaling

pathway [138]. HER2 regulation of fatty acid synthesis does not occur transciptionally via

SREBP-1c, but rather, translationally through activation of mTOR and increased FASN protein

[139]. Inhibition of FASN attenuates HER2 oncogenic signaling and downregulates activation of

AKT and ERK1/2 [124].

A number of studies have linked oxidative stress to prostate cancer development and

progression [140-143]. Hypoxia, a common feature of tumors characterized by lack of blood

supply, and consequently, nutrients and oxygen to the growing tumor, can lead to oxidative

stress. Furuta and colleagues [144] demonstrate that a hypoxic environment or treatment with

hydrogen peroxide induces AKT/HIF-1α-mediated activation of SREBP-1, which increases

transcription of FASN. SREBP-1 can also increase expression of NOX5 [104], a prominent

producer of ROS and regulator of prostate cancer cell growth [145], suggesting another feed

forward mechanism inducing lipogenesis and prostate cancer growth.

Androgen signaling is a major driver of prostate cancer, and androgen ablation is a

primary therapeutic strategy for prostate cancer patients. Activation of the androgen receptor

(AR) by androgen increases expression of lipogenic enzymes in a SREBP-1c-dependent manner

[102, 103], and a positive feedback loop promotes this signaling pathway since binding sites for

SREBP-1 transcription factors are also found in the AR gene of prostate cancer cells [104]. AR

is regulated by β2-microglobulin (β2-M), a component of the housekeeping major

histocompatibility complex class I molecule found on prostate cancer cells, in a MAPK/SREBP-

1-dependent manner. Inhibition of β2-M by a selective antibody decreases the interaction

between SREBP-1 and its binding site in the AR promoter region, resulting in decreased AR

25

expression and lipogenesis by FASN [146]. A new AR splice variant, AR8, was recently

identified and is strongly expressed in castration resistant prostate cancer (CRPC). AR8 is found

at the plasma membrane where it associates with EGFR, which induces nuclear translocation of

AR in response to EGF [147]. Because we know AR activates FASN [102], this may partly

explain how EGF activates FASN [103], and provides a potential mechanism for activation of

AR signaling in CRPC. AR8 also requires FASN for functionality, as palmitoylation appears to

tether it to the membrane [147]. Whether AR8 activates SREBP-1 is unknown. Interestingly,

inhibition of SREBP-1, ACACA, FASN, or SCD-1 results in decreased AR expression and

activity [104, 148, 149]. Concomitant overexpression of AR and FASN in prostate is sufficient

to induce adenocarcinoma in mice [121]. Overall, these results suggest androgen signaling is

strongly associated with activation of fatty acid synthesis and equally, fatty acid synthesis

promotes androgen signaling. Together these signaling pathways may work in concert with other

major oncogenic signaling pathways to promote prostate cancer development and progression.

I.E OVERVIEW

Many cancer types, especially hormone-sensitive cancers, overexpress FASN and depend

on de novo lipogenesis for proliferation, survival, and tumor progression. Fatty acid synthesis

appears to be a very attractive target for the development of cancer therapeutics; however, as

with any therapeutic drug, understanding their potential effects on normal tissue is a critical

component in assessing the cost-benefit of a cancer therapy. Because FASN has now been shown

to play an important role in the normal physiology of select tissue types, such as the intestine,

heart, brain, and endothelium, it is critical to evaluate how the inhibition of fatty acid synthesis

may affect other tissue types. This dissertation outlines the development and characterization of

26

two Fasn knockout mouse models. Chapter II reveals the generation of a mammary gland-

specific Fasn knockout mouse and highlights the requirement of FASN in maintaining the

functional competence of the lactating mammary gland. Chapter III describes the efforts to

knockout Fasn in a prostate cancer mouse model. Using two different genetic approaches we

genetically inhibited FASN in the Pten-null prostate cancer mouse model and describe its role in

prostate cancer progression. Chapter IV will discuss the implications of the findings from these

studies and considerations for the inhibition of FASN in normal physiology and cancer biology.

27

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39

I.G FIGURES AND LEGENDS

Figure 1. Schematic Representation of Palmitate Synthesis by Fatty Acid Synthase.

The fatty acid synthesis by FASN begins with the priming of acetyl-CoA and malonyl-CoA by

the malonyl/acetyl transacylase (MAT) domain. The β-ketoacyl synthase (KS) domain condenses

acetyl-CoA and malonyl-CoA. The condensation of the acyl chain is sequentially performed by

the β-ketoacyl reductase (KR), β-hydroxyacyl dehydratase (DH), and enoyl reductase (ER)

domains. The cycle is then repeated by sequential addition of malonyl-CoA until the final

product is released by the thioester domain (TE). The acyl carrier protein (ACP) shuttles the

growing acyl chain between each enzymatic domain by binding the fatty acid to its 4’-

phophopantetheine prosthetic group.

40

Figure 2. Organization of the Enzymatic Domains of Fatty Acid Synthase.

A. A cartoon representation of the folded protein structure fatty acid synthase. The enzymatic

domains are organized as an “X”-shaped protein homodimer. B. A cartoon representation of the

linear organization of the enzymatic domains of fatty acid synthase as a monomer. These images

are adapted from reference [32].

41

Figure 3. Cancer Cell Metabolism Promotes Fatty Acid Synthesis.

Glucose entering the cell is immediately phosphorylated by Hexokinase 2 (HK2), producing

glucose-6-phosphate (G6P). G6P can enter both glycolysis and the pentose phosphate pathway

(PPP). Expression of pyruvate kinase M2 (PKM2) promotes the ATP-free conversion of

phosphoenolpyruvate (PEP) to pyruvate by a currently unknown enzyme, which may prevent

ATP-mediated inhibition of glycolysis. Citrate is exported from the mitochondrial TCA cycle to

fuel the fatty acid synthesis pathway by conversion to acetyl-CoA by ATP Citrate Lyase

(ACLY). Acetyl-CoA carboxylase 1 (ACACA) initiates the first committed step to fatty acid

synthesis to produce malonyl-CoA. Seven malonyl-CoA molecules are added to acetyl-CoA by

fatty acid synthase (FASN) to produce palmitic acid (16:0). Palmitic acid can be further

elongated to stearic acid (18:0) and desaturated by stearoyl-CoA desaturase (SCD) to produce

oleic acid (18:1). Citrate may also be recycled to produce NADPH for fatty acid synthesis by

malic enzyme (ME1), which produces pyruvate, or by cytosolic aconitase (ACO1) followed by

cytosolic isocitrate dehydrogenase (IDH1) to produce α-ketoglutarate. Glutamine can be

converted to α-ketoglutarate upon entering the cell, to replenish TCA cycle intermediates or fuel

fatty acid synthesis. The reduced form of nicotinamide adenine dinucleotide phosphate

(NADPH), a critical reductant for fatty acid synthase (FASN), is produced from the activity of

the PPP, ME1, and IDH. This figure is adapted from reference [1].

42

43

CHAPTER II

FATTY ACID SYNTHASE IS REQUIRED FOR MAMMARY

GLAND DEVELOPMENT AND MILK PRODUCTION DURING

LACTATION

Janel Suburu, Lihong Shi, Jiansheng Wu, Shihua Wang, Michael Samuel, Michael J. Thomas,

Nancy D. Kock, Guangyu Yang, Steven Kridel, and Yong Q. Chen

The following chapter was published online as an Article in Press in the American Journal of

Physiology – Endocrinology and Metabolism on March 25, 2014.

PMID: 24668799

This study and its experimental design were conceived by Yong Q. Chen and Janel Suburu. Data

collection was performed by Janel Suburu, Lihong Shi, Jiansheng Wu, Shihua Wang, and

Michael Samuel. Data analysis was performed by Janel Suburu, Nancy D. Kock, Guangyu Yang,

Steven Kridel, and Yong Q. Chen. The manuscript was prepared by Janel Suburu and edited by

Michael J. Thomas, Nancy K. Kock, Steven Kridel, and Yong Q. Chen.

44

II.A ABSTRACT

The mammary gland is one of few adult tissues that strongly induce de novo fatty acid

synthesis upon physiological stimulation, suggesting that fatty acid is important for milk

production during lactation. The committed enzyme to perform this function is fatty acid

synthase (FASN). To determine whether de novo fatty acid synthesis is obligatory or dietary fat

is sufficient for mammary gland development and function during lactation, Fasn was

specifically knocked out in mouse mammary epithelial cells. We found that deletion of Fasn

hindered the development and induced premature involution of the lactating mammary gland,

significantly decreased short- and medium-chain fatty acids and total fatty acid contents in the

milk. Consequently, pups nursing from Fasn knockout mothers experienced growth retardation

and pre-weanling death, which was rescued by cross-fostering pups to a lactating wild type

mother. These results demonstrate that FASN is essential for the development, functional

competence, and maintenance of the lactating mammary gland.

45

II.B INTRODUCTION

Fatty acids are necessary components of life. They provide structural integrity in the form

of biological membranes, and act as signaling mediators to support growth, development,

homeostasis, and inflammatory responses. They modify protein localization and activation, and

they act as an energy source. In mammals, fatty acids are acquired from the diet and/or produced

de novo via the fatty acid synthesis pathway.

Fatty acid synthesis is the process of producing de novo fatty acids from carbohydrate

and amino acid-derived carbon sources. Fatty acid synthase (FASN) performs the majority of

enzymatic steps of fatty acid synthesis [1]. It is a multifunctional, polypeptide enzyme that

produces saturated fatty acids. FASN uses one acetyl-CoA and sequentially adds 7 malonyl-CoA

molecules to produce the 16-carbon saturated fatty acid, palmitic acid (16:0). Palmitic acid is the

primary product of FASN, accounting for approximately 90% of the enzyme’s product yield;

however, FASN also produces myristic acid (14:0) and stearic acid (18:0) [2].

In most tissues, fatty acid synthesis is relatively quiescent, and the reason for its

quiescence remains unclear. It is possible that non-proliferating tissues do not demand fatty acid

synthesis, or that dietary fatty acids are sufficient to fulfill the physiological fatty acid

requirement of these tissues. Nonetheless, expression of FASN is observed in a variety of adult

tissues, with the strongest expression in the liver, adipose, lung, and brain, where it is a critical

enzyme for metabolic homeostasis [2, 3]. Whole body knockout of FASN in mice is

embryonically lethal due to impaired blastocyst implantation in the uterus. Even heterozygous

FASN knockout mice exhibit impaired growth and survival, suggesting fatty acid synthesis is

critical for normal development [4]. Tissue-specific knockout of FASN has provided valuable

insight to the role of de novo fatty acid synthesis in the liver, pancreas, brain, macrophage, heart,

46

intestine, and adipose. In these cell types, FASN has been shown to be a critical regulator of

systemic energy homeostasis, atherosclerotic lesions, diet-induced obesity, diabetes,

inflammation, and cardiac stress [5-10]. Overexpression of FASN has also been strongly

associated with many cancer types and is under extensive study as a potential cancer drug target

[11]. Interestingly, although fatty acids may be produced de novo or taken up from the diet, no

study to date has described adequate compensation by dietary fatty acids following FASN

knockout in vivo.

The mammary gland is a unique lipid-metabolizing tissue. Although the mature, resting

mammary gland does not demand fatty acid synthesis, expression of lipogenic genes, and

specifically FASN, is strongly induced upon the morphological changes associated with

pregnancy and lactation [12, 13]. During this time, the mammary gland undergoes four phases of

functional differentiation: early proliferation (early pregnancy), secretory differentiation (mid

pregnancy), secretory activation (parturition), and lactation [14]. During lactation, mice are

estimated to produce a total of about 5 ml of milk per day, of which approximately 30% is

triacylglycerides [15].

Fatty acid synthesis in the mammary gland is quite unique relative to other tissues.

Comparative analysis of fatty acids in the liver and mammary gland shows a shorter average

fatty acid chain length in the mammary gland [16]. Supporting this finding was the identification

and purification of a mammary gland-specific thioesterase II enzyme, which is responsible for

the early cleavage of de novo fatty acids from FASN and, thus, generation of short chain fatty

acids (<16 carbons) [17]. In accordance with its unique lipid synthesizing capabilities, the

mammary gland has been hypothesized to have a distinct program that regulates lipid

metabolism for the preparation and execution of lactation by the concerted efforts of epithelial

47

cells and adipocytes [12, 14, 18, 19]. Gene expression studies revealed that activation of fatty

acid synthesis in the mammary epithelium occurs in response to parturition and the onset of

lactation. This increased expression of lipogenic genes is significantly larger than their relative

expression in the liver, supporting the hypothesis that the mammary gland is undergoing changes

to fulfill a very specific function [13, 20].

De novo fatty acid synthesis in the mammary gland is responsible for producing short-

and medium-chain fatty acids for milk production [21] and provides approximately 15-40% of

the total fatty acid content of milk [16]. Sterol regulatory element binding protein 1 (SREBP-1)

is a well-known transcriptional activator of lipogenic genes [22] and its activation in the

mammary gland is important for the induction of fatty acid synthesis during lactation [13]. In

vivo deletion of SCAP, the activator of SREBP proteins, in the mammary gland significantly

decreases fatty acid synthesis and the growth rate of nursing pups, and feeding mothers with a

high fat diet only partially rescues this effect [23]. Gene expression analysis suggests de novo

lipogenesis in the mammary gland can be modulated by a high fat diet, whereby increased

dietary fat consumption results in a marked decrease in enzymes of the fatty acid synthesis

pathway and fewer short chain fatty acids in milk [13, 16, 23]. Lipid metabolism is clearly an

integral component of mammary gland physiology; however, no study has investigated the

absolute requirement of fatty acid synthesis during mammary gland development and lactation.

We sought to determine the role of FASN in the mammary gland by generating a conditional

mammary gland-specific Fasn knockout mouse (KO) from Fasn floxed mice [5], in which

deletion of Fasn is made specific to the mammary epithelial cells through the activation of a Cre

recombinase gene regulated by the whey acid protein (WAP) promoter [24]. We report that loss

of FASN in mammary epithelial cells decreases the growth and survival of nursing pups, alters

48

the fatty acid profile of milk produced by lactating mothers, and hinders the development and

maintenance of a functional lactating mammary gland.

II.C EXPERIMENTAL PROCEDURES

Transgenic mice

Mammary epithelial cell-specific Fasn knockout mice were generated by crossing floxed

Fasn mice (FasnL/L

, kindly provided by the laboratory of Dr. Clay Semenkovich, Washington

University) [5] with whey acidic protein-Cre recombinase transgenic mice (WAP-CreT) (The

Jackson Laboratory). At 6 weeks, females of either FasnL/L

;WAP-Cre-

(WT) or FasnL/L

;WAP-

CreT

(KO) genotype were bred with FasnL/L

;WAP-Cre- (WT) males. Mothers were subjected to 1,

2, or 3 pregnancies. Following each pregnancy and lactation, mothers were either euthanized and

mammary glands harvested, or separated for at least 14 days before being mated again with a

FasnL/L

;WAP-Cre- (WT) male. Mammary glands were harvested at either lactation day 25 (L25)

or the day at which all pups had deceased. For FasnL/L

;WAP-CreT

(KO) mothers whose

mammary glands were harvested prior to L25, an age-matched FasnL/L

;WAP-Cre- (WT) mouse

was euthanized at the same lactation day and pregnancy number. In cross-fostering experiments,

only mothers that gave birth within 2 days of each other were eligible for growth monitoring

after cross-fostering. Histopathological evaluation of all tissues was performed by board-certified

veterinary pathologists.

All animals were maintained in an isolated environment in barrier cages and fed a

standard chow diet. Animal care was conducted in compliance with the state and federal Animal

Welfare Acts and the standards and policies of the Department of Health and Human Services.

49

The protocol was approved by the Wake Forest University Institutional Animal Care and Use

Committee.

Measurement of pup growth

The date of parturition was considered lactation day 1 (L1). Pups from each pregnancy

were weighed daily between 22 and 26 hours from previous weigh-in from L2 to L25. Litters of

more than 8 pups were culled to 8 pups on L2 to prevent competition among pups for the

mother’s milk supply. To minimize error, up to 6 pups were weighed at once, followed by

sequential measurements of one additional pup, up to 8 pups. For litters of more than six pups,

the 7th

and 8th

pups were distinguished by a tail marking on L8 and were consistently weighed as

the 7th

and 8th

pups for daily measurements. Each measurement was divided by the number of

pups weighed to determine the average pup weight of each litter. The average pup weight for all

litters monitored was then determined.

Milk collection

Mothers were separated from pups for 45 minutes to 2 hours on L2, L10, and L18.

Mothers were anesthetized by intraperitoneal (IP) injection of 6.25 mg per 30 g body weight of

Avertin (Sigma T48402). A 40X solution of Avertin was made by dissolving 500 mg/ml in tert-

amyl alcohol and stored at 4°C. A 1X solution was made with PBS from the 40X solution.

Following anesthesia, mice were injected IP with 200 l of 10 IU/ml (2 IU/mouse) oxytocin

(Sigma O4375) to induce milk release from the mammary gland. Nipples were gently massaged

and secreted milk was collected in a capillary tube. Approximately 10 μl of milk was extracted

from each gland (~100 μl total), pooled together into one microcentrifuge tube, snap frozen in

50

liquid nitrogen, and stored at -80°C until analyzed. All drug solutions were filtered through a

0.22 μm filter and stored at 4°C.

Histochemistry

The right inguinal mammary gland was excised and spread between two microscope

slides, fixed overnight in 10% buffered formalin and processed for embedding in paraffin.

Routine processing and staining was done with hematoxylin and eosin (H&E).

Immunohistochemical (IHC) staining was performed with mouse anti-FASN primary

antibody (BD Bioscience 610962) and guinea pig anti-adipophilin primary antibody (20R-

AP002, Fitzgerald Industries, Flanders, NJ). Primary antibody was incubated at 4°C overnight in

a humidity chamber, followed by HRP-conjugated goat anti-mouse (NA931V, GE Healthcare) or

donkey anti-guinea pig (706-165-148, Jackson Immunoresearch, West Grove, PA) secondary

antibody. DAB chromogen was applied for 3-8 minutes according to the package insert (K3467,

Dako, Carpinteria, CA). TUNEL staining was performed using a DeadEnd Colorimetric Assay

kit (G7360, Promega) according to the manufacturer’s instructions. A positive TUNEL control

included a DNase I digestion that was processed separately from all other samples. A negative

TUNEL control was processed in the absence of any rTdT enzyme.

Immunofluorescent (IF) staining was performed with mouse anti-FASN primary antibody

(BD Bioscience 610962) and guinea pig anti-adipophilin primary antibody (20R-AP002,

Fitzgerald Industries, Flanders, NJ) followed by Alexa Fluor 488 goat anti-mouse (Life

Technologies A-11001) and Alexa Fluor 594 goat anti-guinea pig (Life Technologies A-11076)

secondary antibody. Following this step, DAPI (4′,6-diamidino-2-phenylindole) (Sigma D9542)

51

nuclear stain was incubated for 5 min, followed by mounting with ProLong Gold Antifade

Reagent (Life Technologies P10144).

Mammary gland whole mount

The left inguinal mammary gland was excised and spread between two microscope slides

and fixed overnight in Carnoy’s fixative (60% 200 proof ethanol, 30% chloroform, 10% glacial

acetic acid). Following fixation, mammary glands were subjected to a series of ethanol washes to

rehydrate: twice with 70% ethanol for 15 minutes, twice with 50% ethanol for 10 minutes, twice

with 30% ethanol for 10 minutes, twice with 10% ethanol for 10 minutes, twice with dH2O for 5

minutes. After rehydration, mammary glands were stained overnight in carmine aluminum

staining solution followed by a series of ethanol washes to dehydrate: twice in 70% ethanol for

15 minutes, twice in 95% ethanol for 15 minutes, twice in 100% ethanol for 15 minutes.

Mammary glands were placed in xylene until all fat was cleared and mounted to a microscope

slide using Cytoseal-60 mounting medium. Carmine aluminum staining solution was made by

placing 1 g carmine (Sigma C1022) and 2.5 g aluminum potassium sulfate (Sigma A7167) in 500

ml dH2O and boiling for 20 minutes. Final volume was adjusted to 500 ml with H2O. A thymol

crystal was added as a preservative. Carmine staining solution was stored at 4°C.

Microscopy

All images were captured with either a Hamamatsu Nanozoomer 2.0HT with Olympus

objectives and NDP image software or an Olympus VS110 microscope equipped with Olympus

objectives and VS-ASW FL image software. Image frames were created using Olympus Olyvia

software. Average lumen size was quantified by measuring the largest diameter of 8-10 alveolar

52

lumens at 40X magnification in 4-5 glands of 1-2 mice for each genotype and lactation day.

TUNEL stains were quantified by averaging the number of TUNEL-positive cells per sixteen

random fields of view at 20X magnification for each gland and lactation day.

FAME analysis

Fatty acid methyl ester analysis was performed on milk collected from lactating mothers

as previously described [25]. Results from each milk collection were averaged for each

pregnancy.

Statistics

Statistics were performed using Microsoft Excel, Daniel’s XL Toolbox Plugin 5.05, and

MedCalc Software Version 12.7. A two-tailed student T-test was performed to identify statistical

significance between two groups at a given point of growth curves. A log-rank test was used to

compare statistical significance between Kaplan-Meier survival curves.

II.D RESULTS

Generation of mammary gland-specific Fasn knockout mice

Mice with mammary epithelial cell-specific Fasn knockout (KO) were developed by

crossing mice homozygous for the floxed Fasn alleles [5] with transgenic mice bearing a Cre

recombinase transgene controlled by the WAP promoter (Jackson Laboratory). Knockout of

Fasn in this model occurs only in the mammary epithelial cells, not in mammary adipose tissue,

and only upon pregnancy and lactation. To determine if multiple pregnancies increased the effect

of Fasn KO, female mice were subjected to one, two, or three pregnancies. By lactation day 15

53

of the first pregnancy (L15P1), FASN staining was observed in a large number of epithelial cells

during lactation, suggesting incomplete knockout. However, staining was largely absent by

lactation day 17 of the second pregnancy (L17P2) and decreased further by lactation day 16 of

the third pregnancy (L16P3). In contrast, very strong FASN staining was observed in wild type

(WT) mice during all three pregnancies. Virgin mice also showed positive FASN staining (Fig.

1A).

Despite an obviously gradual deletion of Fasn over the course of multiple pregnancies,

time dependent deletion was also observed over the course of the lactation period. On lactation

day 2 of pregnancy 3 (L2P3) approximately half of the epithelial cells stained strongly positive

for FASN. This number decreased to less than 25% by lactation day 10 (L10P3), and FASN was

nearly completely absent in all epithelial cells by lactation day 16 (L16P3) (Fig. 1B).

Fasn knockout in the lactating mammary gland decreases pup growth and survival

Because FASN is responsible for producing short- and medium-chain fatty acids in milk

[16] and fatty acids are important nutritional components of milk for the mother’s growing pups,

we sought to determine how FASN KO affects the growth of pups nursed by KO mothers for

each pregnancy. No significant differences in the average litter size or the mother’s age at

parturition were observed between WT and KO mothers for all three pregnancies (Table 1). In

the first pregnancy all pups showed similar growth rates regardless of the mother’s genotype

(Fig. 2A). Interestingly, although the growth curves were similar, we observed a significantly

greater pre-weanling death rate for pups from KO mothers compared to pups nursed by WT

mothers. While 75% of pups from WT mothers survived to weaning age (lactation day 25), only

61% of pups from KO mothers survived to weaning age during the first pregnancy (P=0.01)

54

(Fig. 2B). In the second pregnancy, there was a significant decrease in the growth rate of pups

from KO mothers compared to pups from WT mothers (Fig. 2C). No pups from KO mothers

survived beyond lactation day 19, while 75% of pups from WT mothers survived to weaning age

(P<0.0001) (Fig. 2D). In the third pregnancy, pups from KO mothers displayed a similar trend in

the growth of pups compared to the second pregnancy (Fig. 2E). Also, no pups from the third

pregnancy of KO mothers survived beyond lactation day 19, while all pups from WT mothers

survived to weaning age (P<0.0001) (Fig. 2F). In all, the growth and survival trends of pups

from KO mothers over the course of three pregnancies correlates with the progressive deletion of

FASN.

It has been previously reported that both homozygous and heterozygous KO of FASN

leads to developmental complications [4]. To further demonstrate the specificity of the FASN

KO phenotype, pups from the second pregnancy of KO mothers were genotyped to determine

whether growth complications may have possibly been due to unexpected expression of the

WAP-Cre recombinase and premature KO of FASN. As expected, pups demonstrated 50%

Mendelian inheritance of the Cre recombinase transgene (data not shown), providing evidence

that the premature death of KO pups was not a result of erroneous expression of WAP-Cre in

pups.

Deletion of FASN induces premature involution and cell death

Litters from KO mothers typically perished all together in a time frame of 2-3 days (Fig.

3) and mammary glands of KO mothers were immediately harvested following the death of all

their pups. We harvested mammary glands from lactating, age-matched, WT mothers from the

same pregnancy and on the same lactation day as KO mothers to analyze any changes that had

55

occurred in KO mammary glands during the lactation period. Although the difference in

pregnancy 1 appears less dramatic, whole mount mammary gland analysis showed a consistent

and marked difference in the density of glands in KO mothers compared to WT mothers for all

three pregnancies. All age-matched virgin mice showed normal tree-like branching (Fig. 4A).

Closer examination with hematoxylin and eosin (H&E) staining further demonstrated a marked

difference in the alveolar density between KO and WT mammary glands. Adequate formation of

actively secreting alveoli was evident in KO mice on lactation day 15 from the first pregnancy

(L15P1). However, compared to WT mice, KO mice had fewer alveoli and showed a larger

proportion of adipocytes.

In contrast to the first pregnancy, mammary glands from KO mice in the second and third

pregnancies appeared to be undergoing involution at day 16-17, whereas age-matched, WT mice

of the same lactation day and pregnancy displayed large and actively secreting alveoli (Fig. 4B).

To verify that KO glands were experiencing early involution, we compared WT and KO

mammary glands at various time points during each lactation period and performed a TUNEL

assay to analyze cell death. As expected, glands from both WT and KO mice had experienced

involution by lactation day 25 after the first pregnancy. KO glands, in accordance with having

fewer alveoli at lactation day 15, also showed fewer epithelial cells at lactation day 25, compared

to WT mice (Fig. 5A). In the second pregnancy, WT glands had developed mature alveoli and

were abundantly secreting at lactation days 12 and 17, whereas KO glands showed a decreased

number of alveoli at lactation day 12 and showed histological signs of involution at day 17 (Fig.

5B). In the third pregnancy, we analyzed glands at lactation days 2, 10, 16, and 19 to gain a more

comprehensive understanding of the histological progression during the lactation phase in KO

mice. At lactation day 2, both WT and KO mice showed comparable development of actively

56

secreting alveoli. However, by lactation day 10, a discrepancy in the number and size of alveoli

is apparent, and by day 16 and 19, glands are clearly undergoing involution (Fig. 5C), as they

appear very similar to glands of lactation day 25 from the first pregnancy. TUNEL staining

showed no difference in cell death at lactation day 15 of the first pregnancy, (Fig. 5D), nor

lactation days 2 and 10 of the third pregnancy (Fig. 5F). However, in accordance with their

marked histological changes, there was a statistically significant increase in TUNEL positive

cells at lactation day 17 of pregnancy 2 and lactation day 16 of pregnancy 3, validating cell death

and involution at these time points (Fig. 5D-5G). A positive and negative TUNEL control using

a DNase I digestion and no rTdT enzyme, respectively, confirmed the results of the TUNEL

staining (Fig. 5H).

Fasn knockout hinders mammary gland maturation, but not secretory activation during lactation

As we previously mentioned, the deletion of FASN gradually increased over the course

of lactation and multiple pregnancies. Therefore, we questioned whether KO glands were strictly

involuting early, as a consequence of increased FASN deletion, or if they also experienced

developmental deficits during lactation. Glands experiencing involution, either WT or KO, had a

much smaller lumen size than actively lactating glands. However, analysis of lumen

measurements at earlier time points during lactation demonstrated a statistically significant

difference (P<0.001) between WT and KO glands at lactation days 12 and 10, but not at lactation

day 2 (Fig. 6A). Interestingly, a comparison of FASN positive versus FASN negative lumens

showed no difference in lumen size at either lactation day 2 or 10 in KO glands with mosaic

deletion of FASN (Fig. 6B). Despite the differences observed in lumen size, KO glands still

showed ample formation of lipid droplets on the luminal side of epithelial cells at all stages of

57

lactation, as shown by staining for adipophilin (Fig. 6C-E). This was true even in lumens where

FASN had been deleted, as shown by immunofluorescent double staining for adipophilin and

FASN on lactation day 10 of the third pregnancy (Fig. 6F).

Fasn knockout in the lactating mammary gland alters the lipid profile in milk

FASN is responsible for the production of short-chain fatty acids (<16 carbons), as well

as a substantial portion of medium-chain fatty acids (16-18 carbons), while dietary sources are

the major supply for long-chain fatty acids (>18 carbons) [26]. Because we identified a deficit in

the growth and survival of pups nursed by KO mothers, we hypothesized that FASN KO in the

mother’s mammary gland was affecting the fatty acid profile of the milk, and thereby, pups were

succumbing to premature death, in part, by malnutrition. To test this hypothesis, we performed

fatty acid methyl ester (FAME) analysis on milk collected from lactating mothers on lactation

days 2, 10, and 18 during each pregnancy and quantified the presence of each fatty acid.

Milk analysis from the first pregnancy showed significant decreases in 14:0 (P<0.001),

14:1 (P=0.03), 16:0 (P<0.01), 18:0 (P<0.05), 18:2 (P<0.01), and total fatty acid (P<0.01) in milk

from KO mothers compared to WT (Fig. 7A). Milk analysis from the second pregnancy showed

significant decreases in 14:0 (P<0.001), 14:1 (P<0.001), 16:0 (P<0.001), and 22:0 (P=0.03), and

a similarly decreasing trend in the total fatty acid (P=0.09) (Fig. 7B). Finally, milk analysis from

the third pregnancy showed significant decreases in 14:0 (P<0.001), 16:0 (P<0.001), 18:0

(P<0.001), 20:0 (P=0.02), and total fatty acid (P=0.03) (Fig. 7C).

58

Growth and survival deficiencies of pups are rescued by nursing from a wild-type mother

To clearly demonstrate that the functional differences in lactation from impaired

mammary gland development and the consequential growth and survival trends in pups were, in

fact, due to the Fasn KO in the mammary gland of lactating mothers, we swapped the litters of

age-matched KO and WT mothers between lactation day 1 and lactation day 3 and monitored the

growth of pups being nursed by the surrogate mother. Again, there were no significant

differences in litter size or the age of all mothers at parturition between KO and WT mothers for

all three pregnancies (Table 2). Similar to our previous results (Fig. 2), pups that were born to a

WT mother, but nursed by a KO mother, showed a significant decrease in growth and survival.

When mothers were swapped after their first pregnancy, we observed a significant decrease in

the growth and survival (P=0.0003) of pups born to a WT mother and nursed by a KO mother

(Fig. 8A). During the second and third pregnancies, WT pups nursed by a KO mother showed

significantly decreased growth and survival (P<0.0001) (Fig. 8 B,C). Importantly, pups born to a

KO mother and nursed by a WT mother were phenotypically rescued, showing significantly

greater growth and survival (Fig. 8).

II.E DISCUSSION

Fatty acid synthesis is an important aspect of mammary gland function during lactation.

Previous reports have described the importance of lipogenesis for mammary gland milk secretion

and its modulation by dietary fat [14, 16]. However, no study has investigated the requirement of

FASN in the mammary gland. Our study demonstrates that FASN is essential for functional

development and maintenance of the lactating mammary gland. We show that FASN KO mice

develop fewer alveoli, have impaired alveolar development during lactation, undergo premature

59

mammary gland involution, and have an altered milk fatty acid profile. These perturbations

ultimately manifest in significant growth retardation and premature death of nursing offspring.

Deletion of FASN in our model was spatially restricted to mammary epithelial cells and

temporally restricted until late pregnancy and lactation [24]. The mammary gland undergoes

significant apoptosis during involution [27]; however, cells that survive the involution phase may

act as alveolar progenitors that repopulate the epithelium in subsequent pregnancies [28]. Our

data from the second and third pregnancies are very similar, suggesting two pregnancies were

likely sufficient for a phenotypic outcome in our KO mice. Other studies using the WAP-Cre

mouse model to knockout a gene of interest have also reported using multiple pregnancies to

achieve complete knockout and a phenotype [29-31]. The fact that expression of WAP-Cre

begins in late pregnancy, increases sharply following parturition, and is maintained during

lactation [24] likely explains the progressive loss of FASN. While most, but not all, L10 KO

mammary gland samples during the third pregnancy showed a large decrease in FASN staining,

there was even more variability in the extent of FASN staining among L2 KO samples from the

third pregnancy. This suggests activation of Cre was not uniform in all mammary glands

between different KO mice and possibly within the glands of each KO mouse, and instead, was

likely dependent on the progression of lactation. Such variation explains the steady separation

between the growth curves of KO and WT mice as glands progressively lost FASN. Moreover,

the tendency for litters to perish as a whole suggests that only after the majority of glands were

devoid of FASN was the lactating mother unable to support her nursing pups. The fact that

survival of KO pups steadily decreased from lactation day 11 to 17, a rather large, 6-day window

when the majority of pups perished, further demonstrates the varying rates of Fasn deletion

between lactating mice. Concordantly, some mammary glands of KO mothers did not produce

60

sufficient quantities of milk for fatty acid analysis on lactation day 18, which was most likely a

result of premature involution of that gland from Fasn deletion. Finally, because deletion of Fasn

occurs as early as late pregnancy, pups born to WT mothers could possibly have had an early

growth advantage over pups born to KO mothers. This could explain the significant difference in

average pup weight as early as lactation day 2 in pups born to and nursed by a KO mother. It

may also explain the time delay to observe the same growth and survival effects during the third

pregnancy of the cross-fostering experiments as those observed in the second and third

pregnancies of pups born to and nursed by KO mothers. It is important to note that the mothers

monitored for the growth and survival of their pups in figure 2 and the mothers of the cross-

fostering experiments of figure 8 were two different cohorts of mice. Therefore, some of the

variation observed between figures 2 and 8 may also be a result of using different subjects.

The primary function of fatty acid synthesis in the lactating mammary gland is believed

to produce milk fatty acids; however, FASN has also been implicated as a critical regulator of

growth and survival [4]. Despite a clear and often widespread depletion of FASN from

mammary epithelial cells as early as lactation day 2, KO mice still developed tree-like ducts,

acini, and alveoli with grape like clusters. Even though some KO glands developed fewer alveoli

structures, these results suggest that FASN is likely not essential for the overarching structural

development of the gland. Moreover, because adipophilin staining was widespread and appeared

at the luminal side of epithelial cells, even when they are FASN-depleted, FASN is likely not

required for secretory activation. However, there was an apparent deficit in the glandular

maturation of KO mice during lactation, as evidenced by their overall smaller lumen size and

failure to develop alveoli comparable in size to WT mice. This discrepancy was observed in KO

mice regardless of incomplete deletion of FASN. Hence, it is possible that the deletion of FASN

61

affected the development of the lactating gland on a global scale or in a localized manner. The

inhibition of FASN is known to decrease ErbB2 activation and its downstream signaling

mediators PI3K/AKT/mTOR, which are major regulators of cellular growth, proliferation, and

survival [32, 33]. Indeed, transgenic expression of dominant-negative ErbB2 in mice impairs

lobuloalveolar development at parturition, but does not affect milk secretion [34]. These findings

are in agreement with our results that suggest the deletion of FASN may modulate lobuloalveolar

development but does not affect secretory activation.

Despite minor developmental deficits, KO mice were capable of developing lactating

glands. However, functional incompetence of FASN KO mammary glands was evidenced by the

significant decrease in fatty acids and the stunted growth and premature death of pups nursed by

KO mothers. Glands from KO mice during late lactation (L16-17) strongly resembled glands

from WT mice undergoing involution (L25) and had a significant increase in cell death,

suggesting KO mice were experiencing premature onset of involution. Involution typically

occurs in response to the loss of mechanical stimulation from cessation of breast-feeding and

milk stasis [35]. Thus, it is possible that the early involution observed in KO mice was a result of

lost mechanical stimulation from pup weakness, malnutrition, and death. However, KO mothers

showed signs of scabbing and inflammation at the nipples, suggesting pups had not ceased to

nurse, and some glands did not produce a sufficient quantity of milk for collection on lactation

day 18, suggesting glands had already involuted despite the presence of pups. Moreover, the first

signs of adipocyte repopulation following forced involution begin 2 days following the removal

of all pups [12], but mammary glands from KO mothers were harvested immediately following

the death of all their pups and pre-weanling death typically occurred as a whole litter rather than

sporadic individual deaths. Therefore, it is very likely that early activation of involution in KO

62

mice was not a mechanical response, but rather, occurred as a result of the loss in functionality of

the lactating mammary gland. These results suggest that a lack of functional competence from

the loss of FASN can trigger involution and apoptotic cell death in the lactating mammary gland.

Nonetheless, it remains elusive what biological sensors may be triggered in response to the lost

functionality of the gland to elicit involution, and whether these sensors are similar to those

during normal induction of involution.

As evidenced by the change in phenotype from cross-fostering mothers, alterations in the

mammary gland milk were a major contributor to the growth and survival discrepancies

observed during lactation of KO mice. It is very likely that the phenotypic outcomes were a

result of insufficient milk volume due to dysfunctional lipogenesis; however, changes in the milk

fatty acid profile may also have contributed to the impaired growth and survival of pups nursed

by KO mothers. Literature highlights the importance of long chain polyunsaturated fatty acids as

extremely critical for visual and neuronal development [36], but the nutritional significance of

FASN-derived lipids, such as 14:0, 16:0, and 18:0 fatty acids, during development remains

unclear. Our data suggests that perhaps de novo synthesized fatty acids are critical for

development as well. Previous reports have shown unique roles for de novo synthesized fatty

acids in the liver and other tissues including post-translational modification, PPAR activation,

and their incorporation into membrane and signaling phospholipids [5, 6, 9, 37, 38]. Whether

FASN-derived lipids in mammary gland milk fulfill these or similar functions remains unknown.

Few tissues in the body engage de novo fatty acid synthesis during normal physiology

[39]. Nonetheless, a variety of tissue types carry the potential to initiate a lipogenic phenotype

[11]. The intersection between de novo and dietary fatty acids is a growing research interest, as

the activation of de novo fatty acid synthesis has been implicated in various cell types linked to

63

multiple pathophysiologies, including obesity, inflammation, atherosclerosis, and cancer. The

literature suggests a very distinct role for de novo fatty acids in these diseases, for which neither

dietary fatty acids nor preformed fatty acids from adipose can compensate in vivo [5-8, 11].

Functional mammary gland development has been shown to be dependent on interactions

between epithelial cells and adipocytes [18, 19], suggesting the mammary gland is a unique

model system to study the integration of local de novo fatty acid synthesis and other sources of

preformed fatty acids. Large volume production of short chain fatty acids is exclusive to the

mammary gland, as only the mammary tissue expresses the unique thioesterase II enzyme that

prematurely cleaves the growing fatty acyl chain from the FASN enzyme [17]. Previous reports

have shown that short-chain milk fatty acids are exclusively produced de novo, while long-chain

fatty acids are derived from preformed fatty acids, and medium-chain fatty acids are derived

from both [21, 23]. In our model, FASN was specifically deleted in mammary epithelial cells,

but remained intact in mammary adipocytes. Additionally, the fatty acid profile of milk showed a

significant decrease in 16:0 and 18:0 fatty acids and a near depletion of 14:0 fatty acids. These

results suggest that the portion of milk fatty acids produced by de novo fatty acid synthesis in the

mammary epithelial cells was not replaced following the deletion of FASN. It seems that

ultimately, other sources of fatty acid were not able to compensate for this loss, including

standard dietary fatty acid consumption and de novo fatty acid synthesis in adjacent adipocytes

or other tissues such as liver and adipose. Hence, our data further support the notion that de novo

fatty acid synthesis is a unique source of fatty acids and that upregulation of the fatty acid

synthesis pathway characterizes a specialized cellular function that cannot be substituted by

increased uptake of extracellular fatty acids. Nonetheless, it would be interesting to determine

64

whether feeding KO mothers a diet high in saturated fat could rescue the phenotypic outcome of

nursing pups.

Overall, our study highlights FASN as an essential component of mammary gland

physiology during lactation. However, several new questions arise from the findings of this

research that may guide future studies: 1) Why do alveoli with or without FASN in KO mice

seem to experience the same degree of development and secretory activation? 2) At what point

following the deletion of FASN does the mammary gland begin involution and what biological

sensors are regulating this process? 3) Can a high-fat diet rescue the phenotype observed in our

study? An in-depth analysis focusing on the first 10-15 days of the second or third lactation

period will be pertinent to answering these questions.

65

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28. Wagner, K.U., et al., An adjunct mammary epithelial cell population in parous females:

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36. Lauritzen, L., et al., The essentiality of long chain n-3 fatty acids in relation to

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for PPARalpha in liver. Cell, 2009. 138(3): p. 476-88.

38. Swinnen, J.V., et al., Fatty acid synthase drives the synthesis of phospholipids

partitioning into detergent-resistant membrane microdomains. Biochem Biophys Res

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39. Wakil, S.J., J.K. Stoops, and V.C. Joshi, Fatty acid synthesis and its regulation. Annu

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68

II.G ACKNOWLEDGMENTS

We thank Dr. Clay Semenkovich (Washington University School of Medicine, St. Louis,

MO) for generously providing floxed Fasn mice. We thank Brandi Bickford of the Wake Forest

University Virtual Microcopy Core and Crystal Conaway for their technical assistance. We thank

Isabelle Berquin for her assistance editing the manuscript. This study was funded in part by the

National Institutes of Health (R01CA107668, P01CA106742, R01CA163273, T32CA079448

training grant) and The Synergistic Innovation Center for Food Safety and Nutrition, Jiangnan

University.

69

II.H FIGURES AND FIGURE LEGENDS

Figure 1. Epithelial cell-specific deletion of Fasn in the mammary gland.

Right inguinal mammary glands were harvested, sectioned, and stained by IHC for fatty acid

synthase (FASN). A. FASN knockout mothers (KO) showed only partial deletion of FASN

following the first pregnancy, and further deletion in late lactation of the second and third

pregnancies. Age-matched virgin glands showed very strong FASN staining in adipocytes and

positive staining in the epithelium. Wild type mothers (WT) showed strongly positive staining

during all three pregnancies. B. Deletion of FASN was heterogeneous and occurred gradually

over the course of lactation. FASN-positive epithelial cells were present on lactation day 2 and

10 of the third pregnancy. By lactation day 16, nearly all epithelial cells were negative for

FASN, and only adipocytes showed positive staining. All images are shown at 20X resolution.

LXPZ, Lactation day X of pregnancy Z. WT, wild type. KO, Fasn knockout. Virgin glands are

age-matched.

70

71

Figure 2. Deletion of FASN hinders growth and survival of nursing pups.

Average weight of pups and their survival were monitored from lactation day 2 through lactation

day 25 following the first (A), second (B), and third (C) pregnancy. Error bars represent s.e.m.

*P<0.05, **P<0.01, ***P<0.001. P-values for survival curves are indicated in the panel.

72

Figure 3. Pre-weanling death occurs as a whole litter.

Survival of pups was monitored for each litter following the second (A) and third (B)

pregnancies of KO mothers. Each line represents the survival of pups in a single litter from a KO

mother. Typically, litters perished over the course of 2-3 days.

73

Figure 4. KO mammary glands are sparse in alveolar structures.

A. Left inguinal mammary glands were harvested and whole mounted at the indicated lactation

day of each pregnancy. B. Right inguinal mammary glands were harvested and stained with

H&E at the indicated lactation day of each pregnancy. LXPZ, Lactation day X of pregnancy Z.

WT, wild type. KO, Fasn knockout. Virgin glands are age-matched.

74

Figure 5. FASN deletion induces early involution and cell death.

Right inguinal mammary glands were harvested, sectioned, and stained for histology with H&E

(A-C) or for cell death via TUNEL assay (D-F). Red arrows point to TUNEL-positive cells. G.

TUNEL stains were quantified by counting the number of TUNEL-positive cells per mm2 of

tissue. H. A DNase I digestion was used as a positive TUNEL control, and no rTdT enzyme was

used for a negative control. LXPZ, Lactation day X of pregnancy Z. WT, wild type. KO, Fasn

knockout. Error bars represent s.e.m. ***P<0.001.

75

76

Figure 6. Deletion of FASN hinders alveolar development, but not secretory activation

during lactation.

A. Luminal measurements from WT and KO mice were averaged and compared for the

respective lactation days of each pregnancy. B. Measurements of FASN-positive lumens and

FASN-negative lumens in KO mice were averaged and compared for lactation days 2 and 10 of

the third pregnancy. C-E. Mammary gland sections from the first, second, and third pregnancies

were stained for adipophilin as a marker of secretory activation. F. Mammary glands from

lactation day 10 of the third pregnancy were stained for adipophilin (red), FASN (green), and

DAPI (blue). LXPZ, Lactation day X of pregnancy Z. WT, wild type. KO, Fasn knockout. Error

bars represent s.e.m. ***P<0.001.

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78

Figure 7. FASN deletion changes the fatty acid profile of mammary gland milk.

Milk was collected during the lactation period of each pregnancy. Milk was pooled from all 10

glands. Graphs represent fatty acid methyl ester analysis of the first (A), second (B), and third

(C) pregnancy. All fatty acid values were normalized to total protein content. Error bars

represent s.e.m. *P<0.05, **P<0.01, ***P<0.001.

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Figure 8. Phenotypic rescue of offspring by cross-fostering mothers at birth.

Litters from knockout mothers were swapped with litters from wild-type mothers between

lactation day 1 and 3 for the first (A), second (B), and third (C) pregnancies. Average pup weight

and survival were monitored every day beginning on the day after swapping mothers. Error bars

represent s.e.m. *P<0.05, **P<0.01, ***P<0.001. P-values for survival curves are indicated in

the panel.

80

II.I TABLES

Table 1. Litter statistics for growth and survival curves Pregnancy 1 Pregnancy 2 Pregnancy 3

WT KO WT KO WT KO

Total Number of Litters 15 22

10 10

5 4

Average Litter Size 6.3 ± 0.27 7.1 ± 0.21

7.7 ± 0.60 7.6 ± 0.45

7.0 ± 0.95 8.5 ± 0.29

Total Number of Pups 95 156

77 76

35 34

Mother Age at Parturition (wks) 10.0 ± 0.30 10.0 ± 0.26 19.0 ± 0.71 19.2 ± 0.99 31.5 ± 1.41 30.0 ± 2.18

Table 2. Litter statistics for swapped litter growth and survival curves Pregnancy 1 Pregnancy 2 Pregnancy 3

WT Swap KO Swap WT Swap KO Swap WT Swap KO Swap

Total Number of Litters 14 11

10 10

6 6

Average Litter Size 7.3 ± 0.38 7.2 ± 0.44

8.7 ± 0.50 7.9 ± 0.43

8.0 ± 0.53 8.0 ± 0.44

Total Number of Pups 102 79

87 83

48 48

Mother Age at Parturition (wks) 10.5 ± 0.36 10.7 ± 0.41 20.9 ± 0.98 21.0 ± .56 31.5 ± 0.92 31.1 ± 0.81

81

CHAPTER III

FATTY ACID SYNTHASE REGULATES PROSTATE CANCER

PROGRESSION

Janel Suburu, Jiansheng Wu, Lihong Shi, Shihua Wang, Brian Fulp, Shadi A. Qasem, Steven

Kridel, and Yong Q. Chen

This study and its experimental design were conceived by Yong Q. Chen and Janel Suburu. Data

collection was performed by Janel Suburu, Jiansheng Wu, Lihong Shi, Shihua Wang, and Brian

Fulp. Data analysis was performed by Janel Suburu, Shadi A. Qasem, Steven Kridel, and Yong

Q. Chen. The manuscript was prepared by Janel Suburu and edited by Yong Q. Chen.

82

III.A ABSTRACT

Cancerous lesions occur more frequently in the prostate than any another site among men

in the United States, affecting approximately 1 in 7 men. One of the most commonly observed

genetic lesions in prostate cancer is the loss of PTEN, which is known to induce growth and

proliferation of cancer cells through the activation of AKT. A consequence of unrestricted

activation of AKT is the development of a lipogenic phenotype and the expression of fatty acid

synthase (FASN). Expression of FASN is an early event in prostate cancer development and its

increased expression is an indicator of progression and prognostic predictor of poor outcome. A

large body of literature suggests the selective inhibition of FASN is an effective approach to treat

prostate cancer, since FASN knockdown in vitro and pharmacological inhibition in xenograft

models suppresses tumor growth and induces apoptosis. We aimed to validate FASN as a cancer

therapy target by genetically deleting Fasn in the Pten knockout mouse model of prostate cancer.

We report that FASN regulates prostate cancer progression in the setting of Pten deletion.

Inhibition of Fasn in Pten knockout mice hindered the development of neoplastic lesions and

significantly reduced the total prostate weight, proliferation, and activation of AKT. This is the

first study to demonstrate tumor-specific inhibition of FASN by genetic ablation in vivo and

validate the development of FASN-inhibiting drugs for the treatment of prostate cancer.

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III.B INTRODUCTION

Prostate cancer is the most commonly occurring type of cancer, and continues to be the

second most deadly cancer type, among men in the United States. The American Cancer Society

estimated that over 230,000 men will develop prostate cancer in the coming 2014 year,

accounting for nearly 28% of all cancer cases in men. Although most prostate cancers are

detected at an early stage, the survival of advanced prostate cancer patients still remains low at

only 27% [1]. Therefore, it is important to elucidate the molecular mechanisms that drive

progression of prostate cancer to promote the development of new targeted therapies.

Numerous metabolic aberrations drive cancer cell proliferation and survival [2]. In

particular, fatty acid metabolism has acquired great interest, as the enzymes of the fatty acid

synthesis pathway, including ATP-citrate lyase (ACLY), acetyl-CoA carboxylase (ACC1), and

fatty acid synthase (FASN), have all been shown to be up-regulated in numerous tumor types,

especially in hormone-sensitive cancers such as breast and prostate cancer [3, 4]. Androgens, a

major stimulator of prostate cancer, increase fatty acid synthesis through activation of the sterol

response element binding protein 1 (SREBP-1) [5, 6]. The expression of FASN in prostate

cancer is an early event in cancer development and its increased expression is a predictor of

progression and poor prognosis [7, 8]. Forced expression of either SREBP-1 or FASN is

sufficient to enhance prostate cancer cell proliferation and tumor growth in mice [9, 10].

Evidence suggests FASN promotes membrane biogenesis and lipid raft formation in cancer cells

[11]. Selective inhibition of either ACLY or FASN, by pharmacological inhibitors or siRNA

knockdown inhibited prostate tumor growth [12, 13]. Moreover, inhibition of ACC1 or FASN

induced cell death of prostate cancer cells, but not normal prostate cells [14, 15]. Collectively,

84

this evidence suggests inhibiting fatty acid synthesis may be an effective approach for the

treatment of prostate cancer.

Activation of the PI3K/AKT pathway, through a variety of mechanisms, occurs in nearly

all human prostate cancers [16]. Specifically, the deletion of PTEN, a negative regulator of

PI3K/AKT signaling, is among the most commonly observed genetic aberrations and has been

reported in approximately 23% of human neoplasia, 69% of localized prostate cancer, and 86%

of metastatic prostate cancers [17, 18]. Increased activation of AKT, by either deletion of PTEN

or a constitutively active AKT, induces expression of FASN [19]. In 2003, the Pten knockout

prostate cancer mouse model was developed and characterized by Wang and colleagues [20], and

has since become one of the most widely used mouse models of prostate cancer [21]. According

to the National Prostate Pathology Committee, the Pten knockout mouse model is among the best

in vivo models to study prostate cancer because it most accurately recapitulates human disease by

developing prostatic intraepithelial neoplasia (PIN) that eventually progresses to adenocarcinoma

[21].

Using the Pten knockout mouse model, we aimed to validate FASN as a prostate cancer

therapeutic target by genetically deleting Fasn in the mouse prostate. Former reports using

pharmacological inhibitors or xenograft models have shown off-target effects of anti-FASN

drugs or have poorly modeled human disease with the absence of a competent immune system

[22, 23]. Therefore, our approach of genetically inhibiting FASN isolates the discrete role of

FASN in prostate cancer development and progression. Our results show that homozygous

deletion of Fasn using a Cre-loxP approach and the widely appreciated probasin-Cre4 transgene

(Pb-Cre4) [24] is unachievable. However, analysis of Pten-null mice with hemizygous deletion

of Fasn revealed that FASN modulates prostate cancer development and progression.

85

Hemizygous deletion of Fasn significantly decreased tumor growth and histopathological

progression in Pten-null mice at 6 months. Loss of FASN also decreased FASN, pAKT473

, and

proliferation in Pten-null tumors.

III.C EXPERIMENTAL PROCEDURES

Transgenic mice with Pb-Cre4

Prostate-specific deletion of FASN was achieved by crossing mice homozygous for the

floxed Fasn gene (FasnL/L

) (kindly provided by Dr. Clay Semenkovich of Washington University

[25]) with Pb-Cre4 transgenic mice homozygous for the floxed Pten gene (PtenL/L

;CreT) [20].

Triple transgenic males of PtenL/+

;FasnL/+

;CreT or Pten

L/L;Fasn

L/+;Cre

T genotype were mated to

females of PtenL/L

;FasnL/+

;Cre- or Pten

L/L;Fasn

L/L;Cre

- genotype. Males were euthanized at 2 and

6 months of age. Histopathological evaluation of all tissues was performed in a blinded fashion

by a board-certified pathologist.

All animals were maintained in an isolated environment in barrier cages. Animal care

was conducted in compliance with the state and federal Animal Welfare Acts and the standards

and policies of the Department of Health and Human Services. The protocol was approved by the

Wake Forest University Institutional Animal Care and Use Committee.

Transgenic mice with inducible CreER recombinase

Mice homozygous for the floxed Fasn and Pten genes were mated with inducible

probasin-Cre recombinase (CreERT) transgenic mice (kindly provided by Dr. Linda

deGraffenried of University of Texas, Austin). Mice heterozygous for the floxed Pten and Fasn

genes and positive for the CreER transgene were mated with mice heterozygous for the floxed

86

Pten and Fasn genes and negative for the CreER transgene. Activation of Cre recombinase was

induced by intraperitoneal injection of 4-hydroxytamoxifen (4-OHT, Sigma H7904) suspended

in corn oil at a low dose (0.01 mg/day) or a high dose (0.1 mg/day) for 5 consecutive days at the

age of 2 weeks. 4-OHT was stored at -20°C at a concentration of either 0.20 mg/mL (low dose)

or 2.0 mg/mL (high dose). Each mouse was injected with 50 μL of solution using a 30G needle.

All animals were maintained in an isolated environment in barrier cages. Animal care

was conducted in compliance with the state and federal Animal Welfare Acts and the standards

and policies of the Department of Health and Human Services. The protocol was approved by the

Wake Forest University Institutional Animal Care and Use Committee.

Histochemistry

Prostate samples were dissected, fixed overnight in 10% buffered formalin, and

processed for embedding in paraffin. Sections were sliced at 5 µm increments and mounted to

poly-lysine coated glass slides. Routine processing and staining was performed with hematoxylin

and eosin (H&E).

Immunohistochemical (IHC) staining was performed with the following primary

antibodies: mouse anti-Fatty Acid Synthase (BD Bioscience 610962), rabbit anti-pAKT473

(Cell

Signaling 4060), rabbit anti-Ki67 (Abcam 15580), and rabbit anti-Cleaved Caspase 3 (Cell

Signaling 9664). Primary antibody was incubated at 4°C overnight in a humidity chamber,

followed by HRP-conjugated goat anti-mouse (NA931V, GE Healthcare) or goat anti-rabbit

(Jackson Immunoresearch 111-035-003) secondary antibody. DAB chromogen was applied for

3-8 minutes according to the package insert (Dako K3467). TUNEL staining was performed

using a DeadEnd Colorimetric Assay kit (Promega G7360) according to the manufacturer’s

87

instructions. A positive TUNEL control included a DNase I digestion that was processed

separately from all other samples. A negative TUNEL control was processed in the absence of

any rTdT enzyme.

Microscopy

All images were captured with either a Hamamatsu Nanozoomer 2.0HT with Olympus

objectives and NDP image software or an Olympus VS110 microscope equipped with Olympus

objectives and VS-ASW FL image software. Image frames were created using Olympus Olyvia

software. Ki67 staining was quantified by averaging the percentage of positive cells in 10 frames

of each lobe at 80X magnification.

Statistics

Statistics were performed using Microsoft Excel, Daniel’s XL Toolbox Plugin 6.51. A

one-way ANOVA, followed by a Tukey test, was performed to identify statistical significance

for each group.

III.D RESULTS

Generation of prostate-specific Pten and Fasn knockout mice

Male mice homozygous for the floxed Pten gene and positive for the Pb-Cre4 transgene

(PtenL/L

;CreT) [20] were bred with female mice homozygous for the floxed Fasn gene (Fasn

L/L)

[25]. Resultant progeny of the F1 generation were then bred to produce the F2 generation of

desired genotypes: PtenL/L

;FasnL/L

;Cre- (herein referred to as wild-type), Pten

L/L;Fasn

L/L;Cre

T,

PtenL/L

;FasnL/+

;CreT

(herein referred to as Pten-/-

;FasnL/+

), and PtenL/L

;Fasn+/+

;CreT

(herein

88

referred to as Pten-/-

;Fasn+/+

) (Fig. 1A). During the breeding process of generating F2 mice we

observed a noticeable absence of progeny that were homozygous for the floxed Fasn gene and

positive for Cre recombinase, irrespective of gender and inheritance of the floxed Pten gene.

Because the statistical probability of producing a male PtenL/L

;FasnL/L

;CreT mouse from our

early breeding pairs was very low, at only 1 in 64 based on independent Mendelian inheritance,

we increased the probability of producing a PtenL/L

;FasnL/L

;CreT mouse by breeding

PtenL/+

;FasnL/+

;CreT males with Pten

L/L;Fasn

L/+;Cre

- females. This breeding strategy showed

that FasnL/L

;CreT mice were still absent from progeny outcomes. Out of 155 progeny, no mice

were either PtenL/L

;FasnL/L

;CreT or Pten

L/+;Fasn

L/L;Cre

T, even though the expected frequency

(6.3%) for these genotypes was the same as other genotypes that were successful generated: 1

PtenL/+

;Fasn+/+

;Cre- mouse (0.6% of progeny), 23 Pten

L/+;Fasn

+/+;Cre

T mice (14.8% of progeny),

19 PtenL/+

;FasnL/L

;Cre- mice (12.3% of progeny), 3 Pten

L/L;Fasn

+/+;Cre

- mice (1.9% of progeny),

24 PtenL/L

;Fasn+/+

;CreT mice (15.5% of progeny), and 10 Pten

L/L;Fasn

L/L;Cre

- mice (6.5% of

progeny) (Fig. 1B). Next, we further increased the statistical probability of generating a

PtenL/L

;FasnL/L

;CreT mouse by breeding Pten

L/+;Fasn

L/+;Cre

T males to Pten

L/L;Fasn

L/L;Cre

-

females. This breeding strategy also did not generate a PtenL/L

;FasnL/L

;CreT mouse out of 91

total progeny. Even though the expected frequency (12.5%) was the same for all possible

outcomes, 4 mice were PtenL/+

;FasnL/+

;Cre- (4.4% of progeny), 17 mice were

PtenL/+

;FasnL/+

;CreT (18.7% of progeny), 15 mice were Pten

L/+;Fasn

L/L;Cre

- (16.5% of progeny),

4 mice were PtenL/L

;FasnL/+

;Cre- (4.4% of progeny), 22 mice were Pten

L/L;Fasn

L/+;Cre

T (24.2%

of progeny), and 29 mice were PtenL/L

;FasnL/L

;Cre- (31.9% of progeny) (Fig. 1C). Finally, to

verify that generating a PtenL/L

;FasnL/L

;CreT mouse was highly improbable, we bred a

PtenL/L

;FasnL/+

;CreT

male with a PtenL/L

;FasnL/L

;Cre- female. The statistical probability was the

89

same for all possible genotypic outcomes (25%); however, 51.5% of progeny (35 mice) were

PtenL/L

;FasnL/+

;CreT and 48% of progeny (33 mice) were Pten

L/L;Fasn

L/L;Cre

-, and no mice were

PtenL/L

;FasnL/L

;CreT (Fig. 1D).

Hemizygous, prostate-specific deletion of Fasn reduces tumor burden in Pten-null mice

Despite the complications encountered while developing a Fasn knockout mouse using

the Pb-Cre4 transgene, we continued to evaluate the effects of single allele knockout of Fasn in

Pten-null mice. Although not statistically significant (P=0.07), a trend of decreased total prostate

weight was observed between Pten-/-

;Fasn+/+

mice, who had an average prostate weight of 77.7

mg at 2 months, and Pten-/-

;FasnL/+

mice, who showed an average prostate weight of 69.6 mg at 2

months. Wild-type mice had a statistically significant (P<0.001) lower average prostate weight

of 35.9 mg at 2 months compared to both Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice (Fig. 2A).

Interestingly, when stratified by each prostate lobe, there was a statistically significant (P<0.05)

difference in the dorsolateral lobe, but not the ventral or anterior lobe, between Pten-/-

;Fasn+/+

(VP: 12.2 mg, DL: 32.5 mg, AP: 33.9 mg) versus Pten-/-

;FasnL/+

(VP: 10.2 mg, DL: 25.8 mg, AP:

28.2 mg) mice (Fig. 2B), which could be observed in the gross anatomy of the prostate lobes for

each genotype (Fig. 2C). Wild-type mice exhibited a consistently significant (P<0.05) decrease

in prostate weight for each lobe (VP: 7.2 mg, DL: 13.0 mg, AP: 16.6 mg).

Although only a trend in decreased total prostate weight was observed at 2 months, a

single allele knockout of Fasn significantly (P<0.01) decreased the total prostate weight of Pten-

null mice at 6 months (Pten-/-

;Fasn+/+

(157.6 mg) versus Pten-/-

;FasnL/+

(131.8 mg)). Wild-type

mice also continued to show a significantly lower (P<0.001) total prostate weight (57.1 mg) at 6

months (Fig. 2D). Analysis of the individual prostate lobes demonstrated a significant difference

90

in weight between the dorsolateral (P<0.05) and anterior (P=0.01), but not the ventral (P=.37),

prostate lobes between Pten-/-

;Fasn+/+

(VP: 18.7 mg, DL: 41.7 mg, AP: 96.3 mg) and Pten-/-

;FasnL/+

(VP: 22.2 mg, DL: 30.2 mg, AP: 71.2 mg) mice. Wild-type mice showed significantly

decreased weight for the ventral (VP: 10.2 mg) (P<0.01) and anterior (AP: 27.2 mg) (P<0.001)

lobes compared to both Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice. However, the dorsolateral lobe of

wild-type mice (DL: 21.1 mg) was significantly lower than Pten-/-

;Fasn+/+

mice (DL: 41.7 mg)

(P<0.001), but not Pten-/-

;FasnL/+

mice (DL: 30.2 mg) (P=0.11) (Fig. 2E), suggesting the

hemizygous deletion of Fasn had its greatest effects in the dorsolateral lobe. The change in

prostate weight at 6 months was also evident in the gross anatomy of each prostate lobe (Fig.

2F).

Hemizygous, prostate-specific deletion of Fasn attenuates cancer progression in Pten-null mice

To confirm the changes observed in prostate weight, we analyzed the histology of each

prostate lobe of mice. In agreement with the data showing significant changes in the prostate

weight of each lobe, histological changes were also evident at both 2 and 6 months. At 2 months,

22% of Pten-/-

;Fasn+/+

mice had developed hyperplasia, 56% had developed hyperplasia with

atypia, and 22% had developed PIN. In contrast, 22% of Pten-/-

;FasnL/+

mice remained normal,

while 56% developed hyperplasia, and 22% developed PIN. All wild-type mice at 2 months

showed normal histology (Fig. 3A). By 6 months, prostates from Pten-/-

;Fasn+/+

mice had

histologically progressed; 44% of Pten-/-

;Fasn+/+

mice developed hyperplasia with atypia, and

56% developed PIN. In comparison, prostates from Pten-/-

;FasnL/+

mice demonstrated attenuated

tumor progression at 6 months, since 22% showed hyperplasia, 44% developed hyperplasia with

atypia, and 33% developed PIN. Wild-type mice were all normal at 6 months, except 1 mouse

91

(11%) who showed signs of hyperplasia. Of note, Pten-/-

;Fasn+/+

also tended to show evidence of

papillary cystic changes, while Pten-/-

;FasnL/+

mice tended to show stromal hyperplasia (Fig. 3B).

Hemizygous deletion of Fasn reduces FASN expression, activation of AKT, and proliferation at 6

months

Deletion of Pten in the mouse prostate is known to increase pAKT473

activation, and

previous reports have demonstrated reciprocal regulation between AKT and FASN, whereby

inhibition of AKT results in decreased FASN expression and inhibition of FASN leads to

decreased activation of AKT [19, 26]. To determine the effects of hemizygous deletion of Fasn

on AKT signaling in our Pten knockout mouse model, prostate sections were stained by IHC for

pAKT473

and FASN. Pten-/-

;Fasn+/+

mice showed very strong staining of pAKT473

in the

cytoplasm and at the plasma membrane in all prostate lobes, suggesting efficient deletion of Pten

and activation of AKT. In contrast, Pten-/-

;FasnL/+

mice displayed very reduced pAKT473

staining

in the cytoplasm, but similar pAKT473

staining at the plasma membrane. Wild-type mice stained

negative for pAKT473

(Fig. 4A). Similarly, Pten-/-

;Fasn+/+

mice stained strongly for FASN in all

lobes, whereas much weaker FASN staining was apparent in all three prostate lobes of Pten-/-

;FasnL/+

mice, albeit to a lesser extent in the ventral lobe. Wild-type mice showed very little to no

FASN staining in all three lobes (Fig. 4B).

To determine the effects of Fasn deletion on cellular proliferation, we performed a Ki67

immunohistochemical stain, as a nuclear marker of cell cycle activation. In agreement with the

measured weight of each prostate lobe for each genotype, there was a similar trend in

proliferation. A significant difference (P<0.05) in the percentage of Ki67 positive cells was

observed in each lobe and between all three genotypes at 6 months. Wild-type mice showed very

92

little proliferation, Pten-/-

;Fasn+/+

mice showed the greatest amount of proliferation, and Pten-/-

;FasnL/+

expressed a percentage of proliferative cells in between the two. Interestingly, the

dorsolateral lobe appeared to show the greatest difference in proliferation between Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice (Fig. 5).

Hemizygous deletion of Fasn is insufficient to alter cell death in Pten knockout mice

Evidence in vitro suggests inhibiting FASN by small molecule inhibitors or siRNA

promotes growth inhibition and apoptosis of cancer cells [14, 27], and that overexpression of

FASN protects cancer cells from apoptosis due to increased lipid saturation in membrane [10,

28]. To determine what effects inhibiting FASN may have on apoptosis in our prostate cancer

mouse model, we performed cleaved caspase 3 staining on prostate tissues. The results were

similar for both Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice. The ventral lobe showed sparse nuclear

staining in only a few glands. In contrast, the dorsolateral lobe seemed to have intense

cytoplasmic staining in certain glands with preference to the luminal side, and punctate nuclear

staining within these areas of intense cytoplasmic staining. The anterior lobe appeared to display

a staining pattern mixed between that of the ventral and dorsolateral lobes; there was cytoplasmic

staining and minor nuclear staining, but neither were widespread (Fig. 6A).

Because caspases are merely markers of apoptosis and do not necessarily identifying

actively dying cells, we performed a follow up analysis with a TUNEL assay to detect active

DNA fragmentation, a downstream effect of caspase activation and definitive indicator of cell

death. The data, again, showed very similar staining patterns in both Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice. The ventral and dorsolateral lobes of the prostate were extremely sparse in

positively stained cells, having less than 20 positive cells per sample. The anterior lobes of both

93

Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice were also sparse with the exception of very specific areas

within select lumens that seemed to have dense TUNEL staining. Wild-type mice showed very

few positively stained cells in each lobe (Fig. 6B). Although cell death was sparsely scattered in

the majority of each lobe and dense in just a couple areas of the prostate lobes in Pten-/-

;FasnL/+

mice, the hemizygous deletion of Fasn did not appear to alter the extent of cell death compared

to that of Pten-/-

;Fasn+/+

mice, which showed a very similar staining pattern.

Generation and analysis of inducible, prostate-specific Pten and Fasn knockout mice

Because we were unable to achieve a complete knockout of Fasn in the Pten knockout

mouse using the Pb-Cre4 transgene, we generated an inducible knockout mouse model using a

tamoxifen-inducible Cre recombinase transgene that was also regulated by the probasin promoter

(Pb-CreERT). In this model, the Cre transgene is fused to the ligand binding domain of the

estrogen receptor (ER) bearing a glycine to arginine point mutation at residue 525. As a result of

this point mutation, activation of the Cre recombinase is tamoxifen-dependent, and resistant to

activation by endogenous β-estradiol binding [29, 30]. Previous studies using Pb-CreERT mice

reported robust activation of Cre in prostate epithelia, as evidence by Rosa26-lacz and Rosa26-

EYFP reporter strains [31]. When mated with mice homozygously floxed for the Pten gene,

progeny were reported to have developed PIN lesions within 3 months of injecting 2 week old

mice with 4-hydroxytamoxifen (4-OHT), the active metabolite of tamoxifen [32]. Following this

published protocol, we injected mice with 0.10mg/day for 5 days starting at 14 days old and

harvested prostates from mice at 3 months post-injection. We found that this dose of 4-OHT

appeared to increase the prostate weight of control mice, suggesting the 4-OHT was potentially

acting as growth enhancer during the maturation of mice (Fig. 7A). Therefore, we decreased the

94

dose of 4-OHT to 0.01mg/day and harvested prostates from mice at 3 months post-injection. At

the lower 4-OHT dose, PtenL/L

;Fasn+/+

;CreERT mice showed a significant (P<0.01) increase in

total prostate weight compared to PtenL/L

;FasnL/+

;CreERT, Pten

L/L;Fasn

L/L;CreER

T, and

PtenL/L

;FasnL/L

;CreER- mice treated with 4-OHT and Pten

L/L;Fasn

L/L;CreER

T mice treated with

vehicle, but not PtenL/L

;FasnL/L

;CreER-

mice treated with vehicle (Fig. 7B). Upon further

examination, histological analysis of each genotype showed no pathological changes between

any groups (Fig. 8). IHC staining for pAKT473

and FASN showed that only a few glands, located

in the dorsolateral lobe, out of the entire prostate of PtenL/L

;Fasn+/+

;CreERT mice had developed

high activation of AKT and consequentially, strong FASN expression. These results suggest the

approach of using an inducible Cre model was also ineffective at achieving homogenous

prostate-specific deletion of both Pten and Fasn (Fig. 9).

III.E DISCUSSION

Multiple in vitro studies have demonstrated that fatty acid synthesis, particularly FASN,

is an attractive target for therapeutic inhibition of prostate cancer [14, 15, 28, 33]. Previous

studies have used xenograft models and pharmacological inhibitors to model the inhibition of

FASN in prostate cancer in vivo [13, 34]. However, these studies fail to acknowledge important

aspects of tumor biology, such as the immune system and microenvironment-mediated

modulation of prostate cancer [35], as well as off-target drug effects , which can confound results

and conclusions regarding the role of FASN [22, 36]. Our approach of genetically deleting Fasn

in the prostate of immunocompetent mice addresses these concerns. We found that genetically

inhibiting FASN in the widely accepted Pten-knockout prostate cancer mouse model validated a

significant role for FASN in prostate cancer development and progression. Although a complete

95

knockout of Fasn was unachievable using the popular Pb-Cre4 mouse model [24], we have

shown that even hemizygous deletion of Fasn is sufficient to decrease FASN expression and

activation of AKT, and delay prostate cancer development and progression in the setting of Pten

deletion in mice.

The Pb-Cre4 transgenic mouse model has been widely used and accepted for prostate-

specific deletion of genes of interest [20, 37, 38]. In this model the activation of Cre is androgen-

dependent and hence, occurs during the maturation of young, male mice [24]. Despite the

embryonic lethality of deleting Pten in mouse embryos [39], the prostate-specific deletion of

Pten using the Pb-Cre4 transgene was reported to efficiently and reproducibly induce prostate

cancer in male mice, and has been extensively used since its initial report in 2003 [20, 38, 40,

41]. Because the deletion of either Pten or Fasn leads to unviable embryos, one would expect

that the successful, conditional deletion of Pten in the prostate would predict the same for Fasn;

however, this was not true in our study. The fact that we could not generate a homozygous Fasn

knockout mouse, suggests the Pb-Cre4 transgene may exhibit unanticipated activation at an

embryonic stage for which the loss of Fasn is more sensitive than the loss of Pten. A recent

report demonstrated extensive genetic recombination at loxP sites in a variety of tissues when the

Pb-Cre4 transgene was transmitted maternally [42]; however, since our lab has also observed

similar results (Chen et al., unpublished results), we used only male Cre-positive mice for

breeding in our model. Nonetheless, supporting this conclusion is the fact that Pten-/-

embryos

are undetectable following embryonic day 7.5, and Pten+/-

mice are phenotypically

indistinguishable from wild-type littermates [39]. In contrast, Fasn-/-

embryos are undetectable

after embryonic day 3.5, and approximately 70% of expected Fasn+/-

mice die in utero.

Moreover, female Fasn+/-

mice produce even fewer Fasn+/-

progeny within a matter of 12 weeks

96

of active breeding, most likely due to a progressive decline in Fasn- gamete production [43].

Overall, this evidence strongly suggests that Pb-Cre4 mice may have mildly promiscuous

activation of Cre during the embryonic stage, for which recombination events of Pten, but not

Fasn, is permissible to produce viable mice. This could explain the generation of prostate-

specific Pten-null mice and the absence of prostate-specific Fasn-null mice using the Pb-Cre4

transgene.

Despite the deletion of only a single Fasn allele, Pten-null mice with a heterozygous

deletion of FASN showed fewer and lower grade lesions at both 2 and 6 months, and had a

significantly decreased total prostate weight compared to Pten-null mice with wild-type Fasn.

There was an obvious decrease in FASN expression in our Fasn+/-

mice at 6 months, suggesting

the transcription from just one allele directly translated to FASN protein expression, which is not

surprising because expression of FASN is largely regulated at the transcriptional level [44].

Interestingly, there appeared to be a noticeably greater decrease of FASN staining in the

dorsolateral lobe of Pten-/-

;FasnL/+

mice, which among all three lobes most closely resembled the

wild-type expression of FASN. In line with this observation, the pathological review of the

histological changes in prostate samples noted that the greatest distinctions were observed in the

dorsolateral lobe. Moreover, the difference in the total weight of the dorsolateral lobe between

Pten-/-

;FasnL/+

and wild-type mice was insignificant. Collectively, this evidence suggests the

activation of Cre and deletion of Fasn may have been most prominent in the dorsolateral lobe of

prostates, and in agreement with previous reports [8], expression of FASN may be a direct

molecular marker of prostate tumor progression.

Our model also demonstrated a significant decrease in the number of actively

proliferating cells in each prostate lobe. This was, at least in part, likely due to a decrease in the

97

activation of pAKT473

, a known regulator of cell growth and proliferation [45]. Our in vivo

analysis showed that heterozygous loss of FASN was sufficient to hinder activation of AKT and

minimize the cytoplasmic fraction of pAKT473

, while still maintaining strong pAKT473

localization at the plasma membrane. This effect could have been due to decreased saturation in

the plasma membrane and lipid raft-mediated activation of AKT; however, fatty acid methyl

ester analysis showed no significant changes between the saturated or unsaturated fatty acid

profile of prostates from Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice (data not shown). Thus, the

change in pAKT473

was not likely a result of changes in the plasma membrane composition, but

could still be a consequence of alterations in protein-mediated signaling at the plasma membrane.

Finally, despite previous reports showing the induction of apoptosis following inhibition

of FASN [14, 46], our investigation showed that inhibition of FASN by hemizygous deletion

does not induce cell death beyond that of typical tumor biology in vivo. We cannot rule out that

perhaps a complete deletion of Fasn is required to achieve this effect in vivo; nevertheless, our

data imply that increased expression of FASN in Pten knockout mice is more important for

propagating cancer cell growth and proliferation than for preventing apoptosis and cell death.

Multiple reports have described the effects of inhibiting lipogensis in vitro, and thus, a

few hypotheses have arisen to explain the lipogenic phenotype of cancer cells. One of the most

widely accepted hypotheses is that cancer cells require a persistently high demand for fatty acids

to produce new membrane, and that dietary fatty acids are an insufficient supply for this demand.

Supporting this hypothesis, prostate cancer cells have been shown to require fatty acid synthesis

for phospholipid biogenesis and lipid raft formation in membrane [11], and the inhibition of

FASN leads to endoplasmic reticulum (ER) stress and cell death in vitro [47]. Our analysis of the

fatty acids in Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice, which showed no significant differences,

98

suggest this may not hold true in vivo. Because we did not measure the activity of FASN in our

model, it is possible that the decrease in FASN protein is not necessarily an indicator of

decreased fatty acid synthesis. If this is true, then the significant changes we observed in prostate

weight and histology may be accredited to other functions of FASN besides generating fatty

acids.

In vitro and in vivo studies of FASN have also demonstrated its requirement for the

palmitoylation of proteins, which has been shown to occur in both cancer and normal cells, and

can readily modulate the activation and cellular localization of the modified protein [48-50].

Although we did not search for alterations in protein palmitoylation, the seemingly stagnate

localization of pAKT473

at the plasma membrane of FASN-depleted samples implies a missing

piece for activation of pAKT473

at the plasma membrane and its subsequent release into the

cytoplasm where it is known to activate growth and proliferation pathways [45]. It is possible

that FASN is required for the palmitoylation and membrane localization of a specific protein that

allows the cytoplasmic release of pAKT473

from the membrane.

Finally, the increased expression of FASN in cancer cells has also been shown to protect

cells from apoptosis in vitro [10], particularly by increasing membrane lipid saturation and

consequently, reducing the formation of reactive oxygen species (ROS) from lipid peroxidation

of unsaturated fatty acids [28]. It is possible that this phenomenon may also not hold true in vivo,

since we found no changes in the fatty acid profile of phospholipid species between Pten-/-

;Fasn+/+

and Pten-/-

;FasnL/+

mice (data not shown). Moreover, the decrease in FASN did not

appear to alter cleaved caspase 3 activation or cell death beyond that of typical tumor biology in

Pten-null mice. Again, this could be confounded by the fact that FASN was not completed

99

deleted. Nevertheless, it is not surprising to discover that in vitro findings may not translate in

vivo, given the incredible complexity of tumor biology [35].

Overall, the major drawback to this study was, obviously, the unsuccessful generation of

a homozygous deletion of Fasn in Pten-null mice. Although we attempted to overcome this

complication by using an inducible Cre recombinase transgene, the full extent of the requirement

for FASN in prostate tumorigenesis and progression remains unknown. Because 2 month old

Pten-/-

;FasnL/+

mice showed a trending decrease in total prostate weight, and 6 month old Pten-/-

;FasnL/+

mice showed a significant decrease, which both correlated nicely with changes in

histological progression, we can at least confirm that FASN plays an important role in prostate

cancer progression in vivo. It would be interesting to determine whether fatty acid synthesis

activity in Pten-/-

;FasnL/+

mice versus that of Pten-/-

;Fasn+/+

mice is different. If the activity of the

pathway is actually the same, this could explain the absence of a shift in the fatty acid profiles,

despite decreased FASN protein levels. It would also distinguish FASN as more than just a fatty

acid generator for prostate cancer cells, since the decrease in protein still hindered cancer

progression. It would also be interesting to determine whether a diet enriched in saturated fatty

acids could rescue the phenotype of Pten-/-

;FasnL/+

mice. In vitro analysis suggests exogenous

palmitate can rescue the effects of FASN inhibition [10, 13]; however, other in vivo models of

tissue specific FASN deletion suggest diet is insufficient at rescuing the loss of de novo fatty acid

synthesis [51]. Future investigation of FASN in vivo may be possible with a different Cre

recombinase transgene.

100

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III.G ACKNOWLEDGMENTS

We thank Dr. Clay Semenkovich (Washington University School of Medicine, St. Louis, MO)

for generously providing floxed Fasn mice. We thank Linda deGaffenreid for providing the Pb-

CreER mice. We thank Brandi Bickford of the Wake Forest University Virtual Microcopy Core

for technical assistance. This study was funded in part by the National Institutes of Health

(R01CA107668, P01CA106742, R01CA163273, T32CA079448 training grant) and The

Synergistic Innovation Center for Food Safety and Nutrition, Jiangnan University.

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III.H FIGURES AND FIGURE LEGENDS

Figure 1. Genotypic outcomes from animal model breeding.

A. Breeding strategy for generating PtenL/L

;FasnL/L

;CreT mice. B. Expected and observed

genotypes of progeny from breeding a PtenL/+

;FasnL/+

;CreT sire with a Pten

L/L;Fasn

L/+;Cre

- dam.

C. Expected and observed genotypes of offspring from breeding a PtenL/+

;FasnL/+

;CreT sire with

a PtenL/L

;FasnL/L

;Cre- dam. D. Expected and observed genotypes of offspring from breeding a

PtenL/L

;FasnL/+

;CreT sire with a Pten

L/L;Fasn

L/L;Cre

- dam. The number of expected progeny (left

y-axis) was calculated based on the expected Mendelian frequency (right y-axis) for the total

number of progeny generated.

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Figure 2. Heterozygous loss of FASN reduces tumor burden.

A, D. Total prostate weight at 2 months (A) and 6 months (D). ***P<0.001. B, E. Prostate

weight of the ventral (VP), dorsolateral (DL), and anterior (AP) lobes at 2 months (B) and 6

months (E). Letters (a,b,c) represent a statistically significant change within each lobe (P<0.05).

Error bars represent s.e.m. C, F. Gross anatomy of prostate lobes at 2 months (C) and 6 months

(F).

107

Figure 3. Heterozygous loss of FASN hinders prostate cancer progression.

Prostate lobes were harvested, sectioned, and stained with hemotoxylin and eosin at (A) 2

months and (B) 6 months. Histopathological analysis by a board-certified pathologist is graphed

on the right side of each panel. n=9 for each genotype. Ventral prostate (VP), dorsolateral

prostate (DL), and anterior prostate (AP), hyperplasia (HP), prostatic intraepithelial neoplasia

(PIN).

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Figure 4. Heterozygous deletion of Fasn decreases FASN and cytoplasmic pAKT473

.

Prostate lobes from 6 month old mice were harvested, sectioned, and stained by IHC for

pAKT473

(A) and FASN (B). High magnification insets are outlined in black for each image.

Ventral prostate (VP), dorsolateral prostate (DL), and anterior prostate (AP).

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Figure 5. FASN facilitates prostate cancer cell proliferation.

A. Prostate lobes were harvested, sectioned, and stained by IHC for Ki67. B. Cellular

proliferation was quantified by counting the percentage of Ki67-positive cells. Letters (a,b,c)

represent a statistically significant change within each lobe (P<0.05). Error bars represent s.e.m.

Ventral prostate (VP), dorsolateral prostate (DL), and anterior prostate (AP).

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Figure 6. Hemizygous deletion of FASN has no effect on cell death in Pten knockout mice

Prostate lobes were harvested, sectioned, and stained by IHC for cleaved caspase 3 (A) or by

TUNEL assay (B). Lower images outlined in black for each lobe represent the black box

indicated in the panel above. Red arrows indicate TUNEL positive cells. Ventral prostate (VP),

dorsolateral prostate (DL), and anterior prostate (AP).

111

112

Figure 7. Total prostate weight of inducible Pten/Fasn knockout mouse model.

Mice were injected for 5 consecutive days at 14 days old with 4-OHT at either a high dose (A.

0.10 mg/day) or a low dose (B. 0.01 mg/day), or vehicle. Prostates were harvested 3 months

post-injection.

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Figure 8. Histology of inducible Pten/Fasn knockout mouse model.

Prostate lobes of inducible-Cre mice receiving the low dose (0.01mg/day) were harvested,

sectioned, and stained with hematoxylin and eosin for histological analysis. Ventral prostate

(VP), dorsolateral prostate (DL), and anterior prostate (AP).

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Figure 9. Immunohistochemical staining for pAKT473

and FASN in Pb-CreERT mice.

Prostate lobes of inducible-Cre mice receiving the low dose (0.01mg/day) were harvested,

sectioned, and stained for pAKT473

(A) and FASN (B). Ventral prostate (VP), dorsolateral

prostate (DL), and anterior prostate (AP).

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CHAPTER IV:

GENERAL DISCUSSION

Sections of this manuscript were published in a book chapter written by Janel Suburu, et al. [1].

The manuscript was prepared by Janel Suburu and edited by Dr. Yong Q. Chen, whom also acted

as an advisor.

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IV.A BIOLOGICAL IMPLICATIONS OF DE NOVO FATTY ACID SYNTHESIS

The ability to synthesize fatty acids has been well conserved over the course of evolution

from prokaryotes to humans, which speaks volumes about the importance of the enzymes

responsible for this task [2, 3]. The principal enzyme for animal fatty acid synthesis, fatty acid

synthase (FASN), is a highly sophisticated, multi-functional enzyme [4]. Early investigations

over 40 years ago identified FASN as a regulator of metabolic homeostasis in the liver and

adipose of animals during oscillating periods of feeding and fasting, as well as the producer of

milk fat in the lactating mammary gland [5]. Beyond these characteristic roles, FASN was, for

many years, believed to be relatively quiescent in most tissues. However, with the development

of various knockout models in the past decade, the biological significance of de novo fatty acid

synthesis has become clearer. Fatty acid synthesis is absolutely required for embryogenesis, as

evidence by the Fasn and Acc1 knockout mouse models [6, 7]; it has also been shown to regulate

homeostasis for tissues under a variety of specific physiological pressures [8-10]. Because

oscillating periods of fasting and feeding are no longer a concern in developed countries where

food is plentiful, emerging roles for FASN in the development of certain pathologies, including

diet-induced obesity and cancer, identify FASN as a unique target for the development of

therapeutic drugs. Still, a thorough understanding of the effects of inhibiting a drug target in both

pathological and normal cells is a critical aspect of drug development. This dissertation outlines

the development and analysis of two mouse models demonstrating tissue-specific inhibition of

FASN – one in a normal, physiologically-induced setting, and the other in a pathological setting.

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IV.B INHIBITING DE NOVO FATTY ACID SYNTHESIS

Effects in normal biology

Several reports have outlined tissue-specific roles for fatty acid synthase, and its

inhibition often leads to failed homeostasis and/or tissue function, especially when imposed with

specific physiological pressures. For example, the deletion of Fasn in cardiomyocytes impairs the

heart’s ability to overcome cardiovascular stress, and mice deficient in cardiac FASN die from

heart failure when challenged with stress-inducing stimuli [10]. In the brain, FASN is required to

maintain systemic energy homeostasis by modulating feeding behaviors and energy expenditure

[9]. In the intestine, FASN is required to sustain the integrity of the intestinal barrier from the

local microbiome; mice lacking intestinal FASN die from sepsis [11]. In endothelial cells, FASN

is required to prevent vascular permeability and chronic inflammation; mice lacking endothelial

FASN die from infection when challenged with lipopolysaccharide [12]. Overall, these models

have demonstrated that FASN plays a unique role for maintaining functional homeostasis in a

tissue-specific manner.

As described in Chapter II, FASN also appears to play a homeostatic role in the

mammary gland in response to pregnancy and lactation. Although it was previously known that

fatty acid synthesis is strongly induced in the mammary gland during lactation [13], our study

showed that FASN is required to maintain functional competence of the lactating mammary

gland. Deletion of Fasn in the mammary gland hindered gland development and ultimately led to

premature gland involution. The molecular mechanisms involved in this response are yet to be

determined, but evidence of adequate lipid droplet formation suggests FASN is not required for

milk secretion, despite a significant decrease in milk fat content. Nevertheless, it is quite clear

that FASN plays a role in regulating the maturation of the gland to peak lactational competence

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near lactation day 10, as well as preventing early involution of the gland. The premature

involution observed in Fasn KO mice was probably a result of lost functional competence, but

even at lactation day 18 many KO glands produced milk. Thus, it remains in question whether

FASN is more important for milk fat production or the structural/functional integrity of the

gland, or possibly equally important for both.

Future studies investigating the role of FASN in the lactating mammary gland should

focus on understanding the molecular mechanisms triggering the early involution of lactating

mammary glands in Fasn KO mice, and whether a diet enriched in saturated fatty acid can rescue

this phenotype. Involution typically occurs in a two-phase process: the early, reversible stage and

the late, irreversible stage [14]. It is currently unclear exactly which molecules trigger involution,

but the current hypothesis suggests that the signaling molecules are most likely triggered in

response to milk accumulation, either by stretching of the alveolar lumen, or their accumulation

in the milk remaining inside the glands [15]. Our data suggests that milk stasis was not the

leading factor of involution in FASN-depleted mammary glands. In fact, pups were still actively

nursing when involution of glands had already initiated. However, this does not exclude the

possibility that the same signaling molecules are ultimately enforcing involution of the gland.

AKT appears to be a primary regulator of the involution process, as its constitutive activation

suppresses apoptosis and delays involution in mice [16]. AKT activation is triggered within 12

hours of weaning, decreases sharply between 24 and 48 hours, and increases again by 72 hours

[15]. FASN is known to regulate AKT signaling [17], suggesting similar mechanisms may be

regulating involution in both normal and Fasn-deleted mammary glands, but on a different time

scale and in response to different stimuli. Understanding the molecular mechanism by which

FASN deletion induces involution and cell death under these conditions may aide the

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identification of other molecular drug targets besides FASN and the development of drugs to

induce cell death in a pathological setting, such as cancer.

Effects in cancer biology

Inhibiting FASN

Expression and activation of fatty acid synthesis is an integral component of cancer

pathogenesis [18]. A number of studies have attempted to exploit the lipogenic phenotype, by

chemical inhibition of lipogenic enzymes as a means to treat cancer [19-27]. Many in vitro

studies have provided proof-of-concept data showing that inhibition of FASN leads to cancer cell

death and apoptosis [28-31]. Furthermore, increased expression of FASN protects cancer cells

from apoptosis [32]. Because cancer cells seem to depend on FASN for tumor growth and

survival, inhibition of FASN is an attractive therapeutic approach. However, to fully elucidate

the feasibility and efficacy of inhibiting FASN as a cancer treatment, only the tissue-specific

deletion of FASN in an immunocompetent cancer mouse model can definitively establish the

effects of inhibiting FASN on cancer development and progression.

As described in Chapter III of this dissertation, the conditional, genetic ablation of Fasn

in an orthotopic, immunocompetent mouse prostate cancer model validated the role of FASN in

prostate cancer progression. Although a homozygous deletion was unachievable, the hemizygous

deletion of Fasn showed significant changes in prostate histology and molecular markers. No

significant differences were observed in apoptosis, cell death, or fatty acid analysis of prostate

tissues, but significant decreases in the total weight of prostates and a delay in the

histopathological progression were noted upon a blinded review by a board-certified pathologist.

Additionally, a significant decrease in proliferation and reduced cytoplasmic pAKT473

suggest

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FASN promotes prostate cancer progression by stimulating growth and proliferation signaling

pathways.

Our study is the first of its kind to genetically delete Fasn in a cancer mouse model. We

chose a prostate cancer model for our study for several reasons. First, prostate cancer is the most

commonly occurring cancer type in men in the United States, accounting for more than a quarter

of all cancer cases in men, and the identification of new therapeutic options is a priority in

prostate cancer research [33]. Second, prostate cancers tend to be very lipogenic with enhanced

fatty acid synthesis activity [34]. Several reports have shown that prostate cancers exhibit greater

uptake of radiolabeled acetate, as opposed to glucose, which can be used as a tool for the

detection of prostate cancer and expression of FASN as a molecular marker for diagnosis [35-

37]. Third, the deletion of Pten using Cre-LoxP recombination is a widely accepted mouse model

of prostate cancer, and its mechanism of inducing genetic deletion meets the conditional

requirements of deleting Fasn in vivo [6, 38, 39].

No significant differences were observed in the rate of cell death or the total- and

phospholipid fatty acid analyses in our mouse model. These results suggest three possibilities.

First, it’s possible that the hemizygous deletion of Fasn is simply insufficient to change these

features of prostate cancer in the Pten-null mouse model. If this is true, it should be cautioned to

drug developers that inhibitors of FASN will need to be efficient. We saw a large decrease of

FASN protein in mouse prostates with hemizygous deletion of Fasn; however, if even the

reduced amount of FASN is capable of propagating the disease as we observed in our model, the

administration of a FASN inhibitor alone will likely not be sufficient to treat cancer. The second

possibility is that the FASN enzyme of Pten-/-

;FasnL/+

mice had increased activity that

compensated for the decrease in FASN protein expression. While we do not have evidence of

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any changes in enzymatic activity in our mouse model, this has been reported to occur as a result

of HER2-mediated phosphorylation of FASN in breast cancer cells [40]. This possibility would

suggest that post-translational modifiers or other interacting proteins of FASN may be

complimentary targets of inhibition while pharmacologically inhibiting FASN. Supporting this

hypothesis, FASN has been shown to interact with and co-immunoprecipitate with multiple

proteins, including HER2 and regulators of calcium signaling [40, 41]. Finally, it is possible that

the reports of cell death and apoptosis induced by FASN inhibition in vitro may not directly

translate to the in vivo setting. Although this is unlikely given the extent of reproducibility of

data supporting the role of FASN and cancer cell survival [20, 28, 29, 31, 42], if this is true, data

from future studies of FASN in cell culture should be analyzed with caution.

Other mechanisms of inhibiting the fatty acid synthesis pathway

Although FASN is an ideal target due to the numerous metabolic alterations that

converge on this enzyme, inhibition of other enzymes involved in the fatty acid synthesis

pathway may be an alternative approach of targeting pathologically induced fatty acid synthesis.

Inhibition of ACC1, the rate-limiting enzyme of the pathway, in cancer cells has shown very

similar, if not identical, effects as the inhibition of FASN. These effects include decreased

proliferation, induction of apoptosis, and decreased incorporation of de novo fatty acids into

phospholipids. These results suggest that the inhibition of the FASN protein and the synthesis of

fatty acids, as opposed to merely cytotoxic accumulation of FASN substrate, malonyl-CoA, is

responsible for the observed effects of inhibiting FASN [19, 43, 44]. The activation and activity

of ACC1 is negatively regulated by phosphorylation, which means the fatty acid synthesis

pathway can also be shut down by promoting the activation of the ACC1-phosphorylating

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kinase, AMP-activated protein kinase (AMPK). AMPK is also known to inhibit SREBP-1,

making it an attractive target for inhibiting lipogensis in cancer [45].

Inhibitors of fatty acid synthesis

Multiple inhibitors of FASN have been identified, including both natural and synthetic

compounds. The first identified natural inhibitor of FASN, cerulenin, is an antifungal antibiotic

that binds to and inhibits the β-ketoacyl synthase domain of the FASN enzyme [46]. A synthetic

derivative of cerulenin, C75, was the first synthetic molecule developed to inhibit FASN [26].

Early experimentation of pharmacologically inhibiting FASN with the small molecule C75

showed profound effects on food intake and weight loss in mice. In this study, C75 significantly

decreased neuropeptide Y in the hypothalamus in a leptin-independent manner and the authors

purported that accumulation of malonyl-CoA was the culprit for signaling anorexic behaviors

[25]. However, subsequent studies showed that malonyl-CoA alone was not sufficient to induce

anorexia and that increased neuronal glucose metabolism, activation of hypothalamic AMPK,

and central nervous system communication through the sympathetic nervous system to

peripheral tissues were major molecular mechanisms mediating the effects of C75, suggesting

this molecule has many off-target effects [47-51]. Interestingly, a new report addresses the

importance of the chirality of C75, in characterizing its pharmacological properties. Evidence

from this study suggests the (+)-C75 enantiomer is a selective inhibitor of carnitine palmitoyl

transferase-1 (CPT1), and its central or intraperitoneal administration in rats induces hypophagia

and anorexia. In contrast, the (-)-C75 enantiomer is a selective inhibitor of FASN that is capable

of inducing cell death in both breast and ovarian cancer cell lines. Administration of the (-)-C75

enantiomer, either centrally or intraperitoneally, has no effect on feeding behaviors or weight

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loss [52]. This new evidence may provide a rebound opportunity for the (-)-C75 enantiomer as a

potential cancer therapeutic drug.

While C75 remains a commonly used pharmacological inhibitor in vitro, other

compounds with FASN-inhibiting properties have been identified. Another naturally occurring

molecule, epigallocatechin-3-gallate (EGCG), which is a polyphenol found in green tea, was

shown to have anti-FASN properties [53]. A synthetic analogue of EGCG, G28UCM,

demonstrated anti-cancer activity in HER2-positive breast cancer cells, but inhibited the growth

of only 35% of tested breast cancer xengrafts suggesting it has poor inhibiting effects in vivo [54,

55]. Orlistat, an FDA-approved, lipase-inhibiting drug for the treatment of obesity, was shown to

bind and inhibit the thioesterase domain of FASN [31, 56]. A small molecule developed by

GlaxoSmithKline, GSK837149A, also showed anti-FASN properties [27]. Finally, a recently

developed drug, ML356, was designed to inhibit the thioesterase domain of FASN and

demonstrated approximately 50% reduction of palmitate synthesis in prostate cancer cells [57].

Despite the exhaustive efforts to identify and/or generate a viable FASN-inhibiting drug, no

published inhibitor, thus far, has proven effective in vivo, largely because they have either poor

solubility and low bioavailability due to the hydrophobic properties of the FASN active site [58],

or significant off-target effects that alter eating habits, resulting in rapid weight loss and anorexia

[47-49, 51].

In parallel to the quest of identifying FASN inhibitors, several drugs have been described

to inhibit fatty acid synthesis by targeting other enzymes of the pathway. Soraphen A, an

inhibitor of ACC1, possesses anti-lipogenic and anti-tumorigenic properties; however, similar to

FASN inhibitors [47], it is non-selective and activates fatty acid oxidation, while concomitantly

inhibiting fatty acid synthesis [19]. Several small molecules and natural products have been

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shown to either directly or indirectly activate AMPK, including Metformin, a well-recognized

and approved treatment for type II diabetes. Metformin and other natural products known to

activate AMPK are currently being tested in clinical trials to determine their efficacy in treating

various cancers [59].

IV.C DIETARY AND METABOLIC CONSIDERATIONS DURING FASN INHIBITION

Dietary fat and cancer

Although FASN is the only enzyme capable of producing fatty acids from carbohydrate

and protein sources, the consumption of dietary fatty acids should be carefully considered when

pharmacologically inhibiting the fatty acid synthesis pathway. In vitro evidence suggests cancer

cells can utilize exogenous fatty acids to, at least in part, rescue the effects of FASN inhibition

[19, 31, 44]. Cancer cells can also increase their expression of lipoprotein lipase, lipoprotein

receptors, and scavenger fatty acid receptors to increase the uptake of extracellular fatty acids,

which can accelerate the growth of cancer cells [60-62]. Therefore, despite the nuances of

absorption and transport of dietary fatty acids to cells compared to the local availability of de

novo fatty acids, it is important to recognize that the adaptive responsiveness of cancer cells to

their surroundings may allow them to overcome the inhibition of FASN. For this reason, future

studies investigating the effects of FASN inhibition in a cancer setting should experiment with

controlled dietary manipulations, such as a high-saturated fat diet, that could potentially thwart

the efficacy of inhibiting fatty acid synthesis. Of note, dietary polyunsaturated fatty acids

(PUFA) can actually suppress transcriptional activation of lipogenic enzymes, suggesting a

PUFA-enriched diet may be complimentary to a treatment regimen that targets the inhibition of

fatty acid synthesis [63, 64].

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Obesity and cancer

The association between obesity, characterized by excessive fat storage, and cancer has

been recognized by numerous epidemiological studies [65]. Moreover, the cancer mortality due

to obesity is significantly increased by approximately 52% and 88% in men and women,

respectively [66]. Several cancers, such as breast, endometrial, and colorectal cancer, have

particularly strong associations with obesity and the management of obesity is one mechanism

by which cancer can be prevented [65, 67]. Studies of lipid signaling mediators from various

depots of adipose in the body and resident adipocytes within the tumor microenvironment have

provided evidence for a role of fat storage in cancer.

Adipose tissue and cancer

The dysfunctional metabolism and signaling of adipose tissue is believed to be the major

mechanism underlying cancer promotion in obese patients [68]. Adipose is a complex organ

involved in autocrine, paracrine, and endocrine signaling that synchronously modulate systemic

metabolism. Hormones, such as adiponectin and leptin, and other signaling mediators produced

by adipose influence the regulation of feeding and appetite, immune function, and reproduction

[69, 70]. Interestingly, not all fat depots are created equal, as the distribution of fat in the body

can differentially regulate metabolism and chronic inflammation. Among the various locations

for fat accumulation, such as visceral (mesenteric, omental, epiploic) and subcutaneous

(abdominal, gluteal, femoral), research suggests visceral adipose, as opposed to subcutaneous

adipose, has increased lipolysis and greater association with metabolic complications [70].

Abdominal visceral fat was shown to possess a developmental gene signature distinct from that

of gluteal, subcutaneous fat [71]. The de-differentiation of cancer cells and the surrounding

stroma to express developmental genes is a common event in cancer [72]. This evidence points

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to a potential role of adipose tissue depots in the vicinity of a growing tumor. Indeed, recently,

Zhang and colleagues eloquently demonstrated the recruitment of mesenchymal progenitor cells

from white adipose tissue to the tumor site. The white adipose tissue derived cells assimilated

into the tumor stoma and facilitated tumor growth as cancer associated adipocytes [73, 74].

Cancer associated adipocytes

The dysfunctional metabolism of adipose tissue in obese patients closely resembles the

metabolic alterations of cancer cells, a well-recognized hallmark of the disease. Cancer cells

themselves display an altered metabolic profile and the stromal cells of the tumor

microenvironment are also known contributors to the pathological progressions of cancer [75,

76]. However, only very recently has the function of adipocytes as tumor promoting accessories

become a recognized topic in tumor biology [77]. Joining the tumor microenvironment group of

fibroblasts, endothelial cells, and immune cells, cancer associated adipocytes have been shown to

acquire a unique phenotype in the presence of cancer cells characterized by dedifferentiation,

release of lipid stores, and overexpression of inflammatory cytokines that promote the

development of more aggressive cancer [78]. The symbiotic exchange of metabolites between

cancer cells and surrounding stromal cells, including fatty acids for β-oxidation as energy

strongly supports the role of fatty acid enriched adipocytes in cancer progression [79, 80].

Inhibiting FASN in cancer and obesity

Supporting the evidence connecting cancer and obesity, FASN has also been shown to

play a primary role in diet-induced obesity. Mouse models of tissue-specific FASN knockout in

skeletal muscle and adipose tissue have shown that activation of FASN promotes adiposity and

diabetes under conditions of high-fat diet feeding [41, 81]. Translational research supports this

data with similar findings in humans whereby increased expression of FASN in visceral adipose

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tissue is associated with increased visceral fat accumulation and impaired insulin sensitivity [82].

Indeed, many parallels between the pathogenesis of metabolic disease and the lipogenic

phenotype of cancer cells have been revealed, and FASN is certainly a common denominator

[83]. Given the prevalent co-morbidity of obesity in cancer patients and the lipogenic overlap in

the pathological progression of these two diseases, the inhibition of fatty acid synthesis becomes

an even more attractive therapeutic target for the treatment of cancer in obese patients.

IV.D CONCLUDING REMARKS

In summary, emerging evidence suggests FASN contributes to specific functions in both

normal tissue and the pathological progression of cancer. Some studies have highlighted the

importance of FASN in regulating normal physiological processes in certain tissues, whereas

others have shown that only upon a physiological challenge does the absence of FASN disturb

tissue function. This evidence suggests a low, basal expression of FASN is sufficient to maintain

homeostasis in most tissues. Under normal circumstances, the increased expression of FASN

only appears to occur in response to specific stimulation, whereby induction of FASN regulates a

homeostatic response to the stimulus. Because of this explicit regulation of de novo fatty acid

synthesis in normal tissue, and its aberrant expression in tumors, the inhibition of fatty acid

synthesis has become an attractive therapeutic target for the treatment of cancer.

Understanding the role of FASN in normal tissue, as it relates to the enzyme’s function in

responding to physiological stimulation, can guide the understanding of its aberrant activation in

cancer. The studies conducted in Chapter II demonstrate for the first time the requirement of

FASN in mammary gland development and milk fat synthesis during lactation. We found that

128

the loss of FASN led to premature involution of the gland, which truncated the lactation period

and resulted in the death of nursing pups. The fact that FASN was required for the development

and continued functional competence of the lactating mammary gland highlights the importance

of fatty acid synthesis in regulating an induced physiological process, in this case lactation,

which is a natural response to pregnancy. Females with mammary gland-specific deletion of

FASN showed no obvious phenotypic perturbations outside of the pregnancy-lactation cycle,

underscoring the regulated expression of fatty acid synthase as tissue-specific response.

The increased expression of fatty acid synthesis enzymes in cancer was recognized

decades ago and has since gained considerable popularity as an attractive pathway for drug

development. Even still, no study has attempted to conditionally delete Fasn in a cancer mouse

model to validate its potential as a drug target for cancer treatment. Chapter III of this

dissertation describes the development and characterization of a conditional, Fasn-deleted

prostate cancer mouse model. For the first time, we demonstrated, in an immunocompetent

mouse, that inhibition of FASN is an effective approach to prevent prostate cancer development

and progression. FASN-depleted prostate samples exhibited significantly reduced prostate

weight, diminished proliferative capacity, and decreased activation of AKT.

Overall, these studies emphasize the significance of de novo fatty acid synthesis,

specifically Fasn, highlighting its role in regulating both normal and pathological physiology.

Due to the unique expression profile of FASN in normal and cancer cells, the inhibition of FASN

may be an effective therapeutic approach to treating cancer. Nearly all solid tumors overexpress

FASN, and the activation of FASN is a shared characteristic of obesity, a common co-morbidity

of cancer patients. Thus, there exists a large patient population that could benefit from a FASN-

inhibiting therapeutic drug.

129

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135

CURRICULUM VITAE

Janel K. Suburu

Winston-Salem, NC

jsuburu@alumni.ucsd.edu • jsuburu@wakehealth.edu

www.linkedin.com/pub//janel-suburu/b/253/2a7/

EDUCATION

Doctor of Philosophy in Cancer Biology, 2014 (defense: April 16, 2014)

Wake Forest University Graduate School of Arts and Sciences, Winston-Salem, NC

Dissertation: Fatty Acid Synthase Modulates Mammary Gland Lactation and Prostate Cancer

Progression in vivo.

Advisor: Yong Q. Chen, Ph.D.

Bachelor of Science in Biochemistry and Cell Biology, 2006

University of California, San Diego, La Jolla, CA

PROFESSIONAL DEVELOPMENT

2013 – Present Bespoke Therapeutics, LLC

Co-Founder, Member

Created from an international business competition: The Breast Cancer

Start-Up Challenge (http://www.breastcancerstartupchallenge.com/)

Collaborating with science, business, and law students, and relevant

professionals, to launch a start-up biotech company based on intellectual

property developed at the National Cancer Institute

Performed technology evaluation, intellectual property protection analysis,

and preliminary market analyses

Actively seeking start-up funding

2013 – Present Wake Forest University, Winston-Salem, NC

Technology Commercialization Intern, Product Innovation &

Commercialization Services

Developed marketing materials for multiple university commercialization

projects (non-confidential summaries, slide decks, and marketing letters and

fliers)

Evaluated new invention disclosures, including preliminary market analysis

and prior art searches

136

Edited and reviewed non-exclusive licensing agreements for university

intellectual property

2008 – Present Wake Forest University, Winston-Salem, NC

Graduate Research Assistant, Department of Cancer Biology

Validating Fatty Acid Synthase (FASN) as a prostate cancer therapeutic

drug target by its spatiotemporal deletion in the Pten-null prostate cancer

mouse model

Demonstrated the requirement of Fatty Acid Synthase in the mammary

gland during lactation in vivo

Mentored and trained other graduate students and a post-baccalaureate

student

Taught lectures, prepared slide decks, homework assignments, and exams

for first and second year graduate students in physiology, pharmacology,

and cancer biology

2010 Targacept, Inc., Winston-Salem, NC

Summer Intern, Departments of Neurochemistry and Molecular Design

Collaborated between multiple departments and designed experiments and

test the efficacy of small molecules in growth inhibition of non-small cell

lung cancer cells

Identified potential anti-cancer agents targeting nicotinic acetylcholine

receptors through computer generated molecular modeling

Provided pilot data to guide executive decisions in the company

Awarded the Camille & Henry Dreyfus Fellowship

2006 – 2008 Veterans Medical Research Foundation, San Diego, CA

Research Associate, Department of Gastroenterology

Investigated protein signaling during disease onset and progression in a

mouse model of liver fibrosis

Provided proof of concept data for a potential peptide treatment of liver

fibrosis and cirrhosis

2005 – 2006 Torrey Pines Institute for Molecular Studies, La Jolla, CA

Research Technician, Department of Immunology

Investigated unique T-lymphocyte clones responsible for driving disease in

a mouse model of multiple sclerosis

Learned T cell receptor CDR3 spectratyping techniques, aseptic cell culture,

and techniques in murine modeling

137

PEER-REVIEWED PUBLICATIONS

1. Wang S, Wu J, Suburu J, et al. Effect of dietary polyunsaturated fatty acids on castration-

resistant pten-null prostate cancer. Carcinogenesis. 2012 Feb;33(2):404-12

2. Suburu J and Chen Y. Lipids and prostate cancer. Prostaglandins Other Lipid Mediat. 2012

May;98(1-2):1-10. Invited Review

3. Gu Z, Suburu J, Chen H, et al. Mechanisms of Omega-3 Polyunsaturated Fatty Acids in

Prostate Cancer Prevention. Biomed Res Int. 2013 (2013):824563. Invited Review.

4. Suburu J and Chen Y. “Polyunsaturated Fatty Acids and Cancer Prevention.” In

Inflammation, Oxidative Stress, and Cancer: Dietary Approaches for Cancer Prevention, ed.

A-N. T. Kong. Boca Raton, FL: Taylor and Francis Group; 2013: 299-309. Invited Book

Chapter

5. Gu Z, Wu J, Wang S, Suburu J, et al. Polyunsaturated fatty acids affect the localization and

signaling of PIP3/AKT in prostate cancer cells. Carcinogenesis. 2013 Sep;34(9):1968-75.

6. Suburu J, Gu Z, Chen H, et al. Fatty acid metabolism: Implications for diet, genetic

variation, and disease. Food Bioscience. 2013 Dec(4):1-12. Invited Review

7. Suburu J, et al. “Polyunsaturated Fatty Acid Metabolism and Cancer.” In: Polyunsaturated

Fatty Acids: Sources, Antioxidant Properties and Health Benefits, ed. A. Catalá. New York:

Nova Science Publishers; 2013: Chapter 13. Invited Book Chapter

8. Suburu J, Lim K, Calviello G, et al. RE: Serum Phospholipid Fatty Acids and Prostate

Cancer Risk in the SELECT Trial. J Natl Cancer Inst. 2014;106(4).

9. Suburu J, et al. Fatty Acid Synthase is Required for Mammary Gland Development and

Milk Production During Lactation. Am J Physiol Endocrinol Metab. 2014 Mar 25. Article in

Press.

10. Wang S, Wu J, Suburu J, et al. Elevation of Cholesterol and Fatty Acids by Abca1

Knockout and FASN Transgene Promotes Prostate Cancer Development. (Manuscript in

preparation)

11. Suburu J, et al. Fatty Acid Synthase Modulates Prostate Cancer Progression in vivo.

(Manuscript in preparation)

138

ORAL PRESENTATIONS

Janel Suburu, “Fatty Acid Synthase Modulates Mammary Gland Lactation and Prostate Cancer

Progression, in vivo.” Dissertation Defense. Cancer Biology, Wake Forest University, Winston-Salem,

NC

Annual Research Progress Reports:

“The Role of Polyunsaturated Fatty Acids in Prostate Cancer.” June 2, 2009

“Roles for de novo Fatty Acid Synthesis in Prostate Cancer.” October 30, 2009

“Fatty Acid Metabolism and Prostate Cancer.” January 4, 2011

“Fatty Acid Metabolism in Prostate Cancer.” February 14, 2012

“Fatty Acid Synthesis in the Prostate and Mammary Gland.” October 23, 2012

“Product Innovation and Commercialization Services.” Janurary 7, 2014.

Oral Presentations. Department of Cancer Biology, Wake Forest University, Winston-Salem, NC

Janel Suburu, Shihua Wang, Jiansheng Wu, Steven J. Kridel, Massimo Loda, Michael J.

Thomas, Yong Q. Chen. “Roles for de novo Fatty Acid Synthesis.” Department of Cancer

Biology Retreat, 2009. Poster Presentation. Wake Forest University, Winston-Salem, NC

Janel Suburu, Phillip Hammond, Terry Hauser. “Targeting Nicotinic Acetylcholine Receptors in

Non-Small Cell Lung Cancer.” Public Oral Presentation. Targacept Summer Internship Program.

2010. Targacept, Inc., Winston-Salem, NC.

Janel Suburu, Yong Q. Chen. “Validating Fatty Acid Synthase as a Cancer Drug Target by

Conditional Knockout In Vivo.” Oral Presentation. Department of Cancer Biology Retreat, 2011.

Wake Forest University, Winston-Salem, NC

1st Annual Maya Angelou Center for Health Equity (MACHE) Bowl. An interdisciplinary student health

care case competition, March 29, 2012. Oral Presentation. Benton Convention Center, Winston-Salem,

NC.

TEACHING AND LEADERSHIP

2012 Science Fair Mentor, Winston-Salem, NC

Forest Park Elementary School, Grades 5 and 6

2013 Cell Biology of Cancer Lecturer, Winston-Salem, NC

Wake Forest University, Graduate School of Arts and Sciences

“Cell Immortalization and Tumorigenesis”, September 17, 2013.

“The Rational Treatment of Cancer”, October 8, 2013

139

2013 Planning Committee Member, Winston-Salem, NC

TeamWS, a supporting organization for the Michael J. Fox Foundation

We are a local non-profit organization dedicated to raising awareness and

funds for Parkinson’s Disease and research.

Created and manage the website for event information and updates:

www.teamws.org

Recruited donors and sponsors from local businesses to support our lead

annual event.

Recruited volunteers and organized their participation at our lead annual

event.

2010 – 2013 Community Youth Educator, Winston-Salem, NC

Brain Awareness Council, Wake Forest University

Lead interactive learning stations for elementary, middle, and high school

students about neuroscience.

Developed objectives and lesson plans for multiple interactive neuroscience

learning stations.

2011 – 2013 Applied Human Physiology Lecturer, Winston-Salem, NC

Winston-Salem State University, Doctorate of Physical Therapy Program

“Neurons, Action Potentials, and Synaptic Transmission”, May 9, 2013.

“Blood and Hemostasis”, May 30, 2013, June 7, 2012, June 9, 2011.

2013 – 2014 Pharmacology Lecturer, Winston-Salem, NC

Winston-Salem State University, Doctorate of Physical Therapy Program

“Cancer Chemotherapy”, February 18, 2013, February 12, 2014.

“Immunosuppressive Drug Therapy”, February 20, 2013, February 12,

2014.

“Antiviral and Antifungal Drugs”, February 10, 2014.

GRADUATE COURSEWORK

Communicating Science to Lay Audiences

Learned the concepts and techniques of communicating complex scientific data and ideas to lay

audiences through professional conversation and public speaking

Drug Discovery, Design, and Development: Molecules to Medicines

Learned the process of drug target identification and validation, lead compound identification

and optimization, preclinical development, formulation, production up-scaling, clinical

development, bioethics, and marketing of a new prospective drug

140

Lead our “Pharm Team” to organize and present the various aspects of drug discovery, design,

and development on our team’s drug of choice: Gleevec (Imatinib Mesylate).

Drug Discovery VirtuaLaboratory

Learned to operate SYBYL software for receptor-based and ligand-based drug discovery

simulations

Principles of Intellectual Property Development

Learned the comprehensive process of commercializing a scientific invention, including

patenting, regulatory approval, licensing and transfer agreements, and developing a startup

company from a scientific invention

Biomimetics

Learned the concepts of biologically inspired technologies and inventions, and reviewed business

models that drove bio-inspired company products to success

SOFTWARE PROFICIENCIES

Microsoft Office: Word, Excel, PowerPoint, Outlook

Adobe: Photoshop, Illustrator, Lightroom, Acrobat

EndNote

SOCIETY MEMBERSHIPS

American Association for Cancer Research

Association of University Technology Managers

American Medical Writers Association

Wake Forest University Brain Awareness Council

Piedmont Swing Dance Society

PERSONAL INTERESTS

- Planning committee member for TeamWS, a fund raising NPO for the Michael J. Fox Foundation

- Active swing dancer and group leader in the Piedmont Swing Dance Society

- Active soccer player – Co-Captain and organizer for UCSD Woman’s Club Soccer team

(2005/2006)

- Active photographer – I enjoy landscape, family portraiture, and street photography

o Personal photography website: www.janelkathrynphotography.zenfolio.com