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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
I.F REFERENCES
1. Suburu, J. and Y.Q. Chen, Lipids and prostate cancer. Prostaglandins Other Lipid Mediat,
2012. 98(1-2): p. 1-10.
2. Mu, H. and C.E. Hoy, The digestion of dietary triacylglycerols. Prog Lipid Res, 2004.
43(2): p. 105-33.
3. Beare-Rogers, J., A. Dieffenbacher, and J.V. Holm, Lexicon of Lipid Nutrition. Pure
Appl. Chem, 2001. 73(4): p. 685-744.
4. Wassall, S.R. and W. Stillwell, Docosahexaenoic acid domains: the ultimate non-raft
membrane domain. Chem Phys Lipids, 2008. 153(1): p. 57-63.
5. Chapkin, R.S., et al., Docosahexaenoic acid alters the size and distribution of cell surface
microdomains. Biochim Biophys Acta, 2008. 1778(2): p. 466-71.
6. Rabinovich, A.L. and P.O. Ripatti, On the conformational, physical properties and
functions of polyunsaturated acyl chains. Biochim Biophys Acta, 1991. 1085(1): p. 53-
62.
7. Schneider, C., N.A. Porter, and A.R. Brash, Routes to 4-hydroxynonenal: fundamental
issues in the mechanisms of lipid peroxidation. J Biol Chem, 2008. 283(23): p. 15539-43.
8. Mead, J.R., S.A. Irvine, and D.P. Ramji, Lipoprotein lipase: structure, function,
regulation, and role in disease. J Mol Med (Berl), 2002. 80(12): p. 753-69.
9. Guillou, H., et al., The key roles of elongases and desaturases in mammalian fatty acid
metabolism: Insights from transgenic mice. Prog Lipid Res, 2010. 49(2): p. 186-99.
10. Montgomery, M.R., Characterization of fatty acid desaturase activity in rat lung
microsomes. J Lipid Res, 1976. 17(1): p. 12-5.
11. Kuhajda, F.P., et al., Fatty acid synthesis: a potential selective target for antineoplastic
therapy. Proc Natl Acad Sci U S A, 1994. 91(14): p. 6379-83.
12. Jayakumar, A., et al., Human fatty acid synthase: properties and molecular cloning. Proc
Natl Acad Sci U S A, 1995. 92(19): p. 8695-9.
13. Wakil, S.J., J.K. Stoops, and V.C. Joshi, Fatty acid synthesis and its regulation. Annu
Rev Biochem, 1983. 52: p. 537-79.
14. Zehner, Z.E., V.C. Joshi, and S.J. Wakil, Regulation of fatty acid synthetase in perinatal
chicks. Identification of polysomes synthesizing fatty acid synthetase. J Biol Chem, 1977.
252(20): p. 7015-22.
28
15. Lopez, J.M., et al., Sterol regulation of acetyl coenzyme A carboxylase: a mechanism for
coordinate control of cellular lipid. Proc Natl Acad Sci U S A, 1996. 93(3): p. 1049-53.
16. Sim, A.T. and D.G. Hardie, The low activity of acetyl-CoA carboxylase in basal and
glucagon-stimulated hepatocytes is due to phosphorylation by the AMP-activated protein
kinase and not cyclic AMP-dependent protein kinase. FEBS Lett, 1988. 233(2): p. 294-8.
17. Moore, F., J. Weekes, and D.G. Hardie, Evidence that AMP triggers phosphorylation as
well as direct allosteric activation of rat liver AMP-activated protein kinase. A sensitive
mechanism to protect the cell against ATP depletion. Eur J Biochem, 1991. 199(3): p.
691-7.
18. Abu-Elheiga, L., et al., Human acetyl-CoA carboxylase 2. Molecular cloning,
characterization, chromosomal mapping, and evidence for two isoforms. J Biol Chem,
1997. 272(16): p. 10669-77.
19. Abu-Elheiga, L., et al., The subcellular localization of acetyl-CoA carboxylase 2. Proc
Natl Acad Sci U S A, 2000. 97(4): p. 1444-9.
20. Abu-Elheiga, L., et al., Mutant mice lacking acetyl-CoA carboxylase 1 are embryonically
lethal. Proc Natl Acad Sci U S A, 2005. 102(34): p. 12011-6.
21. Choi, C.S., et al., Continuous fat oxidation in acetyl-CoA carboxylase 2 knockout mice
increases total energy expenditure, reduces fat mass, and improves insulin sensitivity.
Proc Natl Acad Sci U S A, 2007. 104(42): p. 16480-5.
22. Abu-Elheiga, L., et al., Acetyl-CoA carboxylase 2 mutant mice are protected against
obesity and diabetes induced by high-fat/high-carbohydrate diets. Proc Natl Acad Sci U S
A, 2003. 100(18): p. 10207-12.
23. Abu-Elheiga, L., et al., Continuous fatty acid oxidation and reduced fat storage in mice
lacking acetyl-CoA carboxylase 2. Science, 2001. 291(5513): p. 2613-6.
24. Abu-Elheiga, L., et al., Acetyl-CoA carboxylase 2-/- mutant mice are protected against
fatty liver under high-fat, high-carbohydrate dietary and de novo lipogenic conditions. J
Biol Chem, 2012. 287(15): p. 12578-88.
25. Mao, J., et al., Liver-specific deletion of acetyl-CoA carboxylase 1 reduces hepatic
triglyceride accumulation without affecting glucose homeostasis. Proc Natl Acad Sci U S
A, 2006. 103(22): p. 8552-7.
26. Mao, J., et al., aP2-Cre-mediated inactivation of acetyl-CoA carboxylase 1 causes growth
retardation and reduced lipid accumulation in adipose tissues. Proc Natl Acad Sci U S A,
2009. 106(41): p. 17576-81.
29
27. Harada, N., et al., Hepatic de novo lipogenesis is present in liver-specific ACC1-deficient
mice. Mol Cell Biol, 2007. 27(5): p. 1881-8.
28. Chirala, S.S., et al., Animal fatty acid synthase: functional mapping and cloning and
expression of the domain I constituent activities. Proc Natl Acad Sci U S A, 1997.
94(11): p. 5588-93.
29. Semenkovich, C.F., Regulation of fatty acid synthase (FAS). Prog Lipid Res, 1997.
36(1): p. 43-53.
30. Stoops, J.K. and S.J. Wakil, Yeast fatty acid synthetase: structure-function relationship
and nature of the beta-ketoacyl synthetase site. Proc Natl Acad Sci U S A, 1980. 77(8): p.
4544-8.
31. Rangan, V.S., A.K. Joshi, and S. Smith, Mapping the functional topology of the animal
fatty acid synthase by mutant complementation in vitro. Biochemistry, 2001. 40(36): p.
10792-9.
32. Witkowski, A., et al., Head-to-head coiled arrangement of the subunits of the animal fatty
acid synthase. Chem Biol, 2004. 11(12): p. 1667-76.
33. Maier, T., M. Leibundgut, and N. Ban, The crystal structure of a mammalian fatty acid
synthase. Science, 2008. 321(5894): p. 1315-22.
34. Ploskon, E., et al., A mammalian type I fatty acid synthase acyl carrier protein domain
does not sequester acyl chains. J Biol Chem, 2008. 283(1): p. 518-28.
35. Nguyen, C., et al., Trapping the dynamic acyl carrier protein in fatty acid biosynthesis.
Nature, 2014. 505(7483): p. 427-31.
36. Hsu, M.H., S.S. Chirala, and S.J. Wakil, Human fatty-acid synthase gene. Evidence for
the presence of two promoters and their functional interaction. J Biol Chem, 1996.
271(23): p. 13584-92.
37. Fukuda, H., et al., Transcriptional regulation of fatty acid synthase gene by
insulin/glucose, polyunsaturated fatty acid and leptin in hepatocytes and adipocytes in
normal and genetically obese rats. Eur J Biochem, 1999. 260(2): p. 505-11.
38. Blake, W.L. and S.D. Clarke, Suppression of rat hepatic fatty acid synthase and S14 gene
transcription by dietary polyunsaturated fat. J Nutr, 1990. 120(12): p. 1727-9.
39. Kersten, S., Mechanisms of nutritional and hormonal regulation of lipogenesis. EMBO
Rep, 2001. 2(4): p. 282-6.
30
40. Yin, D., et al., Somatotropin-dependent decrease in fatty acid synthase mRNA abundance
in 3T3-F442A adipocytes is the result of a decrease in both gene transcription and mRNA
stability. Biochem J, 1998. 331 ( Pt 3): p. 815-20.
41. Dudek, S.M. and C.F. Semenkovich, Essential amino acids regulate fatty acid synthase
expression through an uncharged transfer RNA-dependent mechanism. J Biol Chem,
1995. 270(49): p. 29323-9.
42. Soncini, M., et al., Hormonal and nutritional control of the fatty acid synthase promoter
in transgenic mice. J Biol Chem, 1995. 270(51): p. 30339-43.
43. Fukuda, H., N. Iritani, and T. Noguchi, Transcriptional regulatory regions for expression
of the rat fatty acid synthase. FEBS Lett, 1997. 406(3): p. 243-8.
44. Roder, K., et al., Cooperative binding of NF-Y and Sp1 at the DNase I-hypersensitive
site, fatty acid synthase insulin-responsive element 1, located at -500 in the rat fatty acid
synthase promoter. J Biol Chem, 1997. 272(34): p. 21616-24.
45. Wang, D. and H.S. Sul, Upstream stimulatory factors bind to insulin response sequence
of the fatty acid synthase promoter. USF1 is regulated. J Biol Chem, 1995. 270(48): p.
28716-22.
46. Dang, C.V., MYC on the path to cancer. Cell, 2012. 149(1): p. 22-35.
47. Oskouian, B., V.S. Rangan, and S. Smith, Regulatory elements in the first intron of the
rat fatty acid synthase gene. Biochem J, 1997. 324 ( Pt 1): p. 113-21.
48. Xu, Z.X. and S.A. Rooney, Glucocorticoids increase fatty-acid synthase mRNA stability
in fetal rat lung. Am J Physiol, 1997. 272(5 Pt 1): p. L860-4.
49. Graner, E., et al., The isopeptidase USP2a regulates the stability of fatty acid synthase in
prostate cancer. Cancer Cell, 2004. 5(3): p. 253-61.
50. Semenkovich, C.F., T. Coleman, and R. Goforth, Physiologic concentrations of glucose
regulate fatty acid synthase activity in HepG2 cells by mediating fatty acid synthase
mRNA stability. J Biol Chem, 1993. 268(10): p. 6961-70.
51. Jin, Q., et al., Fatty acid synthase phosphorylation: a novel therapeutic target in HER2-
overexpressing breast cancer cells. Breast Cancer Res, 2010. 12(6): p. R96.
52. An, Z., et al., Nicotine-induced activation of AMP-activated protein kinase inhibits fatty
acid synthase in 3T3L1 adipocytes: a role for oxidant stress. J Biol Chem, 2007. 282(37):
p. 26793-801.
53. Jensen-Urstad, A.P., et al., Nutrient-dependent phosphorylation channels lipid synthesis
to regulate PPARalpha. J Lipid Res, 2013. 54(7): p. 1848-59.
31
54. Zhao, S., et al., Regulation of cellular metabolism by protein lysine acetylation. Science,
2010. 327(5968): p. 1000-4.
55. Semenkovich, C.F., T. Coleman, and F.T. Fiedorek, Jr., Human fatty acid synthase
mRNA: tissue distribution, genetic mapping, and kinetics of decay after glucose
deprivation. J Lipid Res, 1995. 36(7): p. 1507-21.
56. Kusakabe, T., et al., Fatty acid synthase is expressed mainly in adult hormone-sensitive
cells or cells with high lipid metabolism and in proliferating fetal cells. J Histochem
Cytochem, 2000. 48(5): p. 613-22.
57. Buechler, K.F. and R.A. Rhoades, Fatty acid synthesis in the perfused rat lung. Biochim
Biophys Acta, 1980. 619(2): p. 186-95.
58. Chakravarthy, M.V., et al., Brain fatty acid synthase activates PPARalpha to maintain
energy homeostasis. J Clin Invest, 2007. 117(9): p. 2539-52.
59. Pizer, E.S., et al., Expression of fatty acid synthase is closely linked to proliferation and
stromal decidualization in cycling endometrium. Int J Gynecol Pathol, 1997. 16(1): p. 45-
51.
60. Menendez, J.A. and R. Lupu, Fatty acid synthase and the lipogenic phenotype in cancer
pathogenesis. Nat Rev Cancer, 2007. 7(10): p. 763-77.
61. Chirala, S.S., et al., Fatty acid synthesis is essential in embryonic development: fatty acid
synthase null mutants and most of the heterozygotes die in utero. Proc Natl Acad Sci U S
A, 2003. 100(11): p. 6358-63.
62. Chakravarthy, M.V., et al., "New" hepatic fat activates PPARalpha to maintain glucose,
lipid, and cholesterol homeostasis. Cell Metab, 2005. 1(5): p. 309-22.
63. Razani, B., et al., Fatty acid synthase modulates homeostatic responses to myocardial
stress. J Biol Chem, 2011. 286(35): p. 30949-61.
64. Wei, X., et al., De novo lipogenesis maintains vascular homeostasis through endothelial
nitric-oxide synthase (eNOS) palmitoylation. J Biol Chem, 2011. 286(4): p. 2933-45.
65. Wei, X., et al., Fatty acid synthase modulates intestinal barrier function through
palmitoylation of mucin 2. Cell Host Microbe, 2012. 11(2): p. 140-52.
66. Funai, K., et al., Muscle lipogenesis balances insulin sensitivity and strength through
calcium signaling. J Clin Invest, 2013. 123(3): p. 1229-40.
32
67. Lodhi, I.J., et al., Inhibiting adipose tissue lipogenesis reprograms thermogenesis and
PPARgamma activation to decrease diet-induced obesity. Cell Metab, 2012. 16(2): p.
189-201.
68. Schneider, J.G., et al., Macrophage fatty-acid synthase deficiency decreases diet-induced
atherosclerosis. J Biol Chem, 2010. 285(30): p. 23398-409.
69. Leone, T.C., C.J. Weinheimer, and D.P. Kelly, A critical role for the peroxisome
proliferator-activated receptor alpha (PPARalpha) in the cellular fasting response: the
PPARalpha-null mouse as a model of fatty acid oxidation disorders. Proc Natl Acad Sci
U S A, 1999. 96(13): p. 7473-8.
70. Chakravarthy, M.V., et al., Identification of a physiologically relevant endogenous ligand
for PPARalpha in liver. Cell, 2009. 138(3): p. 476-88.
71. Lin, X., et al., Dysregulation of insulin receptor substrate 2 in beta cells and brain causes
obesity and diabetes. J Clin Invest, 2004. 114(7): p. 908-16.
72. Chakravarthy, M.V., et al., Inactivation of hypothalamic FAS protects mice from diet-
induced obesity and inflammation. J Lipid Res, 2009. 50(4): p. 630-40.
73. Johansson, M.E., J.M. Larsson, and G.C. Hansson, The two mucus layers of colon are
organized by the MUC2 mucin, whereas the outer layer is a legislator of host-microbial
interactions. Proc Natl Acad Sci U S A, 2011. 108 Suppl 1: p. 4659-65.
74. Finck, B.N., et al., A potential link between muscle peroxisome proliferator- activated
receptor-alpha signaling and obesity-related diabetes. Cell Metab, 2005. 1(2): p. 133-44.
75. Steinberg, G.R., et al., Tumor necrosis factor alpha-induced skeletal muscle insulin
resistance involves suppression of AMP-kinase signaling. Cell Metab, 2006. 4(6): p. 465-
74.
76. Coupe, C., et al., Lipogenic enzyme activities and mRNA in rat adipose tissue at
weaning. Am J Physiol, 1990. 258(1 Pt 1): p. E126-33.
77. Tontonoz, P., et al., mPPAR gamma 2: tissue-specific regulator of an adipocyte enhancer.
Genes Dev, 1994. 8(10): p. 1224-34.
78. Sears, I.B., et al., Differentiation-dependent expression of the brown adipocyte
uncoupling protein gene: regulation by peroxisome proliferator-activated receptor
gamma. Mol Cell Biol, 1996. 16(7): p. 3410-9.
79. Warburg, O., F. Wind, and E. Negelein, The Metabolism of Tumors in the Body. J Gen
Physiol, 1927. 8(6): p. 519-30.
33
80. Wilson, J.E., Isozymes of mammalian hexokinase: structure, subcellular localization and
metabolic function. J Exp Biol, 2003. 206(Pt 12): p. 2049-57.
81. Mathupala, S.P., Y.H. Ko, and P.L. Pedersen, Hexokinase II: cancer's double-edged
sword acting as both facilitator and gatekeeper of malignancy when bound to
mitochondria. Oncogene, 2006. 25(34): p. 4777-86.
82. Moon, J.S., et al., Androgen stimulates glycolysis for de novo lipid synthesis by
increasing the activities of hexokinase 2 and 6-phosphofructo-2-kinase/fructose-2,6-
bisphosphatase 2 in prostate cancer cells. Biochem J, 2011. 433(1): p. 225-33.
83. Mazurek, S., et al., Pyruvate kinase type M2 and its role in tumor growth and spreading.
Semin Cancer Biol, 2005. 15(4): p. 300-8.
84. Eigenbrodt, E., et al., Double role for pyruvate kinase type M2 in the expansion of
phosphometabolite pools found in tumor cells. Crit Rev Oncog, 1992. 3(1-2): p. 91-115.
85. Christofk, H.R., et al., The M2 splice isoform of pyruvate kinase is important for cancer
metabolism and tumour growth. Nature, 2008. 452(7184): p. 230-3.
86. Anastasiou, D., et al., Inhibition of pyruvate kinase M2 by reactive oxygen species
contributes to cellular antioxidant responses. Science, 2011. 334(6060): p. 1278-83.
87. Vander Heiden, M.G., et al., Evidence for an alternative glycolytic pathway in rapidly
proliferating cells. Science, 2010. 329(5998): p. 1492-9.
88. Koh, H.J., et al., Cytosolic NADP+-dependent isocitrate dehydrogenase plays a key role
in lipid metabolism. J Biol Chem, 2004. 279(38): p. 39968-74.
89. Reitman, Z.J. and H. Yan, Isocitrate dehydrogenase 1 and 2 mutations in cancer:
alterations at a crossroads of cellular metabolism. J Natl Cancer Inst, 2010. 102(13): p.
932-41.
90. Murugan, A.K., E. Bojdani, and M. Xing, Identification and functional characterization
of isocitrate dehydrogenase 1 (IDH1) mutations in thyroid cancer. Biochem Biophys Res
Commun, 2010. 393(3): p. 555-9.
91. Mycielska, M.E., et al., Citrate transport and metabolism in mammalian cells: prostate
epithelial cells and prostate cancer. Bioessays, 2009. 31(1): p. 10-20.
92. Kavanagh, J.P., Isocitric and citric acid in human prostatic and seminal fluid:
implications for prostatic metabolism and secretion. Prostate, 1994. 24(3): p. 139-42.
93. Medrano, A., et al., Utilization of citrate and lactate through a lactate dehydrogenase and
ATP-regulated pathway in boar spermatozoa. Mol Reprod Dev, 2006. 73(3): p. 369-78.
34
94. Mycielska, M.E., et al., Citrate enhances in vitro metastatic behaviours of PC-3M human
prostate cancer cells: status of endogenous citrate and dependence on aconitase and fatty
acid synthase. Int J Biochem Cell Biol, 2006. 38(10): p. 1766-77.
95. Hatzivassiliou, G., et al., ATP citrate lyase inhibition can suppress tumor cell growth.
Cancer Cell, 2005. 8(4): p. 311-21.
96. DeBerardinis, R.J., et al., Beyond aerobic glycolysis: transformed cells can engage in
glutamine metabolism that exceeds the requirement for protein and nucleotide synthesis.
Proc Natl Acad Sci U S A, 2007. 104(49): p. 19345-50.
97. Liu, X., Y.M. Fu, and G.G. Meadows, Differential effects of specific amino acid
restriction on glucose metabolism, reduction/oxidation status and mitochondrial damage
in DU145 and PC3 prostate cancer cells. Oncol Lett, 2011. 2(2): p. 349-355.
98. Mullen, A.R., et al., Reductive carboxylation supports growth in tumour cells with
defective mitochondria. Nature, 2012. 481(7381): p. 385-8.
99. Metallo, C.M., et al., Reductive glutamine metabolism by IDH1 mediates lipogenesis
under hypoxia. Nature, 2012. 481(7381): p. 380-4.
100. Medes, G., A. Thomas, and S. Weinhouse, Metabolism of neoplastic tissue. IV. A study
of lipid synthesis in neoplastic tissue slices in vitro. Cancer Res, 1953. 13(1): p. 27-9.
101. Liang, G., et al., Diminished hepatic response to fasting/refeeding and liver X receptor
agonists in mice with selective deficiency of sterol regulatory element-binding protein-
1c. J Biol Chem, 2002. 277(11): p. 9520-8.
102. Swinnen, J.V., et al., Androgens stimulate fatty acid synthase in the human prostate
cancer cell line LNCaP. Cancer Res, 1997. 57(6): p. 1086-90.
103. Swinnen, J.V., et al., Stimulation of tumor-associated fatty acid synthase expression by
growth factor activation of the sterol regulatory element-binding protein pathway.
Oncogene, 2000. 19(45): p. 5173-81.
104. Huang, W.C., et al., Activation of Androgen Receptor, Lipogenesis, and Oxidative Stress
Converged by SREBP-1 Is Responsible for Regulating Growth and Progression of
Prostate Cancer Cells. Mol Cancer Res, 2012. 10(1): p. 133-42.
105. Swinnen, J.V., et al., Overexpression of fatty acid synthase is an early and common event
in the development of prostate cancer. Int J Cancer, 2002. 98(1): p. 19-22.
106. Milgraum, L.Z., et al., Enzymes of the fatty acid synthesis pathway are highly expressed
in in situ breast carcinoma. Clin Cancer Res, 1997. 3(11): p. 2115-20.
35
107. Gansler, T.S., et al., Increased expression of fatty acid synthase (OA-519) in ovarian
neoplasms predicts shorter survival. Hum Pathol, 1997. 28(6): p. 686-92.
108. Rashid, A., et al., Elevated expression of fatty acid synthase and fatty acid synthetic
activity in colorectal neoplasia. Am J Pathol, 1997. 150(1): p. 201-8.
109. Kuhajda, F.P., Fatty-acid synthase and human cancer: new perspectives on its role in
tumor biology. Nutrition, 2000. 16(3): p. 202-8.
110. Swinnen, J.V., K. Brusselmans, and G. Verhoeven, Increased lipogenesis in cancer cells:
new players, novel targets. Curr Opin Clin Nutr Metab Care, 2006. 9(4): p. 358-65.
111. Fritz, V., et al., Abrogation of de novo lipogenesis by stearoyl-CoA desaturase 1
inhibition interferes with oncogenic signaling and blocks prostate cancer progression in
mice. Mol Cancer Ther, 2010. 9(6): p. 1740-54.
112. Kridel, S.J., et al., Orlistat is a novel inhibitor of fatty acid synthase with antitumor
activity. Cancer Res, 2004. 64(6): p. 2070-5.
113. Swinnen, J.V., et al., Selective activation of the fatty acid synthesis pathway in human
prostate cancer. Int J Cancer, 2000. 88(2): p. 176-9.
114. Brusselmans, K., et al., RNA interference-mediated silencing of the acetyl-CoA-
carboxylase-alpha gene induces growth inhibition and apoptosis of prostate cancer cells.
Cancer Res, 2005. 65(15): p. 6719-25.
115. De Schrijver, E., et al., RNA interference-mediated silencing of the fatty acid synthase
gene attenuates growth and induces morphological changes and apoptosis of LNCaP
prostate cancer cells. Cancer Res, 2003. 63(13): p. 3799-804.
116. Beckers, A., et al., Chemical inhibition of acetyl-CoA carboxylase induces growth arrest
and cytotoxicity selectively in cancer cells. Cancer Res, 2007. 67(17): p. 8180-7.
117. Shah, U.S., et al., Fatty acid synthase gene overexpression and copy number gain in
prostate adenocarcinoma. Hum Pathol, 2006. 37(4): p. 401-9.
118. Wang, S., et al., Prostate-specific deletion of the murine Pten tumor suppressor gene
leads to metastatic prostate cancer. Cancer Cell, 2003. 4(3): p. 209-21.
119. Pearson, H.B., et al., Lkb1 deficiency causes prostate neoplasia in the mouse. Cancer
Res, 2008. 68(7): p. 2223-32.
120. Van de Sande, T., et al., High-level expression of fatty acid synthase in human prostate
cancer tissues is linked to activation and nuclear localization of Akt/PKB. J Pathol, 2005.
206(2): p. 214-9.
36
121. Migita, T., et al., Fatty acid synthase: a metabolic enzyme and candidate oncogene in
prostate cancer. J Natl Cancer Inst, 2009. 101(7): p. 519-32.
122. Swinnen, J.V., et al., Fatty acid synthase drives the synthesis of phospholipids
partitioning into detergent-resistant membrane microdomains. Biochem Biophys Res
Commun, 2003. 302(4): p. 898-903.
123. Zheng, H., et al., Lipid-dependent gating of a voltage-gated potassium channel. Nat
Commun, 2011. 2: p. 250.
124. Alli, P.M., et al., Fatty acid synthase inhibitors are chemopreventive for mammary cancer
in neu-N transgenic mice. Oncogene, 2005. 24(1): p. 39-46.
125. Rysman, E., et al., De novo lipogenesis protects cancer cells from free radicals and
chemotherapeutics by promoting membrane lipid saturation. Cancer Res, 2011. 70(20): p.
8117-26.
126. Fiorentino, M., et al., Overexpression of fatty acid synthase is associated with
palmitoylation of Wnt1 and cytoplasmic stabilization of beta-catenin in prostate cancer.
Lab Invest, 2008. 88(12): p. 1340-8.
127. Zhou, W., et al., Fatty acid synthase inhibition triggers apoptosis during S phase in
human cancer cells. Cancer Res, 2003. 63(21): p. 7330-7.
128. Manning, B.D. and L.C. Cantley, AKT/PKB signaling: navigating downstream. Cell,
2007. 129(7): p. 1261-74.
129. Gray, I.C., et al., Mutation and expression analysis of the putative prostate tumour-
suppressor gene PTEN. Br J Cancer, 1998. 78(10): p. 1296-300.
130. Li, Y., et al., PTEN deletion and heme oxygenase-1 overexpression cooperate in prostate
cancer progression and are associated with adverse clinical outcome. J Pathol, 2011.
224(1): p. 90-100.
131. Friedlander, T.W., et al., Common structural and epigenetic changes in the genome of
castration-resistant prostate cancer. Cancer Res, 2012. 72(3): p. 616-25.
132. Chalhoub, N. and S.J. Baker, PTEN and the PI3-kinase pathway in cancer. Annu Rev
Pathol, 2009. 4: p. 127-50.
133. Van de Sande, T., et al., Role of the phosphatidylinositol 3'-kinase/PTEN/Akt kinase
pathway in the overexpression of fatty acid synthase in LNCaP prostate cancer cells.
Cancer Res, 2002. 62(3): p. 642-6.
134. Wang, H.Q., et al., Positive feedback regulation between AKT activation and fatty acid
synthase expression in ovarian carcinoma cells. Oncogene, 2005. 24(22): p. 3574-82.
37
135. Tomek, K., et al., Blockade of fatty acid synthase induces ubiquitination and degradation
of phosphoinositide-3-kinase signaling proteins in ovarian cancer. Mol Cancer Res, 2011.
9(12): p. 1767-79.
136. Higgins, M.J. and J. Baselga, Targeted therapies for breast cancer. J Clin Invest, 2011.
121(10): p. 3797-803.
137. Minner, S., et al., Low level HER2 overexpression is associated with rapid tumor cell
proliferation and poor prognosis in prostate cancer. Clin Cancer Res, 2010. 16(5): p.
1553-60.
138. Kumar-Sinha, C., et al., Transcriptome analysis of HER2 reveals a molecular connection
to fatty acid synthesis. Cancer Res, 2003. 63(1): p. 132-9.
139. Yoon, S., et al., Up-regulation of acetyl-CoA carboxylase alpha and fatty acid synthase
by human epidermal growth factor receptor 2 at the translational level in breast cancer
cells. J Biol Chem, 2007. 282(36): p. 26122-31.
140. Kumar, B., et al., Oxidative stress is inherent in prostate cancer cells and is required for
aggressive phenotype. Cancer Res, 2008. 68(6): p. 1777-85.
141. Shiota, M., et al., Castration resistance of prostate cancer cells caused by castration-
induced oxidative stress through Twist1 and androgen receptor overexpression.
Oncogene, 2010. 29(2): p. 237-50.
142. Shiota, M., A. Yokomizo, and S. Naito, Oxidative stress and androgen receptor signaling
in the development and progression of castration-resistant prostate cancer. Free Radic
Biol Med, 2011. 51(7): p. 1320-8.
143. Clerkin, J.S., et al., Mechanisms of ROS modulated cell survival during carcinogenesis.
Cancer Lett, 2008. 266(1): p. 30-6.
144. Furuta, E., et al., Fatty acid synthase gene is up-regulated by hypoxia via activation of
Akt and sterol regulatory element binding protein-1. Cancer Res, 2008. 68(4): p. 1003-
11.
145. Brar, S.S., et al., NOX5 NAD(P)H oxidase regulates growth and apoptosis in DU 145
prostate cancer cells. Am J Physiol Cell Physiol, 2003. 285(2): p. C353-69.
146. Huang, W.C., H.E. Zhau, and L.W. Chung, Androgen receptor survival signaling is
blocked by anti-beta2-microglobulin monoclonal antibody via a MAPK/lipogenic
pathway in human prostate cancer cells. J Biol Chem, 2010. 285(11): p. 7947-56.
38
147. Yang, X., et al., Novel membrane-associated androgen receptor splice variant potentiates
proliferative and survival responses in prostate cancer cells. J Biol Chem, 2011. 286(41):
p. 36152-60.
148. Kim, S.J., et al., Stearoyl CoA desaturase (SCD) facilitates proliferation of prostate
cancer cells through enhancement of androgen receptor transactivation. Mol Cells, 2011.
31(4): p. 371-7.
149. Guseva, N.V., et al., TOFA (5-tetradecyl-oxy-2-furoic acid) reduces fatty acid synthesis,
inhibits expression of AR, neuropilin-1 and Mcl-1 and kills prostate cancer cells
independent of p53 status. Cancer Biol Ther, 2011. 12(1): p. 80-5.
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
II.F REFERENCES
1. Martin, D.B., M.G. Horning, and P.R. Vagelos, Fatty acid synthesis in adipose tissue. I.
Purification and properties of a long chain fatty acid-synthesizing system. J Biol Chem,
1961. 236: p. 663-8.
2. Jayakumar, A., et al., Human fatty acid synthase: properties and molecular cloning. Proc
Natl Acad Sci U S A, 1995. 92(19): p. 8695-9.
3. Semenkovich, C.F., T. Coleman, and F.T. Fiedorek, Jr., Human fatty acid synthase
mRNA: tissue distribution, genetic mapping, and kinetics of decay after glucose
deprivation. J Lipid Res, 1995. 36(7): p. 1507-21.
4. Chirala, S.S., et al., Fatty acid synthesis is essential in embryonic development: fatty acid
synthase null mutants and most of the heterozygotes die in utero. Proc Natl Acad Sci U S
A, 2003. 100(11): p. 6358-63.
5. Chakravarthy, M.V., et al., "New" hepatic fat activates PPARalpha to maintain glucose,
lipid, and cholesterol homeostasis. Cell Metab, 2005. 1(5): p. 309-22.
6. Chakravarthy, M.V., et al., Brain fatty acid synthase activates PPARalpha to maintain
energy homeostasis. J Clin Invest, 2007. 117(9): p. 2539-52.
7. Schneider, J.G., et al., Macrophage fatty-acid synthase deficiency decreases diet-induced
atherosclerosis. J Biol Chem, 2010. 285(30): p. 23398-409.
8. Razani, B., et al., Fatty acid synthase modulates homeostatic responses to myocardial
stress. J Biol Chem, 2011. 286(35): p. 30949-61.
9. Wei, X., et al., Fatty acid synthase modulates intestinal barrier function through
palmitoylation of mucin 2. Cell Host Microbe, 2012. 11(2): p. 140-52.
10. Lodhi, I.J., et al., Inhibiting adipose tissue lipogenesis reprograms thermogenesis and
PPARgamma activation to decrease diet-induced obesity. Cell Metab, 2012. 16(2): p.
189-201.
11. Menendez, J.A. and R. Lupu, Fatty acid synthase and the lipogenic phenotype in cancer
pathogenesis. Nat Rev Cancer, 2007. 7(10): p. 763-77.
12. Rudolph, M.C., et al., Functional development of the mammary gland: use of expression
profiling and trajectory clustering to reveal changes in gene expression during pregnancy,
lactation, and involution. J Mammary Gland Biol Neoplasia, 2003. 8(3): p. 287-307.
13. Rudolph, M.C., et al., Metabolic regulation in the lactating mammary gland: a lipid
synthesizing machine. Physiol Genomics, 2007. 28(3): p. 323-36.
66
14. Rudolph, M.C., M.C. Neville, and S.M. Anderson, Lipid synthesis in lactation: diet and
the fatty acid switch. J Mammary Gland Biol Neoplasia, 2007. 12(4): p. 269-81.
15. Anderson, S.M., et al., Key stages in mammary gland development. Secretory activation
in the mammary gland: it's not just about milk protein synthesis! Breast Cancer Res,
2007. 9(1): p. 204.
16. Smith, S., et al., The effect of dietary fat on lipogenesis in mammary gland and liver from
lactating and virgin mice. Biochem J, 1969. 115(4): p. 807-15.
17. Libertini, L.J. and S. Smith, Purification and properties of a thioesterase from lactating rat
mammary gland which modifies the product specificity of fatty acid synthetase. J Biol
Chem, 1978. 253(5): p. 1393-401.
18. Bartley, J.C., J.T. Emerman, and M.J. Bissell, Metabolic cooperativity between epithelial
cells and adipocytes of mice. Am J Physiol, 1981. 241(5): p. C204-8.
19. Landskroner-Eiger, S., et al., Morphogenesis of the developing mammary gland: stage-
dependent impact of adipocytes. Dev Biol, 2010. 344(2): p. 968-78.
20. Vernon, R.G. and D.J. Flint, Control of fatty acid synthesis in lactation. Proc Nutr Soc,
1983. 42(2): p. 315-31.
21. Smith, S. and R. Dils, Factors affecting the chain length of fatty acids synthesised by
lactating-rabbit mammary glands. Biochim Biophys Acta, 1966. 116(1): p. 23-40.
22. Liang, G., et al., Diminished hepatic response to fasting/refeeding and liver X receptor
agonists in mice with selective deficiency of sterol regulatory element-binding protein-
1c. J Biol Chem, 2002. 277(11): p. 9520-8.
23. Rudolph, M.C., et al., Sterol regulatory element binding protein and dietary lipid
regulation of fatty acid synthesis in the mammary epithelium. Am J Physiol Endocrinol
Metab, 2010. 299(6): p. E918-27.
24. Wagner, K.U., et al., Cre-mediated gene deletion in the mammary gland. Nucleic Acids
Res, 1997. 25(21): p. 4323-30.
25. Berquin, I.M., et al., Modulation of prostate cancer genetic risk by omega-3 and omega-6
fatty acids. J Clin Invest, 2007. 117(7): p. 1866-75.
26. Smith, S., Mechanism of chain length determination in biosynthesis of milk fatty acids. J
Dairy Sci, 1980. 63(2): p. 337-52.
27. Lund, L.R., et al., Two distinct phases of apoptosis in mammary gland involution:
proteinase-independent and -dependent pathways. Development, 1996. 122(1): p. 181-93.
67
28. Wagner, K.U., et al., An adjunct mammary epithelial cell population in parous females:
its role in functional adaptation and tissue renewal. Development, 2002. 129(6): p. 1377-
86.
29. Wagner, K.U., et al., Impaired alveologenesis and maintenance of secretory mammary
epithelial cells in Jak2 conditional knockout mice. Mol Cell Biol, 2004. 24(12): p. 5510-
20.
30. Feng, Y., et al., Estrogen receptor-alpha expression in the mammary epithelium is
required for ductal and alveolar morphogenesis in mice. Proc Natl Acad Sci U S A, 2007.
104(37): p. 14718-23.
31. Wagh, P.K., et al., Conditional deletion of beta-catenin in mammary epithelial cells of
Ron receptor, Mst1r, overexpressing mice alters mammary tumorigenesis.
Endocrinology, 2012. 153(6): p. 2735-46.
32. Tomek, K., et al., Blockade of fatty acid synthase induces ubiquitination and degradation
of phosphoinositide-3-kinase signaling proteins in ovarian cancer. Mol Cancer Res, 2011.
9(12): p. 1767-79.
33. Grunt, T.W., et al., Interaction between fatty acid synthase- and ErbB-systems in ovarian
cancer cells. Biochem Biophys Res Commun, 2009. 385(3): p. 454-9.
34. Jones, F.E. and D.F. Stern, Expression of dominant-negative ErbB2 in the mammary
gland of transgenic mice reveals a role in lobuloalveolar development and lactation.
Oncogene, 1999. 18(23): p. 3481-90.
35. Neville, M.C. and M.F. Picciano, Regulation of milk lipid secretion and composition.
Annu Rev Nutr, 1997. 17: p. 159-83.
36. Lauritzen, L., et al., The essentiality of long chain n-3 fatty acids in relation to
development and function of the brain and retina. Prog Lipid Res, 2001. 40(1-2): p. 1-94.
37. Chakravarthy, M.V., et al., Identification of a physiologically relevant endogenous ligand
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
Commun, 2003. 302(4): p. 898-903.
39. Wakil, S.J., J.K. Stoops, and V.C. Joshi, Fatty acid synthesis and its regulation. Annu
Rev Biochem, 1983. 52: p. 537-79.
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
III.F REFERENCES
1. Siegel, R., et al., Cancer statistics, 2014. CA Cancer J Clin, 2014. 64(1): p. 9-29.
2. Vander Heiden, M.G., L.C. Cantley, and C.B. Thompson, Understanding the Warburg
effect: the metabolic requirements of cell proliferation. Science, 2009. 324(5930): p.
1029-33.
3. Swinnen, J.V., et al., Selective activation of the fatty acid synthesis pathway in human
prostate cancer. Int J Cancer, 2000. 88(2): p. 176-9.
4. Milgraum, L.Z., et al., Enzymes of the fatty acid synthesis pathway are highly expressed
in in situ breast carcinoma. Clin Cancer Res, 1997. 3(11): p. 2115-20.
5. Swinnen, J.V., et al., Androgens stimulate fatty acid synthase in the human prostate
cancer cell line LNCaP. Cancer Res, 1997. 57(6): p. 1086-90.
6. Swinnen, J.V., et al., Stimulation of tumor-associated fatty acid synthase expression by
growth factor activation of the sterol regulatory element-binding protein pathway.
Oncogene, 2000. 19(45): p. 5173-81.
7. Swinnen, J.V., et al., Overexpression of fatty acid synthase is an early and common event
in the development of prostate cancer. Int J Cancer, 2002. 98(1): p. 19-22.
8. Shurbaji, M.S., J.H. Kalbfleisch, and T.S. Thurmond, Immunohistochemical detection of
a fatty acid synthase (OA-519) as a predictor of progression of prostate cancer. Hum
Pathol, 1996. 27(9): p. 917-21.
9. Huang, W.C., et al., Activation of Androgen Receptor, Lipogenesis, and Oxidative Stress
Converged by SREBP-1 Is Responsible for Regulating Growth and Progression of
Prostate Cancer Cells. Mol Cancer Res, 2012. 10(1): p. 133-42.
10. Migita, T., et al., Fatty acid synthase: a metabolic enzyme and candidate oncogene in
prostate cancer. J Natl Cancer Inst, 2009. 101(7): p. 519-32.
11. Swinnen, J.V., et al., Fatty acid synthase drives the synthesis of phospholipids
partitioning into detergent-resistant membrane microdomains. Biochem Biophys Res
Commun, 2003. 302(4): p. 898-903.
12. Hatzivassiliou, G., et al., ATP citrate lyase inhibition can suppress tumor cell growth.
Cancer Cell, 2005. 8(4): p. 311-21.
13. Kridel, S.J., et al., Orlistat is a novel inhibitor of fatty acid synthase with antitumor
activity. Cancer Res, 2004. 64(6): p. 2070-5.
101
14. De Schrijver, E., et al., RNA interference-mediated silencing of the fatty acid synthase
gene attenuates growth and induces morphological changes and apoptosis of LNCaP
prostate cancer cells. Cancer Res, 2003. 63(13): p. 3799-804.
15. Beckers, A., et al., Chemical inhibition of acetyl-CoA carboxylase induces growth arrest
and cytotoxicity selectively in cancer cells. Cancer Res, 2007. 67(17): p. 8180-7.
16. Taylor, B.S., et al., Integrative genomic profiling of human prostate cancer. Cancer Cell,
2010. 18(1): p. 11-22.
17. Yoshimoto, M., et al., Interphase FISH analysis of PTEN in histologic sections shows
genomic deletions in 68% of primary prostate cancer and 23% of high-grade prostatic
intra-epithelial neoplasias. Cancer Genet Cytogenet, 2006. 169(2): p. 128-37.
18. Friedlander, T.W., et al., Common structural and epigenetic changes in the genome of
castration-resistant prostate cancer. Cancer Res, 2012. 72(3): p. 616-25.
19. Van de Sande, T., et al., Role of the phosphatidylinositol 3'-kinase/PTEN/Akt kinase
pathway in the overexpression of fatty acid synthase in LNCaP prostate cancer cells.
Cancer Res, 2002. 62(3): p. 642-6.
20. Wang, S., et al., Prostate-specific deletion of the murine Pten tumor suppressor gene
leads to metastatic prostate cancer. Cancer Cell, 2003. 4(3): p. 209-21.
21. Ittmann, M., et al., Animal models of human prostate cancer: the consensus report of the
New York meeting of the Mouse Models of Human Cancers Consortium Prostate
Pathology Committee. Cancer Res, 2013. 73(9): p. 2718-36.
22. Loftus, T.M., et al., Reduced food intake and body weight in mice treated with fatty acid
synthase inhibitors. Science, 2000. 288(5475): p. 2379-81.
23. Pizer, E.S., et al., Increased fatty acid synthase as a therapeutic target in androgen-
independent prostate cancer progression. Prostate, 2001. 47(2): p. 102-10.
24. Wu, X., et al., Generation of a prostate epithelial cell-specific Cre transgenic mouse
model for tissue-specific gene ablation. Mech Dev, 2001. 101(1-2): p. 61-9.
25. Chakravarthy, M.V., et al., "New" hepatic fat activates PPARalpha to maintain glucose,
lipid, and cholesterol homeostasis. Cell Metab, 2005. 1(5): p. 309-22.
26. Wang, H.Q., et al., Positive feedback regulation between AKT activation and fatty acid
synthase expression in ovarian carcinoma cells. Oncogene, 2005. 24(22): p. 3574-82.
27. Kuhajda, F.P., et al., Fatty acid synthesis: a potential selective target for antineoplastic
therapy. Proc Natl Acad Sci U S A, 1994. 91(14): p. 6379-83.
102
28. Rysman, E., et al., De novo lipogenesis protects cancer cells from free radicals and
chemotherapeutics by promoting membrane lipid saturation. Cancer Res, 2011. 70(20): p.
8117-26.
29. Indra, A.K., et al., Temporally-controlled site-specific mutagenesis in the basal layer of
the epidermis: comparison of the recombinase activity of the tamoxifen-inducible Cre-
ER(T) and Cre-ER(T2) recombinases. Nucleic Acids Res, 1999. 27(22): p. 4324-7.
30. Feil, R., et al., Ligand-activated site-specific recombination in mice. Proc Natl Acad Sci
U S A, 1996. 93(20): p. 10887-90.
31. Luchman, H.A., et al., Temporally controlled prostate epithelium-specific gene
alterations. Genesis, 2008. 46(4): p. 229-34.
32. Luchman, H.A., et al., The pace of prostatic intraepithelial neoplasia development is
determined by the timing of Pten tumor suppressor gene excision. PLoS One, 2008.
3(12): p. e3940.
33. Brusselmans, K., et al., RNA interference-mediated silencing of the acetyl-CoA-
carboxylase-alpha gene induces growth inhibition and apoptosis of prostate cancer cells.
Cancer Res, 2005. 65(15): p. 6719-25.
34. Alli, P.M., et al., Fatty acid synthase inhibitors are chemopreventive for mammary cancer
in neu-N transgenic mice. Oncogene, 2005. 24(1): p. 39-46.
35. Hanahan, D. and R.A. Weinberg, Hallmarks of cancer: the next generation. Cell, 2011.
144(5): p. 646-74.
36. Wortman, M.D., et al., C75 inhibits food intake by increasing CNS glucose metabolism.
Nat Med, 2003. 9(5): p. 483-5.
37. Ding, Z., et al., SMAD4-dependent barrier constrains prostate cancer growth and
metastatic progression. Nature, 2011. 470(7333): p. 269-73.
38. Chen, Z., et al., Crucial role of p53-dependent cellular senescence in suppression of Pten-
deficient tumorigenesis. Nature, 2005. 436(7051): p. 725-30.
39. Di Cristofano, A., et al., Pten is essential for embryonic development and tumour
suppression. Nat Genet, 1998. 19(4): p. 348-55.
40. Wang, S., et al., Effect of dietary polyunsaturated fatty acids on castration-resistant Pten-
null prostate cancer. Carcinogenesis, 2012. 33(2): p. 404-12.
41. Berquin, I.M., et al., Modulation of prostate cancer genetic risk by omega-3 and omega-6
fatty acids. J Clin Invest, 2007. 117(7): p. 1866-75.
103
42. Birbach, A., Use of PB-Cre4 mice for mosaic gene deletion. PLoS One, 2013. 8(1): p.
e53501.
43. Chirala, S.S., et al., Fatty acid synthesis is essential in embryonic development: fatty acid
synthase null mutants and most of the heterozygotes die in utero. Proc Natl Acad Sci U S
A, 2003. 100(11): p. 6358-63.
44. Soncini, M., et al., Hormonal and nutritional control of the fatty acid synthase promoter
in transgenic mice. J Biol Chem, 1995. 270(51): p. 30339-43.
45. Manning, B.D. and L.C. Cantley, AKT/PKB signaling: navigating downstream. Cell,
2007. 129(7): p. 1261-74.
46. Zhou, W., et al., Fatty acid synthase inhibition triggers apoptosis during S phase in
human cancer cells. Cancer Res, 2003. 63(21): p. 7330-7.
47. Little, J.L., et al., Inhibition of fatty acid synthase induces endoplasmic reticulum stress
in tumor cells. Cancer Res, 2007. 67(3): p. 1262-9.
48. Fiorentino, M., et al., Overexpression of fatty acid synthase is associated with
palmitoylation of Wnt1 and cytoplasmic stabilization of beta-catenin in prostate cancer.
Lab Invest, 2008. 88(12): p. 1340-8.
49. Wei, X., et al., De novo lipogenesis maintains vascular homeostasis through endothelial
nitric-oxide synthase (eNOS) palmitoylation. J Biol Chem, 2011. 286(4): p. 2933-45.
50. Wei, X., et al., Fatty acid synthase modulates intestinal barrier function through
palmitoylation of mucin 2. Cell Host Microbe, 2012. 11(2): p. 140-52.
51. Chakravarthy, M.V., et al., Brain fatty acid synthase activates PPARalpha to maintain
energy homeostasis. J Clin Invest, 2007. 117(9): p. 2539-52.
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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).
108
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).
109
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).
110
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
126
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
IV.E REFERENCES
1. Suburu, J., et al., Polyunsaturated Fatty Acid Metabolism and Cancer. In:
Polyunsaturated Fatty Acids: Sources, Antioxidant Properties and Health Benefits, A.
Catalá, Editor. 2013. New York: Nova Science Publishers.
2. Chirala, S.S., et al., Animal fatty acid synthase: functional mapping and cloning and
expression of the domain I constituent activities. Proc Natl Acad Sci U S A, 1997.
94(11): p. 5588-93.
3. Semenkovich, C.F., Regulation of fatty acid synthase (FAS). Prog Lipid Res, 1997.
36(1): p. 43-53.
4. Wakil, S.J., Fatty acid synthase, a proficient multifunctional enzyme. Biochemistry,
1989. 28(11): p. 4523-30.
5. Wakil, S.J., J.K. Stoops, and V.C. Joshi, Fatty acid synthesis and its regulation. Annu
Rev Biochem, 1983. 52: p. 537-79.
6. Chirala, S.S., et al., Fatty acid synthesis is essential in embryonic development: fatty acid
synthase null mutants and most of the heterozygotes die in utero. Proc Natl Acad Sci U S
A, 2003. 100(11): p. 6358-63.
7. Abu-Elheiga, L., et al., Mutant mice lacking acetyl-CoA carboxylase 1 are embryonically
lethal. Proc Natl Acad Sci U S A, 2005. 102(34): p. 12011-6.
8. Chakravarthy, M.V., et al., "New" hepatic fat activates PPARalpha to maintain glucose,
lipid, and cholesterol homeostasis. Cell Metab, 2005. 1(5): p. 309-22.
9. Chakravarthy, M.V., et al., Brain fatty acid synthase activates PPARalpha to maintain
energy homeostasis. J Clin Invest, 2007. 117(9): p. 2539-52.
10. Razani, B., et al., Fatty acid synthase modulates homeostatic responses to myocardial
stress. J Biol Chem, 2011. 286(35): p. 30949-61.
11. Wei, X., et al., Fatty acid synthase modulates intestinal barrier function through
palmitoylation of mucin 2. Cell Host Microbe, 2012. 11(2): p. 140-52.
12. Wei, X., et al., De novo lipogenesis maintains vascular homeostasis through endothelial
nitric-oxide synthase (eNOS) palmitoylation. J Biol Chem, 2011. 286(4): p. 2933-45.
13. Smith, S. and S. Abraham, Fatty acid synthetase from lactating rat mammary gland. I.
Isolation and properties. J Biol Chem, 1970. 245(12): p. 3209-17.
14. Lund, L.R., et al., Two distinct phases of apoptosis in mammary gland involution:
proteinase-independent and -dependent pathways. Development, 1996. 122(1): p. 181-93.
130
15. Baxter, F.O., K. Neoh, and M.C. Tevendale, The beginning of the end: death signaling in
early involution. J Mammary Gland Biol Neoplasia, 2007. 12(1): p. 3-13.
16. Schwertfeger, K.L., M.M. Richert, and S.M. Anderson, Mammary gland involution is
delayed by activated Akt in transgenic mice. Mol Endocrinol, 2001. 15(6): p. 867-81.
17. Wang, H.Q., et al., Positive feedback regulation between AKT activation and fatty acid
synthase expression in ovarian carcinoma cells. Oncogene, 2005. 24(22): p. 3574-82.
18. Menendez, J.A. and R. Lupu, Fatty acid synthase and the lipogenic phenotype in cancer
pathogenesis. Nat Rev Cancer, 2007. 7(10): p. 763-77.
19. Beckers, A., et al., Chemical inhibition of acetyl-CoA carboxylase induces growth arrest
and cytotoxicity selectively in cancer cells. Cancer Res, 2007. 67(17): p. 8180-7.
20. Brusselmans, K., et al., Epigallocatechin-3-gallate is a potent natural inhibitor of fatty
acid synthase in intact cells and selectively induces apoptosis in prostate cancer cells. Int
J Cancer, 2003. 106(6): p. 856-62.
21. Fritz, V., et al., Abrogation of de novo lipogenesis by stearoyl-CoA desaturase 1
inhibition interferes with oncogenic signaling and blocks prostate cancer progression in
mice. Mol Cancer Ther, 2010. 9(6): p. 1740-54.
22. Orita, H., et al., Selective inhibition of fatty acid synthase for lung cancer treatment. Clin
Cancer Res, 2007. 13(23): p. 7139-45.
23. Hatzivassiliou, G., et al., ATP citrate lyase inhibition can suppress tumor cell growth.
Cancer Cell, 2005. 8(4): p. 311-21.
24. McFadden, J.M., et al., Application of a flexible synthesis of (5R)-thiolactomycin to
develop new inhibitors of type I fatty acid synthase. J Med Chem, 2005. 48(4): p. 946-61.
25. Loftus, T.M., et al., Reduced food intake and body weight in mice treated with fatty acid
synthase inhibitors. Science, 2000. 288(5475): p. 2379-81.
26. Kuhajda, F.P., et al., Synthesis and antitumor activity of an inhibitor of fatty acid
synthase. Proc Natl Acad Sci U S A, 2000. 97(7): p. 3450-4.
27. Vazquez, M.J., et al., Discovery of GSK837149A, an inhibitor of human fatty acid
synthase targeting the beta-ketoacyl reductase reaction. FEBS J, 2008. 275(7): p. 1556-
67.
28. De Schrijver, E., et al., RNA interference-mediated silencing of the fatty acid synthase
gene attenuates growth and induces morphological changes and apoptosis of LNCaP
prostate cancer cells. Cancer Res, 2003. 63(13): p. 3799-804.
131
29. Migita, T., et al., Fatty acid synthase: a metabolic enzyme and candidate oncogene in
prostate cancer. J Natl Cancer Inst, 2009. 101(7): p. 519-32.
30. Little, J.L., et al., Inhibition of fatty acid synthase induces endoplasmic reticulum stress
in tumor cells. Cancer Res, 2007. 67(3): p. 1262-9.
31. Kridel, S.J., et al., Orlistat is a novel inhibitor of fatty acid synthase with antitumor
activity. Cancer Res, 2004. 64(6): p. 2070-5.
32. Rysman, E., et al., De novo lipogenesis protects cancer cells from free radicals and
chemotherapeutics by promoting membrane lipid saturation. Cancer Res, 2011. 70(20): p.
8117-26.
33. Siegel, R., et al., Cancer statistics, 2014. CA Cancer J Clin, 2014. 64(1): p. 9-29.
34. Swinnen, J.V., et al., Selective activation of the fatty acid synthesis pathway in human
prostate cancer. Int J Cancer, 2000. 88(2): p. 176-9.
35. Oyama, N., et al., 11C-acetate PET imaging of prostate cancer. J Nucl Med, 2002. 43(2):
p. 181-6.
36. Fricke, E., et al., Positron emission tomography with 11C-acetate and 18F-FDG in
prostate cancer patients. Eur J Nucl Med Mol Imaging, 2003. 30(4): p. 607-11.
37. Vavere, A.L., et al., 1-11C-acetate as a PET radiopharmaceutical for imaging fatty acid
synthase expression in prostate cancer. J Nucl Med, 2008. 49(2): p. 327-34.
38. Wang, S., et al., Prostate-specific deletion of the murine Pten tumor suppressor gene
leads to metastatic prostate cancer. Cancer Cell, 2003. 4(3): p. 209-21.
39. Wu, X., et al., Generation of a prostate epithelial cell-specific Cre transgenic mouse
model for tissue-specific gene ablation. Mech Dev, 2001. 101(1-2): p. 61-9.
40. Jin, Q., et al., Fatty acid synthase phosphorylation: a novel therapeutic target in HER2-
overexpressing breast cancer cells. Breast Cancer Res, 2010. 12(6): p. R96.
41. Funai, K., et al., Muscle lipogenesis balances insulin sensitivity and strength through
calcium signaling. J Clin Invest, 2013. 123(3): p. 1229-40.
42. Zhou, W., et al., Fatty acid synthase inhibition triggers apoptosis during S phase in
human cancer cells. Cancer Res, 2003. 63(21): p. 7330-7.
43. Guseva, N.V., et al., TOFA (5-tetradecyl-oxy-2-furoic acid) reduces fatty acid synthesis,
inhibits expression of AR, neuropilin-1 and Mcl-1 and kills prostate cancer cells
independent of p53 status. Cancer Biol Ther, 2011. 12(1): p. 80-5.
132
44. Brusselmans, K., et al., RNA interference-mediated silencing of the acetyl-CoA-
carboxylase-alpha gene induces growth inhibition and apoptosis of prostate cancer cells.
Cancer Res, 2005. 65(15): p. 6719-25.
45. Zadra, G., et al., New strategies in prostate cancer: targeting lipogenic pathways and the
energy sensor AMPK. Clin Cancer Res, 2010. 16(13): p. 3322-8.
46. Funabashi, H., et al., Binding site of cerulenin in fatty acid synthetase. J Biochem, 1989.
105(5): p. 751-5.
47. Landree, L.E., et al., C75, a fatty acid synthase inhibitor, modulates AMP-activated
protein kinase to alter neuronal energy metabolism. J Biol Chem, 2004. 279(5): p. 3817-
27.
48. Wortman, M.D., et al., C75 inhibits food intake by increasing CNS glucose metabolism.
Nat Med, 2003. 9(5): p. 483-5.
49. Cha, S.H., et al., Inhibition of hypothalamic fatty acid synthase triggers rapid activation
of fatty acid oxidation in skeletal muscle. Proc Natl Acad Sci U S A, 2005. 102(41): p.
14557-62.
50. Cha, S.H., et al., Hypothalamic malonyl-CoA triggers mitochondrial biogenesis and
oxidative gene expression in skeletal muscle: Role of PGC-1alpha. Proc Natl Acad Sci U
S A, 2006. 103(42): p. 15410-5.
51. Kim, E.K., et al., C75, a fatty acid synthase inhibitor, reduces food intake via
hypothalamic AMP-activated protein kinase. J Biol Chem, 2004. 279(19): p. 19970-6.
52. Makowski, K., et al., Differential pharmacologic properties of the two C75 enantiomers:
(+)-C75 is a strong anorectic drug; (-)-C75 has antitumor activity. Chirality, 2013. 25(5):
p. 281-7.
53. Wang, X. and W. Tian, Green tea epigallocatechin gallate: a natural inhibitor of fatty-
acid synthase. Biochem Biophys Res Commun, 2001. 288(5): p. 1200-6.
54. Oliveras, G., et al., Novel anti-fatty acid synthase compounds with anti-cancer activity in
HER2+ breast cancer. Ann N Y Acad Sci, 2010. 1210: p. 86-92.
55. Puig, T., et al., A novel inhibitor of fatty acid synthase shows activity against HER2+
breast cancer xenografts and is active in anti-HER2 drug-resistant cell lines. Breast
Cancer Res, 2011. 13(6): p. R131.
56. Pemble, C.W.t., et al., Crystal structure of the thioesterase domain of human fatty acid
synthase inhibited by Orlistat. Nat Struct Mol Biol, 2007. 14(8): p. 704-9.
133
57. Ardecky, R., et al., Selective inhibitors of FAS-TE. In: Probe Reports from the NIH
Molecular Libraries Program [Internet]. 2013 Apr 12 [Updated 2014 Jan 13]. Bethesda
(MD): National Center for Biotechnology Information
http://www.ncbi.nlm.nih.gov/books/NBK189923/.
58. Maier, T., M. Leibundgut, and N. Ban, The crystal structure of a mammalian fatty acid
synthase. Science, 2008. 321(5894): p. 1315-22.
59. Flavin, R., G. Zadra, and M. Loda, Metabolic alterations and targeted therapies in
prostate cancer. J Pathol, 2011(223): p. 283-294.
60. Sekine, Y., et al., Remnant lipoproteins induced proliferation of human prostate cancer
cell, PC-3 but not LNCaP, via low density lipoprotein receptor. Cancer Epidemiol, 2009.
33(1): p. 16-23.
61. He, L., et al., Up-regulated expression of type II very low density lipoprotein receptor
correlates with cancer metastasis and has a potential link to beta-catenin in different
cancers. BMC Cancer, 2010. 10: p. 601.
62. Kuemmerle, N.B., et al., Lipoprotein lipase links dietary fat to solid tumor cell
proliferation. Mol Cancer Ther, 2011. 10(3): p. 427-36.
63. Teran-Garcia, M., et al., Polyunsaturated fatty acid suppression of fatty acid synthase
(FASN): evidence for dietary modulation of NF-Y binding to the Fasn promoter by
SREBP-1c. Biochem J, 2007. 402(3): p. 591-600.
64. Jump, D.B., et al., Coordinate regulation of glycolytic and lipogenic gene expression by
polyunsaturated fatty acids. J Lipid Res, 1994. 35(6): p. 1076-84.
65. Calle, E.E. and R. Kaaks, Overweight, obesity and cancer: epidemiological evidence and
proposed mechanisms. Nat Rev Cancer, 2004. 4(8): p. 579-91.
66. Calle, E.E., et al., Overweight, obesity, and mortality from cancer in a prospectively
studied cohort of U.S. adults. N Engl J Med, 2003. 348(17): p. 1625-38.
67. Anderson, A.S. and S. Caswell, Obesity management--an opportunity for cancer
prevention. Surgeon, 2009. 7(5): p. 282-5.
68. van Kruijsdijk, R.C., E. van der Wall, and F.L. Visseren, Obesity and cancer: the role of
dysfunctional adipose tissue. Cancer Epidemiol Biomarkers Prev, 2009. 18(10): p. 2569-
78.
69. Trujillo, M.E. and P.E. Scherer, Adipose tissue-derived factors: impact on health and
disease. Endocr Rev, 2006. 27(7): p. 762-78.
134
70. Lee, M.J., Y. Wu, and S.K. Fried, Adipose tissue heterogeneity: Implication of depot
differences in adipose tissue for obesity complications. Mol Aspects Med, 2013. 34(1): p.
1-11.
71. Karastergiou, K., et al., Distinct developmental signatures of human abdominal and
gluteal subcutaneous adipose tissue depots. J Clin Endocrinol Metab, 2013. 98(1): p. 362-
71.
72. Nguyen, L.V., et al., Cancer stem cells: an evolving concept. Nat Rev Cancer, 2012.
12(2): p. 133-43.
73. Zhang, Y., et al., White adipose tissue cells are recruited by experimental tumors and
promote cancer progression in mouse models. Cancer Res, 2009. 69(12): p. 5259-66.
74. Zhang, Y., et al., Stromal progenitor cells from endogenous adipose tissue contribute to
pericytes and adipocytes that populate the tumor microenvironment. Cancer Res, 2012.
72(20): p. 5198-208.
75. Hanahan, D. and R.A. Weinberg, Hallmarks of cancer: the next generation. Cell, 2011.
144(5): p. 646-74.
76. Hanahan, D. and L.M. Coussens, Accessories to the crime: functions of cells recruited to
the tumor microenvironment. Cancer Cell, 2012. 21(3): p. 309-22.
77. Nieman, K.M., et al., Adipose tissue and adipocytes support tumorigenesis and
metastasis. Biochim Biophys Acta, 2013.
78. Dirat, B., et al., Cancer-associated adipocytes exhibit an activated phenotype and
contribute to breast cancer invasion. Cancer Res, 2011. 71(7): p. 2455-65.
79. Nakajima, E.C. and B. Van Houten, Metabolic symbiosis in cancer: refocusing the
Warburg lens. Mol Carcinog, 2012. 52(5): p. 329-37.
80. Martinez-Outschoorn, U.E., F. Sotgia, and M.P. Lisanti, Power surge: supporting cells
"fuel" cancer cell mitochondria. Cell Metab, 2012. 15(1): p. 4-5.
81. Lodhi, I.J., et al., Inhibiting adipose tissue lipogenesis reprograms thermogenesis and
PPARgamma activation to decrease diet-induced obesity. Cell Metab, 2012. 16(2): p.
189-201.
82. Berndt, J., et al., Fatty acid synthase gene expression in human adipose tissue: association
with obesity and type 2 diabetes. Diabetologia, 2007. 50(7): p. 1472-80.
83. Menendez, J.A., et al., Fatty acid synthase: association with insulin resistance, type 2
diabetes, and cancer. Clin Chem, 2009. 55(3): p. 425-38.
135
CURRICULUM VITAE
Janel K. Suburu
Winston-Salem, NC
[email protected] • [email protected]
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