# Omega-3 Fatty Acids and PPARγ in Cancer

Authors: Iris J. Edwards; Joseph T. O'Flaherty
Journal: PPAR Research
 (2008)

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## Abstract

Omega-3 (or n-3) polyunsaturated fatty acids (PUFAs) and their metabolites are natural ligands for peroxisome proliferator receptor activator (PPAR)γ and, due to the effects of PPARγ on cell proliferation, survival, and differentiation, are potential anticancer agents. Dietary intake of omega-3 PUFAs has been associated with a reduced risk of certain cancers in human populations and in animal models. In vitro studies have shown that omega-3 PUFAs inhibit cell proliferation and induce apoptosis in cancer cells through various pathways but one of which involves PPARγ activation. The differential activation of PPARγ and PPARγ-regulated genes by specific dietary fatty acids may be central to their distinct roles in cancer. This review summarizes studies relating PUFAs to PPARγ and cancer and offers a new paradigm relating an n-3 PUFA through PPARγ to the expression of the cell surface proteoglycan, syndecan-1, and to the death of cancer cells.


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## Body

## 1. Introduction

The peroxisome proliferator-activated receptor
(PPAR) family of nuclear receptors comprises three distinct gene products, PPARα, β/δ, and γ, that differ in ligand specificity, tissue
distribution, and developmental expression  [1–3]. PPARs demonstrate a
relatively high level of constitutive transcriptional activity which is further
increased upon binding their activating ligands  [4–7]. These ligands are primarily
long chain unsaturated and polyunsaturated fatty acids (PUFAs) and certain
metabolites of these fatty acids  [8–10]. Initially, PPARs were thought
mainly to govern lipid homeostasis by binding fatty acids and their metabolites
to thereby become more active in regulating genes for proteins involved in lipid
metabolism  [8, 10, 11]. Indeed, PPARα is expressed predominantly in tissues with
high fatty acid requirements such as liver, heart, and kidney, while PPARγ isoforms γ1 and γ2 are highly enriched in adipose tissue to regulate
adipocyte differentiation and lipid storage  [3]. However, expression of PPARγ1, as with PPARβ/δ and PPARα, has now been extended to most other tissues
and regulatory roles for PPARs extended to other systemic functions such as
carbohydrate regulation, immune modulation, and the proliferation, survival and
differentiation of cells  [3]. The latter effects have led
to intense interest in the PPARs in relation to cancer.PPARα and its ligand activators regulate fatty
acid and lipoprotein metabolism and promote the development of hepatocellular
carcinoma in rodents and reduce the metastasis of melanoma in hamsters  [12]. These and other of their
effects do not, in general, translate to humans. PPARβ/δ plays a key role in
lipid metabolism of peripheral tissues. Its high expression in colon has been
shown to promote colon cancer  [12, 13], in a mechanism that involves
the stimulation of PPARβ/δ by arachidonic acid, PPARβ/δ-dependent upregulation
of cyclooxygenase (COX)-2 leading to overproduction of prostaglandin (PG)E2,
and PGE2-induced growth of colon cancer cells. There is relatively
little documentation of a role for PPARβ/δ in other cancers  [14]. By contrast, PPARγ has a
broad range of effects on cancer. PPARγ controls fat metabolism by regulating genes
involved in lipogenesis, insulin sensitivity, and adipocyte differentiation  [3, 15]. These effects underlie the
use of thiazolidinediones, which bind and activate PPARγ, to treat insulin-resistant type II
diabetes  [3, 15]. Although PPARγ activators have
been widely shown to inhibit growth in cultured cancer cells, in vivo effects
have proved to be complex: they inhibit but sometimes promote cancer growth  [16] probably due to stimulation of
antiproliferative and apoptotic signaling pathways or proliferative and
antiapoptotic pathways, depending on cellular conditions
[3, 12, 15–18]. These
findings led to the idea of selective PPARγ modulators (SPARMs), drugs analogous to
selective estrogen receptor modulators (SERMs) in which distinct actions of the
modulator depend on the cellular context  [19] and on distinct receptor
conformations, and therefore different gene interactions  [20]. Fatty acids may be considered
as natural SPARMs since their binding does not necessarily lead to PPAR
activation and target gene transcription  [11].The considerations discussed above raise a
possibility that managed alterations in the type of fatty acids in tissues, can
alter the activity of PPARs and thereby the genes they control for therapeutic
benefit. The fatty acid content of tissues is dependent mainly on dietary
intake. Omega-3 PUFAs, docosahexaenoic acid (DHA), and eicosapentaenoic acid
(EPA) are enriched in the diets of many populations that enjoy a low incidence
of cancer  [21]. These diets also obtain some
modest success ameliorating advanced cancer in humans  [22] and have been widely used to
inhibit carcinogenesis and tumor progression in animal models. The ability of
specific fatty acids to differentially activate PPARs and PPAR-regulated genes
may be central to their distinct roles in cancer. This review will focus on
PPARγ, its activation by fatty acids, and functional
results in cancer cells.


## 2. Fatty Acid Metabolism

### 2.1. Fatty Acid Types and Interconversions

Fatty acids are hydrocarbons with a terminal carboxyl group. The carbons of saturated fatty acids are all connected
by single bonds, whereas the chains of monounsaturated and polyunsaturated
fatty acids (PUFAs) contain one or more double bonds, respectively. The n-3 and n-6 designation describes the
position of the double bond closest to the (omega) carbon at the methyl end of
the molecule (Figure 1). Oleic acid (18:1) has a single double bond between
carbons 9 and 10 from the omega carbon and is designated an n-9 or omega-9
monounsaturated fatty acid. Like the saturated fatty acids, oleic acid can be
synthesized de novo in mammalian cells. It can also be obtained from the diet
through intake of oils such as olive and canola. By contrast, PUFAs cannot be
synthesized de novo in mammals and must be obtained from the diet. The shortest
of the n-6 PUFAs is linoleic acid (LA, 18:2, n-6). Its 18 carbon, n-3
counterpart is α-linolenic acid (ALA, 18:3, n-3). Both LA and ALA are metabolized through a series of elongation and
desaturation steps to longer chain PUFAs: LA to arachidonic acid (AA, 20:4, n-6)
and ALA
to
EPA (20:5, n-3) and DHA (22:6, n-3) (Figure 2). The first and rate limiting step
in this pathway is the introduction of a double bond by the Δ6 desaturase (for
review see [23]). For n-3 PUFAs, ALA
is converted to stearidonic
acid (SDA, 18:4, n-3), elongated, and desaturated by Δ5-desaturase to form EPA.
In mammalian cells, the conversion of EPA to DHA follows the Sprecher pathway
in which EPA is elongated to docosapentaenoic acid (DPA, 22:5, n-3), then to
tetracosapentaenoic acid (TPA. 24:5, n-3), and desaturated to tetracosahexaenoic
acid (THA, 24:6, n-3). THA is translocated from the endoplasmic reticulum to
peroxisomes, where β-oxidation results in the loss of 2
carbons to form DHA [24]. The PUFAs are also
metabolized, most importantly for this review, to PPARγ activators (see Section
2.3).Figure 1
Structures of unsaturated fatty acids: oleic acid (n-9 monounsaturated), linoleic acid and
arachidonic acid (n-6 polyunsaturated),α-linoleic acid, eicosapentaenoic acid, and docosahexaenoic
acid (n-3 polyunsaturated ). The “n” numbers are counted from the methyl or omega terminus.Figure 2
The elongation-desaturation pathway for the metabolism of n-6
and n-3 polyunsaturated fatty acids.


### 2.2. Dietary Fatty Acids

The results of both dietary intake
and stable isotope studies have shown that the conversion of ALA to DHA in
humans is extremely inefficient (for review see [25]). Most of the ingested ALA is an immediate
target for β-oxidation to provide energy, leaving an
estimated 8–10% to enter the
elongation-desaturation pathway [26, 27]. A kinetic analysis of 2H-labeled
fatty acids estimated that conversion of ALA to EPA was only 0.2%, EPA to DPA was
0.13%, and DPA to DHA was 0.05% [28]. There is some evidence of
gender-related differences in the activity of the elongation-desaturation
pathway that result in a greater efficiency of conversion of ALA to DHA in
females than in males [25, 27, 29]. Support for a role of sex
hormones in the conversion pathway is provided by data indicating higher DHA in
plasma lipids associated with oral contraceptive use [27] as well in males supplemented
with estrogen during sex-change procedures [30]. Moreover, testosterone
treatment of female-male transsexuals was shown to decrease plasma DHA [30].Because common enzymes in the elongation-desaturation pathway are responsible for conversion of both n-3 and n-6
PUFAs, background diet is also a factor in efficiency of conversion. LA is the
most abundant fatty acid in the Western diet with consumption in US that is ten-fold
that of ALA (reviewed in [31]). Studies have shown that a
high intake of LA is associated with a low conversion of ALA to EPA [26]. In spite of limited metabolism
of ALA to its long chain derivatives in the stable isotope tracer studies,
feeding studies have consistently shown that increased consumption of ALA does
result in higher levels of EPA in plasma or cell lipids [31]. However, there was no
measurable increase in DHA in these pools. Likewise diets supplemented with EPA
do not result in a detectable increase in plasma DHA [32]. Thus, the inefficiency of
this pathway does not appear limited to one step but rather extends throughout
the pathway. The consensus of a number of studies is that the only way to
increase plasma and tissue levels of a specific PUFA is to increase the
consumption of that fatty acid. This may be of particular importance in the
light of recent in vitro studies on the antitumor effects of DHA.


### 2.3. PUFAs Metabolism to PPARγ Activators

Tissues
metabolize PUFAs to oxygenated products that have quite different impacts on
PPARγ than their parent molecules. Moreover, n-3 PUFA inhibit the metabolism
n-6 PUFAs and subplant them from their oxygenation pathways to form products [33–35] that have different effects
on PPARγ than their n-6 PUFAs oxygenated counterparts. It is therefore
important to consider PUFAs oxygenation pathways. LA, AA, and DHA require
>10–30μM to activate
PPARγ but are commonly converted to stronger (>0.1–10μM) activators
in cells. LA is metabolized (Figure 3, upper panel) by 15-lipoxygenases (LOX)-1/2
to 9(S)- and 13(S)-HODE (hydroxy-octadecaenoate) and by cyclooxygenases
(COX)-1/2 to 9(R)- and 13(S)-HODE. The HODEs can be converted to 13-oxo- and
9-oxo-ODE by a dehydrogenase [36–39]. The hydroxy and to a greater
extent oxo LA analogs have greater PPARγ-activating potency than LA [36, 40–42]. AA is metabolized (Figure 3,
center panel) via 5-LOX to 5(S)-HETE (hydroxy-eicosatetra-enoate) and via
15-LOX-1/2 to 15(S)-HETE. These HETEs can be converted to oxo-ETEs and
5-oxo-15(S)-hydroxy-ETE as shown in Figure 3 [39, 43–50]. 15-HETE has weak and 5-HETE essentially
no ability to activate PPARγ. However, their oxo counterparts have appreciable ability
to do so with 5-oxo-15(S)-hydroxy-ETE showing the greatest potency in binding and activating PPARγ [43]. AA is also metabolized (Figure
3, center panel) by COX1/2 to PG (prostaglandin) D2 which as a
consequence of successive dehydrations
and an isomerization, perhaps by nonenzymatic routes, convert to PGJ2,  Δ12-PGJ2,
and 15-deoxyΔ12,14-PGJ2(15-d-Δ12,14-PGJ2);
these PGJ2's have greater ability than PGD2 to activate
PPARγ with 15-d-Δ12,14-PGJ2 being a most potent (>0.1–1μM) naturally occurring PPARγ activator [9, 43, 51–56]. In one study, the Kd's
of 15-d-Δ12,14-PGJ2,
5-oxo-15-OH-ETE, PGJ2, 5-oxo-ETE, and 5(S)-HETE in binding to PPARγ
were 1.4, 11, 37, 81, and >1000μM, respectively; their potency in
activating a cell-based PPARγ reporter paralleled these Kd's [43]. DHA is metabolized (Figure 3,
bottom panel) by 15-LOX or other oxygenase to 17-OH- and 7-OH-DHA, products that
activate PPARγ with greater potency (ED50's in activating a
cell-based PPARγ reporter of ~5μM) than DHA [57]. 4-OH-, and 4-oxo-DHA [53], while not yet shown to be
made by cancer cells, also activate PPARγ with greater potency (ED50's
of 13.4 and 7.8μM in
activating a cellular PPARγ
reporter, resp.) than DHA (ED50>10⁢μm) [53]. Hence, in this DHA series,
similar to the 5-HETE series of AA metabolites, the oxo analog exhibits the
greatest potency. We note that the more potent PPARγ activators, the oxo-PUFAs,
form preferentially in cells undergoing excessive oxidation, free radical, and
NADPH/NADH-depleting reactions [43, 44, 48, 57, 58]. This suggests that PPARγ may
serve as a sensor for oxo-PUFA thereby monitoring cellular oxidative stress and
when this stress is severe, engaging cell death programs [43, 58]. This PPARγ function, we
suggest, could contribute to the necrosis that occurs in tumors particularly
after chemical and radiation treatment [59].The cellular metabolism of LA, AA, and DHA to more potent activators of PPARγ. ODE is
octadecaenoate; HETE is hydroxy-eicosatetraenoate; ETE is eicosatetraenoate; PG is prostaglandin.
(a)(b)(c)Cells process PUFAs in other
relevant ways. They convert them to nitrates, probably in nonenzymatic reactions,
where the nitric oxide made during cell stimulation attacks the PUFAs. Nitrated
LA and AA are stronger PPARγ activators than their precursors [60–62]. Cells also convert PUFAs to cannabinoids such
as anandamide (ethanolamine amide of AA) and arachidonoylglycerol which also
activate PPARγ with greater potency than AA [63–65]. Finally, cells conjugate
glutathione to PUFAs that contain an α,β-unsaturated ketone such as 15-d-Δ12,14-PGJ2 and 5-oxo-ETE [66–68]. Since the conjugates are
rapidly excreted from cells by multidrug-resistance transporters, conjugation inhibits
the ability of α,β-unsaturated ketones to activate PPARγ [66]. Cancer cells excrete
anticancer drugs through these same transporters and become drug-resistant by
overexpressing these transporters [69]. Such mutated cells may also
be resistant to α,β-unsaturated ketone activators of PPARγ.


### 2.4. Low-Density Lipoproteins (LDL) as Deliverers of  PPARγ-Activating n-3 PUFA

LDL carry esterified
PUFAs in glycerolipids and cholesterol. They bind to cell surface LDL receptors
and then internalize in endocytic vesicles which merge with lysosomes to de-esterify
and release the PUFAs into the cytosol  [70]. This route differs from the
direct delivery of PUFA: it bypasses cell surface G protein-coupled fatty acid receptors
(GPR 40 and 120; see Section 4.3), deposits PUFA in cells more slowly, and thereby
avoids stimulation of G protein-coupled receptors and, perhaps, an array of C
domain-bearing proteins which are activated by PUFA. This is also an important
pathway for delivering PUFA to tumor cells because of a significant increase in
LDL receptor activity in neoplastic tissues [71–73]. We have obtained from
monkeys fed special diets, LDL enriched with n-6 PUFA (mostly AA and LA) or n-3
PUFA (mostly DHA and EPA). The n-3 but
not n-6 PUFA-rich LDL mimicked thiazolidinediones and DHA in inhibiting cancer
cell growth [74] and activating PPARγ [75, 76].


## 2.1. Fatty Acid Types and Interconversions

Fatty acids are hydrocarbons with a terminal carboxyl group. The carbons of saturated fatty acids are all connected
by single bonds, whereas the chains of monounsaturated and polyunsaturated
fatty acids (PUFAs) contain one or more double bonds, respectively. The n-3 and n-6 designation describes the
position of the double bond closest to the (omega) carbon at the methyl end of
the molecule (Figure 1). Oleic acid (18:1) has a single double bond between
carbons 9 and 10 from the omega carbon and is designated an n-9 or omega-9
monounsaturated fatty acid. Like the saturated fatty acids, oleic acid can be
synthesized de novo in mammalian cells. It can also be obtained from the diet
through intake of oils such as olive and canola. By contrast, PUFAs cannot be
synthesized de novo in mammals and must be obtained from the diet. The shortest
of the n-6 PUFAs is linoleic acid (LA, 18:2, n-6). Its 18 carbon, n-3
counterpart is α-linolenic acid (ALA, 18:3, n-3). Both LA and ALA are metabolized through a series of elongation and
desaturation steps to longer chain PUFAs: LA to arachidonic acid (AA, 20:4, n-6)
and ALA
to
EPA (20:5, n-3) and DHA (22:6, n-3) (Figure 2). The first and rate limiting step
in this pathway is the introduction of a double bond by the Δ6 desaturase (for
review see [23]). For n-3 PUFAs, ALA
is converted to stearidonic
acid (SDA, 18:4, n-3), elongated, and desaturated by Δ5-desaturase to form EPA.
In mammalian cells, the conversion of EPA to DHA follows the Sprecher pathway
in which EPA is elongated to docosapentaenoic acid (DPA, 22:5, n-3), then to
tetracosapentaenoic acid (TPA. 24:5, n-3), and desaturated to tetracosahexaenoic
acid (THA, 24:6, n-3). THA is translocated from the endoplasmic reticulum to
peroxisomes, where β-oxidation results in the loss of 2
carbons to form DHA [24]. The PUFAs are also
metabolized, most importantly for this review, to PPARγ activators (see Section
2.3).Figure 1
Structures of unsaturated fatty acids: oleic acid (n-9 monounsaturated), linoleic acid and
arachidonic acid (n-6 polyunsaturated),α-linoleic acid, eicosapentaenoic acid, and docosahexaenoic
acid (n-3 polyunsaturated ). The “n” numbers are counted from the methyl or omega terminus.Figure 2
The elongation-desaturation pathway for the metabolism of n-6
and n-3 polyunsaturated fatty acids.


## 2.2. Dietary Fatty Acids

The results of both dietary intake
and stable isotope studies have shown that the conversion of ALA to DHA in
humans is extremely inefficient (for review see [25]). Most of the ingested ALA is an immediate
target for β-oxidation to provide energy, leaving an
estimated 8–10% to enter the
elongation-desaturation pathway [26, 27]. A kinetic analysis of 2H-labeled
fatty acids estimated that conversion of ALA to EPA was only 0.2%, EPA to DPA was
0.13%, and DPA to DHA was 0.05% [28]. There is some evidence of
gender-related differences in the activity of the elongation-desaturation
pathway that result in a greater efficiency of conversion of ALA to DHA in
females than in males [25, 27, 29]. Support for a role of sex
hormones in the conversion pathway is provided by data indicating higher DHA in
plasma lipids associated with oral contraceptive use [27] as well in males supplemented
with estrogen during sex-change procedures [30]. Moreover, testosterone
treatment of female-male transsexuals was shown to decrease plasma DHA [30].Because common enzymes in the elongation-desaturation pathway are responsible for conversion of both n-3 and n-6
PUFAs, background diet is also a factor in efficiency of conversion. LA is the
most abundant fatty acid in the Western diet with consumption in US that is ten-fold
that of ALA (reviewed in [31]). Studies have shown that a
high intake of LA is associated with a low conversion of ALA to EPA [26]. In spite of limited metabolism
of ALA to its long chain derivatives in the stable isotope tracer studies,
feeding studies have consistently shown that increased consumption of ALA does
result in higher levels of EPA in plasma or cell lipids [31]. However, there was no
measurable increase in DHA in these pools. Likewise diets supplemented with EPA
do not result in a detectable increase in plasma DHA [32]. Thus, the inefficiency of
this pathway does not appear limited to one step but rather extends throughout
the pathway. The consensus of a number of studies is that the only way to
increase plasma and tissue levels of a specific PUFA is to increase the
consumption of that fatty acid. This may be of particular importance in the
light of recent in vitro studies on the antitumor effects of DHA.


## 2.3. PUFAs Metabolism to PPARγ Activators

Tissues
metabolize PUFAs to oxygenated products that have quite different impacts on
PPARγ than their parent molecules. Moreover, n-3 PUFA inhibit the metabolism
n-6 PUFAs and subplant them from their oxygenation pathways to form products [33–35] that have different effects
on PPARγ than their n-6 PUFAs oxygenated counterparts. It is therefore
important to consider PUFAs oxygenation pathways. LA, AA, and DHA require
>10–30μM to activate
PPARγ but are commonly converted to stronger (>0.1–10μM) activators
in cells. LA is metabolized (Figure 3, upper panel) by 15-lipoxygenases (LOX)-1/2
to 9(S)- and 13(S)-HODE (hydroxy-octadecaenoate) and by cyclooxygenases
(COX)-1/2 to 9(R)- and 13(S)-HODE. The HODEs can be converted to 13-oxo- and
9-oxo-ODE by a dehydrogenase [36–39]. The hydroxy and to a greater
extent oxo LA analogs have greater PPARγ-activating potency than LA [36, 40–42]. AA is metabolized (Figure 3,
center panel) via 5-LOX to 5(S)-HETE (hydroxy-eicosatetra-enoate) and via
15-LOX-1/2 to 15(S)-HETE. These HETEs can be converted to oxo-ETEs and
5-oxo-15(S)-hydroxy-ETE as shown in Figure 3 [39, 43–50]. 15-HETE has weak and 5-HETE essentially
no ability to activate PPARγ. However, their oxo counterparts have appreciable ability
to do so with 5-oxo-15(S)-hydroxy-ETE showing the greatest potency in binding and activating PPARγ [43]. AA is also metabolized (Figure
3, center panel) by COX1/2 to PG (prostaglandin) D2 which as a
consequence of successive dehydrations
and an isomerization, perhaps by nonenzymatic routes, convert to PGJ2,  Δ12-PGJ2,
and 15-deoxyΔ12,14-PGJ2(15-d-Δ12,14-PGJ2);
these PGJ2's have greater ability than PGD2 to activate
PPARγ with 15-d-Δ12,14-PGJ2 being a most potent (>0.1–1μM) naturally occurring PPARγ activator [9, 43, 51–56]. In one study, the Kd's
of 15-d-Δ12,14-PGJ2,
5-oxo-15-OH-ETE, PGJ2, 5-oxo-ETE, and 5(S)-HETE in binding to PPARγ
were 1.4, 11, 37, 81, and >1000μM, respectively; their potency in
activating a cell-based PPARγ reporter paralleled these Kd's [43]. DHA is metabolized (Figure 3,
bottom panel) by 15-LOX or other oxygenase to 17-OH- and 7-OH-DHA, products that
activate PPARγ with greater potency (ED50's in activating a
cell-based PPARγ reporter of ~5μM) than DHA [57]. 4-OH-, and 4-oxo-DHA [53], while not yet shown to be
made by cancer cells, also activate PPARγ with greater potency (ED50's
of 13.4 and 7.8μM in
activating a cellular PPARγ
reporter, resp.) than DHA (ED50>10⁢μm) [53]. Hence, in this DHA series,
similar to the 5-HETE series of AA metabolites, the oxo analog exhibits the
greatest potency. We note that the more potent PPARγ activators, the oxo-PUFAs,
form preferentially in cells undergoing excessive oxidation, free radical, and
NADPH/NADH-depleting reactions [43, 44, 48, 57, 58]. This suggests that PPARγ may
serve as a sensor for oxo-PUFA thereby monitoring cellular oxidative stress and
when this stress is severe, engaging cell death programs [43, 58]. This PPARγ function, we
suggest, could contribute to the necrosis that occurs in tumors particularly
after chemical and radiation treatment [59].The cellular metabolism of LA, AA, and DHA to more potent activators of PPARγ. ODE is
octadecaenoate; HETE is hydroxy-eicosatetraenoate; ETE is eicosatetraenoate; PG is prostaglandin.
(a)(b)(c)Cells process PUFAs in other
relevant ways. They convert them to nitrates, probably in nonenzymatic reactions,
where the nitric oxide made during cell stimulation attacks the PUFAs. Nitrated
LA and AA are stronger PPARγ activators than their precursors [60–62]. Cells also convert PUFAs to cannabinoids such
as anandamide (ethanolamine amide of AA) and arachidonoylglycerol which also
activate PPARγ with greater potency than AA [63–65]. Finally, cells conjugate
glutathione to PUFAs that contain an α,β-unsaturated ketone such as 15-d-Δ12,14-PGJ2 and 5-oxo-ETE [66–68]. Since the conjugates are
rapidly excreted from cells by multidrug-resistance transporters, conjugation inhibits
the ability of α,β-unsaturated ketones to activate PPARγ [66]. Cancer cells excrete
anticancer drugs through these same transporters and become drug-resistant by
overexpressing these transporters [69]. Such mutated cells may also
be resistant to α,β-unsaturated ketone activators of PPARγ.


## 2.4. Low-Density Lipoproteins (LDL) as Deliverers of  PPARγ-Activating n-3 PUFA

LDL carry esterified
PUFAs in glycerolipids and cholesterol. They bind to cell surface LDL receptors
and then internalize in endocytic vesicles which merge with lysosomes to de-esterify
and release the PUFAs into the cytosol  [70]. This route differs from the
direct delivery of PUFA: it bypasses cell surface G protein-coupled fatty acid receptors
(GPR 40 and 120; see Section 4.3), deposits PUFA in cells more slowly, and thereby
avoids stimulation of G protein-coupled receptors and, perhaps, an array of C
domain-bearing proteins which are activated by PUFA. This is also an important
pathway for delivering PUFA to tumor cells because of a significant increase in
LDL receptor activity in neoplastic tissues [71–73]. We have obtained from
monkeys fed special diets, LDL enriched with n-6 PUFA (mostly AA and LA) or n-3
PUFA (mostly DHA and EPA). The n-3 but
not n-6 PUFA-rich LDL mimicked thiazolidinediones and DHA in inhibiting cancer
cell growth [74] and activating PPARγ [75, 76].


## 3. PPARγ

### 3.1. Structural Considerations

PPARγ1 and γ2 originate from the PPARγ gene through separate promoters and 5′ exons.  Compared to the ubiquitously expressed PPARγ1, PPARγ2, which is limited mainly to adipose tissue, has 30 additional amino acids at its NH2 terminus and is a more potent transcription activator  [77]. Because they appear to have
the same targets, however, the two isoforms are here considered together under
the term PPARγ. PPARγ is comprised of four functional parts: the NH2-terminal
A/B region bears a ligand-independent transcription-activating motif AF-1; C
region binds response elements (PPREs with a DR-1 consensus half-sequence of
AGGTCA); D region binds various transcription cofactors; and E/F region has an
interface for dimerizing with 9-cis retinoic acid receptors (RXRs), an AF-2 ligand-dependent
transcription-activating motif, and a ligand-binding domain (LBD) [3, 12, 15, 17]. The LBD has a spacious cavity
that binds ligands having a polar head group extending from a hydrophobic tail
such as diverse PUFAs and PUFA metabolites [7, 77].


### 3.2. PPARγ Regulation by Other Signaling Pathways

PPARγ is
phosphorylated by extracellular signal-regulated kinases (ERK)-1/2 and C-Jun
N-terminal kinase; when so phosphorylated, it has less ligand-binding affinity
and gene-regulating activity [3, 78, 79]. The phosphorylation and attendant
decrease in activity of PPARγ occur in cells treated with PPARγ activators and may
cause the activators to show little or no ability to stimulate PPARγ [3, 79–81]. ERK pathways impact PPARγ in
another way: the ERK-activating enzyme, MEK, when activated, binds with PPARγ's
AF-2 motif. This causes PPARγ to release from PPRE complexes and, bound to MEK
and directed by MEK's nuclear export signal, to exit the nucleus [81, 82]. It is important to note that
PUFAs and PUFA metabolites can activate the MEK/ERK pathway (see Section 4.3)
and therefore may have biphasic effects: they not only directly activate PPARγ
but also entrain events inhibiting PPARγ.PPARγ is
targeted for degradation by ubiquitylation and sumoylation. Ligand binding,
certain protein kinases, and some transcription cofactors (e.g., p300) promote
ubiquitin-dependent degradation of PPARγ in proteasomes [3]. Sumoylation occurs on K107
of PPARγ2 in a ligand-independent fashion to inhibit AF-1 function and on K365
of PPARγ in a ligand-dependent fashion to promote PPARγ's binding of nuclear
receptor corepressor [83, 84]. Sumoylation of PPARγ causes its
proteasomal degradation. ERK phosphorylation promotes K107 sumoylation. This
reaction represents yet another means by which ERKs can inhibit PPARγ [84].


### 3.3. PPARγ Transcriptional Cofactors

PPARs bind
a specific DNA sequence termed peroxisome proliferator response element (PPRE)
in the 5′-flanking region of target genes as a heterodimer
with RXR. Studies using various techniques [3, 85, 86] suggest the following model:
PPARγ•RXR
complexes (the interaction is ligand-independent) exist in nuclei as
macrocomplexes associated with various transcription corepressors [3, 87]. Some complexes, ligand-bound
or not, may associate with transcription coactivators to control the basal
expression of genes. In any event, PPARγ•RXR
complexes are highly mobile, rapidly scanning chromatin, although this scanning
does not involve their DNA binding domain [86]. Ligands trigger PPARγ•RXR to localize
at their cognate PPREs and to exchange corepressors for coactivators such as
cyclic AMP response element binding protein (CREB) and p300 [3, 16, 87, 88]. At some gene sites,
activators cause PPARγ•RXR to
recruit corepressors and thereby cause gene repression [3, 89, 90]. However, the availability of
cofactors differs between cell types and within cells over time depending on
the cell's history and the association of the cofactors to other genes [3, 15, 16], for example, activation of
PPARγ deprives T cell factor/lymphoid enhancing factor (TCF/LEF) of cofactors to
thereby inhibit oncogenic signaling by the Wnt pathway [16]. Thus, the effects of PPARγ activation vary
depending on context and cofactor availability at each genetic site. It seems
at least possible that the PUFA ligands for PPARγ will have differential
effects in impacting its interactions with these transcriptional cofactors in a
manner similar to the SPARMs model [19].


## 3.1. Structural Considerations

PPARγ1 and γ2 originate from the PPARγ gene through separate promoters and 5′ exons.  Compared to the ubiquitously expressed PPARγ1, PPARγ2, which is limited mainly to adipose tissue, has 30 additional amino acids at its NH2 terminus and is a more potent transcription activator  [77]. Because they appear to have
the same targets, however, the two isoforms are here considered together under
the term PPARγ. PPARγ is comprised of four functional parts: the NH2-terminal
A/B region bears a ligand-independent transcription-activating motif AF-1; C
region binds response elements (PPREs with a DR-1 consensus half-sequence of
AGGTCA); D region binds various transcription cofactors; and E/F region has an
interface for dimerizing with 9-cis retinoic acid receptors (RXRs), an AF-2 ligand-dependent
transcription-activating motif, and a ligand-binding domain (LBD) [3, 12, 15, 17]. The LBD has a spacious cavity
that binds ligands having a polar head group extending from a hydrophobic tail
such as diverse PUFAs and PUFA metabolites [7, 77].


## 3.2. PPARγ Regulation by Other Signaling Pathways

PPARγ is
phosphorylated by extracellular signal-regulated kinases (ERK)-1/2 and C-Jun
N-terminal kinase; when so phosphorylated, it has less ligand-binding affinity
and gene-regulating activity [3, 78, 79]. The phosphorylation and attendant
decrease in activity of PPARγ occur in cells treated with PPARγ activators and may
cause the activators to show little or no ability to stimulate PPARγ [3, 79–81]. ERK pathways impact PPARγ in
another way: the ERK-activating enzyme, MEK, when activated, binds with PPARγ's
AF-2 motif. This causes PPARγ to release from PPRE complexes and, bound to MEK
and directed by MEK's nuclear export signal, to exit the nucleus [81, 82]. It is important to note that
PUFAs and PUFA metabolites can activate the MEK/ERK pathway (see Section 4.3)
and therefore may have biphasic effects: they not only directly activate PPARγ
but also entrain events inhibiting PPARγ.PPARγ is
targeted for degradation by ubiquitylation and sumoylation. Ligand binding,
certain protein kinases, and some transcription cofactors (e.g., p300) promote
ubiquitin-dependent degradation of PPARγ in proteasomes [3]. Sumoylation occurs on K107
of PPARγ2 in a ligand-independent fashion to inhibit AF-1 function and on K365
of PPARγ in a ligand-dependent fashion to promote PPARγ's binding of nuclear
receptor corepressor [83, 84]. Sumoylation of PPARγ causes its
proteasomal degradation. ERK phosphorylation promotes K107 sumoylation. This
reaction represents yet another means by which ERKs can inhibit PPARγ [84].


## 3.3. PPARγ Transcriptional Cofactors

PPARs bind
a specific DNA sequence termed peroxisome proliferator response element (PPRE)
in the 5′-flanking region of target genes as a heterodimer
with RXR. Studies using various techniques [3, 85, 86] suggest the following model:
PPARγ•RXR
complexes (the interaction is ligand-independent) exist in nuclei as
macrocomplexes associated with various transcription corepressors [3, 87]. Some complexes, ligand-bound
or not, may associate with transcription coactivators to control the basal
expression of genes. In any event, PPARγ•RXR
complexes are highly mobile, rapidly scanning chromatin, although this scanning
does not involve their DNA binding domain [86]. Ligands trigger PPARγ•RXR to localize
at their cognate PPREs and to exchange corepressors for coactivators such as
cyclic AMP response element binding protein (CREB) and p300 [3, 16, 87, 88]. At some gene sites,
activators cause PPARγ•RXR to
recruit corepressors and thereby cause gene repression [3, 89, 90]. However, the availability of
cofactors differs between cell types and within cells over time depending on
the cell's history and the association of the cofactors to other genes [3, 15, 16], for example, activation of
PPARγ deprives T cell factor/lymphoid enhancing factor (TCF/LEF) of cofactors to
thereby inhibit oncogenic signaling by the Wnt pathway [16]. Thus, the effects of PPARγ activation vary
depending on context and cofactor availability at each genetic site. It seems
at least possible that the PUFA ligands for PPARγ will have differential
effects in impacting its interactions with these transcriptional cofactors in a
manner similar to the SPARMs model [19].


## 4. Targets of PPARγ Relevant to Cancer

### 4.1. Gene Targets of PPARγ

Most known target
genes of PPARγ regulate lipid metabolism and transport [15]with few cancer-related genes
having been confirmed as induced by PPARγ. PPARγ does induce G0/G1 switch gene 2 whose product causes growth arrest in 3T3-L1 cells [91, 92]. PPARγ also binds the NFκB promoter of p53 to
stimulate expression of p53 and, in consequence, p21WAF1/Cip1. It
also binds to a promoter in the Fas ligand gene to induce the expression of
this member of the extrinsic apoptosis pathway. These effects appear
responsible for slowing growth and causing apoptosis in MCF7 breast cancer [93], human umbilical vein
endothelial [94], and possibly Reh [95]  cells. Recent studies have
identified the heparan sulfate proteoglycan, syndecan 1, as a target for PPARγ in human breast [75, 76] and prostate [96] cancer cells. The upregulation
of syndecan 1 by PPARγ resulted in apoptosis induction [76].


### 4.2. Other Targets of PPARγ

PPARγ impacts
many growth-promoting elements through its secondary actions that, while ligand-dependent,
do not directly involve its gene promoters. It interacts with nuclear factor of
activated T cells, phosphorylated signal transducer, and activator of
transcription (STAT)-3, and nuclear factor κB (NFκB) to block signaling through
these pathways [3]. It binds transcription
cofactors to alter these cofactors' availability to other transcription factors:
ligand bound-PPARγ deprives NFκB of AP-1; deprives STAT-1 of CREB binding
protein; and releases SMRT to render it available to repress STAT-3's
transcriptional activity [3, 16, 17, 97]. PPARγ activation is also associated
with the activation of ERK1/2, protein kinases C, protein kinase A,
AMP-activated protein kinase α [17]; induction of p16, p18, and
p21 cyclin-dependent kinase inhibitors [3, 17, 18]; decreased expression of
cyclooxygenase 2, cmyc, cmyb, D1, and D3 cell cycle control
genes, and regenerating gene 1A [17, 18]; decreased secretion of
cytokines and growth factors [17, 98]; depression of the Akt
survival pathway by upregulating PTEN and inhibiting the phosphorylation of Akt
and mTOR [3, 17]; inhibiting retinoblastoma
protein (Rb) activity to repress the activities of cyclins D3 and E [3]; and regulating a host of
other elements involved in the growth and death of cells [3, 12, 16–18]. It is not clear which if any
of these effects are due to the action of PPARγ or PPARγ activators. PUFAs impact many of these same targets but
can do so not only by PPARγ-dependent but also PPARγ-independent routes (see
the next section).


### 4.3. Targets of PPARγ-Activating Ligands

Studies of
PPARγ function depend on challenging cells with PPAR-activating ligands that have
numerous side effects impacting cell growth. 15-d-Δ12,14-PGJ2 has a reactive α,β-unsaturated ketone (Figure 3) that covalently binds to cysteine
sulfur on PPARγ; this renders its PPARγ binding irreversible [58, 68]. 15-d-Δ1,14-d-PGJ2 also binds to cysteines in the IKKβ subunit of IκB kinase, thereby inhibiting
NFκB activation [99, 100]. Other ligands with an α,β
unsaturated ketone (e.g., oxo-ODEs and oxo-ETEs; see Figure 3) have this chemical
reactivity [58] and along with 15-d-Δ1,14-d-PGJ2 may exert anticancer effects by covalently attaching to signal molecules like
IKKβ [58, 99, 101] or elements needed for expressing
the epidermal growth factor receptor (EGFR) and JAK [102, 103].Naturally
occurring ligands have other PPARγ-independent effects. The D and J series of PGs
including 15-d-Δ12,14-PGJ2 bind to PGD2 receptors [104], 5-oxo- and 5-oxo-15-hydroxy-ETE
bind to the OXE receptor [105], and AA, EPA, and DHA bind to
GPR40 and GPR120 receptors [106, 107]. These G protein-coupled
receptors regulate signal pathways that effect cancer cell growth. For example,
5-oxo-15-hydroxy ETE acts on OXE to stimulate cells to activate ERK and Akt and
proliferate; this stimulation counters its antigrowth activity in various cancer
cell types. Indeed, HEK293 cells lack OXE receptors and in contrast to OXE
receptor-bearing breast, prostate and ovarian cancer cell lines respond to
5-oxo-ETE and 5-oxo-15-oxo-ETE only by slowing, not speeding, their
proliferation [43]. PUFAs activation of GPR120
also causes ERK and Akt activation to increase the survival of serum-starved
STC-1 cells [108]. Finally, PUFAs are also
metabolized to products that act on G protein receptors to promote cell growth,
for example, prostate cancer cells convert AA to PGE2, which acts
through its receptors to stimulate the NFκB pathway and thereby the expression
of various cytokines and growth factors [109]. The G protein
receptor-dependent actions of PPARγ ligands may explain reports that these
ligands have biphasic effects in stimulating proliferation and antiproliferation
in cancer cells [110].Thiazolidinediones
stimulate cells to activate ERK1/2, p38, and JNK [111–113] by discharging Ca2+  from the ER to evoke an ER stress response; this activates Ca2+/calmodulin
kinase II, proline-rich tyrosine kinase 2, protein kinases C, c-Src, EGFR, the ERK1/2 and JNK pathways,
the double stranded RNA-activated protein kinase, and p38 [111]. Double stranded RNA-activated
protein kinase inactivates eukaryotic initiation factor-2 to depress protein
translation [111, 114]. Since EPA has recently been shown
to have similar effects on ER calcium discharge [111, 115], it seems likely that various
other PUFAs activate the ER stress pathway. Nonetheless, PPARγ activators often
show very different side effects [42, 103, 116–120]. For example, among three
PPARγ agonists, ciglitazone, 9-HODE, and 13-HODE, only 9-HODE induced apoptosis
in U937 cells  [38], 15d-Δ12,14-PGJ2,
but not various other PPARγ ligands, reduced EGFR expression in squamous carcinoma cells
[99], 15d-Δ12,14-PGJ2,
but not troglitazone, inhibited the stimulated induction of MHC class II
molecules in retinal pigmented epithelial cells  [112], and DHA, but not EPA, stimulated
the target gene, syndecan 1 to inhibit the proliferation and induce apoptosis
in breast and prostate cancer cell lines [75, 76, 96]. Numerous other examples of differential
effects among PPARγ agonists exist (e.g.,  [113–116]), but it is worth stressing
that n-3 PUFAs inhibit the metabolism of n-6 PUFAs to products that promote the
growth of cancer cells such as PGE2, 5-HETE, and leukotriene B4
[33–35, 45, 113]. This inhibitory effect may
make an important contribution to the anticancer effects of n-3 PUFAs.


## 4.1. Gene Targets of PPARγ

Most known target
genes of PPARγ regulate lipid metabolism and transport [15]with few cancer-related genes
having been confirmed as induced by PPARγ. PPARγ does induce G0/G1 switch gene 2 whose product causes growth arrest in 3T3-L1 cells [91, 92]. PPARγ also binds the NFκB promoter of p53 to
stimulate expression of p53 and, in consequence, p21WAF1/Cip1. It
also binds to a promoter in the Fas ligand gene to induce the expression of
this member of the extrinsic apoptosis pathway. These effects appear
responsible for slowing growth and causing apoptosis in MCF7 breast cancer [93], human umbilical vein
endothelial [94], and possibly Reh [95]  cells. Recent studies have
identified the heparan sulfate proteoglycan, syndecan 1, as a target for PPARγ in human breast [75, 76] and prostate [96] cancer cells. The upregulation
of syndecan 1 by PPARγ resulted in apoptosis induction [76].


## 4.2. Other Targets of PPARγ

PPARγ impacts
many growth-promoting elements through its secondary actions that, while ligand-dependent,
do not directly involve its gene promoters. It interacts with nuclear factor of
activated T cells, phosphorylated signal transducer, and activator of
transcription (STAT)-3, and nuclear factor κB (NFκB) to block signaling through
these pathways [3]. It binds transcription
cofactors to alter these cofactors' availability to other transcription factors:
ligand bound-PPARγ deprives NFκB of AP-1; deprives STAT-1 of CREB binding
protein; and releases SMRT to render it available to repress STAT-3's
transcriptional activity [3, 16, 17, 97]. PPARγ activation is also associated
with the activation of ERK1/2, protein kinases C, protein kinase A,
AMP-activated protein kinase α [17]; induction of p16, p18, and
p21 cyclin-dependent kinase inhibitors [3, 17, 18]; decreased expression of
cyclooxygenase 2, cmyc, cmyb, D1, and D3 cell cycle control
genes, and regenerating gene 1A [17, 18]; decreased secretion of
cytokines and growth factors [17, 98]; depression of the Akt
survival pathway by upregulating PTEN and inhibiting the phosphorylation of Akt
and mTOR [3, 17]; inhibiting retinoblastoma
protein (Rb) activity to repress the activities of cyclins D3 and E [3]; and regulating a host of
other elements involved in the growth and death of cells [3, 12, 16–18]. It is not clear which if any
of these effects are due to the action of PPARγ or PPARγ activators. PUFAs impact many of these same targets but
can do so not only by PPARγ-dependent but also PPARγ-independent routes (see
the next section).


## 4.3. Targets of PPARγ-Activating Ligands

Studies of
PPARγ function depend on challenging cells with PPAR-activating ligands that have
numerous side effects impacting cell growth. 15-d-Δ12,14-PGJ2 has a reactive α,β-unsaturated ketone (Figure 3) that covalently binds to cysteine
sulfur on PPARγ; this renders its PPARγ binding irreversible [58, 68]. 15-d-Δ1,14-d-PGJ2 also binds to cysteines in the IKKβ subunit of IκB kinase, thereby inhibiting
NFκB activation [99, 100]. Other ligands with an α,β
unsaturated ketone (e.g., oxo-ODEs and oxo-ETEs; see Figure 3) have this chemical
reactivity [58] and along with 15-d-Δ1,14-d-PGJ2 may exert anticancer effects by covalently attaching to signal molecules like
IKKβ [58, 99, 101] or elements needed for expressing
the epidermal growth factor receptor (EGFR) and JAK [102, 103].Naturally
occurring ligands have other PPARγ-independent effects. The D and J series of PGs
including 15-d-Δ12,14-PGJ2 bind to PGD2 receptors [104], 5-oxo- and 5-oxo-15-hydroxy-ETE
bind to the OXE receptor [105], and AA, EPA, and DHA bind to
GPR40 and GPR120 receptors [106, 107]. These G protein-coupled
receptors regulate signal pathways that effect cancer cell growth. For example,
5-oxo-15-hydroxy ETE acts on OXE to stimulate cells to activate ERK and Akt and
proliferate; this stimulation counters its antigrowth activity in various cancer
cell types. Indeed, HEK293 cells lack OXE receptors and in contrast to OXE
receptor-bearing breast, prostate and ovarian cancer cell lines respond to
5-oxo-ETE and 5-oxo-15-oxo-ETE only by slowing, not speeding, their
proliferation [43]. PUFAs activation of GPR120
also causes ERK and Akt activation to increase the survival of serum-starved
STC-1 cells [108]. Finally, PUFAs are also
metabolized to products that act on G protein receptors to promote cell growth,
for example, prostate cancer cells convert AA to PGE2, which acts
through its receptors to stimulate the NFκB pathway and thereby the expression
of various cytokines and growth factors [109]. The G protein
receptor-dependent actions of PPARγ ligands may explain reports that these
ligands have biphasic effects in stimulating proliferation and antiproliferation
in cancer cells [110].Thiazolidinediones
stimulate cells to activate ERK1/2, p38, and JNK [111–113] by discharging Ca2+  from the ER to evoke an ER stress response; this activates Ca2+/calmodulin
kinase II, proline-rich tyrosine kinase 2, protein kinases C, c-Src, EGFR, the ERK1/2 and JNK pathways,
the double stranded RNA-activated protein kinase, and p38 [111]. Double stranded RNA-activated
protein kinase inactivates eukaryotic initiation factor-2 to depress protein
translation [111, 114]. Since EPA has recently been shown
to have similar effects on ER calcium discharge [111, 115], it seems likely that various
other PUFAs activate the ER stress pathway. Nonetheless, PPARγ activators often
show very different side effects [42, 103, 116–120]. For example, among three
PPARγ agonists, ciglitazone, 9-HODE, and 13-HODE, only 9-HODE induced apoptosis
in U937 cells  [38], 15d-Δ12,14-PGJ2,
but not various other PPARγ ligands, reduced EGFR expression in squamous carcinoma cells
[99], 15d-Δ12,14-PGJ2,
but not troglitazone, inhibited the stimulated induction of MHC class II
molecules in retinal pigmented epithelial cells  [112], and DHA, but not EPA, stimulated
the target gene, syndecan 1 to inhibit the proliferation and induce apoptosis
in breast and prostate cancer cell lines [75, 76, 96]. Numerous other examples of differential
effects among PPARγ agonists exist (e.g.,  [113–116]), but it is worth stressing
that n-3 PUFAs inhibit the metabolism of n-6 PUFAs to products that promote the
growth of cancer cells such as PGE2, 5-HETE, and leukotriene B4
[33–35, 45, 113]. This inhibitory effect may
make an important contribution to the anticancer effects of n-3 PUFAs.


## 5. Dietary Fatty Acids and Cancer

## 5.1. Human Studies

Although there are inconsistencies [121], human population studies
have shown that consumption of a diet enriched in n-3 PUFAs may offer
protection against a number of cancers including those of breast [122–124], prostate [125, 126], and colon [127–129]. Although many of these
studies have relied on dietary intake data from self-reported questionnaires or
estimates based on national consumption, a few have used the fatty acid composition
of tissues as a measure of exposure to dietary fats. The EURAMIC study is one
of the largest to provide evidence that the balance between n-3 and n-6 PUFA
may play a role in breast cancer [130]. Adipose tissue aspirates
from breast cancer patients and controls demonstrated that the ratio of long
chain n-3 to n-6 PUFAs was inversely associated with breast cancer in four of
five centers studied. In human prostate tissue, lower EPA and DHA as well as lower
n-3 to n-6 PUFAs ratios were associated with cancer compared to benign prostate
hyperplasia [131] and with advanced stage
compared to organ confined disease [132]. This inverse association of n-3
PUFAs and prostate cancer is supported by analyses of fatty acids in serum and
red-cell membranes of patients with prostate disease [133, 134].


## 5.2. Animal Studies

Animal studies provide convincing
evidence of a negative relationship with n-3 PUFA diets and a positive
relationship with n-6 PUFA diets for breast, prostate, and colon cancer. In
studies of breast cancer induced by chemical carcinogens in rats [135–137], and human cancer cell
xenografts in nude mice [138–140], tumor growth rate, size, and
metastases were all suppressed by dietary n-3 PUFA supplementation. Likewise
for colon cancer, antitumor properties of n-3 PUFA diets have been shown in transplantable
mouse tumors [141–143] as well as in chemically
induced rat tumors [144–151]. Although there have been
fewer animal studies of PUFAs in prostate cancer, they are consistent with
those in breast and colon cancer. In xenograft models of prostate cancer, n-3
PUFAs enriched diets inhibited tumor growth compared to n-6 PUFA diets [152–154]. Recently, a prostate-specific Pten knockout
mouse model was used to demonstrate that a dietary ratio of n-6 to n-3 PUFA
lower than 5 was effective in suppressing tumor growth, and extending animal
lifespan [155].


## 5.3. Cell Culture Studies

Insight into the mechanism(s)
responsible for the anticancer properties of n-3 PUFAs have been provided by
animal studies as well as by in vitro investigations using human cancer cell
lines. A major focus for such studies has been the competitive inhibition between
n-6 and n-3 PUFAs for the enzymes involved in their metabolism. The
desaturation and elongation of LA to AA were shown to be decreased in the
presence of high n-3 PUFAs due to enzyme preference for the n-3 substrates [156]. AA and EPA compete for the COX
and LOX enzymes, again with preferential n-3 utilization that results in a
reduction in the highly reactive eicosanoids generated from AA [157, 158] in favor of less inflammatory
n-3 eicosanoids [159]. The decreased growth of
prostate xenograft tumors was shown to involve inhibition of COX 2 and PGE2 in the tissues [154]. Thus, the combined human,
animal, and cell culture studies indicate that diet is an important regulator
of the levels of n-3 versus n-6 PUFAs in tissues, including those that are cancerous.
High levels of n-3 PUFAs may directly evoke antitumor events, become
metabolized to products with antitumor activity, or suppress the production of
tumor-promoting metabolites such as those formed by n-6 PUFAs.


## 6. N-3 Pufa Regulation of Syndecan-1

Increasing evidence implicates PPARγ in the divergent effects of n-3 and n-6
PUFAs in cancer cells and point to a growth inhibitory role for PPARγ [160–164]. We recently found that n-3
PUFAs—but not n-6 PUFAs—enriched LDL,
inhibited the proliferation, and induced apoptosis in human breast cancer cells [74–76]. The n-3 LDL delivered both
EPA and DHA to the cells. When these individual fatty acids were delivered to
cells by albumin, DHA but not EPA proved effective in stimulating apoptosis in
a pathway that involved activation of PPARγ [75]. The molecular target for
both DHA and PPARγ in these cells was shown to be the
heparan sulfate proteoglycan, syndecan-1. Syndecan-1 itself was effective in
apoptosis induction and when syndecan-1 was silenced, the ability of DHA to
induce apoptosis was completely blocked as it was in the presence of a dominant negative
PPARγ [76]. Moreover, syndecan-1 siRNA
was effective in blocking troglitizone-induced apoptosis. Thus, a novel pathway
linking n-3 PUFAs to apoptosis in tumor cells is as follows: DHA activates PPARγ, which results in transcriptional
upregulation of the syndecan-1 target gene, and the syndecan-1 protein induces
apoptosis (Figure 4). This novel pathway has been confirmed in human prostate
cancer cells [96].Figure 4
The syndecan-1 pathway for n-3 PUFA induction of apoptosis. Dashed lines
indicate that effects may be indirect with involvement of other metabolites and signaling
molecules.Although PPARγ was not a target for EPA in breast and
prostate cancer cells, a recent report has demonstrated that EPA was an
effective PPARγ transactivator in HT-29 human colon
cancer cells [165]. In contrast, both EPA and
DHA were shown to reduce PPRE reporter activity in an HCT-116 colon cancer
cells [166]. DHA has recently been shown
to reduce the growth of human lung cancer cells in a process that was
associated with increased PPARγ protein [167]. These conflicting reports
are consistent with data showing selective modulation of PPARγ by different ligands in different cells [168]. Several other reasons may be proposed for the
differential response to DHA and EPA in the breast and prostate tumor cells
including (1) PPARγ activation may be mediated by a unique
DHA metabolite rather than DHA itself; (2) there may be a difference in the
bioavailability of the two fatty acids following cellular uptake; (3) EPA may
be a ligand for or metabolized to a ligand (e.g., 5(S)-hydroxy-eicosapentaenoic
acid) for a G protein-coupled receptor that activates ERK and thereby inactivates
or in some other way counteracts PPARγ; (4) EPA may directly, or after being
metabolized, activate other pathways that counteract PPARγ signaling.The identification of syndecan-1 as
a target gene for PPARγ in the breast and prostate cancer cells
was a novel but not unexpected finding. The syndecan-1 promoter contains a DR-1
element that is recognized by a several members of the nuclear hormone receptor
superfamily including PPARγ. Although there are conflicting reports
of a role for syndecan-1 in cancer, the importance of these studies is the
identification of a PPARγ molecular target that is regulated by
PUFAs and results in functional response in the tumor cells. As more such
targets emerge, we may be able to understand how different dietary fatty acids
play divergent roles in cancer.



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*Source: 358052-2008-08-27.xml*