Post on 12-Feb-2022
University of KentuckyUKnowledge
University of Kentucky Doctoral Dissertations Graduate School
2007
MODULATION OF ENDOTHELIAL CELLACTIVATION BY OMEGA-6 AND OMEGA-3FATTY ACIDSLei WangUniversity of Kentucky, leiwang210@gmail.com
Click here to let us know how access to this document benefits you.
This Dissertation is brought to you for free and open access by the Graduate School at UKnowledge. It has been accepted for inclusion in University ofKentucky Doctoral Dissertations by an authorized administrator of UKnowledge. For more information, please contact UKnowledge@lsv.uky.edu.
Recommended CitationWang, Lei, "MODULATION OF ENDOTHELIAL CELL ACTIVATION BY OMEGA-6 AND OMEGA-3 FATTY ACIDS" (2007).University of Kentucky Doctoral Dissertations. 573.https://uknowledge.uky.edu/gradschool_diss/573
ABSTRACT OF DISSERTATION
Lei Wang
The Graduate School
University of Kentucky
2007
MODULATION OF ENDOTHELIAL CELL ACTIVATION BY OMEGA-6 AND OMEGA-3 FATTY ACIDS
ABSTRACT OF DISSERTATION
A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the
Graduate Center for Nutritional Sciences at the University of Kentucky
By
Lei Wang
Lexington, Kentucky
Advisor: Dr. Bernhard Hennig, Professor of Nutrition and Toxicology
Lexington, Kentucky
2007
Copyright © Lei Wang 2007
ABSTRACT OF DISSERTATION
MODULATION OF ENDOTHELIAL CELL ACTIVATION BY OMEGA-6 AND OMEGA-3 FATTY ACIDS
Endothelial activation is considered to be an early and critical event in the pathology of atherogenesis which can be modified by environmental factors such as diet, pollutants, and lifestyle habits. Dietary ω-6 and ω-3 fatty acids have been reported to either amplify or diminish inflammatory responses related to atherosclerosis development. However, the interactions of ω-6 and ω-3 fatty acids with inflammatory cytokines or organic pollutants on endothelial cell activation are not well understood. The studies presented in this dissertation tested the hypothesis that ω-6 and ω-3 fatty acids alone, or in varying ratios can differently modulate pro-atherogenic mediators and inflammatory responses that are initiated by tumor necrosis factor- α (TNF-α) or polychlorinated biphenyls (PCBs) in endothelial cells. Exposure to TNF-α induced oxidative stress, p38 MAPK, NF-κB, COX-2 and PGE2, which was amplified by pre-enrichment with linoleic acid but blocked or reduced by α-linolenic acid. Furthermore, TNF-α-induced caveolin-1 up-regulation and the co-localization of TNF receptor-1 with caveolin-1 was markedly increased in the presence of linoleic acid and diminished by α-linolenic acid. Silencing of the caveolin-1 gene completely blocked TNF-α-induced production of COX-2 and PGE2 and significantly reduced the amplified response of linoleic acid plus TNF-α. These data suggest that omega-6 and omega-3 fatty acids can differentially modulate TNF-α-induced inflammatory stimuli and that caveolae and its fatty acid composition play a regulatory role in these observed metabolic events. Besides cytokines, lipophilic environmental contaminants such as PCBs can also trigger inflammatory events in endothelial cells. Our data suggest that increasing the relative amount of α-linolenic acid to linoleic acid can markedly decrease oxidative stress and NF-κB-responsive genes. The inhibitor study revealed that the modulation effect of ω-6 and ω-3 fatty acids on PCB toxicity was mainly through the oxidative stress sensitive transcription factor, NF-κB. In conclusion, our studies demonstrate that different dietary fats can selectively modulate vascular cytotoxicity caused by TNF-α as well as by persistent organic pollutants such as PCBs. We also demonstrated the important relevance of substituting dietary ω-3 fatty
acids such as α-linolenic acid for ω-6 fatty acid such as linoleic acid in reducing cardiovascular diseases.
KEYWORDS: Atherosclerosis, fatty acids, inflammation, TNF-α, PCB
_ Lei Wang _____________
_________12/06/2007_________________
MODULATION OF ENDOTHELIAL CELL ACTIVATION BY OMEGA-6 AND OMEGA-3 FATTY ACIDS
By
Lei Wang
______ Bernhard Hennig__________ Director of Dissertation
_______ Steven Post ___________
Director of Graduate Studies
________12/06/2007_____________ Date
RULES FOR THE USE OF DISSERTATIONS Unpublished dissertations submitted for the Doctor’s degree and deposited in the University of Kentucky Library are as a rule open for inspection, but are to be used only with due regard to the rights of the authors. Bibliographical references may be noted, but quotations or summaries of parts may be published only with the permission of the author, and with the usual scholarly acknowledgements. Extensive copying or publication of the dissertation in whole or in part also requires the consent of the Dean of the Graduate School of the University of Kentucky. A library that borrows this dissertation for use by its patrons is expected to secure the signature of each user. Name Date
DISSERTATION
Lei Wang
The Graduate School
University of Kentucky
2007
MODULATION OF ENDOTHELIAL CELL ACTIVATION BY OMEGA-6 AND OMEGA-3 FATTY ACIDS
DISSERTATION
A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the
Graduate Center for Nutritional Sciences at the University of Kentucky
By
Lei Wang
Lexington, Kentucky
Advisor: Dr. Bernhard Hennig, Professor of Nutrition and Toxicology
Lexington, Kentucky
2007
Copyright © Lei Wang 2007
Acknowledgments
First of all, I want to sincerely thank my advisor Dr. Bernhard Hennig, who
provided me the great opportunity to join the Graduate Center for Nutritional Sciences
for my training as a Ph.D. student in 2003. It was my most valuable four years’ studying
time at the University of Kentucky. Dr. Bernhard Hennig not only gave me advice in
academic aspect but also taught me how to handle challenges and frustrations in an
optimal attitude in all the other aspects of life. All of these will company me in my future
career. I would also like to express my gratitude to all the members of my Ph.D.
committee: Drs. Steve Post, Michal Toborek, and Ming Gong who were very generous in
sharing their expertise and support to any research related questions I had during my time
as a graduate student. In general, I would also like to thank all the faculties, staffs,
research fellows, and students on the fifth floor of C.T.Wethington building for creating
such a supportive and enjoyable working environment.
In addition, I would like to thank all my wonderful laboratory coworkers: Drs.
Meerarani Purushothaman, Viswanathan Saraswathi, Gudrun Reiterer, Eun Jin Lim,
Xabier Arzuaga for their guidance and advice on my research. Elizabeth Oesterling and
Zuzana Majkova for their valuable comments and editing of all my conference abstracts,
poster presentations, and manuscripts. Drs. Sung Yong Eum, Ibolya Andras, and Yu
Zhong from Dr. Toborek’s laboratory for giving me advice on experimental design or
answering any of my technical questions at any time.
Last but not least I would like to acknowledge my family: my parents Chengshan
Wang and Haiyu Zhu, grandparents Tingyun Zhu and Peiwen Gao. Despite the long
distance, their encouragement and support were very valuable to me. Especially I would
also like to dedicate this work to my husband Xiaohu Feng. It had been such a wonderful
time to meet him at the University of Kentucky, and could study and graduate together
from UK.
The research reported in this dissertation has been supported by grants from
NIH/NIEHS (P42 ES 07380), and the University of Kentucky Agricultural Experiment
Station.
iii
TABLE OF CONTENTS
Acknowledgments.............................................................................................................. iii List of Tables ..................................................................................................................... vi List of Figures ................................................................................................................... vii Chapter One : Introduction ................................................................................................. 1
Endothelial Cell Activation and Atherosclerosis........................................................ 1 Dietary Fatty Acids and Atherosclerosis .................................................................... 3 Pro-Inflammatory Cytokine TNF-α-Stimulated Endothelial Activation.................... 4 Significance of Caveolae in Fatty acids and TNF-α Mediated Endothelial Cell Activation.................................................................................................................... 6 PCB-Induced Endothelial Cell Activation.................................................................. 9 Hypothesis and Significance of the Current Study ................................................... 10
Chapter Two : Effects of ω-6 and ω-3 Fatty Acids on TNF-α-Induced Endothelial Cell Activation.......................................................................................................................... 15
Synopsis .................................................................................................................... 15 Introduction............................................................................................................... 16 Materials and Methods.............................................................................................. 19 Results....................................................................................................................... 23 Discussion................................................................................................................. 44
Chapter Three : Involvement of Caveolae in Regulation of Endothelial Cell Activation Mediated by Fatty Acids and TNF-α................................................................................ 47
Synopsis .................................................................................................................... 47 Introduction............................................................................................................... 48 Materials and Methods.............................................................................................. 49 Results....................................................................................................................... 52 Discussion................................................................................................................. 65
Chapter Four : Changing Ratios of ω-6 to ω-3 Fatty Acids Can Differentially Modulate Polychlorinated Biphenyl Toxicity in Endothelial Cells .................................................. 68
Synopsis .................................................................................................................... 68 Introduction............................................................................................................... 68 Materials and Methods.............................................................................................. 71 Results....................................................................................................................... 72 Discussion................................................................................................................. 90
Chapter Five : Summary ................................................................................................... 94 Conclusion ................................................................................................................ 94 Future Directions ...................................................................................................... 97
Appendix: Protocols........................................................................................................ 100 Primary Culture of Porcine Pulmonary Artery Endothelial Cells .......................... 100 Freeze Endothelial Cells ......................................................................................... 101 Start a Frozen Endothelial Cell Line....................................................................... 102 Cell Culture and Experimental Media .................................................................... 102 Preparation of Fatty Acid Enriched Media ............................................................. 103 Oxidative Stress Measurement ............................................................................... 104 PGE2 EIA kit........................................................................................................... 105 Immunofluorescence............................................................................................... 108 Confocal Microscopy.............................................................................................. 110
iv
Detergent-free Purification of Caveolin Enriched Membrane Fractios (Sucrose Gradient) ................................................................................................................. 111
References:...................................................................................................................... 113 Vita.................................................................................................................................. 127
v
List of Tables
Table 1.1 Major dietary polyunsaturated fatty acids and sources............................ 12
vi
List of Figures
Figure 1.1 Major TNF receptor-mediated cellular responses .................................. 13 Figure 1.2 Organization of lipid rafts and caveolae membranes ............................. 14 Figure 2.1 Optimal TNF-α concentration and incubation time for E-selectin
expression ......................................................................................................... 28 Figure 2.2 Immunofluorescence microscopy study of TNF-α-induced E-selectin
expression ......................................................................................................... 28 Figure 2.3 TNF-α-induced VCAM-1 and COX-2 expression................................. 29 Figure 2.4 Optimal linoleic acid incubation time for VCAM-1 gene expression.... 30 Figure 2.5 Effect of linoleic acid and α-linolenic acid on TNF-α-induced E-selectin
gene expression................................................................................................. 31 Figure 2.6 Effect of linoleic acid and α-linolenic acid on TNF-α-induced COX-2
gene expression................................................................................................. 32 Figure 2.7 Effect of linoleic acid and α-linolenic acid on TNF-α-induced COX-2
protein expression ............................................................................................. 33 Figure 2.8 Effect of linoleic acid and α-linolenic acid on TNF-α-induced oxidative
stress.................................................................................................................. 34 Figure 2.9 Effect of linoleic acid and α-linolenic acid on TNF-α-induced activation
of NF-κB........................................................................................................... 35 Figure 2.10 Effect of linoleic acid and α-linolenic acid on TNF-α-induced p38
MAPK activation .............................................................................................. 36 Figure 2.11 Effect of p38 MAPK inhibitor on TNF-α-induced COX-2 up-regulation
........................................................................................................................... 37 Figure 2.12 Effect of p38 MAPK inhibitor on TNF-α- and linoleic acid-induced
VCAM-1 up-regulation..................................................................................... 38 Figure 2.13 Effect of TNF-α on cPLA2 activation .................................................. 39 Figure 2.14 Effect of linoleic acid and α-linolenic acid on PGE2 production......... 40 Figure 2.15 Optimal TNF-α incubation time for PGE2 production......................... 41 Figure 2.16 Effect of linoleic acid and α-linolenic acid on TNF-α-induced PGE2
production ......................................................................................................... 42 Figure 2.17 Proposed mechanism for fatty acid-mediated modulation of endothelial
cell activation induced by TNF-α ..................................................................... 43 Figure 3.1 Effect of linoleic acid and α-linolenic acid on caveolin-1 expression ... 56 Figure 3.2 Immunofluorescence microscopy study of TNF-α-induced caveolin-1
up-reguation. ..................................................................................................... 57 Figure 3.3 Effect of linoleic acid and α-linolenic acid on TNF-α-induced TNFR-1
and caveolin-1 localization ............................................................................... 58 Figure 3.4 Effect of linoleic acid and α-linolenic acid on TNF-α-induced caveolin-1
expression ......................................................................................................... 59 Figure 3.5 Effects of fatty acids on TNF-α-induced caveolin-1 redistribution ....... 60 Figure 3.6 Effects of fatty acids on TNF-α-induced caveolin-1 redistribution ....... 61 Figure 3.7 Effect of caveolin-1 silencing on TNF-α-induced caveolin-1 up-
regulation .......................................................................................................... 62
vii
viii
Figure 3.8 Effect of caveolin-1 silencing on linoleic acid and TNF-α-induced caveolin-1 expression........................................................................................ 62
Figure 3.9 Effect of caveolin-1 silencing on linoleic acid and TNF-α-induced COX-2 expression ...................................................................................................... 63
Figure 3.10 Effect of caveolin-1 silencing on linoleic acid and TNF-α-induced PGE2 production ............................................................................................... 64
Figure 4.1 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-induced oxidative stress .................................................................................... 76
Figure 4.2 Effect of different ratios of linoleic acid to α-linolenic acid on the activation of NF-κB .......................................................................................... 77
Figure 4.3 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-induced activation of NF-κB ............................................................................ 78
Figure 4.4 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-induced VCAM-1 gene expression................................................................... 79
Figure 4.5 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-induced VCAM-1 protein expression ............................................................... 80
Figure 4.6 Effect of PCB77 on cPLA2 phosphorylation.......................................... 81 Figure 4.7 Effect of PCB77 on COX-2 expression.................................................. 81 Figure 4.8 Effect of PCB77 on PGE2 synthase expression...................................... 82 Figure 4.9 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced COX-2 gene expression ...................................................................... 83 Figure 4.10 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced COX-2 protein expression................................................................... 84 Figure 4.11 Optimal PCB77 incubation time for PGE2 production......................... 85 Figure 4.12 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
stimulated release of PGE2................................................................................ 86 Figure 4.13 Effect of NF-κB inhibitor on PCB77- or linoleic acid-induced VCAM-1
expression ......................................................................................................... 87 Figure 4.14 Effect of NF-κB inhibitor on PCB77- or linoleic acid-induced COX-2
expression ......................................................................................................... 88 Figure 4.15 Proposed mechanism for fatty acid-mediated modulation of endothelial
cell activation induced by PCB77..................................................................... 89 Figure 5.1 Summery................................................................................................. 99 Figure 5.2 Future direction ...................................................................................... 99
Chapter One : Introduction
Endothelial Cell Activation and Atherosclerosis
Cardiovascular disease is the number one killer worldwide, including in the
United States. For example, from the United States Chronic Disease Overview (1999),
cardiovascular disease is responsible for more than 30 percent of all the deaths in the
United States, which is more than all forms of cancer which is 23 percent of deaths.
Cardiovascular disease can refer to many different types of heart or blood vessel
problems and atherosclerosis is the generally the cause of all these problems.
Atherosclerosis is an inflammatory disease of the vessel wall which predominantly
involves endothelium and vascular smooth muscle layers of the arterial wall. Vascular
inflammation is a critical event in the pathogenesis of many human diseases, including
atherosclerosis, hypertension, autoimmune diseases [1-3]. Progression of atherosclerosis
can lead to the development of a plaque that is vulnerable to rupture and would then
produce acute coronary syndromes, such as arterial thrombosis and myocardial infarction
or stroke [4].
Numerous risk factors associated with atherosclerosis have been identified, such
as hypertension, diabetes mellitus, and dyslipidemia. These particular cardiovascular risk
factors commonly occur in obese individuals as components of the metabolic syndrome
[5]. Furthermore, smoking, obesity, insulin resistance, hypertriglyceridemia, elevated
levels of plasma cholesterol, low high-density lipoprotein, shear stress, and oxidative
stress can also increase the risk of atherosclerosis [6, 7]. A growing body of literature
also indicates that environmental factors such as diet, lifestyle habit, and nutrition factors
can play an important role in the development of atherosclerosis. Substantial evidence
from epidemiological studies suggests that cardiovascular diseases are linked to
environmental pollution. For example, there was a significant increase in mortality from
cardiovascular diseases among Swedish capacitor manufacturing workers exposed to
polychlorinated biphenyls (PCBs) for at least five years [8], and most excess deaths were
1
due to cardiovascular disease in power workers exposed to phenoxy herbicides and PCBs
in waste transformer oil [9].
Atherosclerosis depends critically on altered behavior of the intrinsic cells of the
artery wall, in particular the endothelial cells. Endothelial cells are constantly exposed to
blood flow and are directly in contact with blood components. These components
include various kinds of cells, cytokines, toxic wastes, and dietary nutrients which might
be accompanied with environmental pollutant. In healthy condition, endothelial cell
surface of the lumen is non-adhesive and anti-thrombogenic. But in certain diseases,
endothelial cell activation has been considered to trigger the initial development of
atherosclerosis which involves the development of numerous oxidative and inflammatory
reactions and will result in lymphocyte and monocyte adhesion to the vascular
endothelium [10]. Oxidized LDL is one of the important triggers for the formation of the
atherosclerotic lesion. It may enhance migration and localization of inflammatory cells
including T lymphocytes, monocytes, and macrophages [11]. Up-regulation of adhesion
molecules on the surface of endothelial cells is prominent when they are exposed to pro-
inflammatory molecules such as tumor necroses factor-α (TNF-α), interleukin-1β (IL-
1β), interferon-γ (IFN-γ), platelet-derived growth factor (PDGF), and vascular endothelial
growth factor (VEGF) [12, 13]. In sites of inflammation, leukocytes will migrate across
activated endothelium. This is a three-step process: the first step is the rolling of
leukocytes on the endothelium which is mediated by selectins and addressins [14]; the
second step is the attachment of leukocytes to the endothelium which is mediated by
vascular cell adhesion molecule-1 (VCAM-1) and intercellular adhesion molecule-1
(ICAM-1); the third step is the transmigration of leukocytes across the thin layer of
endothelial cells [15, 16]. Accumulation of monocytes in vascular subendothelial spaces
and their conversion into lipid-laden “foam cells” is an early and important event in
atherosclerosis [17]. Severe endothelial cell activation and injury can lead to necrotic and
apoptotic cytotoxicity, and ultimately to disruption of endothelial integrity.
Overall, the inflammatory reaction in endothelial cells is mediated by complex
interactions between circulating nutrients, cytokines and environmental pollutants.
Research in the current dissertation is trying to address the effects of dietary essential
2
omega-6 or omega-3 fatty acids on pro-inflammatory cytokine tumor necrosis factor-α
(TNF-α) and environmental contaminant PCB77 on the activation of endothelial cells.
Dietary Fatty Acids and Atherosclerosis
Prospective and cross-sectional studies indicate that the dyslipidemia, insulin
resistance and type 2 diabetes is an important contributor to atherosclerotic disease [18].
The chronic elevation of serum free fatty acids, also referred as albumin-bound non-
esterified fatty acids is important component of the dyslipidemia and have been found to
be associated with metabolic risk markers for coronary heart disease [19] and increased
risk of cardiovascular diseases [20] . The composition of intracellular non-esterified fatty
acids pool is affected by the concentration of exogenous fatty acids. The albumin-bound
fatty acid can originate from distal lipolysis of triglycerides from adipose tissue and focal
lipolysis of the triglycerides of VLDL and chylomicrons. During the postprandial period
albumin-bound free fatty acids mainly come from hydrolysis of triglyceride-rich
lipoproteins of intestinal origin by the lipoprotein lipase.
Diets high in omega-6 fatty acids have been shown to increase the risk of
cardiovascular diseases. In contrast, due to the low rates of coronary heart disease in
Greenland Eskimos and Japanese who are exposed to a diet rich in fish oil, the beneficial
effects of omega-3 fatty acids on cardiovascular health have been well documented. The
major dietary sources of omega-6 and omega-3 fatty acids are listed in Table 1. Linoleic
acid, the major omega-6 fatty acids in the American diet, is considered to be pro-
inflammatory and pro-atherogenic as it favors oxidative modification of LDL cholesterol,
increases platelet response to aggregation, and suppresses the immune system [21].
While, omega-3 fatty acids from fish oil have been shown to markedly reduce mortality
from cardiovascular diseases [22-24]. Furthermore, a higher intake of α-linolenic acid
which is enriched in vegetable oil has also been shown to inversely associate with risk of
myocardial infarction [25] and carotid atherosclerotic disease [26]. In vitro studies also
have shown that omega-3 fatty acids can down-regulate the expression of vascular
adhesion molecules in endothelial cells [27]. There is evidence that intake of omega-3
fatty acids are reflected in the concentration of these fatty acids not only in serum
3
phospholipids [28], but also in serum non-esterified fatty acids [28, 29]. In addition,
accumulation of linoleic acid can affect endothelial cell lipid pools [30, 31]. However the
primary mechanisms of the beneficial effects of dietary omega-3 especially α-linoleic
acid on endothelial cell activation and the interaction of omega-3 with omega-6 fatty
acids have not been well studied yet. The human body can convert 18 carbon fatty acids
into their 20 or longer chain derivatives via elongation and desaturation reactions. In the
current study, we focused on the 18-carbon linoleic acid and α-linolenic acid because
they are the major parental essential fatty acids for the omega-6 and omega-3 fatty acid
families, respectively, and are more sufficient in the human diet than their longer chain
derivatives.
Pro-Inflammatory Cytokine TNF-α-Stimulated Endothelial Activation
Tumor necrosis factor (TNF) was isolated in 1984 and was named after its ability
to cause tumor necrosis or regression in patients whose tumor has become infected with a
bacterial infection [32, 33]. TNF is also referred to as TNF-α, cachectin, or
differentiation-inducing factor (DIF) [34]. TNF-α can cause necrotic cell death, and may
also cause apoptotic cell death, cellular proliferation, differentiation and inflammation.
TNF-α is produced mostly by activated macrophages and monocytes, but also by many
other cell types including B lymphocytes, T lymphocytes and fibroblasts [35]. Fig.1.1
adapted from MacEwan et al [35], shows TNF mediated cellular responses.
TNF-α, a trimeric 17-kd cytokine, is one of the most potent pro-inflammatory
cytokines [36, 37] which is highly expressed throughout the full spectrum of
atherosclerotic development [38]. TNF-α has numerous pro-inflammatory and
procoagulant effects on endothelial cells [39]. It can stimulate production of reactive
oxygen species (ROS) through induction of enzymes such as xanthine oxidase, COX, and
nicotinamide adenine dinucleotide phosphate hydrogen (NADPH) oxidase as well as
other sources [40]. TNF-α increases recruitment of inflammatory cells to vascular lesion
sites [39], and promotes vascular smooth muscle cell adverse remodeling and plaque
4
rupture [41]. Further, it promotes matrix degradation [42], thus facilitate influx of
inflammatory cells to the vessel wall.
TNF-α effects on endothelial cells include protein synthesis-independent changes
in cell shape and motility that may contribute to vascular leakage at sites of inflammation
as well as expression of new proteins that may regulate other parameters of the
inflammatory response such as vasoregulation (e.g., COX-2 and GTP cyclohydrolase I),
leukocyte adhesion (e.g., E-selectin, ICAM-1, VCAM-1), leukocyte activation
(chemokines such as IL-8 or MCP-1), and coagulation (tissue factor and PAI-1). TNF-α
may also selectively turn off synthesis and eventually expression of some proteins such
as thrombomodulin or eNOS.
TNF-α engagement of TNF receptor-1 (TNFR-1) recruites the adaptor proteins
TRADD, TRAF-2 into lipid rafts and activates NF-κB and MAPK pathways [43, 44].
Two specific transcription factors, NF-κB and activator protein-1 (AP-1) are essential for
TNF-α induction of adhesion molecules. E-selectin, ICAM-1, and VCAM-1 genes each
contain DNA sequences in their 5’ flanking regions that bind various forms of NF-κB
and AP-1 and that mutations of these sequences reduce TNF-caused these adhesion
molecule up-regulation [45]. NF-κB is a DNA binding factor that can be activated by
several inflammatory mediators such as TNF-α, interferon (IFN)-β, interleukin (IL)-8,
IL-1β, IL-2, and IL-6 [46, 47]. The NF-κB complex consists of two heterodimers, p50
and p65. The activation of NF-κB requires the phosphorylation of NF-κB-inhibitor IκBα
and then free NF-κB dimmers can translocate into nucleus, bind to the promoter regions
of target genes, and induce transcription [47].
Besides the induction of adhesion molecules, most of cellular, tissue, and
systemic responses to TNF-α can be ascribed to TNF-α-induced PGE2 release.
Prostanoids are lipid-derived mediators involved in various homeostatic and
inflammatory processes through out the body. The biosynthesis of prostanoids involves
three enzymatic reactions including phospholipase A2 (PLA2) which liberates arachidonic
acid (AA) from the sn-2 position of cellular membrane phospholipids, cyclooxygenase
(COX) which converts AA to the endoperoxide intermediate Prostaglandin H2 (PGH2).
PGH2 is subsequently converted to prostanoids by the action of cell-specific synthases
5
which are also called terminal prostaglandin synthases [48]. After the formation,
prostanoids are released from the cells and act on their cognate receptors on cell surfaces
to exert their biological actions. Prostaglandin E2 (PGE2) which PGE synthase is the most
common prostanoids that can be produced by a wide variety of cells and tissues and has a
broad range of bioactivity. There are four types of PGE receptor, EP1, EP2, EP3, and
EP4 [49].
In the current study, we tested the hypothesis that omega-6 and omega-3 fatty
acids can differentially modulate TNF-α-induced endothelial cell activation via oxidative
stress-sensitive transcription factor NF-κB signaling pathway, thus influence the
inflammatory mediator PGE2 synthesis.
Significance of Caveolae in Fatty acids and TNF-α Mediated Endothelial Cell
Activation
In the 1950s the membrane invaginated vesicles of 50 to 100 nm were observed
by electron microscopy, which were given the name of caveolae due to their cave-like
appearance [50]. After the identification of caveolin, the signature protein of caveolae,
caveolae have been implicated to be important in a variety of cellular functions including
endocytic processes, cholesterol and lipid homeostasis, signal transduction, and tumor
suppression [51]. Fig.1.2 shows the organization of caveolae membranes as adapted
from Razani et al [51].
Caveolae have been considered a specialized form of lipid raft [52]. But certain
proteins have been found to preferentially partition into lipid rafts or caveolae but not
both [53]. Furthermore, the localization caveolin proteins to caveolae distinguishes these
membrane domains from lipid rafts [51]. Forty years after the definition of caveolae, the
22-kDa protein caveolin was isolated and subsequently cloned [54, 55]. There are three
gene family members of caveolin: caveolin-1, -2, and -3. Adipocytes, endothelial cells,
pneumocytes, and fibroblasts have the highest levels of caveolin-1 and -2 [56, 57],
whereas caveolin-3 expression is limited to muscle cell types [58].
Numerous roles have been attributed to caveolae proteins: vesicular transport [59],
cellular cholesterol homeostasis [60], oncogenes and tumorigenesis [61], and signal
6
transduction mechanisms [62, 63]. Several lines of research suggest that caveolin family
members function as scaffolding proteins [64] to concentrate and organize specific lipids,
such as cholesterol and sphingolipids [60, 65] and lipid-modified signaling molecules
within caveolae membranes [66-68].
Cholesterol and glycosphingolipids are essential components of caveolae. Sterol-
binding agents can bind to cholesterol and disrupt caveolar structure [69]. Also, utilize
cholesterol oxidase which reduces cellular cholesterol can cause the translocation of
caveolin-1 from the plasma membrane to intracellular membrane compartments [70].
Consequently, changes in the lipid component of caveolae membranes can potentially
result in profound changes in caveolar functioning. Thus, caveolae lipids are important
for organizing and regulating the molecular interactions of multiple signaling pathways.
Gafencu et al [71] investigated fatty acid composition of caveolae from
endothelial cells and indicated that palmitic acid (16:0), palmitoleic (16:1), stearic acid
(18:0), and oleic acid (18:1) were most prevalent which compose the 80% of total fatty
acids, linoleic acid (18:2) was about 8 to 10% of total fatty acids. Omega-3 fatty acids
have been shown to alter caveolae microenvironment, thereby modifying location and
function of proteins in caveolae. Eicosapentaenoic acid (EPA) induces displace caveolin-
1 from caveolae in endothelial cells [72] and omega-3 fatty acid feeding reduces both
cholesterol and caveolin-1 in mouse colon [73]. EPA and docosahexaenoic acid (DHA)
have also been shown to decrease lipid raft sphingomyelin, cholesterol content [74] and
to selectively alter subcellular distribution of cytosolic proteins, including Ras isoforms,
by modifying membrane lipid composition [75]. Furthermore, H-Ras has been shown to
target to the plasma membrane and caveolin is crucial for the function of H-Ras [76].
Caveolin-1 can bind to many types of plasma membrane receptor proteins or
membrane associated enzymes and concentrate these molecules within caveolae [51].
Fatty acid translocase (FAT/CD36), a major protein associated with fatty acid uptake [77],
is preferentially located in the caveolae at the plasma membrane level [78]. TNFR-1 has
been reported to concentrated within the caveolae of a human endothelial cell line [79].
Interleukin-1β-induced COX-2 in human fibroblasts is colocalized with caveolin-1 [80].
Previous studies demonstrated that in the absence of ligand, PDGF is associated with
rafts, and in the presence of ligand, it transiently associates with caveolae [81].
7
It has been widely accepted that binding of TNF-α to monomeric receptors results
in receptor trimerization and that this ligand-induced receptor clustering causes
recruitment of specific adapter signaling proteins to intracellular domains of receptors
that have been incorporated into these ligand-receptor complexes [82]. There are two
types of TNF receptor: TNFR-1 and TNFR-2. The TNFR1 is expressed on all cell types
including endothelial cells and is better understood, while the TNFR2 is mainly
expressed on cells of the immune system. TNFR-1 contains a death domain, termed
TNFR-associated death domain (TRADD) which is recruitment at the plasma membrane,
and this region is required for TNF-α-induced pro-inflammatory cellular responses, such
as activation of nuclear factor κB (NF-κB) [34]. In promonocyte cell line U937, if the
caveolae like domains are disrupted, the surface expression of TNFR-1 and CD36 are
significantly reduced [83]. The p38 MAPK pathway is activated by hydrogen peroxide
and UV light [84, 85]. p38MAPK, but not p42/44 is a critical component of the oxidant
stress (H2O2)-sensitive signaling pathways [85]. Phosphorylation of caveolin-1 occurs
specifically through activation of the p38 MAPK [86]. Activation of MAPKs has also
been shown to associate with subcellular redistribution of caveolin and cholesterol [87]
Caveolin-1 has been proposed to be essential in a variety of cellular functions.
Caveolin-1 knockout mice revealed that caveolin-1 is not essential for maintaining life
but plays an important role in vascular functions [88, 89]. Both in vivo and in vitro
experiments demonstrated that caveolin-1 is essential for caveolae formation and
interacts with a variety of signaling molecules in endothelial cells, suggesting that
caveolin-1 may be an essential signaling platform. In the current study, we tested the
hypothesis that omega-6 and omega-3 fatty acids can differentially modulate TNF-α-
induced endothelial cell activation and that functional caveolae are required for
endothelial cell activation. TNF-α-induced proinflammatory signaling pathways were
analyzed under experimental conditions with intact caveolae and with cells in which
caveolin-1 was silenced by siRNA.
8
PCB-Induced Endothelial Cell Activation
Polychlorinated biphenyls (PCB) are produced by chlorination of biphenyl and
comprise a class of 209 individual compounds, depending on the position of chlorine
atom substituents. The commercial use of PCBs started in the late 1920s including fluids
in transformers, capacitors and hydraulic transfer systems, plasticizers in paints, dyes,
plastics and rubber products etc. [90]. The total amount produced world-wide is
estimated at 1.5 million tons. PCBs have non-flammability, chemical stability and high
dielectric constants.
More than 90% of human PCBs exposure derives from food and PCBs are poorly
metabolized and tend to accumulate in animal tissues, including humans. The
accumulation particularly in tissues and organs rich in lipids appears to be higher in the
case of penta and more highly chlorinated biphenyls. Studies have revealed PCBs in
human fat tissue and blood plasma. [91-93]. Scientific Cooperation (EU SCOOP Task
3.2.5 June 2000) reported that the main contributors to the daily dietary exposure dioxins
appear to be fish and fish products (11-63%); meat and meat products (6-32%), milk and
diary products (16-39%). Following exposure to animals, PCB will accumulate in meat,
liver and particularly in fat tissues. PCBs build up (bioaccumulation) in the environment,
increasing in concentration as you move up the food chain. This is of special concern in
areas where fish are exposed to PCB contamination and may be consumed by humans (as
in the Hudson River).
PCBs were banned by the EPA in 1979, and are classified as a probable human
carcinogen by numerous national and international health-protective organizations, such
as the EPA, The Agency for Toxic Substances and Disease Registry (an arm of the U.S
Public Health Service) and the World Health Organization. Research also links PCB
exposure to developmental problems.
Numerous experimental and epidemiological studies have demonstrated that
environmental pollution is strongly involved in the pathogenesis of atherosclerosis.
Endothelial cells directly contact with blood substances which include environmental
pollutant. Certain polyhalogenated aromatic hydrocarbons can cause endothelial cell
activation, thus can be implicated in atherosclerotic lesion formation [94, 95]. Coplanar
PCBs, such as PCB77 can induce oxidative stress through the aryl hydrocarbon receptor
9
(AhR)-cytochrome P450 1A1 (CYP1A1) pathway [96]. Induction of CYP1A1 is
considered cardiotoxic through generating reactive oxygen species (ROS).
Previous studies have shown that environmental contaminants such as PCBs can
be implicated in atherosclerotic lesion formation [97, 98]. We have previously reported
that coplanar polychlorinated biphenyl (PCBs) such as PCB77 can induce inflammatory
genes VCAM-1 and IL-6 in vascular endothelial cells [96, 99]. PCBs can significantly
activate CYP1A1 enzyme [100]. PCB77, a coplanar PCB, is a 3-methylcholanthrene-
type cytochrome P450 inducer and an aryl hydrocarbon receptor (AhR) ligand.
Mechanisms of the interaction of ω-6 or ω-3 fatty acids and PCBs in endothelial
cell activation are not clear. Exposure to PCBs can change fatty acid composition of
tissues like the liver with a proportional decrease in arachidonic acid and increase in oleic
and linoleic acid [101, 102]. Long term exposure to PCBs have been shown to increase
plasma triglycerides and decrease HDL-cholesterol [103]. In the current study, we tested
the hypothesis that different types of dietary polyunsaturated fatty acids can selectively
modify PCB-induced vascular endothelial cell activation and the mechanisms of PCB77-
induced inflammation can be modified by different ratios of omega-6 to omega-3 fatty
acids.
Hypothesis and Significance of the Current Study
Previous studies have demonstrated that increasing levels of circulating cytokines
such as TNF-α, environmental contaminants such as PCBs, or certain dietary nutrients
such as omega-6 fatty acids can cause endothelial activation and promote an
inflammatory response [6, 104, 105]. Furthermore, our previous data have shown that
linoleic acid could potentiate inflammatory events induced by both TNF-α and PCBs
[106, 107]. Most of these inflammatory events involve oxidative stress and result in the
activation of oxidative stress sensitive transcription factors NF-κB or AP-1 which can
consequently stimulate the expression of cytokines and adhesion molecules [108, 109].
But the anti-atherogenic and anti-inflammatory effects of ω-3 fatty acids, in particular α-
linolenic acid, have not been investigated under these stimulations. And the modulation
effect of changing the relative amounts of omega-6 and omega-3 fatty acids on
10
environmental pollutant-induced endothelial cell activation has not been well elucidated
yet. It is possible that changes in the lipid (fatty acid) environment can alter the
membrane lipid rafts, inflammatory mediators, cell signal transduction, and gene
expression.
This study investigated the effects of omega-6 and omega-3 fatty acids on TNF-α
and PCB77-induced PGE2 release in endothelial cells. This was accomplished by
examining points of potential intervention in the cellular signaling cascades. We
explored the impact of these fatty acids alone, or at different ratios on TNF-α binding to
TNFR-1, cytoplasmic phospholipase A2 activation, and COX-2 expression. In addition,
we explored their impact on the binding activity of NF-κB, and signaling pathways such
as p38 MAPK.
Research described in this dissertation provides a better understanding of the
differential modulation effects by omega-6 and omega-3 fatty acids alone, or by changing
the ratios of omega-6 to omega-3 fatty acids on endothelial cell activation induced by
pro-inflammatory cytokine TNF-α or PCB77 and the underlying molecular mechanisms.
There is a great need to further explore this nutritional paradigm in environmental
toxicology and to improve our understanding of the relationship between nutrition and
lifestyle, exposure to environmental toxins and disease. Nutrition may provide the most
sensible means to develop primary intervention and prevention strategies of diseases
associated with many inflammatory and environmental toxic insults.
11
Table 1.1 Major dietary polyunsaturated fatty acids
Dietary Fatty acids
ω-6 or ω-3 families
Source
linoleic acid (LA; 18:2) ω-6 fatty acid soy, corn, safflower, and
sunflower oils
arachidonic acid (AA; 20:4) ω-6 fatty acid animal products
α-linolenic acid (ALA; 18:3) ω-3 fatty acid canola, walnut, and
flaxseed oils
eicosapentaenoic acid (EPA; 20:5) ω-3 fatty acid fish oil
docosahexaenoic acid (DHA; 22:6) ω-3 fatty acid fish oil
12
Figure 1.1 Major TNF receptor-mediated cellular responses
13
Figure 1.2 Organization of lipid rafts and caveolae membranes
A, lipid rafts: the liquid-ordered phase is dramatically enriched in cholesterol and
exoplasmic oriented sphingolipids (sphingomyelin and glycosphingolipids). The lipid-
disordered phase is composed essentially of phospholipids (phosphatidylcholine,
phosphatidylethanolamine, and phosphatidylserine). B, caveolae: upon integration of the
caveolin-1 protein, liquid-ordered domains form small flask-shaped invaginations called
caveolae. Caveolin-1 monomers assemble into discrete homo-oligomers containing ~14
to 16 individual caveolin molecules. Adjacent homo-oligomers are thought to pack side-
by-side within caveolae membranes thereby providing the structural meshwork for
caveolae invagination. Adapted from Razani et al. [51].
Copyright © Lei Wang 2007
14
Chapter Two : Effects of ω-6 and ω-3 Fatty Acids on TNF-α-Induced Endothelial
Cell Activation
Synopsis
TNF-α, an important pro-inflammatory mediator of endothelial cell activation,
can lead to the formation of lipid-derived prostaglandins. Different ω-6 and ω-3 fatty
acids have been implicated to have pro- or anti- inflammatory actions, respectively.
However, the mechanisms of ω-6 and ω-3 fatty acids mediated TNF-α-induced
production of the major inflammatory mediator prostaglandin E2 (PGE2) and the
underlying signaling mechanism remains unclear. The present study focused on the
effects of ω-6 and ω-3 fatty acids on TNF-α-induced oxidative stress and the
inflammatory signaling that involves in PGE2 production. Porcine pulmonary-arterial
endothelial cells were first enriched with linoleic acid (LA; 18:2 ω-6) or α-linolenic acid
(ALA; 18:3 ω-3), followed by exposure to TNF-α. TNF-α-induced oxidative stress, p38
MAPK activation and activation of nuclear factor κB (NF-κB) were markedly increased
in the presence of linoleic acid and diminished by pretreatment with α-linolenic acid.
Similar protective effects of α-linolenic acid were observed by measuring COX-2
expression and the production of PGE2. Treatment with p38 MAPK specific inhibitor
significantly decreased TNF-α-induced COX-2 and VCAM-1 protein expression. Taken
together, our data suggest that in endothelial cells, linoleic acid and α-linolenic acid can
generate distinct signaling outcomes in response to inflammatory stimuli caused by TNF-
α and p38 MAPK plays an important role in fatty acid mediated TNF-α-induced
inflammatory response.
15
Introduction
Vascular endothelial dysfunction is thought to be involved in the initiation of
atherosclerotic lesions. Endothelial cells lying at the blood-tissue interface play an
essential role in homeostasis of the circulation and vessels. At normal state, endothelial
cells have antithrombotic and anticoagulant activities [110]. But the “activation” of
endothelial cells can cause changes to the hemodynamic equilibrium and the
development of numerous oxidative and inflammatory reactions, which result in
lymphocyte and monocyte adhesion to the vascular endothelium [10, 111]. This is
considered to involve in the very early stage of atherogenesis. Leukocyte adhesion to the
endothelium is facilitated by the appearance of adhesion molecules at the surface of
lymphocytes, monocytes, and endothelial cells. These molecules are mainly selectins
(e.g. E-selectins are specific to endothelial cells), vascular cell adhesion molecule type 1
(VCAM-1), and intercellular adhesion molecule type 1 (ICAM-1) [112].
One of the common mediators of endothelial cell activation is macrophage-
derived cytokines, such as tumor necrosis factor alpha (TNF-α) [113]. Patients with
clinical cardiovascular disease had significantly higher levels of TNF-α compared with
subjects without cardiovascular disease [114]. High levels of circulating TNF-α can
cause endothelial cell activation resulting in the development of numerous oxidative and
inflammatory reactions and facilitating lymphocyte and monocyte adhesion to the
vascular endothelium [10, 111]. Furthermore, TNF-α profoundly affects eicosanoids
metabolic pathways leading to the accumulation of one or more pro-inflammatory
prostaglandins.
The mechanisms by which selected fatty acids induce endothelial cell activation,
oxidative stress and inflammation are not fully understood. Oxidative stress-induced
transcription factors, which regulate inflammatory cytokine and adhesion molecule
production, are important regulatory elements in the induction of inflammatory responses.
One of these transcription factors, nuclear factor κB (NF-κB), plays a significant role in
these regulatory processes [115]. Binding sites for NF-κB and related transcription
factors were identified in the promoter regions of a variety of inflammatory genes [116,
16
117] such as interleukin 6 (IL-6), vascular cell adhesion molecule-1 (VCAM-1) or
cyclooxygenase-2 (COX-2), all of which are up-regulated by TNF-α [118, 119].
Numerous risk factors for the development of atherosclerosis have been identified,
including obesity [120] and hypertriglyceridemia [121]. Increased circulating free fatty
acid levels are associated with hypertriglyceridemia and obesity [122], and high plasma
free fatty acids may contribute to an environment of increased oxidative stress and
inflammation in the vasculature and especially in vascular endothelial cells [123, 124].
Dietary balance of long-chain fatty acids may influence processes involving leukocyte-
endothelium interactions, such as atherogenesis and inflammation [125]. Even though
diets high in omega-6 fatty acids may lead to a decrease in serum cholesterol [126],
replacing saturated with unsaturated omega-6 rich lipids may not be desirable because of
their ability to easily oxidize. High intake of linoleic acid-rich oils or fats will lead to an
increase in cellular oxidative stress and can elicit an inflammatory response [127], events
which have been implicated in most chronic diseases. Omega-6 fatty acids, and
especially linoleic acid can cause endothelial cell dysfunction as well as potentiate tumor
necrosis factor-α (TNF-α)-mediated endothelial injury [128]. We have recently
demonstrated that both the extracellular signal regulated kinase (ERK1/2) and
phosphoinositide-3 kinase/ amino kinase terminal (PI3K/Akt) signaling pathways can
contribute to the effect of linoleic acid on nuclear factor-kappa B (NF-κB)-dependent
transcription and endothelial cell activation [105].
In contrast to omega-6 fatty acids, omega-3 fatty acids can influence
cardiovascular disease pathology by beneficially modulating inflammation.
Epidemiological and interventional studies have shown a dose-dependent decrease in risk
of cardiovascular disease endpoints with increased dietary consumption of moderate
amounts of omega-3 fatty acids, either plant or marine derived [126]. Current estimates
indicate that over 90% of the omega-3 consumed by U.S. citizens is in the form of α-
linolenic acid, not the longer chain omega-3 fatty acids found in fish oils [129].
Independent of their dietary source, omega-3 fatty acids contribute to cardio-protective
properties, which include down-regulation of proinflammatory and proatherogenic genes,
including adhesion molecules and cytokines, during early atherogenesis and possibly also
during later stages of plaque development and plaque rupture [130]. For example, an α-
17
linolenic acid-rich oil decreased oxidative stress and CD40 ligand in patients with mild
hypercholesterolemia [131], reduced levels of soluble cell adhesion molecules in plasma
[132] and recurrence of coronary heart disease [133]. Also, by partially replacing omega-
6 analogues in membrane phospholipids with omega-3 fatty acids, it is possible to
decrease the transcriptional activation of inflammatory and pro-atherogenic genes
involved in endothelial cell activation and atherosclerosis [134].
Previous studies in our laboratory have shown that linoleic acid potentiated TNF-
α–mediated pro-inflammatory responses in endothelial cells [107]. Linoleic acid may
further be atherogenic by causing activation of NF-κB which in turn increases the
expression of E-selectin and VCAM-1 [128, 135]. On the other hand, lipids are crucial
structural and functional components of cells. Even more importantly, many lipid species
have distinct cellular functions. For example, ceramides, eicosanoids, diacylglycerol
(DAG) and lysolipids are all second messengers which participate in various cellular
events such as inflammation, growth, proliferation, differentiation and cell death.
However the primary mechanism of the modification of these different metabolism
pathways by the availability of different series of omega-6 and omega-3 PUFAs in the
presence of TNF-α is not well known.
The present study was conducted to test the effects of linoleic acid and α-linolenic
acid on TNF-α-induced prostaglandin E2 (PGE2) release in primary porcine pulmonary
artery endothelial cells. Three markers of inflammation, oxidative stress, p38 MAPK,
COX-2 that have been recognized to play a role in atherogenesis, along with transcription
factor NF-κB, were assessed.
In the current study, we demonstrate that ω-3 fatty acid inhibits the ability of pro-
inflammatory cytokine TNF-α to induce the expression of adhesion molecules associated
with endothelial cell activation. We also show here that ω-3 fatty acid were associated
with the inhibition of p38 MAPK signal transduction pathways. These results provide
new insights into the modulation of TNF-α-induced inflammatory responses by dietary
polyunsaturated fatty acids.
18
Materials and Methods
Cell culture and experimental media
Endothelial cells were isolated from porcine pulmonary arteries and cultured as
previously described [30] (see detailed protocol in Appendix B). Arteries obtained
during routine slaughter were donated from the College of Agriculture, University of
Kentucky. The basic culture medium consisted of medium 199 (M-199) (GIBCO
Laboratories, NY) containing 10% (v/v) fetal bovine serum (FBS, HyClone Laboratories,
UT). The experimental media were composed of M-199 enriched with 5% (v/v) FBS and
with linoleic acid (LA) or α-linolenic acid (ALA) (> 99% pure; Nu-Chek Prep, MN).
Preparation of experimental media with LA and ALA were performed as described
earlier [136] (see detailed protocol in Appendix B). Metabolic studies [137] indicated
that the molar ratio of free fatty acid to albumin is the main factor controlling free fatty
acid availability to tissues. In plasma, albumin concentration is about 600 μM, and total
free fatty acid is 180 to 2000 μM thus the range of ratios of fatty acid to albumin is 0.3 to
4. In the experimental media, 20 μM fatty acids were enriched in 5% media with which
the albumin concentration is about 50 to 60 μM. So the ratio of free fatty acids to
albumin is within physiological and metabolic relevance and not toxic to the cells.
Human TNF-α was purchased from Sigma (Sigma-Aldrich, MO). In the p38
MAPK inhibitor study, SB203580 was dissolved in DMSO (Sterile-filtered, Sigma-
Aldrich, MO) to a stock solution of 10 mM. After getting confluent, cells were first
incubated with 10 μM of SB203580 (a thousand times dilution from stock solution) for
30 min followed by exposure to TNF-α. The control culture contained the same volume
of DMSO.
E-selectin, VCAM-1, and COX-2 gene expression studies
RT-PCR:
Total cellular RNA was extracted from culture plates according to the RNA-
STAT-60 protocol (TEL-TEST, Friendswood, TX). RNA concentration was
spectrometrically determined by reading the absorbance at 260 nm. cDNA first strand
was synthesized from total RNA by RT reaction. The reaction mixture contained 1 μg
19
RNA. Resulting cDNA was amplified by the polymerase chain reaction with enzyme
nucleotides and buffers from Quiagen (Valencia, CA). The following primers for E-
selectin, VCAM-1, and COX-2 were employed in the PCR. E-selectin: forward 5’-
GACTCGGGCAAGTGGAATGATGAG-3’; reverse 5’-CATCACCATTCTGAGGATG
GCCGAC-3’. VCAM-1: forward 5’- GTTTACCCGGTTGAAAAGTTGGAG-3’;
reverse 5’- CACCGTGTCG CCTGTCTCT-3’. COX-2: forward 5’-GGAGAGACAGGC
ATAAACTGC-3’, reverse 5’-GTGTGTTAAACTCAGCAGCA-3’. β-actin was used as
a house keeping gene: forward 5’- GGGACCTGACCGACTACCTC-3’; reverse 5’-GGG
CGATGATCTTG ATCTTC-3’. The PCR products were electrophoresed on a 2% (w/v)
EDTA agarose gel. Then stained with SYBR Green ( Molecular Probes, Eugene, OR)
and visualized by using phosphorimaging technology (FLA-5000; Fuji, Stamford, CT).
Real-time PCR:
Total RNA was extracted from endothelial cells using RNA-STAT-60 (TEL-
TEST, Friendswood, TX) according to the manufacturer’s protocol. Reverse
transcription was performed using the AMV reverse transcription system (Promega,
Madison, WI). The levels of mRNAs and the PCR-product were then assessed by real-
time PCR using 7300 Real Time PCR System (Applied Biosystems). Real-time PCR
samples were mixed with SYBR Green Master Mix (Applied Biosystems) and VCAM-1
or COX-2 specific primers. The sequences for porcine VCAM-1 and COX-2 gene were
designed by Primer Express Software 3.0 for real-time PCR (Applied Biosystems).
VCAM-1 sequences: sense, 5’-TGGAAAGACATGGCTGCCTAT-3’; antisense, 5’-
ACACCACCCCAGTCACCATA TC-3’. COX-2 sequences: sense, 5’-TGCTGAAGCC-
CTATCGATCA-3’; antisense, 5’-TACAGCTCCATGGCATCAATG-3’. β-actin was
used as a housekeeping gene in both VCAM-1 and COX-2 studies. β-actin sequences:
sense, 5’-TCATCACCATCGGCAACG-3’; antisense, 5’-TTCCTGATGTCCACGTCG-
3’.
Immunofluorescence study of E-selectin expression
Confluent endothelial cells were grown on glass culture slides and incubated with
or without TNF-α for 4 h. After three washing in phosphate-buffered saline (PBS), cells
were fixed by 70% ethyl alcohol for 10 min and air-dried. After 30 min of blocking of
20
non-specific binding with PBS containing 3% BSA, cells were incubated with E-selectin
primary antibody for 1 h on ice at a dilution of 1:50. Then slides were washed twice in
PBS then incubated simultaneously with FITC-labeled secondary antibody (1 h at room
temperature, dilution 1:5; Chemicon, CA). Negative controls were prepared by
incubation of the cells with anti-IgG antibody. The cells were washed twice in PBS and
mounted in aqueous mounting medium before being observed using an epifluorescence
Nikon Eclipse E600 microscope. The images were capture using a Spot charge coupled
device camera system (Nikon, Melville, NY).
Measurement of cellular oxidative stress
Cellular oxidation was determined by 2', 7'-dichlorofluorescein (DCF)
fluorescence as described earlier [138]. This method is based on the conversion of 2’, 7’-
dichlorofluorescin into fluorescent 2’,7’-dichlorofluorescein by oxygen reactive species,
primarily peroxyl radicals and peroxides. Please find the detailed protocol in Appendix B.
After treatment cells were washed with Hanks buffer to remove serum. Then KREBS-
Ringer-Glucose buffer containing DCF (100 μmol/L) (Molecular Probes, Inc., Eugene,
OR) were added to microplates and incubate for 30 minutes. Blank well also received
KREBS-Ringer-Glucose buffer without DCF probe. Wrap plate in aluminum foil to
protect from the light. After incubation washed wells 2 to 3 times with HEPES buffer
and then added HEPES to each well (1 ml for 12 well plates and 0.5 ml for 24 well plates.
Wrap plate in aluminum foil before reading. A multi-well fluorescent plate reader
(Molecular Devices, Sunnyvale, CA) was utilized for the imaging study. Excitation and
emission wavelengths were 490 and 520 nm respectively.
Transcription factor NF-κB activation studies: electrophoretic mobility shift assay
(EMSA)
Nuclear extracts containing active proteins were prepared from cells according to
the method of Beg et al. [139]. Binding reactions were performed in a 20 μL volume
containing 7 μg of nuclear protein extracts. Nuclear extracts were incubated for 25 min
with 32P-end-labeled oligonucleotide probes containing enhancer DNA element NF-κB
(5’ AGTTGAGGGGACTTTCCCAGGC 3’) (Santa Cruz, Santa Cruz, CA). Following
21
binding, the protein-DNA complexed and uncomplexed DNA in the mixture were
resolved on native 5% polyacryamide gels using 0.5x TBE buffer (50 mM Tris-Cl, 45
mM boric acid, 0.5 mM EDTA, pH 8.4) and visualized by autoradiography. Control
reactions using 200-fold molar excess of unlabeled oligonucletide probes or a supershift
assay were performed to demonstrate the specificity of the shifted DNA-protein
complexes for NF-κB.
Measurement of VCAM-1 or COX-2 protein levels, and phosphorylation of p38MAPK or
cPLA2 by western blot
Cells were harvested using cell lysis buffer as previously described [105]. Protein
samples were resolved by SDS-PAGE using 10% gradient gels and transferred
electrophoretically to nitrocellulose membranes using a Bio-Rad immunoblot transfer
apparatus (Bio-Rad Laboratories, Hercules, CA) according to the manufacturer’s
instructions. The nonspecific sites on the membrane were blocked 1 h at room
temperature with 5% nonfat dry milk in TBST followed by incubation with β-actin
(1:5000 dilution in 5% nonfat milk), VCAM-1 (1:2000 dilution in 5% nonfat milk),
COX-2 (1:2000 dilution in 5% nonfat milk), total p38 MAPK (1:1000 dilution in 5%
nonfat milk), phosphor-p38 MAPK (1:1000 dilution in 5% nonfat milk), or phosphor-
cPLA2 (1:500 dilution in 5% nonfat milk) primary antibodies (Santa Cruz Biotech, CA)
overnight at 4 °C. Bands were visualized using the appropriate horseradish peroxidase-
conjugated secondary antibodies followed by ECL immunoblotting detection reagents
(Amersham Biosci, UK). Detection and quantitative analysis were performed using a
digitizing system (UN-SCAN-IT, Silk Scientific Corporation). To determine the optimal
amounts of samples for linear area detection, a titration of different amounts of sample
were loaded. For COX-2 and VCAM-1, 30 μg protein were loaded; for p38 MAPK, 35
μg protein were loaded; for phospho-cPLA2, 40 μg protein were loaded.
PGE2 determination
Cells were seeded in 60 mm petri dishes (Becton Dickinson Labware, NJ) and
grown to confluence. After treatment, supernatants of cell cultures were collected into
microcentrifuge tubes (Isc BioExpress, UT), centrifuged at 4 ℃ to remove cellular debris
22
and stored at -80 ℃. PGE2 levels were assessed using a PGE2-specific enzyme
immunoassay (EIA) (Cayman Chemicals, MI) following the manufacturer’s protocol
with minor modification (see detailed protocol in Appendix B). A microplate
spectrophotometer SpectraMaxPro M2 (Molecular Devices Corporation, CA) was used to
read the plate at 405 nm.
Statistical analysis
Values are reported as mean ± standard error of the mean (SEM) of at least three
independent groups. Data were analyzed using Sigma Stat software (Jandel Corp., Wan
Rafael, CA). One way ANOVA followed by post hoc least significant difference (LSD)’s
pairwise multiple comparison procedure were used for statistical analysis of the original
data. A statistical probability of p< 0.05 was considered significant. In figures 2.4, 2.13,
and 2.15, * indicates significant difference from control group. In all the other figures,
different letters represent significant differences among treatment groups. Capital or
small letters represent with or without TNF-α treatment respectively as were compared
separately by one way ANOVA.
Results
Determination of optimal TNF-α concentration and incubation time for the stimulation of
endothelial cell activation
TNF-α is arguably the most potent inducer of a power pro-inflammatory cytokine.
Previous epidemiological studies indicated that the plasma concentrations of TNF-α are
increased in various pathological situations, and TNF-α levels vary widely not only
among different diseases but also among different patients with the same disease. For
example, in diabetic patients TNF-α can reach 19.3±7.5 pg/ml in comparison with
healthy controls (11.1±5.8 pg/ml) [140], or 228 pg/ml vs. 4.2 pg/ml [141]. Another
study reported a median TNF-α level at 26 pg/ml in patients with sepsis, ranging from 0
to 1000 pg/ml [142]. In patients with lymphoma, the TNF-α levels were between 5 to
23
380 pg/ml with the median of 20 pg/ml, and the healthy controls were between 4 to 9
pg/ml with the median of 7 pg/ml [143]. Even when the plasma levels of TNF-α are
known, local concentrations of inflammatory cytokines in the endothelial
microenvironment are not well defined. Some in vitro studies used relatively high
concentration of TNF-α from 10 ng/ml [144] to 50 ng/ml [145]. For some apoptosis
studies, TNF-α concentration also has been used at 50 ng/ml [146].
In the current study, as we were concerning about the effects of fatty acids on
inflammatory responses caused by TNF-α, it was necessary to avoid challenging cells
with an overwhelming amount of TNF-α. In our in vitro endothelial cell model, the
binding of TNF-α to its receptor on endothelial cells is followed by rapid translocation of
the transcription factor NF-κB from the cytoplasm to the nucleus. This translocation is
important for stimulated transcription of the adhesion molecules E-selectin, VCAM-1,
and inducible enzyme COX-2 which all possess NF-κB promoter/enhancer elements.
Thus, to determine the condition of TNF-α-induced endothelial cell activation, we first
did time and concentration course studies for E-selectin, VCAM-1, and COX-2.
Endothelial cells were incubated in the presence of TNF-α concentrations ranging
from 0.1 to 0.5 ng/ml for 2, 4, or 6 h. Expression of E-selectin (Fig. 2.1) was assessed by
RT-PCR. We observed that as low as 0.2 ng/ml of TNF-α, there was a dramatic increase
of E-selectin at 4 h. When endothelial cells were treated with 0.5 ng/ml TNF-α, the
expression of E-selectin rose within 2 h. But the increase of E-selectin went back to
basal level in all the treatment groups after 6 h of incubation. We confirmed by
immunofluorescence study that 0.5 ng/ml TNF-α increased cell surface expression of E-
selectin on endothelial cells (Fig. 2.2). Furthermore, 0.5 ng/ml TNF-α at 4 h also
increased VCAM-1 and COX-2 (Fig. 2.3) genes expression which were measured by RT-
PCR. Thus TNF-α at the concentration of 0.5 ng/ml was determined to activate
endothelial cells in the current study.
Determination of optimal linoleic acid incubation time for endothelial cell activation
Nutritional habits can result in modified levels of circulating metabolites,
including free fatty acids. The dietary different fatty acids have been shown to be
24
efficient for modifying inflammatory diseases. Dietary polyunsaturated free fatty acid
concentration in plasma has been reported to be 20 -30 μΜ [147] and a concentration
range of 180-2,5000 μM has also been reported [148]. We next investigated the effect of
physiological level of linoleic acid (20 μM) alone on endothelial cell activation. As
shown in Fig. 2.4, expression of VCAM-1 mRNA rose at 18 h and continuous increasing
at 24 h. Thus for inflammatory gene expression study, cells were pretreated with fatty
acids for 24 h, followed by TNF-α treatment for another 4 h.
Linoleic acid and α-linolenic acid modulate TNF-α-induced E-selectin and COX-2
expression
To examine the effects of ω-6 and ω-3 fatty acids on TNF-α-induced endothelial
activation, cells were pretreated with either linoleic acid or α-linolenic acid for 24 h,
followed by exposure to TNF-α for another 4 h. As indicated in Fig. 2.5, 2.6,
pretreatment with α-linoleic acid significantly decreased TNF-α-induced both E-selectin
and COX-2 gene expression. For E-selectin expression, pre-incubation with linoleic acid
followed by TNF-α didn’t show any different from TNF-α treatment alone (Fig. 2.5).
But COX-2 mRNA expression was further amplified in the presence of linoleic acid
compared with TNF-α treatment alone (Fig. 2.6). We confirmed the COX-2 data by
measuring protein levels of COX-2 by western blot (Fig. 2.7).
Linoleic acid and α-linolenic acid modulate cellular oxidative stress and NF-κB activity
induced by TNF-α
The NF-κB family of transcription factors plays a crucial role in inflammatory
events induced by TNF-α. The transcription of adhesion molecules E-selectin, VCAM-1,
and inducible enzyme COX-2 all possess NF-κB promoter/enhancer elements [149].
Furthermore, NF-κB is respond directly to oxidative stress [150]. We further assess
effects fatty acids on TNF-α-induced oxidative stress and NF-κB activation. Exposure to
TNF-α alone significantly increased cellular oxidative stress (ROS) (Fig.2.8) and NF-κB
DNA binding activity (Fig. 2.9). Pretreatment with linoleic acid following by exposure
to TNF-α further induced oxidative stress and activation of NF-κB compared to cultures
25
treated only with TNF-α. In contrast, pretreatment with α-linolenic acid blocked the
TNF-α-induced both oxidative stress and subsequent induction of NF-κB.
Linoleic acid and α-linolenic acid modulate TNF-α-induced activation of p38 MAPK
In mammalian cells, MAPKs are strongly activated by growth factors,
environmental stresses, and inflammatory cytokines [151]. As shown in Fig. 2.10, p38
MAPK was significantly activated by TNF-α at 10 min and pre-treatment with linoleic
acid followed by exposure to TNF-α further increased p38 MAPK activation, this event
was blocked by pre-enrichment with α-linolenic acid.
In order to further evaluate the role of p38 MAPK in TNF-α-induced
inflammatory gene expression, endothelial cells were pretreated with the specific p38
MAPK inhibitor SB203580, followed by exposure to TNF-α for an additional 8 h. As
shown in Fig. 2.11 and 2.12, SB203580 significantly decreased TNF-α-induced both
COX-2 and VCAM-1 protein expression. These results indicate that p38 MAPK is
crucially involved in ω-6 and ω-3 fatty acids mediated TNF-α-induced COX-2
expression.
Induction of cPLA2 phosphorylation by TNF-α
The biosynthesis of prostaglandin E2 (PGE2), a major lipid-derived inflammatory
mediator, involves three enzymatic reactions including phospholipase A2 (PLA2),
cyclooxygenase (COX), and PGE2 synthase. Cytosolic phospholipase A2 (cPLA2) is the
most characterized phospholipase members of PLA2 family which are responsible for
remarkable selectivity toward arachidonic acid at the sn-2 position of phospholipids [152].
To assess the effect of TNF-α on PGE2 production, we first measured the activation of
cPLA2. As shown in Fig. 2.13, TNF-α induced significant increase of cPLA2
phosphorylation at 8 h treatment. cPLA2 mRNA expression was not changed after TNF-
α stimulation measured by Real-time PCR.
26
Linoleic acid and α-linolenic acid modulate TNF-α-induced PGE2 production
We further investigated cellular PGE2 levels after different fatty acids or TNF-α
treatments. As shown in Fig. 2.14, an 18 h exposure to linoleic acid dramatically induced
PGE2 production. In contrast, α-linolenic acid did not have much induction effect.
To access the optimal TNF-α incubation time for PGE2 production, cells were
treated with TNF-α for 6, 18, and 24 h. As indicated in Fig. 2.15, a 24 h exposure to
TNF-α significantly induced PGE2 production. Compared to the TNF-α treatment group,
enriching endothelial cells with linoleic acid further enhanced TNF-α-induced PGE2
production. In contrast, cellular exposure to α-linolenic acid markedly reduced the
proinflammatory effect of TNF-α (Fig. 2.16).
27
Figure 2.1 Optimal TNF-α concentration and incubation time for E-selectin expression
Cells were treated with indicated concentration of TNF-α for 2, 4, or 6 h. E-selectin gene
expression was measured by RT-PCR. β-actin was used as housekeeping gene.
Figure 2.2 Immunofluorescence microscopy study of TNF-α-induced E-selectin
expression
Cells were treated with 0.5 ng/ml TNF-α for 4 h. Immunofluorescence staining for E-
selectin (FITC green fluorescence) and nuclear (DAPI blue fluorescence) were indicated.
The merged images were shown to better illustrate the expression of E-selectin for each
individual cell.
28
Figure 2.3 TNF-α-induced VCAM-1 and COX-2 expression
Cells were exposed to control culture (C), or 0.5 ng/ml TNF-α for 4 h. VCAM-1 and
COX-2 gene expression were measured by RT-PCR. β-actin was used as housekeeping
gene.
29
Figure 2.4 Optimal linoleic acid incubation time for VCAM-1 gene expression
Cells were incubated with or without 20 μM linoleic acid (LA) for indicated time.
VCAM-1 gene expression was measured by real-time PCR. β-actin was used as
housekeeping gene. * indicates significant difference from control group without linoleic
acid.
30
Figure 2.5 Effect of linoleic acid and α-linolenic acid on TNF-α-induced E-selectin gene
expression
Cells were treated with 0.5 ng/ml TNF-α alone for 4 h, or pretreated with 20 μM
linoleic acid (LA)/α-linolenic acid (ALA) for 24 h followed by exposure to TNF-α for
additional 4 h. E-selectin gene expression was measured by real-time PCR. β-actin was
used as housekeeping gene. Experiments were repeated three times. Different letters
represent significant differences among treatment groups. Capital or small letters
represent with or without TNF-α treatment respectively as were compared separately by
one way ANOVA.
31
Figure 2.6 Effect of linoleic acid and α-linolenic acid on TNF-α-induced COX-2 gene
expression
Cells were treated with 0.5 ng/ml TNF-α alone for 4 h, or pretreated with 20 μM
linoleic acid (LA)/α-linolenic acid (ALA) for 24 h followed by exposure to TNF-α for
additional 4 h. COX-2 gene expression was measured by real-time PCR. β-actin was
used as housekeeping gene. Experiments were repeated three times. Values are means±
SEM. Different letters represent significant differences among treatment groups. Capital
or small letters represent with or without TNF-α treatment respectively as were compared
separately by one way ANOVA.
32
Figure 2.7 Effect of linoleic acid and α-linolenic acid on TNF-α-induced COX-2 protein
expression
Cells were pretreated with 20 μM of linoleic acid or α-linolenic acid for 24 h and
then exposed to 0.5 ng/ml TNF-α for an additional 8 h. COX-2 protein levels were
measured by western blot. β-Actin was used as a housekeeping gene. Experiments were
repeated three times, and the blots shown are representative of one of the experiments.
Values are means ± SEM. Different letters represent significant difference among
treatment groups.
33
Figure 2.8 Effect of linoleic acid and α-linolenic acid on TNF-α-induced oxidative stress
Cultures were pretreated in media supplemented with 20 μM linoleic acid (LA) or α-
linolenic acid (ALA) for 24 h followed by exposure to 0.5 ng/ml TNF-α for additional 3
h. Values are means ± SEM (n = 3). Different letters represent significant differences
among the treatment groups.
34
Figure 2.9 Effect of linoleic acid and α-linolenic acid on TNF-α-induced activation of
NF-κB
Cells were treated with 20 μM of linoleic acid (LA) or a-linolenic acid (ALA) for 24 h
prior to exposure to 0.5 ng/ml TNF-α for an additional 6 h. Experiments were repeated
three times, and the blots shown are a representative of one of the experiments. The bar
graph shows the corresponding densitometric analysis of the blots. Values are means ±
SEM. Different letters represent significant differences among treatment groups.
35
Figure 2.10 Effect of linoleic acid and α-linolenic acid on TNF-α-induced p38 MAPK
activation
Cells were pretreated with 20 μM of linoleic acid (LA) or a-linolenic acid (ALA) for 24 h
and then exposed to 0.5 ng/ml TNF-α for 10 min. Phosphorylated and total p38 MAPK
were detected by western blot using specific dually phosphorylated p38 MAPK antibody
(Thr 180/Tyr 182) or anti-total p38 MAPK antibody respectively. Experiments were
repeated three times and the blots shown are a representative of one of the experiments.
Values are means ± SEM. Different letters represent significant difference among
treatment groups.
36
Figure 2.11 Effect of p38 MAPK inhibitor on TNF-α-induced COX-2 up-regulation
Cells were exposed to 0.5 ng/ml TNF-α for 8 h, or first pre-enriched with 10 μM
SB203508 for 30 min followed by co-exposure to TNF-α for additional 8 h. COX-2
protein levels were measured by western blot. β-Actin was used as a housekeeping gene.
Experiments were repeated three times and the blots shown are a representative of one of
the experiments. Different letters represent significant differences among treatment
groups.
37
Figure 2.12 Effect of p38 MAPK inhibitor on TNF-α- and linoleic acid-induced VCAM-
1 up-regulation
Cells were exposed to 0.5 ng/ml TNF-α for 8 h or linoleic acid (LA) for 24 h, or first pre-
enriched with 10 μM SB203508 for 30 min followed by co-exposure to TNF-α or LA.
VCAM-1 protein levels were measured by western blot. β-Actin was used as a
housekeeping gene.
38
Figure 2.13 Effect of TNF-α on cPLA2 activation
Cells were treated with 0.5 ng/ml TNF-α for 4, 6, or 8 h. cPLA2 phosphorylation was
measured by western blot using specific p-cPLA2 antibody. β-actin was used as
housekeeping gene. Experiments were repeated three times. Values are means±SEM.
Different letters represent significant difference among treatment groups. * indicates
significant difference from other treatment groups.
39
Figure 2.14 Effect of linoleic acid and α-linolenic acid on PGE2 production
Cells were exposed to 20 μM linoleic acid (LA) or α-linolenic acid (ALA) 18 and 24 h.
Supernatants of cell cultures were collected and PGE2 levels were measured by enzyme
immunoassay (EIA). Bars represent means ± SEM from three independent experiments.
40
Figure 2.15 Optimal TNF-α incubation time for PGE2 production
Cells were exposed to 0.5 ng/ml TNF-α for 6, 18 and 24 h. Supernatants of cell cultures
were collected and PGE2 levels were measured by enzyme immunoassay (EIA). Bars
represent means ± SEM from three independent experiments. * represents significant
difference compared with the control treatment.
41
Figure 2.16 Effect of linoleic acid and α-linolenic acid on TNF-α-induced PGE2
production
Cells were pretreated with 20 μM of LA or ALA for 24 h and then exposed to 0.5 ng/ml
TNF-α for additional 24 h. Supernatants of cell cultures were collected and PGE2 levels
were measured by EIA. Values are means ± SEM. Different letters represent significant
difference among treatment groups.
42
Figure 2.17 Proposed mechanism for fatty acid-mediated modulation of endothelial cell
activation induced by TNF-α
Omega-6 or omega-3 fatty acids can either enhance or reduce ΤΝF-α-induced p38
MAPK activation, oxidative stress (ROS), and NF-κB activation. Relative activation of
NF-κB will further regulate VCAM-1, as well as COX-2 enzyme activity and subsequent
PGE2 release from endothelial cells.
43
Discussion
Atherosclerosis is considered an inflammatory disease, which involves the
interplay of prooxidative activities, induction of inflammatory cytokine, activation of
vascular endothelial cells and activation of adhesion molecules, all events which promote
vascular leukocyte infiltration and plaque development [6]. Inflammatory events also
include the cyclooxygenase and subsequent eicosanoid pathways [153, 154]. For
example, inflammatory cytokines like interleukin-1beta (IL-1-β) can induce COX-2
expression and the release of PGE2, events which may be mediated through activation of
p42/44 and p38 MAPKs, and the NF-κB pathway [155]. Our data demonstrate the TNF-
α can markedly induce oxidative stress, p38 MAPK, NF-κB, COX-2 and PGE2.
Sever hypertriglyceridemia has been identified as an independent risk factor for
atherosclerosis, and increased circulating free fatty acid levels are associated with
hypertriglyceridemia and obesity [122]. High plasma free fatty acids may contribute to
an environment of increased oxidative stress and inflammation in the vasculature [123,
124], and we have previously shown that linoleic acid can markedly amplify a TNF-α-
mediated endothelial inflammatory response [156]. We also have recently demonstrated
that both the extracellular signal regulated kinase (ERK1/2) and phosphoinositide-3
kinase/ amino kinase terminal (PI3K/Akt) signaling pathways can contribute to the effect
of linoleic acid on NF-κB-dependent transcription and endothelial cell activation [105].
In the current study, exposure to TNF-α induced oxidative stress, p38 MAPK, NF-κB,
COX-2 and PGE2, which was amplified by pre-enrichment with linoleic acid. This
clearly supports our hypothesis that diets high in omega-6 fatty acids, and especially in
linoleic acid, are proinflammatory and thus may contribute to the early pathology of
atherosclerosis.
In contrast to omega-6 fatty acids, epidemiological and interventional studies
have shown a dose-dependent decrease in risk of cardiovascular disease endpoints with
increased dietary consumption of moderate amounts of omega-3 fatty acids, either plant
or marine derived [126]. Our data clearly demonstrate the anti-inflammatory properties
of omega-3 fatty acids. In fact, our observed TNF-α induced oxidative stress, p38
44
MAPK, NF-κB, COX-2 and PGE2 in endothelial cells was blocked or reduced when cells
were enriched relatively more with α-linolenic acid than with linoleic acid. Mechanisms
of vascular inflammation are largely regulated through p38 MAPK signaling [157-160],
and our data suggest that p38 MAPK is an important player in different ω-6 and ω-3 fatty
acid modified TNF-α-induced inflammatory signaling cascade. The p38 MAPK inhibitor
SB203580 significantly decreased TNF-α-mediated induction of COX-2 protein
expression, suggesting a regulatory aspect through p38 MAPK signaling. This was
supported by the evidence that p38 MAPK increases NF-κB recruitment in cells exposed
to inflammatory stimuli [161].
Reactive oxygen intermediates have been reported to function as second
messengers in the activation of intracellular signaling cascades, particularly the activation
of the transcription factors NF-κB and AP-1 [162]. The MAPK signaling cascades are
involved in the regulation and activation of these transcription factors [105]. Superoxide
dismutase (SOD) has been shown to inhibit TNF-α-induced activation of p38 MAPK,
NF-kB, and AP-1, thus reduces expression of VCAM-1 and ICAM-1 [144]. Implied that
antioxidants have beneficial effect in pathological events involving leukocyte adhesion.
Components of the MAPK pathway have been implicated as mediators of
phosphorylation of intracellular substrates such as protein kinases and transcription
factors. In mammalian cells, at least three different subfamilies of MAPK have been
identified: extracellular signal-regulated kinases (ERKs) with isoforms p44 MAPK
(ERK1) and p42 MAPK (ERK2); stress-activated protein kinases also called c-Jun N-
terminal kinases (JNKs), and the p38 MAPKs. It has been demonstrated that both JNK
and p38 MAPK cascades are activated preferentially by the inflammatory cytokines such
as TNF-α, as well as by a wide variety of cellular stresses such as ultraviolet light, heat
shock, oxidative stress, etc. [163, 164]. Whereas activation of ERK group of MAPK is
not currently considered a general event associated with TNF-α stimulation and is
actually primarily involved in control of growth and differentiation [165, 166]. In
general, the ERKs are mostly activated by mitogenic and proliferative stimli, whereas the
JNKs and p38 MAPKs respond to environmental stress, including inflammatory
cytokines [160]. Inhibition of the p38 MAPK is considered to be a promising candidate
45
for treatment of chronic inflammation [167]. p38 MAPK plays an important role in the
regulation of NF-κB recruitment to a subset of genes activated in cells exposed to
inflammatory stimuli, and a proposed mechanism is that p38 MAPK can modulate the
accessibility of selected NF-κB sites in the genome [161]. P38 MAPK pathway has also
been shown to involve in the induction of several other inflammatory molecules, such as
COX-2, implicated in the regulation of inflammatory mediators, including PGE2 [167].
P38 MAPK has been shown to involve in TNF-α-stimulated NF-κB activation and
interleukin-6 expression [168]. Our data showed that pharmacologic inhibition of p38
MAPK by SB203508 inhibited TNF-α-induced stimulation of VCAM-1, COX-2 and
subsequently PGE2 synthesis. Furthermore, MAPK signaling appears to be involved in
regulating omega-6 fatty acid and TNF-α-induced endothelial cell activation and
modification by omega-3 fatty acids (Fig. 2.8).
Thus, data from the current study provide evidence that an increase in dietary
omega-3 fatty acids contributes to an anti-inflammatory environment, which may involve
cellular signaling through local repletion of selected eicosanoid species.
Copyright © Lei Wang 2007
46
Chapter Three : Involvement of Caveolae in Regulation of Endothelial Cell
Activation Mediated by Fatty Acids and TNF-α
Synopsis
TNF-α transduces inflammatory responses through its receptor TNF receptor-1
(TNFR-1) which has been indicated to be enriched in plasma membrane microdomains
caveolae. Hypertriglyceridemia and associated high circulating free fatty acids are
important risk factors of atherosclerosis. In contrast to omega-3 fatty acids, linoleic acid,
the major omega-6 unsaturated fatty acid in the American diet, may be atherogenic by
amplifying an endothelial inflammatory response. We hypothesize that omega-6 and
omega-3 fatty acids can differentially modulate TNF-α-induced endothelial cell
activation and that functional called caveolae are required for endothelial cell activation.
Caveolae are particularly abundant in endothelial cells and play a major role in
endothelial trafficking and the regulation of signaling pathways associated with the
pathology of vascular diseases. To test our hypothesis, endothelial cells were pre-
enriched with either linoleic acid or α-linolenic acid prior to TNF-α-induced endothelial
activation. Measurements included oxidative stress and NF-κB-dependent induction of
COX-2 and PGE2 under experimental conditions with intact caveolae and with cells in
which caveolin-1 was silenced by siRNA. Image overlay demonstrated TNF-α-induced
co-localization of TNF receptor type 1 (TNFR-1) with caveolin-1. Caveolin-1 was
significantly induced by TNF-α, which was further amplified by linoleic acid and
blocked by α-linolenic acid. Furthermore, silencing of the caveolin-1 gene completely
blocked TNF-α-induced production of COX-2 and PGE2 and significantly reduced the
amplified response of linoleic acid plus TNF-α. These data suggest that omega-6 and
omega-3 fatty acids can differentially modulate TNF-α-induced inflammatory stimuli and
that caveolae and its fatty acid composition play a regulatory role during TNF-α-induced
endothelial cell activation and inflammation.
47
Introduction
TNF-α exerts its cellular and pathological effects by binding to TNF receptor
(TNFR). There are two distinct subtypes of TNFR: TNFR1 and TNFR2 [169]. TNF
receptors themselves have no metabolic capability and must bind intracellular proteins to
control the signaling of its target cell’s biochemistry.
Dietary ω-3 fatty acids have been shown to markedly alter the lipid composition
of colonic caveolae/lipid rafts in mice fed with fish oil, relative to mice fed with ω-6 fatty
acids [73]. Caveolae are 50- to 100-nm cell-surface plasma membrane invaginations that
are particularly abundant in endothelial cells, where they are believed to play a major role
in the regulation of endothelial vesicular trafficking and signal transduction. Caveolin-1-
deficient mice suggested an important role for caveolin-1 in the pathogenesis of
atherosclerosis, and vascular disease [170].
Furthermore, TNF signaling is initiated at the plasma membrane which is not
homogenous. Recent evidence indicates that TNFR1 is enriched in caveolae [79] and it
can form complex with caveolin-1 [171]. Caveolae are flask-shaped invaginations of
plasma membrane which are composed of caveolin-1, -2 and cholesterol and
glycosphingolipids. These organelles were originally discovered in the early 1950s in
bladder epithelium [172], and have since been observed in adipocytes, fibroblasts,
smooth and striated muscle cells, and endothelial cells show the greatest abundance [54].
Caveolae have been well accepted to function as signaling platforms. Caveolin-1 is the
structural protein for caveolae which are the functional lipid rafts organelles abundant in
endothelial cells. Recently, it has been more accepted that caveolae can also play a key
role in cell signaling, because caveolae especially caveolin-1 has been shown to interact
with many types of plasma membrane receptor proteins and concentrate these molecules
with the caveolae [51].
There is evidence that fatty acids can alter localization and function of caveolae-
associated signaling proteins in mouse colonic mucosa [73]. Besides their role in cellular
uptake of lipophilic substances, including fatty acids [173], caveolae house an array of
cell signaling molecules, and numerous genes involved in endothelial cell dysfunction
and inflammation are associated with caveolae [170]. Examples of caveolae facilitated
48
targeting of proteins involved in pro-inflammatory signal transduction include activators
of p44/p42 (ERK) MAPK pathway, H-Ras [76, 174], non-receptor tyrosine kinase c-Src
[174], and the upstream regulator of NF-κB, IKK [175]. Furthermore, caveolins have
been reported to co-localize with cyclooxygenase, suggesting that caveolins play a role in
regulating the function of this enzyme [176, 177].
A major objective of the current study was to explore specific mechanisms
involved in fatty acid-mediated activation of endothelial cells. Our current data support
our hypothesis that omega-6 and omega-3 fatty acids can differentially modulate TNF-α-
induced endothelial cell activation and that regulatory mechanisms are associated within
caveolae and linked to caveolae function and associated gene inductions.
Materials and Methods
Measurement of caveolin-1 protein level by western blot
Cells were harvested using cell lysis buffer as previously described [105]. 15 μg
protein samples were resolved by SDS-PAGE using 15% gradient gels and transferred
electrophoretically to nitrocellulose membranes using a Bio-Rad immunoblot transfer
apparatus (Bio-Rad Laboratories, Hercules, CA) according to the manufacturer’s
instructions. The nonspecific sites on the membrane were blocked 1 h at room
temperature with 5% nonfat dry milk in TBST followed by incubation with caveolin-1
primary antibody at a dilution of 1:20,000 overnight at 4 °C. Caveolin-1 primary
antibody was diluted 10 times from the original before using filtered PBS. β-actin was
used as a house keeping gene. Bands were visualized using the appropriate horseradish
peroxidase-conjugated secondary antibodies followed by ECL immunoblotting detection
reagents (Amersham Biosci, UK). Hyperfilm (Amersham) was used for the visualization.
Detection and quantitative analysis of the density of the bands were performed using a
digitizing system (UN-SCAN-IT, Silk Scientific Corporation). To determine the optimal
amounts of samples for linear area detection, a titration of different amounts of sample
were loaded. For caveolin-1, 15 μg protein were loaded and for COX-2, 30 μg protein
were loaded.
49
Immunofluorescence microscopy
This technique was adapted from previous protocols [178]. Cells were plated on
LAB-TEKII chamber slides (Nalge Nunc International, Naperville, IL) and grown to
confluence. After experimental treatments, cells were fixed in 4% (vol/vol)
paraformaldehyde (in PBS) for 1 h at room temperature. After permeabilization with
0.1% (vol/vol) Triton X-100 in PBS for 5 min, cells were washed three times with PBS.
Nonspecific binding sites were blocked with 5% (vol/vol) donkey serum in PBS. Cells
were then incubated with anti-caveolin-1 antibody (Affinity BioReagents, Golden, CO)
and anti-TNFR-1 antibody (R&D Systems, Minneapolis, MN), followed by incubation
with Alexa Fluor 488-labeled donkey anti-mouse IgG antibody and Alexa Fluor 546-
labeled donkey anti-goat IgG antibody respectively (Invitrogen-Molecular Probes,
Carlsbad, CA). Cell nuclei were stained using Hoechst (AnaSpec Inc., San Jose, CA).
Slides were mounted in Aqueous Mounting Medium with Anti-fading Agents (Biomeda
Corp., Foster City, CA) and covered with coverslips. Images were captured digitally by
Fluoview 300 Confocal Microscopy (Olympus America Inc., PA). In order to avoid
artificial effects, the detection of the control group was set at a lower level than in Fig.
3.2. For all the other three treatment groups the strength of individual lasers, the level of
contrast, and brightness were kept at the same condition as used for the control group.
Sucrose density gradient fraction of caveolin-1 protein
Three 100 mm culture dishes were used for each treatment group. Sucrose
gradient ultracentrifugation was utilized as described in Appendix B. In brief, the
sucrose gradient was created by gently overlaying lower concentrations of sucrose on
higher concentrations in a centrifuge tube. Cells were washed twice with PBS and lysed
with ice-cold MES-buffered saline. Sucrose was added to the lysate (final concentration
of sucrose 45%), and the lysate was placed at the bottom of the ultracentrifuge tube and
overlaid with lysis buffer containing 35 and 5% sucrose, respectively. After
centrifugation at 39,000 rpm for 16 h, different particles travel through the gradient until
they reach the point in the gradient at which their density matches that of the surrounding
sucrose. Twelve 1 ml fractions from the top to the bottom were collected. Each fraction
50
was analyzed by Western blot for its caveolin-1 contents using specific caveolin-1
antibody.
Detergent-free caveolae membrane purification
Ten 100 mm culture dishes were combined for each treatment group. Detergent-
free caveolae membrane purification was followed Dr. Smart’s method [179] with minor
adjustment. This procedure generates a highly purified plasma membrane microdomain
that is free from intracellular markers and bulk plasma membrane markers and has ben
used extensively to characterize caveolae membrane signaling events [180, 181]. After
multiple ultracentrifugations with 5 to 15% OptiPrep gradient, the band at the 5%
interface was collected and designated caveolae membranes. The cytosol fraction is
obtained by collecting the overlaid material on the Percoll gradient, centrifuging again,
and collecting the resulting supernatant. The plasma membranes are obtained by pooling
the bottoms of both OptiPrep gradients. Each fraction was analyzed by Western blot for
its caveolin-1 contents using specific caveolin-1 antibody. 1 ng protein was loaded for
each sample.
Caveolin-1 siRNA design and transfection
Caveolin-1 gene silencing was adapted from Salani et al [182, 183]. The silence
of caveolin-1 gene (GenBank Accession No. NM001753) was performed using a mix of
two siRNAs directed against the following two target sequences: 5’-
AACAGGGCAACATCTACAAGC-3’ and 5’-AACCAGAAGGGACACACAGTT-3’.
The sequence of control siRNA was as followed: 5’-AAAGAGCGACTTTACACACTT-
3’. Cells were grown in 60 mm plates until 100% confluent and transfected with control
siRNA or caveolin-1 siRNA at a final concentration of 40 nM using GeneSilencer
(Genlantis, San Diego, CA) in Optimem I medium (Invitrogen, Carlsbad, California).
Cells were incubated with 1 ml/plate serum free transfection mixtures for 4 hr, and then
add 1 ml/plate of 20% FBS OptiMEM to achieve the final concentration of 10% FBS in
the medium. 48 h after transfection, replace the medium by 5% FBS M199 overnight and
then treated with TNF-α in the presence or absence of linoleic acid.
51
PGE2 determination
Cells were seeded in 60 mm culture dishes (Becton Dickinson Labware, NJ) and
grown to confluence. After transfected with caveolin-1 siRNA or control siRNA, cells
were pretreated with linoleic acid followed by TNF-α exposure. After treatment,
supernatants of cell cultures were collected into microcentrifuge tubes (Isc BioExpress,
UT), centrifuged at 4 °C to remove cellular debris and stored at -80 °C. PGE2 levels were
assessed using a PGE2-specific enzyme immunoassay (EIA) (Cayman Chemicals, MI)
following the manufacturer’s protocol. Absorbance at 405 nm was detected using a
microplate spectrophotometer SpectraMaxPro M2 (Molecular Devices Corporation, CA).
Statistical analysis
Values are reported as mean ± standard error of the mean (SEM) of at least three
independent experiments. Data were analyzed using Sigma Stat software (Jandel Corp.,
Wan Rafael, CA). One way ANOVA was followed by post hoc comparisons using the
least significant difference (LSD) method. A statistical probability of p< 0.05 was
considered significant. Different letters represent significant differences among treatment
groups. Capital or small letters represent caveolin-1 silencing or control silencing
respectively as were compared separately by one way ANOVA.
Results
Fatty acids and TNF-α modulated caveolin-1 expression
There is increasing evidence that lipid raft proteins and lipids play an important
role in health and disease [184]. A major subclass of lipid rafts are caveolae, i.e.,
membrane domains which have been implicated in the pathology of atherosclerosis [170].
The lack of the caveolin-1 gene may provide protection against the development of
atherosclerosis [185]. This may be important in understanding mechanisms of
atherosclerosis, because caveolae are particularly abundant in endothelial cells, where
they are believed to play a major role in the regulation of endothelial vesicular trafficking
as well as the uptake of lipids and related lipophilic compounds [186]. To assess the
52
effects of linoleic acid and α-linolenic acid on caveolin-1 expression, endothelial cells
were incubated with either linoleic acid or α-linolenic acid for 24 h. As indicated in Fig.
3.1, caveolin-1 protein level was significantly down-regulated by α-linolenic acid
compared with the control culture, while there was no different in the linoleic acid
treatment from control group. We next investigated the effect of TNF-α on caveolin-1
expression. As shown in Fig. 3.2, TNF-α increased caveolin-1 expression at 4 h
measured by immunofluorescence study. We can also observe the basal level of
caveolin-1 expression in the control culture.
Linoleic acid and α-linolenic acid modulate TNF-α-induced caveolin-1 up-regulation
and the co-localization of TNFR-1and caveolin-1
The first step in the TNF-α-induced inflammatory response is the binding of the
cytokine to its cognate receptor. TNF receptor type 1 (TNFR-1) has been reported to be
enriched in caveolae [79]. Caveolin-1 has also been shown to form a complex with the
TNF receptor [171]. We further pre-incubated endothelial cells with either linoleic acid
or α-linolenic acid, followed by activation with TNF-α. The immunofluorescence
images in Fig. 3.3 demonstrated that TNF-α can increase caveolin-1 and TNFR-1
expression compared with the control group, and that pretreated cells with linoleic acid
followed by TNF-α can further increase these effects compared with TNF-α treatment
alone. Cells pretreated with α-linolenic acid followed by exposure to TNF-α diminished
TNF-α-induced both caveolin-1 and TNFR-1 up-regulation. Similar effects of linoleic
acid and α-linolenic acid on TNF-α-induced caveolin-1 expression were observed by
measuring caveolin-1 protein levels by western blot (Fig. 3.4).
Linoleic acid and α-linolenic acid modulate TNF-α-induced caveolin-1 re-distribution
To further establish the caveolar localization of TNFR-1, after treated cells with
different fatty acids for 24 h followed by 4 h TNF-α treatment, we used sucrose density
gradient centrifugation. 12 fractions were collected and transferred to nitrocellulose
membrane by Western blot. Caveolin-1 protein levels were analyzed by caveolin-1
specific antibody. As indicated in Fig. 3.5, in the control culture, caveolin-1 protein were
53
most enriched in fraction 4 and 5 (containing 5% sucrose, represent detergent-resistant
membranes). TNF-α induced a shift of caveolin-1 protein into fraction 3 and linoleic
acid further amplified this effect. In contrast, α-linolenic acid diminished TNF-α-
induced the shift of caveolin-1 into fraction 3, and even decreased caveolin-1 in fraction
5 compared with control treatment. To confirm that the observation of this shift was not
due to uneven transfer or other artificial effects, we analyzed fractions 3, 4, and 5 from all
the four treatment groups on one western blot gel and similar results were observed (Fig.
3.5, lower panel). In fractions 7 to 9, we also observed a shift of caveolin-1 protein from
fraction 8 and 9 into fraction 7 after TNF-α treatment. In the presence of linoleic acid,
the TNF-α-induced shift into fraction 7 was amplified. But in the presence of α-linolenic
acid, there was no change of the TNF-α-induced shift compared with TNF-α treatment
alone. In fractions 10 to 12 (containing 45% sucrose) we didn’t observe the association
of caveolin-1 protein.
Furthermore, the results of detergent-free caveolae membrane purification
indicated that only in the presence of linoleic acid, caveolin-1 was enriched in cytosol
membrane proportion (Fig. 3.6). These date suggested that linoleic acid or α-linolenic
acid had profound modulation effects on caveolin-1 distribution.
Caveolin-1 silencing decreases TNF-a-induced caveolin-1 and COX-2 expression,
subsequently diminishes PGE2 production
In order to determine the role of caveolin-1 in fatty acids-modulated TNF-α-
induced endothelial cell activation, we utilized small interfering RNA (siRNA) to
specifically down-regulate caveolin-1. As indicated in Fig. 3.7, after 48 h transfection of
caveolin-1 siRNA or control siRNA, cells were exposed to TNF-α for 4 h, in both control
and TNF-α treatment groups, caveolin-1 siRNA specifically reduced caveolin-1 protein
expression by approximately 80%. Similar decreasing effect by siRNA was also
observed after 24 h treatment with TNF-α in the presence or absence of linoleic acid (Fig.
3.8). TNF-α-induced COX-2 expression was totally blocked by caveolin-1 silencing (Fig.
3.9) and caveolin-1 silencing also diminished the combined COX-2 induction induced by
both linoleic acid and TNF-α. Similar to the COX-2 data, caveolin-1 silencing blocked
54
TNF-α-induced PGE2 production and also diminished TNF-α-induced PGE2 production
in the presence of linoleic acid (Fig. 3.10). The effect of linoleic acid-induced COX-2
expression and PGE2 production were not changed after caveolin-1 silencing. Clearly,
the functional caveolae are important for TNF-α-induced up-regulation of COX-2 and
subsequently production of PGE2. Linoleic acid-induced COX-2 expression and PGE2
production were not caveolae dependent, indicated that an alternative signaling pathway
might be involved.
55
Figure 3.1 Effect of linoleic acid and α-linolenic acid on caveolin-1 expression
Cells were treated with 20 μM of linoleic acid (LA) or α-linolenic acid (ALA) for 24 h.
Experiments were repeated three times, and the blots shown are a representative of one of
the experiments. The bar graph shows the corresponding densitometric analysis of the
blots. Values are means ± SEM. Different letters represent significant differences
among treatment groups.
56
Figure 3.2 Immunofluorescence microscopy study of TNF-α-induced caveolin-1 up-
reguation.
Cells were treated with 0.5 ng/ml TNF-α for 4 h. FITC green fluorescence was used for
immunofluorescence staining of caveolin-1. Experiment were repeated three times and
the epifluorescence micrographs shown are representative fields of one of the
experiments (original magnification, X 400).
57
Figure 3.3 Effect of linoleic acid and α-linolenic acid on TNF-α-induced TNFR-1 and
caveolin-1 localization
Cells were treated with 0.5ng/ml TNF-α alone for 4 h, or pretreated with 20 μM linloleic
acid (LA) or α-linolenic acid (ALA) for 24 h followed by exposure to TNF-α for 4 h.
Immunofluorescence staining for caveolin-1 (FITC green fluorescence), TNF Receptor-1
(TNFR-1) (Texas red fluorescence), and nuclear (DAPI blue fluorescence). The merged
images were shown to better illustrate the partial colocalization between caveolin-1 and
TNFR-1 in the same cells. Experiments were repeated three times and the
epifluorescence micrographs shown are representative fields of one of the experiments
(original magnification, X 400).
58
Figure 3.4 Effect of linoleic acid and α-linolenic acid on TNF-α-induced caveolin-1
expression
Cells were treated with 0.5ng/ml TNF-α alone for 4 h, or pretreated with 20μM linoleic
acid (LA) or α-linolenic acid (ALA) for 24 h followed by exposure to TNF-α for 4 h.
Caveolin-1 protein levels were detected by western blot. β-Actin was used as a
housekeeping gene in the measurement. Experiments were repeated three times, and the
blots shown are a representative of one of the experiments. The bar graph shows the
corresponding densitometric analysis of the blots. Values are means ± SEM. Different
letters represent significant differences among treatment groups.
59
Figure 3.5 Effects of fatty acids on TNF-α-induced caveolin-1 redistribution
Cells were treated with 0.5ng/ml TNF-α alone for 4 h, or pretreated with 20 μM linoleic
acid (LA) or α-linolenic acid (ALA) for 24 h followed by exposure to TNF-α for 4 h.
Caveolin-1 distribution was measured by glucose sucrose gradient. Fractions 1 to 3
containing 5% sucrose, fractions 4 to 8 containing 35% sucrose, fractions 9 to 12
containing 45% sucrose.
60
Figure 3.6 Effects of fatty acids on TNF-α-induced caveolin-1 redistribution Cells were treated with 0.5ng/ml TNF-α alone for 4 h, or pretreated with 20 μM linoleic
acid (LA) or α-linolenic acid (ALA) for 24 h followed by exposure to TNF-α for 4 h.
Different cellular membranes were isolated using detergent-free caveolae membrane
purification. This method has been well established, and different isolated membranes
have been confirmed by specific markers.
61
Figure 3.7 Effect of caveolin-1 silencing on TNF-α-induced caveolin-1 up-regulation Cells were first transfected with siRNA for caveolin-1 (Cav-1 siRNA) or with scrambled
control siRNA for 48 h. Then cells were treated with 0.5ng/ml TNF-α for 4 h. Whole
cell lysate was collected and caveolin-1 protein levels were measured by western blot.
Figure 3.8 Effect of caveolin-1 silencing on linoleic acid and TNF-α-induced caveolin-1
expression
Cells were first transfected with siRNA for caveolin-1 (Cav-1 siRNA) or with scrambled
control siRNA, then treated with 0.5ng/ml TNF-α or linoleic acid (LA) alone for 24 h, or
pretreated with 20μM LA for 24 h followed by exposure to TNF-α for additional 24 h.
Whole cell lysate was collected and caveolin-1 protein levels were measured by western
blot. β-Actin was used as a housekeeping gene. Experiments were repeated three times
and the blots shown are a representative of one of the experiments.
62
Figure 3.9 Effect of caveolin-1 silencing on linoleic acid and TNF-α-induced COX-2
expression
Cells were first transfected with siRNA for caveolin-1 (Cav-1 siRNA) or with
scrambled control siRNA. Then cells were treated with 0.5ng/ml TNF-α alone for 24 h,
or pretreated with 20μM LA for 24 h followed by exposure to TNF-α for additional 24 h.
COX-2 protein levels were measured by western blot under the same experimental
conditions as in Fig. 3.5B. β-Actin was used as a housekeeping gene. Experiments were
repeated three times, and the blots shown are representative of one of the experiments.
Values are means ± SEM. Different letters represent significant difference among
treatment groups. Capital or small letters represent caveolin-1 silencing or control
silencing respectively as were compared separately by one way ANOVA.
63
Figure 3.10 Effect of caveolin-1 silencing on linoleic acid and TNF-α-induced PGE2
production
Cells were first transfected with siRNA for caveolin-1 (Cav-1 siRNA) or with
scrambled control siRNA. Then cells were treated with 0.5ng/ml TNF-α or linoleic acid
(LA) alone for 24 h, or pretreated with 20μM LA for 24 h followed by exposure to TNF-
α for additional 24 h. Supernatants of cell cultures were collected and PGE2 levels were
measured by EIA under the same experimental conditions as in Fig. 3.8 and 3.9.
Experiments were repeated three times and the blots shown are a representative of one of
the experiments. Values are means ± SEM. Different letters represent significant
differences among treatment groups. Capital or small letters represent caveolin-1
silencing or control silencing respectively as were compared separately by one way
ANOVA.
64
Discussion
Caveolae, specific microdomains of the plasma membrane, can concentrate
different membrane receptors, signaling molecules and membrane transporters to vary
extents. Caveolins are thought to function as scaffolding proteins that organize and
concentrate cholesterol, glycosphingolipids and lipid-modified signaling molecules, such
as endothelial nitric oxide synthase, H-ras, Src-like kinases, G protein and several
hormone receptors within caveolae [187, 188]. The binding of both free cholesterol and
protein by caveolae involves the same, short sequence in caveolin (residues 61 to 101)
[187]. This implies that there might be a competition between lipids and proteins for the
same site, within a highly ordered assembly of multiple caveolin molecules within a
single caveola, in which individual polypeptides have assigned roles in the binding of
either free cholesterol or protein. Several proteins of cellular lipid homeostasis are found
in caveolae, including annexin II, lipoprotein scavenger receptors and CD36 [189].
These proteins do not have a caveolin-binding scaffold consensus sequence, suggesting
that their primary interaction is with the lipid, not protein, moiety of caveolae [190].
Our data presented in Chapter 2 suggest that exposure to TNF-α can cause
endothelial cell activation and that fatty acids (and in particular linoleic acid) can amplify
TNF-α-induced endothelial activation [156], mechanisms of this interaction may be
multi-layered and are not known. We further hypothesized that caveolae play a critical
role in facilitating the cellular uptake and trafficking of fatty acids and trigger the initial
cell signaling induced by TNF-α and lipids. We utilized siRNA-induced down-
regulation of caveolin-1 which is an effective experimental approach to determine the
role of caveolae in cellular signal transduction. Our data suggest that functional caveolae
are necessary for TNF-α-mediated activation of endothelial cells. TNF-α markedly
induced both caveolin-1 and TNFR-1, and both expressions were further induced when
cells were first enriched with linoleic acid. Furthermore, silencing caveolin-1 totally
blocked both TNF-α-induced COX-2 expression and cellular release of PCE2. In human
foreskin fibroblasts, a significant proportion of induced COX-2 was present in DRM
fractions concomitantly with caveolin-1 [80].
65
It is very likely that selected fatty acids either stabilize or perturb caveolae
functions, thus leading to modifications of caveolae-dependent signaling. There is
evidence that fatty acids can alter localization and function of caveolae-associated
signaling proteins in mouse colonic mucosa [73]. Our data suggest that omega-3 fatty
acids may in part be anti-inflammatory by decreasing caveolin-1 expression or by causing
dysfunctional caveolae, leading to down-regulation of caveolin-dependent down-stream
signaling. Using human breast cancer cells, others have shown recently that omega-3
fatty acids can alter lipid raft composition and a decrease in epidermal growth factors
[74], suggesting that selected fatty acids can modify caveolae-associated cell functions.
There is evidence that caveolae play a role in lipid trafficking and uptake and transport of
fatty acids [184]. Furthermore, preferential uptake of fatty acids via the FAT/CD36, a
receptor exclusively located in lipid rafts such as caveolae in the plasma membrane, may
explain the fatty acid effects we observed. In fact, working with mouse embryonic
fibroblasts, it was recently demonstrated that mistargeting of FAT/CD36 in these cells
lacking the caveolin-1 gene resulted in reduced fatty acid uptake compared with the wild-
type cells [191]. Long-chain omega-3 fatty acids, such as DHA, have been shown to
induce caveolar cholesterol reduction, this might due to the poor incorporation of
cholesterol into phospholipids bilayers be explained by in a highly unsaturated lipid
environment, [192]. In addition, α-linolenic acid has also been shown to modulate
cholesterol partitioning into model membranes [193].
Our observations that linoleic acid can amplify the TNF-α-mediated induction of
both caveolin-1 and COX-2 is significant because of the link between caveolins (caveolae)
and the pathology of atherosclerosis [170]. Caveolin-1 has been reported to co-localize
with interleukin-1beta-induced COX-2 [194], suggesting the dependence of COX-2
induction on functional caveolae. Recent evidence suggests that high-fat diets can up-
regulate caveolin-1 expression in aorta of diet-induced obese rats [195], suggesting that
our fatty acid data may mimic an in vivo response by activating COX-2. High-fat diets
contribute to hypertriglyceridemia, and the vascular endothelium can be exposed to
significant levels of free fatty acids derived from lipoprotein lipase-mediated hydrolysis
of triglyceride-rich lipoproteins [128].
66
In summary, we provide novel data demonstrating that omega-6 and omega-3
fatty acids can differentially modulate TNF-α-induced inflammatory stimuli and that
these events require functional caveolae. Furthermore, functional changes of caveolae
associated with modifications by dietary fatty acids appear to affect critical phases of
induction of oxidative stress-sensitive transcription factors and inducible inflammatory
parameters during endothelial cell activation. Because caveolae and caveolins have been
implicated in several human diseases and in particular vascular diseases, our data may
have implications in understanding novel mechanisms of inflammatory diseases
modulated by dietary lipids.
Copyright © Lei Wang 2007
67
Chapter Four : Changing Ratios of ω-6 to ω-3 Fatty Acids Can Differentially
Modulate Polychlorinated Biphenyl Toxicity in Endothelial Cells
Synopsis
Exposure to persistent organic pollutants, such as polychlorinated biphenyls
(PCBs) can cause endothelial cell activation by inducing pro-inflammatory signaling
pathways. Our previous studies indicated that linoleic acid (LA, 18:2), a major omega-6
unsaturated fatty acid in the American diet, can potentiate PCB77-mediated inflammatory
responses in ECs. In addition, omega-3 fatty acids (such as α-linolenic acid, ALA, 18:3)
are known for their anti-inflammatory properties. We tested the hypothesis that
mechanisms of PCB-induced endothelial cell activation and inflammation can be
modified by different ratios of omega-6 to omega-3 fatty acids. Porcine pulmonary
arterial endothelial cells were pretreated with linoleic acid, α-linolenic acid, or different
ratios of these fatty acids, followed by exposure to PCB77. PCB77-induced oxidative
stress and activation of the oxidative stress sensitive transcription factor nuclear factor κB
were markedly increased in the presence of linoleic acid and diminished by increasing the
relative amount of α-linolenic acid to linoleic acid. Similar protective effects by
increasing α-linolenic acid were observed by measuring NF-κB-responsive genes, such
as vascular cell adhesion molecule-1 (VCAM-1) and cyclooxygenase-2 (COX-2). COX-
2 catalyzes the rate limiting step of the biosynthesis of prostaglandin E2 (PGE2). PCB77
exposure also increased PGE2 levels, which were down-regulated with relative increasing
amounts of α-linolenic acid to linoleic acid. Inhibitor studies suggest that NF-κB is a
critical player in the regulation of PCB-induced inflammatory markers as modulated by
omega-6 and omega-3 fatty acids.
Introduction
Substantial evidence from epidemiological studies suggests that cardiovascular
diseases are linked to environmental pollution. For example, there was a significant
68
increase in mortality from cardiovascular diseases among Swedish capacitor
manufacturing workers exposed to polychlorinated biphenyls (PCBs) for at least five
years [8], and most excess deaths were due to cardiovascular disease in power workers
exposed to phenoxy herbicides and PCBs in waste transformer oil [9]. Furthermore, an
increase in hospitalization rates was reported for coronary heart disease in populations
residing near areas contaminated with persistent organic pollutants [196]. A recent study
still found excessive concentrations of PCBs in the serum of Yusho victims, 35 years
after accidental poisoning with PCBs in Nagasaki, Japan [197]. Interestingly, lipid
peroxidation was markedly enhanced in these victims as well. These studies suggest that
populations near contaminated sites are at increased risk to develop cardiovascular
diseases, and in particular in the presence of additional risk factors, such as
hypertriglyceridemia and an associated persistent state of oxidative stress. There is
evidence linking the aryl hydrocarbon receptor (AhR) with mechanisms associated with
cardiovascular diseases [198] and that AhR ligands may be atherogenic by disrupting the
functions of endothelial cells in blood vessels.
A growing body of literature demonstrated that dietary components can modulate
the inflammatory response in humans, thereby affecting cardiovascular risk. Evidence
suggests that nutrition can influcence the lipid milieu, oxidative stress and antioxidant
status within cells, and thus modulate mechanisms of cytotoxicity mediated by
environmental pollutants [199]. For example, certain dietary fats may increase the risk to
environmental insult induced by PCBs, while fruits and vegetables, rich in antioxidant
and anti-inflammatory nutrients or bioactive compounds, may provide protection [200].
Specific fatty acids rich in plant oils, such as linoleic acid (the parent omega-6
fatty acid), can amplify PCB toxicity in vascular endothelial cells [106]. There is also
evidence that elevated levels of linoleic acid may enhance the cellular availability of
PCBs [201]. The average American diet comprises of an excess of ω-6 fatty acids
compared with ω-3 fatty acids, the ratio can reach 10 to 20:1, and it has been suggested
that this imbalance could potentially promote a prothrombotic state [202]. Furthermore,
coplanar PCBs can suppress delta 5 and 6 desaturase activities, thus disrupting the
synthesis of fatty acid precursors for eicosanoid metabolism [203]. Our own data from
plasma and livers of LDL receptor-deficient mice support the hypothesis that treatment
69
with PCBs can facilitate clearance of linoleic acid from plasma into vascular tissues [108].
Such a change in lipid milieu could exacerbate fatty acid- and/or PCB-induced oxidative
stress and a vascular inflammatory response.
In contrast to omega-6 fatty acids, omega-3 fatty acids can influence
cardiovascular disease pathology by beneficially modulating inflammation.
Epidemiological and interventional studies have shown a dose-dependent decrease in risk
of cardiovascular disease endpoints with increased dietary consumption of moderate
amounts of omega-3 fatty acids, either plant or marine derived [126]. Cardio-protective
properties of omega-3 fatty acids include down-regulation of proinflammatory and
proatherogenic genes, including adhesion molecules and cytokines, during early
atherogenesis and possibly also during later stages of plaque development and plaque
rupture [130]. For example, an α-linolenic acid-rich oil decreased oxidative stress and
CD40 ligand in patients with mild hypercholesterolemia [131], reduced levels of soluble
cell adhesion molecules in plasma [132] and recurrence of coronary heart disease [133].
Randomized trials have reported that after α-linolenic acid supplementation reached the
ω-6 to ω-3 fatty acids ratio 1.3:1, a significant reduction of c-reactive protein and
interleutin-6 levels were observed [204].
The mechanisms by which cytokines and environmental chemicals induce
endothelial cell activation, oxidative stress and inflammation are not fully understood.
Oxidative stress-induced transcription factors, which regulate inflammatory cytokine and
adhesion molecule production, play critical roles in the induction of inflammatory
responses. One of these transcription factors, nuclear factor κB (NF-κB), plays a
significant role in these regulatory processes [115]. Binding sites for NF-κB and related
transcription factors were identified in the promoter regions of a variety of inflammatory
genes [116, 117] such as interleukin 6 (IL-6), vascular cell adhesion molecule-1 (VCAM-
1) or cyclooxygenase-2 (COX-2), all of which are up-regulated during PCB toxicity [104,
197, 205, 206]
Of increasing recognition is the paradigm that nutrition can modulate the toxicity
of environmental pollutants and thus affect health and disease outcome associated with
chemical insult [200]. The current study was designed to test the hypothesis that PCB-
induced endothelial cell inflammation can be enhanced by omega-6 fatty acids and
70
antagonized by omega-3 fatty acids. We focused on omega-6 and omega-3 fatty acids,
which are most commonly consumed in the average U.S. diet [129].
Materials and Methods
Cell culture and experimental media
Endothelial cells were isolated from porcine pulmonary arteries and cultured as
previously described [30]. Arteries obtained during routine slaughter were donated from
the College of Agriculture, University of Kentucky. The basic culture medium consisted
of medium 199 (M-199) (GIBCO Laboratories, NY) containing 10% (v/v) fetal bovine
serum (FBS, HyClone Laboratories, UT). The experimental media were composed of M-
199 enriched with 5% (v/v) FBS and with different ratios of linoleic acid (LA) to α-
linolenic acid (ALA) (> 99% pure; Nu-Chek Prep, MN). Preparation of experimental
media with LA and ALA were performed as described earlier [136]. Different ratios of
LA to ALA were 2:1, 1:1, and 1:2 (v/v), and the total concentration of fatty acids in all
the cultures did not exceed 20 µM. Following the pretreatment with fatty acids for 18 h,
cells were exposed to coplanar PCB77 for indicated times (6 to 24 h). In NF-κB
inhibition studies, cells were first pre-enriched with 50 μM PDTC (EMD Biosciences,
Inc., CA) for 4 h, then treated with PCB77 or LA.
PCB77 solution
PCB77 was solubilized in DMSO (sterile-filtered, Sigma-Aldrich, MO) and the
final concentration in the cell culture media was 3.4 μM. This level was chosen because
it has been reported in serum after acute exposure to PCBs [207, 208]. The final
concentration of DMSO in the culture media did not exceed 0.03%. All vehicle controls
and PCB treated cultures contained the same amount of DMSO. On the other hand,
controls for fatty acid experiments did not contain DMSO.
Analysis of PGE2 synthase gene expression by real-time PCR
Total RNA was extracted from endothelial cells using RNA-STAT-60 (TEL-
TEST, Friendswood, TX) according to the manufacturer’s protocol. Reverse
71
transcription was performed using the AMV reverse transcription system (Promega,
Madison, WI). The levels of mRNAs and the PCR-product were then assessed by real-
time PCR using 7300 Real Time PCR System (Applied Biosystems). Real-time PCR
samples were mixed with SYBR Green Master Mix (Applied Biosystems) and PGE2
specific primers. The sequences for porcine PGE2 gene were designed by Primer Express
Software 3.0 for real-time PCR (Applied Biosystems). PGE2 sequences: sense, 5’-GCC-
TGAGCGTCTGTGTTTTTTA-3’; antisense, 5’-GCACCTGAATCCTCAAGGACAT-3’.
β-actin was used as a housekeeping gene. β-actin sequences: sense, 5’-
TCATCACCATCGGCAACG-3’; antisense, 5’-TTCCTGATGTCCACGTCG-3’.
Statistical analysis
Values are reported as mean ± standard error of the mean (SEM) of at least three
independent groups. Data were analyzed using Sigma Stat software (Jandel Corp., Wan
Rafael, CA). One way ANOVA followed by post hoc least significant difference (LSD)’s
pairwise multiple comparison procedure were used for statistical analysis of the original
data. A statistical probability of p< 0.05 was considered significant. In figure 4.8, *
represent significant differences among treatment groups. In all the other figures,
different letters represent significant differences among treatment groups. Capital or
small letters represent with or without PCB77 respectively as were compared separately
by one way ANOVA.
Results
Different ratios of linoleic acid to α-linolenic acid modulate cellular oxidative stress
induced by exposure to PCB77
To assess the effects of changing relative amounts of linoleic acid (LA) to α-
linolenic acid (ALA) on oxidative stress generation in the absence and presence of
PCB77, endothelial cells were incubated with different ratios of LA to ALA for 24 h
without PCB77 exposure, or pre-incubated with different ratios of LA to ALA for 18 h
and then exposed to PCB77 for an additional 6 h. This 6 h incubation time was
72
determined from our previous study [106]. As indicated in Fig. 4.1, in the absence of
PCB77, LA significantly induced oxidative stress as observed by DCF fluorescence
compared with control cells. When increasing the relative amount of ALA, a significant
decrease in oxidative stress was observed in all groups containing more ALA than LA.
In the PCB77 treatment group, oxidative stress was increased significantly compared
with the control group, and pretreated cells with LA followed by PCB77 further induced
oxidative stress compared with cells that were treated only with PCB77 or LA.
Replacing LA with relative increasing amounts of ALA significantly reduced the additive
effect of both LA and PCB on oxidative stress. Furthermore, cells pretreated with ALA
followed by exposure to PCB77 blocked the oxidative stress induced with PCB77
treatment alone.
Different ratios of linoleic acid toα-linolenic acid modulate PCB77-induced NF-κB DNA
binding activity
Oxidative stress can alter gene expression via the activation of redox sensitive
transcription factors such as NF-κB [209]. Previous studies in our lab have demonstrated
that treatment of PCB77 for 3 h can activate NF-κB, but the highest activation at
approximately 6 h [106]. In the current study, we further investigated the effects of
changing the relative amounts of LA to ALA on PCB77-induced NF-κB activation.
First, endothelial cells were incubated with different ratios of LA to ALA for 6 h. As
shown in Fig. 4.2, LA significantly induced NF-κB activation. When introducing ALA
into the media, the activity of NF-κB significantly decreased compared with cells treated
with only LA. As indicated in Fig. 4.3, cells were also pretreated first with different
ratios of LA to ALA for 18 h, followed by exposure to PCB77 for an additional 6 h.
Compared to the control group, exposure to PCB77 significantly increased NF-κB DNA
binding activity and this effect of PCB77 was further increased in the presence of
supplemental LA. When introducing ALA into the media, the activity of NF-κB
significantly decreased compared with cells treated with only LA plus PCB77. When
cultures were pretreated only with ALA, a PCB77-induced increase in NF-κB DNA
binding was completely blocked when compared with the control group.
73
Different ratios of linoleic acid toα-linolenic acid modulate PCB77-induced VCAM-1
expression
NF-κB is critical in the regulation and expression of inflammatory genes, such as
adhesion molecule VCAM-1. Expression of VCAM-1 on endothelial cells represents one
of the early pathological changes in immune and inflammatory diseases, such as
atherosclerosis [210]. We investigated whether PCB77-induced VCAM-1 expression
was mediated by changing the ratios of LA to ALA. As indicated in Fig. 4.4, VCAM-1
gene expression, measured by real-time PCR, was up-regulated after exposure to PCB77.
LA pre-treatment alone followed by PCB77 increased VCAM-1 gene expression, which
was reduced with a relative increase in ALA. This real-time PCR data was confirmed by
measuring VCAM-1 protein levels by western blot (Fig. 4.5).
PCB77 modulate three enzymes participating in PGE2 synthesis
The biosynthesis of prostaglandin E2 (PGE2), a major lipid-derived inflammatory
mediator, involves three enzymatic reactions including phospholipase A2 (PLA2), COX,
and PGE2 synthase. To assess the effect of PCB77 on PGE2 production, we first
measured the activation of cPLA2. As shown in Fig. 4.6, PCB77 induced an increase of
cPLA2 phosphorylation at 8 h treatment. Similarly, COX-2 which catalyzes the rate
limiting step for PGE2 production was also increased by PCB77 at 8 h (Fig. 4.7).
Furthermore, the gene expression of PGE2 synthase was significantly increased after 24 h
of PCB77 treatment (Fig. 4.8).
Different ratios of linoleic acid toα-linolenic acid modulate PCB77-induced COX-2
expression and subsequent PGE2 production
COX-2 is an inducible pro-inflammatory enzyme controlled by NF-κB [211]. To
assess whether different ratios of LA to ALA can modulate PCB77-induced COX-2 up-
regulation, real time RT-PCR was performed on total cell RNA. Similar to the NF-κB
data, PCB77-induced COX-2 mRNA levels were attenuated by increasing relative
amounts of ALA (Fig. 4.9). These data were confirmed by measuring protein levels of
COX-2 by western blot (Fig. 4.10).
74
The inducible COX-2 isoform plays a key role in inflammation and is the rate-
limiting enzyme for prostaglandin synthesis. In order to assess COX-2 activity, we
measured PGE2 production. As shown in Fig. 4.11, PCB77 significantly induced PGE2
production in a time-dependent manner. We also pretreated cells with different ratios of
LA to ALA for 18 h followed by exposure to PCB77 for an additional 24 h. As indicated
in Fig. 4.12, compared to the control group, LA significantly increased PGE2 production,
which was further enhanced by PCB77 exposure. Substituting relative amounts of LA
with ALA markedly decreased the PCB-induced PGE2 production.
Inhibition of NF-κB decreases PCB77- and linoleic acid-induced VCAM-1 and COX-2
expression
In order to confirm that NF-κB is a critical transcription factor in PCB77-induced
endothelial inflammatory response as modulated by fatty acids, we studied the effect of
pyrrolidine dithiocarbamate (PDTC), an antioxidant, markedly inhibited the activation of
NF-κB, on both PCB77- and LA-mediated VCAM-1 and COX-2 expression by real time
RT-PCR. As shown in Fig. 4.13 and 4.14, PDTC significantly decreased PCB77- and
LA-induced VCAM-1 and COX-2 expression, respectively. These results clearly
indicate that NF-κB is crucially involved in linoleic acid and PCB77 modulated
endothelial cell activation.
75
Figure 4.1 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced oxidative stress
Cultures were incubated in media supplemented with 20 μM fatty acids, either all linoleic
acid (LA), all α-linolenic acid (ALA), or different ratios of LA to ALA (R2:1, R1:1, R1:2)
for 24 h. Some cultures were first pre-treated with these different ratios of fatty acids for
18 h, followed by co-exposure to 3.4 μM PCB77 for additional 6 h. Values are means ±
SEM (n = 3). Statistical comparisons of the two experimental settings (without and with
added PCB77) are indicated by small and capital letters. Different letters represent
significant differences among the treatment groups.
76
Figure 4.2 Effect of different ratios of linoleic acid to α-linolenic acid on the activation
of NF-κB
Cells were treated with 20 μM of linoleic acid (LA), α-linolenic acid (ALA), or different
ratios of LA to ALA (R2:1, R1:1, R1:2) for 6 h. Experiments were repeated three times,
and the blots shown are a representative of one of the experiments. The bar graph shows
the corresponding densitometric analysis of the blots. Values are means ± SEM.
Different letters represent significant differences among treatment groups.
77
Figure 4.3 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced activation of NF-κB
Cells were treated with 20 μM of linoleic acid (LA), α-linolenic acid (ALA), or different
ratios of LA to ALA (R2:1, R1:1, R1:2) for 18 h prior to exposure to 3.4 μM PCB77 for
an additional 6 h. Experiments were repeated three times, and the blots shown are a
representative of one of the experiments. The bar graph shows the corresponding
densitometric analysis of the blots. Values are means ± SEM. Different letters represent
significant differences among treatment groups.
78
Figure 4.4 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced VCAM-1 gene expression
Endothelial cells were pretreated with 20 μM of linoleic acid (LA), α-linolenic acid
(ALA), or different ratios of LA to ALA (R2:1, R1:1, R1:2) for 18 h and then exposed to
3.4 μM PCB77 for additional 6 h for VCAM-1 mRNA measurement. β-actin was used
as a housekeeping gene. Experiments were repeated three times. Values are means ±
SEM. Different letters represent significant differences among treatment groups.
79
Figure 4.5 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced VCAM-1 protein expression
Endothelial cells were pretreated with 20 μM of linoleic acid (LA), α-linolenic acid
(ALA), or different ratios of LA to ALA (R2:1, R1:1, R1:2) for 18 h and then exposed to
3.4 μM PCB77 for additional 8 h. β-actin was used as a housekeeping gene.
Experiments were repeated three times, and the blots shown are a representative of one of
the experiments. Values are means ± SEM. Different letters represent significant
differences among treatment groups.
80
Figure 4.6 Effect of PCB77 on cPLA2 phosphorylation
Cells were exposed to 3.4 μM PCB77 for indicated time. Phosphorylated cPLA2 was
measured by phosph-cPLA2 specific antibody. β-Actin was used as a housekeeping gene.
Experiment was repeated twice as shown.
Figure 4.7 Effect of PCB77 on COX-2 expression
Cells were exposed to 3.4 μM PCB77 for 6 h. COX-2 protein levels were measured by
western blot. β-Actin was used as a housekeeping gene. Experiment was repeated twice
as shown.
81
Figure 4.8 Effect of PCB77 on PGE2 synthase expression
Cells were exposed to 3.4 μM PCB77 for indicated time. PGE2 synthase gene
expression was measured by real-time PCR. β-Actin was used as a housekeeping gene.
Experiments were repeated three times. Values are means ± SEM. * represents
significant difference among treatment groups.
82
Figure 4.9 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced COX-2 gene expression
Cells were pretreated with 20 μM of linoleic acid (LA), α-linolenic acid (ALA), or
different ratios of LA to ALA (R2:1, R1:1, R1:2) for 18 h and then exposed to 3.4 μM
PCB77 for additional 6 h. β-Actin was used as a housekeeping gene. Experiments were
repeated three times. Values are means ± SEM. Different letters represent significant
difference among treatment groups.
83
Figure 4.10 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
induced COX-2 protein expression
Cells were pretreated with 20 μM of linoleic acid (LA), α-linolenic acid (ALA), or
different ratios of LA to ALA (R2:1, R1:1, R1:2) for 18 h and then exposed to 3.4 μM
PCB77 for additional 8 h. β-Actin was used as a housekeeping gene. Experiments were
repeated three times, and the blots shown are representative of one of the experiments.
Values are means ± SEM. Different letters represent significant difference among
treatment groups.
84
Figure 4.11 Optimal PCB77 incubation time for PGE2 production
Cells were exposed to 3.4 μM PCB77 for 6, 18 and 24 h. Supernatants of cell cultures
were collected and PGE2 levels were measured by enzyme immunoassay (EIA). Bars
represent means ± SEM from three independent experiments. Different letters represent
significant difference among treatment groups.
85
Figure 4.12 Effect of different ratios of linoleic acid to α-linolenic acid on PCB77-
stimulated release of PGE2
Cells were incubated in media supplemented with 20 μM fatty acids, either all linoleic
acid (LA), all α-linolenic acid (ALA), or different ratios of LA to ALA (R2:1, R1:1, R1:2)
for 18 h. Some cultures were first pre-treated with these different ratios of fatty acids for
18 h, followed by co-exposure to 3.4 μM PCB77 for additional 24 h. PGE2 levels were
measured by enzyme immunoassay. Bars represent means ± SEM from three
independent experiments. Statistical comparisons of the two experimental settings
(without or with added PCB77) are indicated by small or capital letters.
86
Figure 4.13 Effect of NF-κB inhibitor on PCB77- or linoleic acid-induced VCAM-1
expression
Cells were exposed to 3.4 μM PCB77 or 20 μM linoleic acid (LA) alone for 6 h, or first
pre-enriched with 50 μM PDTC for 4 h followed by coexposure to PCB77 or LA for
additional 6 h. PCB77- or LA-induced VCAM-1 mRNA levels, as measured by real-time
RT-PCR. β-Actin was used as a housekeeping gene. Experiments were repeated three
times. Values are means ± SEM. Different letters represent significant difference among
treatment groups.
87
Figure 4.14 Effect of NF-κB inhibitor on PCB77- or linoleic acid-induced COX-2
expression
Cells were exposed to 3.4 μM PCB77 or 20 μM linoleic acid (LA) alone for 6 h, or first
pre-enriched with 50 μM PDTC for 4 h followed by coexposure to PCB77 or LA for
additional 6 h. PCB77- or LA-induced COX-2 mRNA levels, as measured by real-time
RT-PCR. β-Actin was used as a housekeeping gene. Experiments were repeated three
times. Values are means ± SEM. Different letters represent significant difference among
treatment groups.
88
Figure 4.15 Proposed mechanism for fatty acid-mediated modulation of endothelial cell
activation induced by PCB77
Omega-6 or omega-3 fatty acids can either enhance or reduce PCB77-induced up-
regulation of oxidative stress and activation of NF-κB. Relative activation of NF-κB will
further regulate VCAM-1, as well as COX-2 enzyme activity and subsequent PGE2
release from endothelial cells.
89
Discussion
Exposure to environmental toxicants such as persistent organic pollutants can
significantly compromise heath, and there is evidence that PCBs are proatherogenic. In
fact, epidemiological studies with humans demonstrate a link between cardiovascular
diseases and exposure to environmental pollutants. For example, an increase in
hospitalization rates was reported for coronary heart disease in populations residing near
areas contaminated with persistent organic pollutants [196]. Endothelial cells which line
the inner layer of blood vessels are an important cell type involved in the regulation of
metabolic events associated with the pathology of atherosclerosis. Dysfunction of the
vascular endothelium is considered to be a critical underlying cause of the initiation of
cardiovascular diseases such as atherosclerosis [6]. Environmental toxicants, once
absorbed, distribute themselves to tissues, especially adipose, where they are in dynamic
equilibrium with the blood. Thus, risk factors of pollutants such as PCBs are chronic and
can continuously amplify pathologies of diseases that are associated with endothelial
dysfunction. Data from our present study confirm that endothelial exposure to PCB77
provides a prooxidative cellular environment, sufficient to induce oxidative stress-
sensitive transcription factors such as NF-κB and associated inflammatory events
characteristic of early events in the pathology of atherosclerosis.
The diet is a major route of exposure to environmental toxic pollutants, such as
persistent organic pollutants, including PCBs. Since many of these pollutants are fat
soluble, fatty foods are not only unhealthy risk factors by themselves, but they usually
contain higher levels of persistent organics than vegetable matter [109]. Thus, high-fat
foods may pose as multiple risk factors in contributing to diet-derived proatherogenic
lipids as well as environmental toxicants. Since omega-6 rich oils are prooxidative and
can amplify PCB-mediated dysfunction of endothelial cells [106], and since omega-3 rich
oils exhibit cardio-protective properties [126, 130], we were interested in comparing the
effect of linoleic acid (the parent omega-6 fatty acid) and α-linolenic acid (the parent
omega-3 fatty acid) on modulating PCB-mediated endothelial inflammatory events.
Linoleic acid is the major fatty acid in common vegetable oils, and current estimates
indicate that over 90% of the omega-3 consumed by U.S. citizens is in the form of α-
90
linolenic acid, not the longer chain omega-3 fatty acids found in fish oils [129].
Nevertheless, many types of fish which are an excellent source of long-chain omega-3
fatty acids are often contaminated with persistent organic pollutants such as PCBs.
Whether the cardio-protective omega-3 fatty acids in fish neutralize or markedly decrease
the potential health risks associated with exposure to persistent organic pollutants is not
known. There appears to be some consensus that regular fish consumption should be
encouraged for its overwhelming cardio-protective properties [212, 213].
Using an established cell culture model, our data clearly demonstrate that
increasing the relative amount of α-linolenic acid (ALA) over linoleic acid (LA) protects
against PCB-mediated endothelial activation. We have previously observed that linoleic
acid can amplify PCB-mediated dysfunction of vascular endothelial cells [106]. We also
have demonstrated that antioxidants such as vitamin E can protect against PCB-mediated
endothelial cell activation [99]. There is increasing evidence that nutrition can dictate the
lipid milieu, oxidative stress and antioxidant status within cells, and thus modulate
mechanisms of cytotoxicity mediated by environmental pollutants [199]. Many
environmental pollutants induce signaling pathways that respond to oxidative stress and
lead to inflammatory events; and many of these same pathways are associated with the
etiology and early pathology of many chronic diseases [214].
Our data support the hypothesis that the lipid milieu within the vascular
endothelium can either upregulate or downregulate inflammatory events induced by
PCB77. We have shown previously that 18-carbon fatty acids differing in degree of
unsaturation selectively induce an inflammatory environment in human endothelial cells
[215]. Of the fatty acids studied, linoleic acid stimulated NF-κB transcriptional
activation the most. In addition, treatment with this fatty acid markedly enhanced mRNA
levels of TNF-α, monocyte chemoattractant protein 1 (MCP-1) and adhesion molecules
such as VCAM-1 [215]. In contrast, treatment with α-linolenic acid had minimal effects
on these inflammatory markers. Interestingly, in the current study, a relative increase in
α-linolenic acid over linoleic acid markedly reduced inflammatory markers induced by
exposure to PCB77.
Mechanisms of the lipid and PCB interactions on endothelial inflammatory events
are not clear, but our data suggest that NF-κB is a critical player in the overall regulation
91
of these events. We have previously demonstrated that PCB77-mediated activation of
NF-κB can be blocked by pretreatment with pyrrolidine dithiocarbamate (PDTC),
suggesting regulatory functions of PCB-induced endothelial cell activation through NF-
κB signaling [99]. In the present study, we demonstrated that inhibition of NF-κB
markedly reduced both PCB and linoleic acid -mediated endothelial inflammation. NF-
κB binds to and affects the function of several genes encoding proteins mediating
inflammation, and inhibition of NF-κB by PDTC resulted in reduced neutrophil
infiltration in lungs, liver and hearts of rabbits infused with lipopolysaccharide (LPS)
[216]. A recent study demonstrated that inhibition of NF-κB in aged vessels significantly
attenuated inflammatory gene expression and inhibited monocyte adhesiveness [217].
Protective mechanisms of omega-3 fatty acids on down-regulation of PCB-
induced vascular inflammation are not well understood. PCBs have been reported to
induce arachidonic acid release in human platelets [218], and in rat amnion fibroblast
cells [219] with a subsequent increase in eicosanoid formation. Our data suggest that
linoleic acid can enhance and α-linolenic acid inhibit these events. One inhibitory
mechanism may be a relative decrease in substrate availability for PGE2 formation during
cellular enrichment with α-linolenic acid [220]. Others have reported that
eicosapentaenoic acid can inhibit TNF-α induced endothelial cell inflammation via
PI3K/Akt signaling pathways [221], pathways which also can regulate NF-κB DNA
binding [105]. Omega-3 fatty acids also have been shown to inhibit NF-κB activation via
peroxisome proliferators-activated receptor alpha signaling [222].
A recent human study demonstrated that if intake of ω-3 fatty acid (such as ALA)
is sufficient, the ω-6 fatty acid intake can be increased without causing pro-inflammatory
response [223]. ALA is the most commonly consumed ω-3 fatty acid in Western diets.
Thus, strategies to increase ALA in the diet are important for the perspective of vascular
inflammation to reduce atherosclerosis.
In summary, the current study demonstrates that diet-derived lipids can modulate
PCB77 stimulated activation of endothelial cells by affecting oxidative stress sensitive
signaling pathways (Fig. 4.15), including activation of NF-κB and up-regulation of
inflammatory genes. Our data contribute to the paradigm that nutrition can modulate the
92
toxicity of environmental pollutants and thus affect health and disease outcome
associated with chemical insult [200]. Whether an increase in the consumption of
omega-3 polyunsaturated fatty acids can be used therapeutically against inflammation
that is mediated in part by environmental pollutants warrants further study.
Copyright © Lei Wang 2007
93
Chapter Five : Summary
Conclusion
Overall, the initiation of atherosclerosis by endothelial cell activation is mediated
by complex interactions between circulating nutrients, cytokines and environmental
pollutants. In the current work we studied the effects of different dietary omega-6 and
omega-3 fatty acids on cytokine and pollutant-induced endothelial cell activation. We
have previously shown that linoleic acid, the major omega-6 fatty acid in the American
diet potentiated both TNF-α and PCB-induced endothelial cell activation [106, 107, 156].
In contrast, omega-3 fatty acids from fish oil have been shown to have anti-atherogenic
effects. But the beneficial effect of α-linolenic acid which is actually the major omega-3
fatty acid in the diet has not been well studied yet. Considering these findings the
hypothesis was tested that the induction of endothelial cell activation may be modulated
differentially by linoleic or α-linolenic acids during both TNF-α and PCB77 stimulation.
Activation of endothelial cells and associated inflammatory events are critical
underlying causes of the initiation of cardiovascular diseases such as atherosclerosis [6].
The mechanisms by which selected fatty acids induce endothelial cell activation,
oxidative stress and inflammation are not fully understood. Oxidative stress-induced
transcription factors, which regulate inflammatory cytokine and adhesion molecule
production, are important regulatory elements in the induction of inflammatory responses.
One of these transcription factors, NF-κB, plays a significant role in these regulatory
processes [115]. Binding sites for NF-κB and related transcription factors were identified
in the promoter regions of a variety of inflammatory genes [116, 117].
Our data in chapters 2 and 4 indicate that both TNF-α and PCB77 can induce up-
regulation of oxidative stress, NF-κB activation and NF-κB-dependent induction of
VCAM-1, COX-2, and a subsequent increase in PGE2 production. All of these events
were amplified by pre-enrichment with linoleic acid but blocked or reduced by α-
linolenic acid. Prostanoids represent a group of lipid mediators. PGE2 is the most
common prostanoid which is generated at sites of inflammation [224]. The biosynthesis
of PGE2 involves three enzymatic reactions including phospholipase A2 (PLA2), COX,
94
and terminal PGE2 synthase. We demonstrated that both TNF-α and PCB77 can activate
cPLA2 at 8 hour, a similar time point when COX-2 up-regulation was observed. Both
cPLA2 and COX-2 have been shown to locate predominantly in the nuclear envelope
[225, 226], these permit efficient functional coupling between cPLA2 and downstream
enzyme COX-2. Furthermore, the time course studies indicated that after 24 hours
treatment, TNF-α and PCB77 induced significant PGE2 production. This later response
might partially due to the late activation of terminal PGE2 synthase, as shown by our real-
time PCR data that PGE2 synthase expression was significantly increased by PCB77 after
24 hours treatment.
In chapter 3, we further explored the role of caveolae in fatty acids mediated
TNF-α-induced inflammatory responses in endothelial cells. Because a growing body of
evidence indicates that dietary fatty acids can alter caveolae microenvironment, thereby
modifying location and function of proteins in caveolae. Previous study in endothelial
cell fatty acid composition of caveolae has shown that palmitic acid (16:0), palmitoleic
(16:1), stearic acid (18:0), together with oleic acid (18:1) compose the 80% of total fatty
acids, whereas linoleic acid (18:2) composes of 10% of total fatty acids in caveolae [71].
EPA has been shown to displaces caveolin-1 from caveolae in endothelial cells [72].
Moreover, caveolin-1, the functional protein of caveolae can bind to many types of
plasma membrane receptor proteins or membrane associated enzymes and concentrate
these molecules within caveolae [227]. TNFR-1 has also been shown to concentrate
within the caveolae of a human endothelial cell line [79].
Interestingly, our data indicated that functional caveolae are necessary for TNF-α-
mediated activation of endothelial cells. TNF-α markedly induced caveolin-1 and
TNFR-1, and both expressions were further induced when cells were first enriched with
linoleic acid. In contrast, α-linolenic acid diminished these effects. These data indicated
that different fatty acids may modulate both TNFR-1 and caveolin-1 expression, thus
altering the interaction of the two. These will in turn profoundly influence TNFR-1
dependent downstream signaling. Moreover, silencing caveolin-1 totally blocked TNF-
α-induced COX-2 up-regulation and PGE2 biosynthesis. Previous studies have shown
that COX-2 may be localized to caveolae-like structures [228]. COX-2 in human
foreskin fibroblasts stimulated by IL-1β was localized to plasma membrane in addition to
95
nuclear envelope, and image overlay showed colocalization of COX-2 with caveolin-1
[80]. These results indicated that caveolae are important cellular platform for TNF-α
receptor-mediated down stream COX-2 activity. In contrast, linoleic acid-induced COX-
2 up-regulation and PGE2 production were not altered by caveolin-1 silencing. Taken
together, our data indicated that lipid microdomains, and in particular caveolae, may be a
novel modality mediating the effects of linoleic and α-linolenic acids in response to TNF-
α-mediated inflammatory stimulations, but the inflammatory responses induced by
linoleic acid appear to be independent of caveolae protein. It is very likely that selected
fatty acids either stabilize or perturb caveolae functions, thus leading to modifications of
caveolae-dependent signaling.
There is evidence that fatty acids can alter localization and function of caveolae-
associated signaling proteins in mouse colonic mucosa [73]. These data suggests that
omega-3 fatty acids may in part be anti-inflammatory by decreasing caveolin-1
expression or by causing dysfunctional caveolae, leading to down-regulation of caveolin-
dependent down-stream signaling. Moreover, perturbation of membrane raft structural
integrity with cholesterol-sequestering compounds had been shown to cause
delocalization of eNOS from caveolae and to inhibit TNF-α-induced ROS production and
protein tyrosine nitration [229].
Furthermore, we recently investigated caveolin-1-regulated mechanisms
associated with PCB-induced markers of peroxynitrite formation and DNA binding of
NF-κB [230] and demonstrated that caveolin-1 silencing abolished the PCB-stimulated
NF-κB DNA binding, suggesting a regulatory role of caveolae in PCB-induced
endothelial cell activation. Our unpublished data have also shown that PCB treatment
increased formation of caveolae and PCB77 accumulated mainly in the caveolae-rich
fraction and AhR, the main molecular target of PCB77 was associated with caveolin-1,
and the association was increased after PCB77 treatment. Thus, the broad mediation
effects of fatty acids appear to be largely due to their role as structural components of
cellular membranes and are involved in many important cell-signaling pathways. This
suggests that a common fundamental mechanism might be involved in the inflammatory
cytokine TNF-α and environmental pollutant PCB77-triggered pro-inflammatory
cascades.
96
In summary (Fig. 5.1), we provide novel data demonstrating that omega-6 and
omega-3 fatty acids can differentially modulate TNF-α and PCB77-induced
inflammatory stimuli and that these events require functional caveolae. Furthermore,
functional changes of caveolae associated with modifications by dietary fatty acids
appear to affect critical phases of induction of oxidative stress-sensitive transcription
factors and inducible inflammatory parameters during endothelial cell activation.
Because caveolae and caveolins have been implicated in several human diseases and in
particular vascular diseases, our data may have implications in understanding novel
mechanisms of inflammatory diseases modulated by dietary lipids. Furthermore,
research findings present in this dissertation contribute to the paradigm that nutrition can
modulate the toxicity of environmental pollutants and thus affect health and disease
outcome associated with chemical insult [200]. Whether an increase in the consumption
of omega-3 polyunsaturated fatty acids can be used therapeutically against inflammation
that is mediated in part by environmental pollutants warrants further study.
Future Directions
The current study demonstrated the critical role of functional caveolae in fatty
acid-mediated endothelial activation. Since caveolae contain sphingomyelin and initiate
the conversion of sphingomyelin to ceramide which is a novel sphingomyelin-derived
second messenger mediating cellular signals, further studies are needed to clarify the
effects of fatty acids on caveolae composition and to explore further their role in
ceramide metabolism (Fig. 5.2).
A lipid raft membrane is composed of sphingolipids and cholesterol in the
exoplasmic leaflet, linked to glycerophospholipids and cholesterol in the cytoplasmic
leaflet of the lipid bilayer [231]. It is possible that long chain fatty acid substituents of
sphingolipids extend from the outer to the inner leaflet where they interdigitate with the
saturated side chains of glycerophospholipids and establish a close association [232]. In
caveolin-1 knockout mice, the transport of glycosylphosphatidylinositol-anchored
proteins to the cell surface was inefficient, consistent with a role for caveolin in the
transport of these lipid raft-associated proteins to the plasma membrane [233]. Caveolin-
97
1 null mice showed a dramatic increase in serum triacylglycerols and non-ester fatty acid
levels [227] indicated that caveolae might have a primary role in regulating the lipid
balance by stimulating fatty acid uptake.
Lipids are crucial structural and functional components of cells. Even more
importantly, many lipid species have distinct cellular functions. For example, ceramides,
eicosanoids, diacylglycerol (DAG) and lysolipids are all second messengers which
participate in various cellular events such as inflammation, growth, proliferation,
differentiation and cell death. Sphingomyelin is one of the major lipids in cell
membranes and plasma lipoproteins. In clinical studies, sphingomyelin plasma levels
have been shown to correlate with the occurrence of coronary heart disease independent
of plasma cholesterol levels [234]. Previous studies indicated that the endothelium
contributes to ceramide-induced relaxation possibly through an interaction between
sphingomyelin hydrolysis and endothelial NO synthase within caveolae [235].
Furthermore, TNF-α has also been shown to profoundly affect sphingolipid metabolic
pathways leading to the accumulation of one or more sphingolipids. For example, TNF-
α transiently increases cellular ceramide by increasing the hydrolysis of sphingomyelin.
Increased ceramide levels activate ERK [236, 237], JNK [237], MAPKs and NF-κB
translocation [238]. Sphingomyelin and ceramide both contain one fatty acyl moiety that
is linked to the sphingosine backbone by an amide bond. Whether the type of this fatty
acyl moiety can be altered by dietary fatty acids in endothelial cells has not yet been
studied. In our study the different polyunsaturated fatty acid-induced modification in
response to TNF-α and/or PCB may be due to the changes of the microdomain
polarization within sphingolipids modified by dietary fatty acids. However, the effects of
PCB on ceramide signaling pathways have not been explored yet.
Thus, future studies of a more detailed involvement of caveolae in the cellular
regulation of fatty acid-mediated modulation of a vascular inflammatory response may be
considered.
98
Figure 5.1 Summery
Figure 5.2 Future direction
Copyright © Lei Wang 2007
99
Appendix: Protocols
Primary Culture of Porcine Pulmonary Artery Endothelial Cells
Obtain tissue: A. Supplies:
1. Sterile: tweezers, scissors, razor blades, sponge gauze, specimen containers, gloves, Other supplies: 3 beakers (250 ml);
2. Media and solutions: M199+3xP/S (warm up to 37℃), 70% ethanol B. Obtain tissue:
1. Take the heart of porcine during routine slaughter at the College of Agriculture. Find pulmonary artery which is next to aorta. Gently separate aorta and pulmonary artery using the other side of scissors. Try not to squeeze vessels too hard.
2. After separation, use intestinal forceps to clamp artery at the end nearest the heart. Cut the vessel off at the edge of clamp, then with the clamps still on, dip artery in the 70% ethanol solution and then dip into M199+3xP/S.
3. Take off the clamps, put artery into a sterile specimen cup with enough M199+3xP/S to cover the whole artery. Label the container, and place on ice carry back to the lab and put in 4 ℃.
Culture cells (should be done completely inside the hood): A. Supplies
1. Sterile: culture dishes, tweezers, allis intestinal forceps, sponge gauze, centrifuge tubes, scapel, scissors, plugged Pasteur piptes, plastic transfer pipets, measuring pipets, cotton swabs with wooden stick, T25 tissue culture flasks.
2. Media and solutions: M199+3xP/S and M199+10% FBS (both warmed up to 37℃). Fill two 250 ml beakers with 70% ethanol and one 250 ml beaker with M199+3xP/S.
3. Collagenase solution: will need 2 ml for each artery. Add 1 mg of collagenase per ml of Hanks buffer (warm up to 37℃). Vortex the solution and centrifuge at 2500 rpm for 10 min. Filter supernatant using a syringe filter unit (0.22 μM).
B. Isolation of endothelail cells 1. Take artery from specimen container and place on Hanks moisturized sterile
sponge gauze (handle artery with steriled tweezers, don’t touch it with hands). Use a Pasteur pipet to rinse the inside of the artery with M199 to get off blood clots and blood cells and avoid touching the inside of the artery. Then use a forcep and tweezers to clamp one end of the artery. Use a pasteur pipet fill from the open end of the artery with collagenase solution and clamp it up with another forcep. Wait for 5 min.
2. During the waiting time, add 2 ml of M199+10% FBS into each of two 30 mm tissue culture dishes (swab twice for each artery), and label with cell line, date, and number of swab (1 or 2). When the 5 min are over, release forceps and dry collagenase solution into sterile sponges. Rinse inside of artery with a pasteur pipet full of M199+10%FBS and place artery on a dry, sterile gauze pad. Cut
100
artery open length wise with a pair of sharp sterile scissors, avoid touching other area of the artery besides the cutting line.
3. Wet a sterile cotton swab in the media from the 30 mm tissue culture dish labeled swab1. Very gently rub the swab over the inside surface of the artery. Do not put any pressure on the swab and avoid swabbing where the forceps clamped or over the same place twice or at the edge of the artery. Make about two rounds swabbing and then stir the swab around in the media and press it against the sides to release the endothelial cells into the media. Repeat this with a new swab and put into the dish labeled with swab 2.
4. Place culture dishes into the incubator. 4 h later, use the aspirator to remove the media and replace it with 2 ml of new 37℃ M199+10%FBS media. Change media again at 24 or 48 h after this depends on how many cells are attached.
5. 4 or 5 days later, when the cells get confluent, pass one 30 mm dish into one T25 tissue culture flask (passage 1). When it gets confluent, pass one T25 into one T75 flask (passage 2). Later pass one T75 into one T225 or three T75 (passage 3). Usually freeze cells at passage 4 or 5. Isolated primary endothelial cells can be used up to 15 passages.
Freeze Endothelial Cells
One confluent T-75 flask can be frozen into 2 cryotubes. One T-225 flask can be frozen into 6 cryotubes.
1. For one T-75 flask: rinse cells with 10ml hanks then add 1 ml trypsin. Can do two T-75 at one time. When cells are detached, add 10ml 10% M199 and transfer into a 15ml conical tubes.
2. For one T-225 : Trypsinized cells with 1.5 ml trypsin. add 30 ml 10% M199 and transfer into three 15 ml conical tubes.
3. Centrifuge at 1200rpm for 10 min (at 37℃). Decant supernatant (gently) and add 2 ml freezing cocktail* into each conical tube. Gently mix the cells with freezing cocktail. Transfer to 2 cryotubes with 1 ml each (tubes should be properly labeled with passage, date and operator).
4. Keep tubes between two foam layers in -20℃ overnight or in “Slow Freeze Apparatus (isopropano jacketed container)” for 24 h, then put in -80℃. After at least 18 h in -80℃, put cryotubes in liquid nitrogen tank. *Freezing cocktail must be made fresh each time: Every T-75 flask needs 2 ml of freezing cocktail, prepare a little extra (plus 2ml). Warm up everything to 37 C before using. 1ml For 8 T75
prepare 18 ml For 9 T75
prepare 20 ml For 18 T75
prepare 38 ml 28% DMSO (ml) 0.25 4.5 5 9.5
FBS (ml) 0.2 3.6 4 7.6 M199 (ml) 0.55 9.9 11 20.9
Preparation of 28% DMSO in M199: 1 ml 4.5 ml 5 ml 10 ml DMSO (ml) 0.28 1.26 1.4 2.8 M199 (ml) 0.72 2.24 3.6 7.2
101
Start a Frozen Endothelial Cell Line
1. Warm up M199+10%FBS media to 37℃and set centrifuge to 37 ℃ in ahead. (To achieve the temperature quicker, can set the temperature to 40℃, speed to 3000 rpm, and let run for 10 min)
2. Take one cryotube of frozen cells from liquid N2 tank or -80℃ freezer at one time. 3. Add 9 ml media into a 15 ml conical tube for one cryotube. Use a 5 ml pipet get 1 ml
media from the conical tube put it quickly up and down into cryotube to thaw the 1 ml freezing cocktail with endothelial cells. Put the resolved part into the conical tube and get some fresh and warm media from the same tube, up and down again until get all the liquid out from the cryotube.
4. Centrifuge at 2000 rpm for 5 min immediately. Don’t leave cells with freezing cocktail at room temperature for too long. Then use the aspirator to remove the media and freezing cocktail, put 1 ml of fresh media and gently scatter the pellet of cells. Add 9 ml of media to conical tube and centrifuge for 5 min again. Remove the media after centrifuge, repeat the scatter and mix cells with 10 ml of media. Place the cell suspension into one T75 cell culture flask. Change media 4 h after revival or change media in the following morning depends on how many cells attached.
Cell Culture and Experimental Media
A. Cell culture Pass the cells:
1. It will usually take two days for cells to grow confluent. When cells are ready, use the aspirator to remove old media. Add 6 ml of Hanks for a T75 flask and 10 ml for T225. Gentely tilt the flask to rinse all the cells. Remove Hanks buffer 1 after 1 min incubation at room temperature.
2. Add trypsin (1 ml for the T75 and 2 ml for the T225 flask). Place in the incubator for 2 to 3 minutes. Then strike the side of flask to detach the cells and check under the microscope to see whether all cells are detached as little round circle. .
3. While cells are in the trypsin incubating, add medium to new flasks (7 ml for the T75 and 20 for the T225), and label with “P#” for the passage, and date.
4. After the cells are detached gentlely add 12 ml of medium (for T75) or 30ml of medium (for T225), up and down avoid air bubble. Usualy one T75 can be passed into 3 T75 or one T225. One T225 can be passed into 3 T225. Add 4 ml (for T75) or 10 ml (for T225) of the re-suspended cells to each new flask.
5. Place flasks in the incubator. Plating the cells:
1. Ratio of cell suspension and media for plates and flasks 100 mm culture dishes can hold up a total volume of 10 ml (3 ml of the cell suspension plus 7 ml of medium) 60 mm dishes can hold a total volume of 3 ml (1 ml of the cell suspension plus 2 ml of medium). After plating, it usually takes about 48 h for cells to get confluent.
102
T-75 flask can be passed into three T-75, one T-225, four 100 mm plates, or twelve 60 mm plates. T-225 flask can be passed into three T-225, twelve 100 mm plates, or thirty six 60 mm plates. 2. Plating cells on multiple well plates No matter the multiple-well plate has how many wells, the whole plate usually can hold up to 12 ml of total media volume. This will need 4 ml of the cell suspension plus 8 ml of media. Usually use T-75 flask to plate. Always make the 12 ml of the cell and medium mixture first before divided into each well. For a 6 well plate: 12 divided by 6, so put 2 ml of the mixture into each well. For a 12 well plate: put 1 ml into each well. For a 24 well plate: put 0.5 ml et al.
B. Preparation of Media and Hanks 1. M199+AA:
Add 2 pouches of the media (M199, Gibco Cat No 31100-035) Add 4.4g of NaHCO Add 2 L of autoclaved (dI H2O) Add 2ml of NaOH (6N same as 6M) Add 40 ml of amino acids (premade solution from Gibco) Mix well and adjust pH to 7.4
2. Hanks KCl: 0.8g KH2PO4: 0.12g NaCl: 16g NaHCO3: 0.7g Na2HPO4 * 7H2O: 0.18g D glucose: 2.0g Phenol Red: 0.02g Add 2 L of autoclaved H2O adjust pH to 7.4 and then filter.
3. Trypsin Remove 40ml of Hanks (new un-used bottle that has 400 ml total) Add 40 ml of trypsin to the remaining 360 ml of Hanks. Swirl and make 12 ml aliquots
4. 10 % or 5% FBS M199 For 10% media: Take out 52 ml of the M199 media from 400ml into another bottle and add 4ml of P/S (penicillin streptomycin, Cat No 15140-122), 4 ml of L-glutamine (Cat No 25030-156), 4 ml of MEM amino acids (GIBCO Cat. No. 11130-051), and 40 ml of FBS (fetal bovine serum). For 5% media: Take out 32 ml of the M199 media and add 4ml of P/S (penicillin streptomycin, Cat No 15140-122), 4 ml of l-glutamine (Cat No 25030-156), 4ml of MEM amino acids (GIBCO Cat. No. 11130-051) and 20 ml of FBS (fetal bovine serum)
Preparation of Fatty Acid Enriched Media
A. Preparation of fatty acids stock solution
103
The pure fatty acids are purchased from NuCheck (http://www.nu-chekprep.com /home.htm). The stock solution is made in Hexane.
1. Get fatty acid from the commercial container (100mg). Record the weight of fatty acid and the molecular weight. Make a 5 ml solution in graduated flask and then calculate the final concentration: N1 X mg × 1000 5 ml × MW
Result is in unit as mM. 2. Fill the graduated flask with gas N2 and carefully sealed with parafilm.
B. Fatty acids Media preparation
Fatty acid will be prepared in M199 enriched with 5% (v/v) FBS. Assuming the 100% serum has an average concentration of albumin at 600 μM, then the 5% FBS will give us up to 30 μM albumin. The ratio of free fatty acids to albumin should not exceed 3 to 1, otherwise fatty acid is toxic to cells. So, for a 5% FBS media, the highest fatty acid concentration can achieve is around 90 μM. 1. For 10ml plate, if the final conc. of fatty acid is 20 μM (0.02 mM), we need to
calculate how much fatty acid solution in hexane we will need. V1 Set 1 as stock solution, 2 as experimental solution: V1N1=V2N2 X×N1 =10×0.02 10×0.02 X= N1
X is in the unit of μl 2. For 12 100 ml plates, we will need 120ml media. Always prepare extra 5 ml,
because we will need to filter the media and later which can cause the lost of some volume.
After finishing the calculation, take a glass tube put NaOH 30 times exceed of FA concentration. Volume of 6 N NaOH: V3 FA conc.×30× V2 6
Put NaOH (V3) in a culture glass tube, then add FA (V1). Carefully blow in the gas N2 for 15 min to 30 min until it totally dried (white powder). 3. Wash out FA from glass tubes into the media (V2).
Adjust pH of the media to 7.4 with HCl. 4. Filtration through the syringe filter or bottle filter (0.02 μm) then add to the cells.
Oxidative Stress Measurement
1. Grow cells in microplates (12, 24 or 48) well plates. Include a control groups to be used as negative and positive controls. 2. Wash cells at least two or three times with Hanks buffer to remove serum. 3. Add KREBS-Ringer-Glucose buffer containing DCF (100 μmol/L) to microplates. Use 1ml for 12 wells and 0.5 ml for 24 wells and incubate for 30 minutes. Blank well
104
receives KREBS-Ringer-Glucose buffer without DCF probe. Wrap plate in aluminum foil to protect from the light. 4. Wash 2 to 3 times with HEPES buffer. 5. Add HEPES to each well (1 ml for 12 well plates and 0.5 ml for 24 well plates. Wrap plate in aluminum foil. Optional: Positive control receives 0.1 mmol/L H2O2 in HEPES buffer after DCF staining. 6. Set plate reader excitation to λ 490 nm and emission to λ 520 and/or 530 nm. Light sensitive reactions: protect stocks from light by wrapping in aluminum foil when adding DCF media on cells, switch off the light in the hood. Buffers and solutions: HEPES buffer (500ml pH 7.4):
NaCl: 4.237g KCl: 186.4mg MgCl2: 50.8mg D-glucose: 901.0mg HEPES: 1.1915g CaCl2: 110.25mg
KREBS-Ringer-Glucose Buffer: Components To make 1 L To make 500 ml
118 mM NaCl 6.9 g 3.448 g 4.7 mM KCl 0.35 g 0.175 g
1.3 mM CaCl2 0.191 g 0.096 g 12.0 mM MgCl2 2.375 g 1.187 g
1 mM NaH2PO4H2O 0.138 g 0.069 g 25 mM NaHCO3 2.1 g 1.050 g 11mM Glucose 1.982 g 0.991 g
DCF stock solution: To make the DCF stock solution mix 9.7458 mg of DCF with 1 ml of DMSO. The DCF concentration in the stock is 20 mM. Note on DCF stock: 20mM in EtOH or DMSO, MW of DCF 487.29g. To make the DCF working solution, add 125 μl of the stock and complete volume to 25 ml with KREBS buffer. The DCF concentration in the working solution should be 100 μM.
Fluorescent plate reader settings: Softmax Pro 3.1.2 Set up wavelength: Sensitivity: 15 Auto Calibrate: on Strip Template: set blank at 0, or set all samples.
PGE2 EIA kit
Plate cells:
105
Small plates will be used, so that the protein level of COX-2 and mPGE synthase can be measured.
Supernatants of cell cultures will be mixed well then collected 1 ml directly into Eppendorf’s tubes. Store at -80 C. In the day of measurement, thaw the samples, mix well then centri at 4C 12000rpm for 10 min. Using the Kit:
Condition preparation: for the first time to use the kit. Make standard curve dilution in 5% medium (the four highest ones). And one control and one TNF-α treatment sample to check whether it’s within the range of standard curve. Key terms:
Blank: background absorbance caused by Ellman’s reagent. Total activity: total enzymatic activity of the AchE-linked tracer. Non-specific binding (NSB): non-immunological binding of the tracer to the well. Maximum binding (B0): maximum amount of the tracer that the antibody can bind in the absence of free analyte. % bound/ maximum bound (%B/B0): ratio of the absorbance of a particular sample or standard well to that of the maximum binding. Standard curve: a plot of the %B/B0 values versus concentration of a series of wells containing various known amounts of analyte.
A. Pre-assay preparation: Buffer preparation (all stored at 4C, will be stable for ~2 months) 1. EIA buffer: dilute one vial of EIA buffer (vial #4) with 90 ml of Ultrapure water.
Rinse the vial well. 2. Wash buffer: dilute 5 ml vial of Wash buffer (vial #5) to a total volume of 2 liters
with Ultrapure water and add 1 ml of Tween 20 (vial #5a) use a syringe.
Wash buffer (vial #5) Ultrapure water (1:400 of wash buffer)
Tween 20 (vial #5a) (0.5ml/liter wash buffer)
5ml 2 L 1ml 12.5ml 5 L 2.5ml
Collecting samples
Cell culture supernatants may be assayed directly. If the PGE2 concentration in the medium is high enough to dilute the sample 10-fold with EIA buffer, the assay can be performed without any modifications. When assaying less concentrated samples (where samples cannot be diluted with EIA buffer), dilute the standard curve in the same culture medium as that used in the experiment. This will ensure that the matrix for the standards is comparable to the samples. A standard curve is recommended to be run first to ensure that the assay will perform in a particular medium.
B. Preparation of assay-specific reagents 1. PGE2 standard
106
Reconstitute the contents of the PGE2 standard with 1 ml of EIA buffer. The concentration of this solution (the bulk standard) will be 10 ng/ml. Stored at 4C; this standard will be stable for up to 4 weeks. NOTE: if assaying culture medium samples that have not been diluted with EIA buffer, culture medium should be used in place of EIA buffer for dilution of the standard curve. The diluted standards should not be stored for more than 24 hr.
2. PGE2 AChE tracer: Will be used twice in the two days measurement. Always keep the rest.
Reconstitute the 100 dtn PGE2 tracer (vial #2) with 6 ml EIA buffer. Store the reconstituted PGE2 tracer at 4C (DO NOT FREEZE!) and use within 2 weeks.
Tracer Dye Instructions: this dye may be added to the tracer, if desired, to aid in visualization of tracer-containing wells. Add the dye to the reconstituted tracer at a final dilution of 1:100 (add 60 μl of dye to 6 ml tracer or add 300 μl of dye to 30 ml of tracer).
3. PGE2 monoclonal antibody: Reconstitute the 100 dtn PGE2 antibody (vial #1) with 6 ml EIA buffer. Store the
reconstituted PGE2 antibody at 4 C. it will be stable for at least 4 weeks. Antiserum dye instructions: this dye may be added to the antiserum, if desired, to aid
in visualization of antiserum-containing wells. Add the dye to the reconstituted antiserum at a final dilution of 1:100 (add 60 μl of dye to 6 ml antiserum).
C. Performing the assay
1. Plate set up It is not necessary to rinse the plate prior to adding the reagents. If strip plate, place
the unused strips back in the plate packet and store at 2-4C. Be sure the packet is sealed with the desiccant inside.
Each set of strips must contain a minimum of two blanks (Blk), two non-specific
binding wells (NSB), two maximum binding wells (B0), and an eight point standard curve run in duplicate. Each sample should be assayed at two dilutions and each dilution should be assayed in duplicate. For statistical purposes, we recommend assaying samples in triplicate.
2. Pipet the reagents: Before pipetting each reagent, equilibrate the pipet tip in that reagent (slowly fill the
tip and gently expel the contents, repeat several times). EIA buffer:
Add 100 μl EIA buffer to Non-specific binding (NSB) wells. Add 50 μl EIA buffer to Maximum binding (B0) wells. Or: If culture medium was used to dilute the standard curve, substitute 50 μl of
culture medium for EIA buffer in the NSB and B0 wells: Add 50 μl culture medium and 50 μl of EIA buffer to NSB wells. Add 50 μl culture medium to B0 wells.
PGE2 standard: Add 50 μl from tube #8 to both of the lowest standard wells (S8). Add 50 μl from tube #7 to each of the next two standard wells (S7). Continue with this procedure until all the standards are aliquoted.
107
The same pipet tip should be used to aliquot all the standards. Before pipetting each standard, be sure to equilibrate the pipet tip in that standard. Samples:
Add 50 μl of sample per well. Each sample should be assayed at a minimum of two dilutions. Each dilution should be assayed in duplicate (triplicate recommended). PGE2 AChE tracer:
Add 50 μl to each well except the total activity (TA) and the blank (Blk) wells. PGE2 Monoclonal antibody:
Add 50 μl to each well except the Total Activity (TA), the Non-Specific Binding (NSB), and the Blank (Blk) wells. Incubate the plate:
Cover each plate with plastic film and incubate 18 hr at 4C. Develop the plate:
When ready to develop the plate, reconstitute one 100 dtn vial of Ellman’s Reagent (vial #8) with 20 ml of UltraPure water. Reconstitute Ellman’s Reagent is unstable and should be used the same day it is prepared; protect the Ellman’s Reagent from light when not in use.
Empty the wells and rinse five times with Wash Buffer. Add 200 ml of Ellman’s Reagent to each well and 5 ml of tracer to the Total Activity wells. Cover the plate with plastic film.
Optimum development is obtained by using an orbital shaker equipped with a large, flat cover to allow the plates to develop in the dark. This assay typically develops (i.e., B0 well ≥ 0.3 A.U.) in 60-90 minutes. Read the plate:
Read the plate at a wavelength between 405-420 nm (405 was used). Before reading, wipe the bottom of the plate with a clean tissue to remove finger prints. The plate may be checked periodically until the B0 wells have reached a minimum of 0.3 A.U. the plate should be read when the absorbance of the B0 wells is n the range of 0.3-1.0 A.U. If the absorbance of the wells exceeds 1.5, wash the plate, add fresh Ellman’s Reagent and let it develop again.
Immunofluorescence
Plate cells (100μl cells plus 300μl Medium)onto glass chamber slides leaving room for 3 extra controls (IgG, no 1°, no 1° or 2°), grow to confluence, serum starve overnight. Then start the treatment. 1) Remove media from wells. 2) Depends on antigen of interest, could fix with:
a). 500 μL of 50:50 acetone/methanol solution for 30 minutes at RT; b). 500 μL of methanol for 15 min at RT; c). 500 μL of 4% paraformaldehyde in PBS for 1hr. Would need to quench with 0.3M/ (made a 0.2M) glycine for 10 min. During the fixation, turn on the heater to 37C on the shaker.
3) Wash with 1x PBS 3x for 5 minutes at 37°C shaking.
108
4) Depending on antigen, permealize with 0.2% Triton X 100 in 1x PBS for 15 min at 37°C shaking (could use less if unsure 0.01%)
5) Wash with 1x PBS 3x for 5 minutes at 37°C shaking 6) At this step you can put the slide in the fridge, wrapped in parafilm, before going on. The second day: 7) Heat slide back up if stopped. 8) Block with 500 μL of a solution of 15 μL/mL of the serum of the secondary in 1%
BSA (0.1g BSA in 9.9ml 1x PBS) in 1x PBS for 45 minutes at 37°C shaking.
Protein of interest Secondary Ab Serum of the 2nd Ab Dye (color) Cav-1 (mouse) Donkey anti-mouse Donkey (7.5μL in
500μL/well) Green
TNF-receptor 1 (goat) Donkey anti-goat Donkey (7.5μL in 500μL/well)
Orange
9) Without washing, put the respective primaries on the indicated wells diluted in 1x
PBS. Add corresponding IgG to the control well allotted at the same concentration as your primary (ex. 5 μg/mL) for 30 minutes at 37°C shaking.
Protein of interest Primary Ab conc. (dilution in 1x PBS) Cav-1 (mouse) Mouse-200μg/ml.
(need a 40x dilution: 1.25μl into 500μl PBS) TNF-receptor 1 (goat) Goat-100μg/ml.
(need a 20x dilution: 25μl into 500μl PBS) 10) Wash the wells 2x in 1x PBS, then an additional 1x for 5 minutes at 37°C shaking 11) Avoiding light, add the secondary antibody (ex. 1/200) diluted in 1x PBS for 15
minutes at 37°C shaking covered in aluminum foil.
Protein of interest 2nd Ab conc. (dilution in 1x PBS) Dye
(color)
Cav-1 (mouse) Alexa Fluor 488 Donkey anti-mouse: 2mg/ml (need a 1/200 dilution: 2.5μl into 500μl PBS)
Green
TNFRI (goat) AlexaFluor 546 donkey anti-goat: 2mg/ml (need a 1/200 dilution: 2.5μl into 500μl PBS)
Orange
12) Wash the well 2x in 1x PBS, and then an additional 1x for 5 minutes at 37°C shaking 13) Add Hoechst stain at a concentration of 1 μg/mL (add 0.5μl into 500μl from the
1mg/ml Hoechst solution) for 5-10 minutes at 37°C shaking. 14) Wash the wells 3x in 1x PBS, then pull off the sides of the wells. Mount with one
drop of VectaMount and covered with large cover slides avoid bubbles. Use nail polish to seal the sides. Store at 4°C in foil.
109
Confocal Microscopy
A. Turn on the machine: From the right side to the 1eft side and then right again, last is the Middle. 1. Right side: Burner 2. Left: Laser black switch and turn the key, then turn on the Green and Red lasers. 3. Right: LD laser 4. Middle: Up LG-PS2 Middle: BX-UCB Lower: FV5 Let the machine warm up for 10 min before start experiment. Only when the FV5-PSU on, the laser can be turn on. FV5-LDPSU has a safety knob
switch, need to be on all the time. B. Adjust the focus of samples and check the dye by switch light path 1 through 5 by
microscope. Our system is having 2 channels and 4 filters. Switch the filter on the left side of the machine. If use 60X, must use a drop of oil between the projector and the slide. Use the up baton on the right lower side of the microscope to make the length at the proper position. When check different dye, use the respective filter. Shut off the shutter after each check in order to avoid the quencher of the slide.
C. After ready for taking pictures, switch the light path to 1 (clear light). And pull out
the camera light path on the right side of microscope.
First clear the dye selection then add preferred dyes on the middle of the screen. APPLY The lower mirror set to an empty one. Active the laser filters on the left lower part of the screen. 1. DAPI: blue color. Dark pink laser. DAPI is Hoechst, Blue color. Channel 1, A in B out. (LD405) 2. FITC: green. Blue laser. FITC is green. (AR488) 3. TEX red. On the left side of the channel and filter switch, a black bar is for the setting of how much light goes to each channel. For DAPI set the black bar to the middle position and for FITC, pull the black bar all the way out. There is a round bigger turning, can adjust the signal. Try the ONCE scanning first to see how the picture is. To clean up the image, always keep the Gain low at 1 and increase the PMT (voltage) to get stronger signals. Don’t increase the laser adjusting on the left lower part of the screen, because increase the density of laser is bad for the sample.
110
Can also try the XYREPEAT and adjust the focus at the same time to choose better image. The type of scanning: I. Normal, can give only one picture each time. II. Sequence: can add channel, and do a continuous scanning with different dye settings. Optical Syst. Group 4. Add channel, click on optical system, add DPSI or FITC. Click on the icon: --Δ Δ--- which is “scan unit configuration”. Set FITC (green) and Mito (red) to group I, and DAPI to group II. Sequence: once. The three-color bar can be used to change the image from different channel. One group can include 2 channels.
Detergent-free Purification of Caveolin Enriched Membrane Fractios (Sucrose Gradient)
1. Cells were plated in 4 100 mm plate/treatment. Cells were washed with 1x PBS and lysed with 2 ml of ice-cold MES-buffered saline (MBS; 25mM MES (morpholineethanesulfonic acid, pH 6.5) 150 mM NaCl) containing 500 mM Na2CO3. For the preparation: 25 mM MES: 2.4405 g in 500 ml water. Adjust PH to 6.5. Then add 4.383g NaCl into MES, achieve MBS. Take 50 ml MBS, add 2.65g Na2CO3, get 500mM. this is lysis buffer. 90% Sucrose: 45g sucrose and make final volume 50 ml. Heat in water bath to dissolve. 35% Sucrose: 17.5 g sucrose and final volume 50 ml. 5% sucrose: 2.5 g sucrose and final volume 50 ml.
2. Homogenation was carried out with 10 strokes of a loose-fitting Dounce homogenizer (7ml). After homogenized, get samples into 15ml cornival tubes for sonication: 30 sec. for 10 times on ice. The setting is 4 and 60. The frequency should be under 20.
3. After sonication, centrifuge at 3000 rpm for 10 min at 4C. 4. Collect superant into centrifuge tubes (14x89mm), if its not exactly 2 ml, add more
MBS to reach 2 ml. 5. The homogenate was adjusted to 45% sucrose by addition of 2 ml of 50ml 90%
sucrose prepared in MBS and placed at the bottom of an ultracentrifuge tube, and overlaid with a discontinuous sucrose gradient Add 2 ml 90% sucrose to make the final 45% sucrose (cut off the top of the tip), add into tube slowly against walls, then vortex to mix well.
6. Add 5 ml of 35% sucrose. Use 1 ml peppit five times. The first 1 and 2 ml must be very slow, add drop by drop to avoid interrupt layers. After the 3rd ml, can be a little faster.
7. Add 3 ml of 5% sucrose. Then the tube is pretty full.
111
8. Measure weight of the 2 or 4 tubes (depends on sample number), put the one with more volume first on the scale, set to 0. then put the less volume ones. Achieve 0 with 5% sucrose. At least the 3 digital.
9. Get the rotor from 4C. Centrifuge 39000 rpm for 16 h. 10. The second day morning, collect fractions into 1.5ml clear tubes. Label 1 to 12. can
see a white fraction which is Cav-1 enriched faction. Sometime the 12th fraction has a little bit less or more volume.
11. Run the gel. Jin: 16 μl + 4 μl 5x loading buffer. Lei: 15 μl + 3 μl 6x loading buffer. Run the gel for 12 fractions on each western blot gel.
112
References:
1. Springer, T.A., Adhesion receptors of the immune system. Nature, 1990. 346(6283): p. 425-34.
2. Libby, P., M. Aikawa, and M.K. Jain, Vascular endothelium and atherosclerosis. Handb Exp Pharmacol, 2006(176 Pt 2): p. 285-306.
3. Schiffrin, E.L., Beyond blood pressure: the endothelium and atherosclerosis progression. Am J Hypertens, 2002. 15(10 Pt 2): p. 115S-122S.
4. Blum, A. and H.I. Miller, The role of inflammation in atherosclerosis. Isr J Med Sci, 1996. 32(11): p. 1059-65.
5. Cheung, B.M., The cardiovascular continuum in Asia--a new paradigm for the metabolic syndrome. J Cardiovasc Pharmacol, 2005. 46(2): p. 125-9.
6. Ross, R., Atherosclerosis--an inflammatory disease. N Engl J Med, 1999. 340(2): p. 115-26.
7. Meerarani, P., et al., Metabolic syndrome and diabetic atherothrombosis: implications in vascular complications. Curr Mol Med, 2006. 6(5): p. 501-14.
8. Gustavsson, P. and C. Hogstedt, A cohort study of Swedish capacitor manufacturing workers exposed to polychlorinated biphenyls (PCBs). Am J Ind Med, 1997. 32(3): p. 234-9.
9. Hay, A. and J. Tarrel, Mortality of power workers exposed to phenoxy herbicides and polychlorinated biphenyls in waste transformer oil. Ann N Y Acad Sci, 1997. 837: p. 138-56.
10. Ross, R., The pathogenesis of atherosclerosis: a perspective for the 1990s. Nature, 1993. 362(6423): p. 801-9.
11. Kher, N. and J.D. Marsh, Pathobiology of atherosclerosis--a brief review. Semin Thromb Hemost, 2004. 30(6): p. 665-72.
12. Morisaki, N., et al., Platelet-derived growth factor is a potent stimulator of expression of intercellular adhesion molecule-1 in human arterial smooth muscle cells. Biochem Biophys Res Commun, 1994. 200(1): p. 612-8.
13. Rothlein, R., et al., Induction of intercellular adhesion molecule 1 on primary and continuous cell lines by pro-inflammatory cytokines. Regulation by pharmacologic agents and neutralizing antibodies. J Immunol, 1988. 141(5): p. 1665-9.
14. McEver, R.P., Selectins: lectins that initiate cell adhesion under flow. Curr Opin Cell Biol, 2002. 14(5): p. 581-6.
15. Cook-Mills, J.M. and T.L. Deem, Active participation of endothelial cells in inflammation. J Leukoc Biol, 2005. 77(4): p. 487-95.
16. Poston, R.N., et al., Expression of intercellular adhesion molecule-1 in atherosclerotic plaques. Am J Pathol, 1992. 140(3): p. 665-73.
17. Lusis, A.J., Atherosclerosis. Nature, 2000. 407(6801): p. 233-41. 18. Lehto, S., et al., Dyslipidemia and hyperglycemia predict coronary heart disease
events in middle-aged patients with NIDDM. Diabetes, 1997. 46(8): p. 1354-9. 19. Kooner, J.S., et al., Abdominal obesity, impaired nonesterified fatty acid
suppression, and insulin-mediated glucose disposal are early metabolic
113
abnormalities in families with premature myocardial infarction. Arterioscler Thromb Vasc Biol, 1998. 18(7): p. 1021-6.
20. Carlsson, M., et al., High levels of nonesterified fatty acids are associated with increased familial risk of cardiovascular disease. Arterioscler Thromb Vasc Biol, 2000. 20(6): p. 1588-94.
21. Simopoulos, A.P., Evolutionary aspects of diet, the omega-6/omega-3 ratio and genetic variation: nutritional implications for chronic diseases. Biomed Pharmacother, 2006. 60(9): p. 502-7.
22. Das, U.N., Beneficial effect(s) of n-3 fatty acids in cardiovascular diseases: but, why and how? Prostaglandins Leukot Essent Fatty Acids, 2000. 63(6): p. 351-62.
23. Kromhout, D., Fatty acids, antioxidants, and coronary heart disease from an epidemiological perspective. Lipids, 1999. 34 Suppl: p. S27-31.
24. Oomen, C.M., et al., Fish consumption and coronary heart disease mortality in Finland, Italy, and The Netherlands. Am J Epidemiol, 2000. 151(10): p. 999-1006.
25. Hu, F.B., et al., Dietary intake of alpha-linolenic acid and risk of fatal ischemic heart disease among women. Am J Clin Nutr, 1999. 69(5): p. 890-7.
26. Djousse, L., et al., Dietary linolenic acid and carotid atherosclerosis: the National Heart, Lung, and Blood Institute Family Heart Study. Am J Clin Nutr, 2003. 77(4): p. 819-25.
27. De Caterina, R., et al., Structural requirements for inhibition of cytokine-induced endothelial activation by unsaturated fatty acids. J Lipid Res, 1998. 39(5): p. 1062-70.
28. Conquer, J.A., et al., Effect of supplementation with dietary seal oil on selected cardiovascular risk factors and hemostatic variables in healthy male subjects. Thromb Res, 1999. 96(3): p. 239-50.
29. Yli-Jama, P., et al., Selective mobilisation of fatty acids from human adipose tissue. Eur J Intern Med, 2001. 12(2): p. 107-115.
30. Hennig, B., et al., Exposure to free fatty acid increases the transfer of albumin across cultured endothelial monolayers. Arteriosclerosis, 1984. 4(5): p. 489-97.
31. Spector, A.A., et al., Utilization of arachidonic and linoleic acids by cultured human endothelial cells. J Clin Invest, 1981. 68(4): p. 1003-11.
32. Old, L.J., Tumor necrosis factor (TNF). Science, 1985. 230(4726): p. 630-2. 33. Idriss, H.T. and J.H. Naismith, TNF alpha and the TNF receptor superfamily:
structure-function relationship(s). Microsc Res Tech, 2000. 50(3): p. 184-95. 34. Aggarwal, B.B., Tumour necrosis factors receptor associated signalling
molecules and their role in activation of apoptosis, JNK and NF-kappaB. Ann Rheum Dis, 2000. 59 Suppl 1: p. i6-16.
35. MacEwan, D.J., TNF ligands and receptors--a matter of life and death. Br J Pharmacol, 2002. 135(4): p. 855-75.
36. Hakoshima, T. and K. Tomita, Crystallization and preliminary X-ray investigation reveals that tumor necrosis factor is a compact trimer furnished with 3-fold symmetry. J Mol Biol, 1988. 201(2): p. 455-7.
37. Aggarwal, B.B., et al., Human tumor necrosis factor. Production, purification, and characterization. J Biol Chem, 1985. 260(4): p. 2345-54.
38. Barath, P., et al., Detection and localization of tumor necrosis factor in human atheroma. Am J Cardiol, 1990. 65(5): p. 297-302.
114
39. Pober, J.S. and R.S. Cotran, Cytokines and endothelial cell biology. Physiol Rev, 1990. 70(2): p. 427-51.
40. Terry, C.M., et al., TNF-alpha and IL-1alpha induce heme oxygenase-1 via protein kinase C, Ca2+, and phospholipase A2 in endothelial cells. Am J Physiol, 1999. 276(5 Pt 2): p. H1493-501.
41. Galis, Z.S., et al., Cytokine-stimulated human vascular smooth muscle cells synthesize a complement of enzymes required for extracellular matrix digestion. Circ Res, 1994. 75(1): p. 181-9.
42. Rajavashisth, T.B., et al., Membrane type 1 matrix metalloproteinase expression in human atherosclerotic plaques: evidence for activation by proinflammatory mediators. Circulation, 1999. 99(24): p. 3103-9.
43. Legler, D.F., et al., Recruitment of TNF receptor 1 to lipid rafts is essential for TNFalpha-mediated NF-kappaB activation. Immunity, 2003. 18(5): p. 655-64.
44. Doan, J.E., D.A. Windmiller, and D.W. Riches, Differential regulation of TNF-R1 signaling: lipid raft dependency of p42mapk/erk2 activation, but not NF-kappaB activation. J Immunol, 2004. 172(12): p. 7654-60.
45. Collins, T., et al., Transcriptional regulation of endothelial cell adhesion molecules: NF-kappa B and cytokine-inducible enhancers. Faseb J, 1995. 9(10): p. 899-909.
46. Welters, I.D., et al., NF-kappaB, nitric oxide and opiate signaling. Med Hypotheses, 2000. 54(2): p. 263-8.
47. Blackwell, T.S., et al., In vivo antioxidant treatment suppresses nuclear factor-kappa B activation and neutrophilic lung inflammation. J Immunol, 1996. 157(4): p. 1630-7.
48. DeWitt, D. and W.L. Smith, Yes, but do they still get headaches? Cell, 1995. 83(3): p. 345-8.
49. Sugimoto, Y., S. Narumiya, and A. Ichikawa, Distribution and function of prostanoid receptors: studies from knockout mice. Prog Lipid Res, 2000. 39(4): p. 289-314.
50. Palade, G.E., An electron microscope study of the mitochondrial structure. J Histochem Cytochem, 1953. 1(4): p. 188-211.
51. Razani, B., S.E. Woodman, and M.P. Lisanti, Caveolae: from cell biology to animal physiology. Pharmacol Rev, 2002. 54(3): p. 431-67.
52. Simons, K. and D. Toomre, Lipid rafts and signal transduction. Nat Rev Mol Cell Biol, 2000. 1(1): p. 31-9.
53. Liu, J., et al., Organized endothelial cell surface signal transduction in caveolae distinct from glycosylphosphatidylinositol-anchored protein microdomains. J Biol Chem, 1997. 272(11): p. 7211-22.
54. Rothberg, K.G., et al., Caveolin, a protein component of caveolae membrane coats. Cell, 1992. 68(4): p. 673-82.
55. Glenney, J.R., Jr., The sequence of human caveolin reveals identity with VIP21, a component of transport vesicles. FEBS Lett, 1992. 314(1): p. 45-8.
56. Scherer, P.E., et al., Identification, sequence, and expression of caveolin-2 defines a caveolin gene family. Proc Natl Acad Sci U S A, 1996. 93(1): p. 131-5.
115
57. Scherer, P.E., et al., Cell-type and tissue-specific expression of caveolin-2. Caveolins 1 and 2 co-localize and form a stable hetero-oligomeric complex in vivo. J Biol Chem, 1997. 272(46): p. 29337-46.
58. Tang, Z., et al., Molecular cloning of caveolin-3, a novel member of the caveolin gene family expressed predominantly in muscle. J Biol Chem, 1996. 271(4): p. 2255-61.
59. Schnitzer, J.E., et al., Filipin-sensitive caveolae-mediated transport in endothelium: reduced transcytosis, scavenger endocytosis, and capillary permeability of select macromolecules. J Cell Biol, 1994. 127(5): p. 1217-32.
60. Murata, M., et al., VIP21/caveolin is a cholesterol-binding protein. Proc Natl Acad Sci U S A, 1995. 92(22): p. 10339-43.
61. Glenney, J.R., Jr. and L. Zokas, Novel tyrosine kinase substrates from Rous sarcoma virus-transformed cells are present in the membrane skeleton. J Cell Biol, 1989. 108(6): p. 2401-8.
62. Sargiacomo, M., et al., Signal transducing molecules and glycosyl-phosphatidylinositol-linked proteins form a caveolin-rich insoluble complex in MDCK cells. J Cell Biol, 1993. 122(4): p. 789-807.
63. Lisanti, M.P., et al., Characterization of caveolin-rich membrane domains isolated from an endothelial-rich source: implications for human disease. J Cell Biol, 1994. 126(1): p. 111-26.
64. Sargiacomo, M., et al., Oligomeric structure of caveolin: implications for caveolae membrane organization. Proc Natl Acad Sci U S A, 1995. 92(20): p. 9407-11.
65. Fra, A.M., et al., De novo formation of caveolae in lymphocytes by expression of VIP21-caveolin. Proc Natl Acad Sci U S A, 1995. 92(19): p. 8655-9.
66. Li, S., et al., Baculovirus-based expression of mammalian caveolin in Sf21 insect cells. A model system for the biochemical and morphological study of caveolae biogenesis. J Biol Chem, 1996. 271(45): p. 28647-54.
67. Li, S., et al., Evidence for a regulated interaction between heterotrimeric G proteins and caveolin. J Biol Chem, 1995. 270(26): p. 15693-701.
68. Garcia-Cardena, G., et al., Targeting of nitric oxide synthase to endothelial cell caveolae via palmitoylation: implications for nitric oxide signaling. Proc Natl Acad Sci U S A, 1996. 93(13): p. 6448-53.
69. Orlandi, P.A. and P.H. Fishman, Filipin-dependent inhibition of cholera toxin: evidence for toxin internalization and activation through caveolae-like domains. J Cell Biol, 1998. 141(4): p. 905-15.
70. Liu, P., et al., Presence of oxidized cholesterol in caveolae uncouples active platelet-derived growth factor receptors from tyrosine kinase substrates. J Biol Chem, 2000. 275(41): p. 31648-54.
71. Gafencu, A., et al., Protein and fatty acid composition of caveolae from apical plasmalemma of aortic endothelial cells. Cell Tissue Res, 1998. 293(1): p. 101-10.
72. Li, Q., et al., Eicosapentaenoic acid modifies lipid composition in caveolae and induces translocation of endothelial nitric oxide synthase. Biochimie, 2007. 89(1): p. 169-77.
73. Ma, D.W., et al., n-3 PUFA alter caveolae lipid composition and resident protein localization in mouse colon. Faseb J, 2004. 18(9): p. 1040-2.
116
74. Schley, P.D., D.N. Brindley, and C.J. Field, (n-3) PUFA alter raft lipid composition and decrease epidermal growth factor receptor levels in lipid rafts of human breast cancer cells. J Nutr, 2007. 137(3): p. 548-53.
75. Seo, J., et al., Docosahexaenoic acid selectively inhibits plasma membrane targeting of lipidated proteins. Faseb J, 2006. 20(6): p. 770-2.
76. Roy, S., et al., Dominant-negative caveolin inhibits H-Ras function by disrupting cholesterol-rich plasma membrane domains. Nat Cell Biol, 1999. 1(2): p. 98-105.
77. Abumrad, N.A., et al., Cloning of a rat adipocyte membrane protein implicated in binding or transport of long-chain fatty acids that is induced during preadipocyte differentiation. Homology with human CD36. J Biol Chem, 1993. 268(24): p. 17665-8.
78. Kolleck, I., et al., Cellular cholesterol stimulates acute uptake of palmitate by redistribution of fatty acid translocase in type II pneumocytes. Biochemistry, 2002. 41(20): p. 6369-75.
79. D'Alessio, A., et al., Caveolae participate in tumor necrosis factor receptor 1 signaling and internalization in a human endothelial cell line. Am J Pathol, 2005. 166(4): p. 1273-82.
80. Liou, J.Y., et al., Colocalization and interaction of cyclooxygenase-2 with caveolin-1 in human fibroblasts. J Biol Chem, 2001. 276(37): p. 34975-82.
81. Matveev, S.V. and E.J. Smart, Heterologous desensitization of EGF receptors and PDGF receptors by sequestration in caveolae. Am J Physiol Cell Physiol, 2002. 282(4): p. C935-46.
82. Madge, L.A. and J.S. Pober, TNF signaling in vascular endothelial cells. Exp Mol Pathol, 2001. 70(3): p. 317-25.
83. Ko, Y.G., et al., TNF-alpha-mediated apoptosis is initiated in caveolae-like domains. J Immunol, 1999. 162(12): p. 7217-23.
84. Huang, C., et al., p38 kinase mediates UV-induced phosphorylation of p53 protein at serine 389. J Biol Chem, 1999. 274(18): p. 12229-35.
85. Ushio-Fukai, M., et al., p38 Mitogen-activated protein kinase is a critical component of the redox-sensitive signaling pathways activated by angiotensin II. Role in vascular smooth muscle cell hypertrophy. J Biol Chem, 1998. 273(24): p. 15022-9.
86. Volonte, D., et al., Cellular stress induces the tyrosine phosphorylation of caveolin-1 (Tyr(14)) via activation of p38 mitogen-activated protein kinase and c-Src kinase. Evidence for caveolae, the actin cytoskeleton, and focal adhesions as mechanical sensors of osmotic stress. J Biol Chem, 2001. 276(11): p. 8094-103.
87. Ballard-Croft, C., et al., Regional myocardial ischemia-induced activation of MAPKs is associated with subcellular redistribution of caveolin and cholesterol. Am J Physiol Heart Circ Physiol, 2006. 291(2): p. H658-67.
88. Drab, M., et al., Loss of caveolae, vascular dysfunction, and pulmonary defects in caveolin-1 gene-disrupted mice. Science, 2001. 293(5539): p. 2449-52.
89. Razani, B., et al., Caveolin-1 null mice are viable but show evidence of hyperproliferative and vascular abnormalities. J Biol Chem, 2001. 276(41): p. 38121-38.
90. La Rocca, C. and A. Mantovani, From environment to food: the case of PCB. Ann Ist Super Sanita, 2006. 42(4): p. 410-6.
117
91. Yobs, A.R., Levels of Polychlorinated Biphenyls in Adipose Tissue of the General Population of the Nation. Environ Health Perspect, 1972. 1: p. 79-81.
92. Finklea, J., et al., Polychlorinated biphenyl residues in human plasma expose a major urban pollution problem. Am J Public Health, 1972. 62(5): p. 645-51.
93. Kimbrough, R.D., The toxicity of polychlorinated polycyclic compounds and related chemicals. CRC Crit Rev Toxicol, 1974. 2(4): p. 445-98.
94. Toborek, M., et al., Exposure to polychlorinated biphenyls causes endothelial cell dysfunction. J Biochem Toxicol, 1995. 10(4): p. 219-26.
95. Stegeman, J.J., et al., Induction of cytochrome P4501A1 by aryl hydrocarbon receptor agonists in porcine aorta endothelial cells in culture and cytochrome P4501A1 activity in intact cells. Mol Pharmacol, 1995. 47(2): p. 296-306.
96. Ramadass, P., et al., Dietary flavonoids modulate PCB-induced oxidative stress, CYP1A1 induction, and AhR-DNA binding activity in vascular endothelial cells. Toxicol Sci, 2003. 76(1): p. 212-9.
97. Hansen, E.S., International Commission for Protection Against Environmental Mutagens and Carcinogens. ICPEMC Working Paper 7/1/2. Shared risk factors for cancer and atherosclerosis--a review of the epidemiological evidence. Mutat Res, 1990. 239(3): p. 163-79.
98. Lovati, M.R., et al., Increased plasma and aortic triglycerides in rabbits after acute administration of 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicol Appl Pharmacol, 1984. 75(1): p. 91-7.
99. Slim, R., et al., Antioxidant protection against PCB-mediated endothelial cell activation. Toxicol Sci, 1999. 52(2): p. 232-9.
100. Li, Y., et al., Polychlorinated biphenyls, cytochrome P450 1A1 (CYP1A1) polymorphisms, and breast cancer risk among African American women and white women in North Carolina: a population-based case-control study. Breast Cancer Res, 2005. 7(1): p. R12-8.
101. Azais-Braesco, V., et al., Effects of two prototypic polychlorinated biphenyls (PCBs) on lipid composition of rat liver and serum. J Nutr Biochem, 1990. 1(7): p. 350-4.
102. Matsusue, K., et al., A highly toxic PCB produces unusual changes in the fatty acid composition of rat liver. Toxicol Lett, 1997. 91(2): p. 99-104.
103. Bell, F.P., et al., Long-term effects of Aroclor 1254 (PCBs) on plasma lipid and carnitine concentrations in rhesus monkey. Toxicology, 1994. 89(2): p. 139-53.
104. Hennig, B., et al., Proinflammatory properties of coplanar PCBs: in vitro and in vivo evidence. Toxicol Appl Pharmacol, 2002. 181(3): p. 174-83.
105. Hennig, B., et al., Linoleic acid induces proinflammatory events in vascular endothelial cells via activation of PI3K/Akt and ERK1/2 signaling. J Nutr Biochem, 2006. 17(11): p. 766-72.
106. Hennig, B., et al., Linoleic acid amplifies polychlorinated biphenyl-mediated dysfunction of endothelial cells. J Biochem Mol Toxicol, 1999. 13(2): p. 83-91.
107. Toborek, M., et al., Linoleic acid potentiates TNF-mediated oxidative stress, disruption of calcium homeostasis, and apoptosis of cultured vascular endothelial cells. J Lipid Res, 1997. 38(10): p. 2155-67.
118
108. Hennig, B., et al., Dietary fat interacts with PCBs to induce changes in lipid metabolism in mice deficient in low-density lipoprotein receptor. Environ Health Perspect, 2005. 113(1): p. 83-7.
109. Hennig, B., et al., Polychlorinated biphenyls, oxidative stress and diet, in Reviews in food and nutrition toxicity, V.R. Preedy and R.R. Watson, Editors. 2005, CRC Press: Boca Raton, FL. p. 93-128.
110. Quyyumi, A.A., Endothelial function in health and disease: new insights into the genesis of cardiovascular disease. Am J Med, 1998. 105(1A): p. 32S-39S.
111. Steinberg, D., et al., Beyond cholesterol. Modifications of low-density lipoprotein that increase its atherogenicity. N Engl J Med, 1989. 320(14): p. 915-24.
112. Christon, R.A., Mechanisms of action of dietary fatty acids in regulating the activation of vascular endothelial cells during atherogenesis. Nutr Rev, 2003. 61(8): p. 272-9.
113. Pober, J.S. and W. Min, Endothelial cell dysfunction, injury and death. Handb Exp Pharmacol, 2006(176 Pt 2): p. 135-56.
114. Cesari, M., et al., Inflammatory markers and cardiovascular disease (The Health, Aging and Body Composition [Health ABC] Study). Am J Cardiol, 2003. 92(5): p. 522-8.
115. de Winther, M.P., et al., Nuclear factor kappaB signaling in atherogenesis. Arterioscler Thromb Vasc Biol, 2005. 25(5): p. 904-14.
116. Kunsch, C. and R.M. Medford, Oxidative stress as a regulator of gene expression in the vasculature. Circ Res, 1999. 85(8): p. 753-66.
117. Muller, J.M., R.A. Rupec, and P.A. Baeuerle, Study of gene regulation by NF-kappa B and AP-1 in response to reactive oxygen intermediates. Methods, 1997. 11(3): p. 301-12.
118. Libby, P., et al., Cytokines regulate vascular functions related to stability of the atherosclerotic plaque. J Cardiovasc Pharmacol, 1995. 25 Suppl 2: p. S9-12.
119. Meager, A., Cytokine regulation of cellular adhesion molecule expression in inflammation. Cytokine Growth Factor Rev, 1999. 10(1): p. 27-39.
120. Romero-Corral, A., et al., Association of bodyweight with total mortality and with cardiovascular events in coronary artery disease: a systematic review of cohort studies. Lancet, 2006. 368(9536): p. 666-78.
121. Cullen, P., Evidence that triglycerides are an independent coronary heart disease risk factor. Am J Cardiol, 2000. 86(9): p. 943-9.
122. Mostaza, J.M., et al., Abnormal metabolism of free fatty acids in hypertriglyceridaemic men: apparent insulin resistance of adipose tissue. J Intern Med, 1998. 243(4): p. 265-74.
123. Austin, M.A., et al., Cardiovascular disease mortality in familial forms of hypertriglyceridemia: A 20-year prospective study. Circulation, 2000. 101(24): p. 2777-82.
124. Malloy, M.J. and J.P. Kane, A risk factor for atherosclerosis: triglyceride-rich lipoproteins. Adv Intern Med, 2001. 47: p. 111-36.
125. De Caterina, R., J.K. Liao, and P. Libby, Fatty acid modulation of endothelial activation. Am J Clin Nutr, 2000. 71(1 Suppl): p. 213S-23S.
126. Psota, T.L., S.K. Gebauer, and P. Kris-Etherton, Dietary omega-3 fatty acid intake and cardiovascular risk. Am J Cardiol, 2006. 98(4A): p. 3i-18i.
119
127. Grimble, R.F., Dietary lipids and the inflammatory response. Proc Nutr Soc, 1998. 57(4): p. 535-42.
128. Hennig, B., M. Toborek, and C.J. McClain, High-energy diets, fatty acids and endothelial cell function: implications for atherosclerosis. J Am Coll Nutr, 2001. 20(2 Suppl): p. 97-105.
129. Trumbo, P., et al., Dietary reference intakes for energy, carbohydrate, fiber, fat, fatty acids, cholesterol, protein and amino acids. J Am Diet Assoc, 2002. 102(11): p. 1621-30.
130. De Caterina, R., et al., Nutritional mechanisms that influence cardiovascular disease. Am J Clin Nutr, 2006. 83(2): p. 421S-426S.
131. Alessandri, C., et al., Alpha-linolenic acid-rich wheat germ oil decreases oxidative stress and CD40 ligand in patients with mild hypercholesterolemia. Arterioscler Thromb Vasc Biol, 2006. 26(11): p. 2577-8.
132. Thies, F., et al., Influence of dietary supplementation with long-chain n-3 or n-6 polyunsaturated fatty acids on blood inflammatory cell populations and functions and on plasma soluble adhesion molecules in healthy adults. Lipids, 2001. 36(11): p. 1183-93.
133. Mozaffarian, D., Does alpha-linolenic acid intake reduce the risk of coronary heart disease? A review of the evidence. Altern Ther Health Med, 2005. 11(3): p. 24-30; quiz 31, 79.
134. De Caterina, R. and M. Massaro, Omega-3 fatty acids and the regulation of expression of endothelial pro-atherogenic and pro-inflammatory genes. J Membr Biol, 2005. 206(2): p. 103-16.
135. Meerarani, P., et al., Zinc protects against apoptosis of endothelial cells induced by linoleic acid and tumor necrosis factor alpha. Am J Clin Nutr, 2000. 71(1): p. 81-7.
136. Toborek, M., et al., Measurement of inflammatory properties of fatty acids in human endothelial cells. Methods Enzymol, 2002. 352: p. 198-219.
137. Spector, A.A., Lipids, hormones, and atherogenesis. The transport and utilization of free fatty acid. Ann N Y Acad Sci, 1968. 149(2): p. 768-83.
138. Mattson, M.P., et al., Calcium, free radicals, and excitotoxic neuronal death in primary cell culture. Methods Cell Biol, 1995. 46: p. 187-216.
139. Beg, A.A., et al., Tumor necrosis factor and interleukin-1 lead to phosphorylation and loss of I kappa B alpha: a mechanism for NF-kappa B activation. Mol Cell Biol, 1993. 13(6): p. 3301-10.
140. Lechleitner, M., et al., Tumour necrosis factor-alpha plasma level in patients with type 1 diabetes mellitus and its association with glycaemic control and cardiovascular risk factors. J Intern Med, 2000. 248(1): p. 67-76.
141. Pickup, J.C., et al., Plasma interleukin-6, tumour necrosis factor alpha and blood cytokine production in type 2 diabetes. Life Sci, 2000. 67(3): p. 291-300.
142. Casey, L.C., R.A. Balk, and R.C. Bone, Plasma cytokine and endotoxin levels correlate with survival in patients with the sepsis syndrome. Ann Intern Med, 1993. 119(8): p. 771-8.
143. Salles, G., et al., Elevated circulating levels of TNFalpha and its p55 soluble receptor are associated with an adverse prognosis in lymphoma patients. Br J Haematol, 1996. 93(2): p. 352-9.
120
144. Lin, S.J., et al., Superoxide dismutase inhibits the expression of vascular cell adhesion molecule-1 and intracellular cell adhesion molecule-1 induced by tumor necrosis factor-alpha in human endothelial cells through the JNK/p38 pathways. Arterioscler Thromb Vasc Biol, 2005. 25(2): p. 334-40.
145. Zauli, G., et al., Tumor necrosis factor-related apoptosis-inducing ligand (TRAIL) sequentially upregulates nitric oxide and prostanoid production in primary human endothelial cells. Circ Res, 2003. 92(7): p. 732-40.
146. Pham, C.G., et al., Ferritin heavy chain upregulation by NF-kappaB inhibits TNFalpha-induced apoptosis by suppressing reactive oxygen species. Cell, 2004. 119(4): p. 529-42.
147. Sperling, R.I., R.A. Lewis, and K.F. Austen, Regulation of 5-lipoxygenase pathway product generation in human neutrophils by n-3 fatty acids. Prog Lipid Res, 1986. 25(1-4): p. 101-4.
148. Fredrickson, D.S. and R.S. Gordon, Jr., Transport of fatty acids. Physiol Rev, 1958. 38(4): p. 585-630.
149. Sun, X.F. and H. Zhang, NFKB and NFKBI polymorphisms in relation to susceptibility of tumour and other diseases. Histol Histopathol, 2007. 22(12): p. 1387-98.
150. Wang, T., X. Zhang, and J.J. Li, The role of NF-kappaB in the regulation of cell stress responses. Int Immunopharmacol, 2002. 2(11): p. 1509-20.
151. Chen, Z., et al., MAP kinases. Chem Rev, 2001. 101(8): p. 2449-76. 152. Clark, J.D., et al., A novel arachidonic acid-selective cytosolic PLA2 contains a
Ca(2+)-dependent translocation domain with homology to PKC and GAP. Cell, 1991. 65(6): p. 1043-51.
153. Vila, L., Cyclooxygenase and 5-lipoxygenase pathways in the vessel wall: role in atherosclerosis. Med Res Rev, 2004. 24(4): p. 399-424.
154. Bishop-Bailey, D., J.A. Mitchell, and T.D. Warner, COX-2 in cardiovascular disease. Arterioscler Thromb Vasc Biol, 2006. 26(5): p. 956-8.
155. Yang, C.M., et al., Interleukin-1beta-induced cyclooxygenase-2 expression is mediated through activation of p42/44 and p38 MAPKS, and NF-kappaB pathways in canine tracheal smooth muscle cells. Cell Signal, 2002. 14(11): p. 899-911.
156. Toborek, M., et al., Linoleic acid and TNF-alpha cross-amplify oxidative injury and dysfunction of endothelial cells. J Lipid Res, 1996. 37(1): p. 123-35.
157. Li, Y.P., et al., TNF-alpha acts via p38 MAPK to stimulate expression of the ubiquitin ligase atrogin1/MAFbx in skeletal muscle. Faseb J, 2005. 19(3): p. 362-70.
158. Clark, J.E., N. Sarafraz, and M.S. Marber, Potential of p38-MAPK inhibitors in the treatment of ischaemic heart disease. Pharmacol Ther, 2007. 116(2): p. 192-206.
159. Gustin, J.A., et al., Tumor necrosis factor activates CRE-binding protein through a p38 MAPK/MSK1 signaling pathway in endothelial cells. Am J Physiol Cell Physiol, 2004. 286(3): p. C547-55.
160. Pearson, G., et al., Mitogen-activated protein (MAP) kinase pathways: regulation and physiological functions. Endocr Rev, 2001. 22(2): p. 153-83.
121
161. Saccani, S., S. Pantano, and G. Natoli, p38-Dependent marking of inflammatory genes for increased NF-kappa B recruitment. Nat Immunol, 2002. 3(1): p. 69-75.
162. Roebuck, K.A., Oxidant stress regulation of IL-8 and ICAM-1 gene expression: differential activation and binding of the transcription factors AP-1 and NF-kappaB (Review). Int J Mol Med, 1999. 4(3): p. 223-30.
163. Kyriakis, J.M. and J. Avruch, Protein kinase cascades activated by stress and inflammatory cytokines. Bioessays, 1996. 18(7): p. 567-77.
164. Han, J., et al., A MAP kinase targeted by endotoxin and hyperosmolarity in mammalian cells. Science, 1994. 265(5173): p. 808-11.
165. Yao, B., et al., Phosphorylation of Raf by ceramide-activated protein kinase. Nature, 1995. 378(6554): p. 307-10.
166. Muller, G., et al., Regulation of Raf-1 kinase by TNF via its second messenger ceramide and cross-talk with mitogenic signalling. Embo J, 1998. 17(3): p. 732-42.
167. Kumar, S., J. Boehm, and J.C. Lee, p38 MAP kinases: key signalling molecules as therapeutic targets for inflammatory diseases. Nat Rev Drug Discov, 2003. 2(9): p. 717-26.
168. Beyaert, R., et al., The p38/RK mitogen-activated protein kinase pathway regulates interleukin-6 synthesis response to tumor necrosis factor. Embo J, 1996. 15(8): p. 1914-23.
169. MacEwan, D.J., TNF receptor subtype signalling: differences and cellular consequences. Cell Signal, 2002. 14(6): p. 477-92.
170. Frank, P.G., et al., Caveolin, caveolae, and endothelial cell function. Arterioscler Thromb Vasc Biol, 2003. 23(7): p. 1161-8.
171. Feng, X., et al., Caveolin-1 associates with TRAF2 to form a complex that is recruited to tumor necrosis factor receptors. J Biol Chem, 2001. 276(11): p. 8341-9.
172. Yamada, E., The fine structure of the gall bladder epithelium of the mouse. J Biophys Biochem Cytol, 1955. 1(5): p. 445-58.
173. Pohl, J., et al., FAT/CD36-mediated long-chain fatty acid uptake in adipocytes requires plasma membrane rafts. Mol Biol Cell, 2005. 16(1): p. 24-31.
174. Stan, R.V., et al., Immunoisolation and partial characterization of endothelial plasmalemmal vesicles (caveolae). Mol Biol Cell, 1997. 8(4): p. 595-605.
175. Hunter, I. and G.F. Nixon, Spatial compartmentalization of tumor necrosis factor (TNF) receptor 1-dependent signaling pathways in human airway smooth muscle cells. Lipid rafts are essential for TNF-alpha-mediated activation of RhoA but dispensable for the activation of the NF-kappaB and MAPK pathways. J Biol Chem, 2006. 281(45): p. 34705-15.
176. Cha, S.H., et al., Evidence for cyclooxygenase-1 association with caveolin-1 and -2 in cultured human embryonic kidney (HEK 293) cells. IUBMB Life, 2004. 56(4): p. 221-7.
177. Kwak, J.O., et al., Evidence for cyclooxygenase-2 association with caveolin-3 in primary cultured rat chondrocytes. J Korean Med Sci, 2006. 21(1): p. 100-6.
178. Kosswig, N., et al., Class A scavenger receptor-mediated adhesion and internalization require distinct cytoplasmic domains. J Biol Chem, 2003. 278(36): p. 34219-25.
122
179. Smart, E.J., et al., A detergent-free method for purifying caveolae membrane from tissue culture cells. Proc Natl Acad Sci U S A, 1995. 92(22): p. 10104-8.
180. Smart, E.J., C. Mineo, and R.G. Anderson, Clustered folate receptors deliver 5-methyltetrahydrofolate to cytoplasm of MA104 cells. J Cell Biol, 1996. 134(5): p. 1169-77.
181. Smart, E.J., et al., A role for caveolin in transport of cholesterol from endoplasmic reticulum to plasma membrane. J Biol Chem, 1996. 271(46): p. 29427-35.
182. Salani, B., et al., Glimepiride activates eNOS with a mechanism Akt but not caveolin-1 dependent. Biochem Biophys Res Commun, 2005. 335(3): p. 832-5.
183. Repetto, S., et al., Insulin and IGF-I phosphorylate eNOS in HUVECs by a caveolin-1 dependent mechanism. Biochem Biophys Res Commun, 2005. 337(3): p. 849-52.
184. Michel, V. and M. Bakovic, Lipid rafts in health and disease. Biol Cell, 2007. 99(3): p. 129-40.
185. Frank, P.G., et al., Genetic ablation of caveolin-1 confers protection against atherosclerosis. Arterioscler Thromb Vasc Biol, 2004. 24(1): p. 98-105.
186. Matveev, S., et al., The role of caveolae and caveolin in vesicle-dependent and vesicle-independent trafficking. Adv Drug Deliv Rev, 2001. 49(3): p. 237-50.
187. Smart, E.J., et al., Caveolins, liquid-ordered domains, and signal transduction. Mol Cell Biol, 1999. 19(11): p. 7289-304.
188. Lisanti, M.P., M. Sargiacomo, and P.E. Scherer, Purification of caveolae-derived membrane microdomains containing lipid-anchored signaling molecules, such as GPI-anchored proteins, H-Ras, Src-family tyrosine kinases, eNOS, and G-protein alpha-, beta-, and gamma-subunits. Methods Mol Biol, 1999. 116: p. 51-60.
189. Anderson, R.G., The caveolae membrane system. Annu Rev Biochem, 1998. 67: p. 199-225.
190. Harris, J., et al., Caveolae and caveolin in immune cells: distribution and functions. Trends Immunol, 2002. 23(3): p. 158-64.
191. Ring, A., et al., Caveolin-1 is required for fatty acid translocase (FAT/CD36) localization and function at the plasma membrane of mouse embryonic fibroblasts. Biochim Biophys Acta, 2006. 1761(4): p. 416-23.
192. Raza Shaikh, S., et al., Acyl chain unsaturation in PEs modulates phase separation from lipid raft molecules. Biochem Biophys Res Commun, 2003. 311(3): p. 793-6.
193. Niu, S.L. and B.J. Litman, Determination of membrane cholesterol partition coefficient using a lipid vesicle-cyclodextrin binary system: effect of phospholipid acyl chain unsaturation and headgroup composition. Biophys J, 2002. 83(6): p. 3408-15.
194. Moller, D.E. and K.D. Kaufman, Metabolic syndrome: a clinical and molecular perspective. Annu Rev Med, 2005. 56: p. 45-62.
195. Yang, N., et al., High-fat diet up-regulates caveolin-1 expression in aorta of diet-induced obese but not in diet-resistant rats. Cardiovasc Res, 2007.
196. Sergeev, A.V. and D.O. Carpenter, Hospitalization rates for coronary heart disease in relation to residence near areas contaminated with persistent organic pollutants and other pollutants. Environ Health Perspect, 2005. 113(6): p. 756-61.
123
197. Shimizu, K., et al., Lipid peroxidation is enhanced in Yusho victims 35 years after accidental poisoning with polychlorinated biphenyls in Nagasaki, Japan. J Appl Toxicol, 2007. 27(2): p. 195-7.
198. Savouret, J.F., A. Berdeaux, and R.F. Casper, The aryl hydrocarbon receptor and its xenobiotic ligands: a fundamental trigger for cardiovascular diseases. Nutr Metab Cardiovasc Dis, 2003. 13(2): p. 104-13.
199. Hennig, B., et al., Introductory comments: nutrition, environmental toxins and implications in prevention and intervention of human diseases. J Nutr Biochem, 2007. 18(3): p. 161-2.
200. Hennig, B., et al., Using nutrition for intervention and prevention against environmental chemical toxicity and associated diseases. Environ Health Perspect, 2007. 115(4): p. 493-5.
201. Doi, A.M., et al., Effect of micelle fatty acid composition and 3,4,3', 4'-tetrachlorobiphenyl (TCB) exposure on intestinal [(14)C]-TCB bioavailability and biotransformation in channel catfish in situ preparations. Toxicol Sci, 2000. 55(1): p. 85-96.
202. Simopoulos, A.P., Essential fatty acids in health and chronic disease. Am J Clin Nutr, 1999. 70(3 Suppl): p. 560S-569S.
203. Matsusue, K., et al., A highly toxic coplanar polychlorinated biphenyl compound suppresses Delta5 and Delta6 desaturase activities which play key roles in arachidonic acid synthesis in rat liver. Chem Res Toxicol, 1999. 12(12): p. 1158-65.
204. Rallidis, L.S., et al., Dietary alpha-linolenic acid decreases C-reactive protein, serum amyloid A and interleukin-6 in dyslipidaemic patients. Atherosclerosis, 2003. 167(2): p. 237-42.
205. Kwon, O., et al., Expression of cyclooxygenase-2 and pro-inflammatory cytokines induced by 2,2',4,4',5,5'-hexachlorobiphenyl (PCB 153) in human mast cells requires NF-kappa B activation. Biol Pharm Bull, 2002. 25(9): p. 1165-8.
206. Choi, W., et al., PCB 104-induced proinflammatory reactions in human vascular endothelial cells: relationship to cancer metastasis and atherogenesis. Toxicol Sci, 2003. 75(1): p. 47-56.
207. Jensen, A.A., Polychlorobiphenyls (PCBs), polychlorodibenzo-p-dioxins (PCDDs) and polychlorodibenzofurans (PCDFs) in human milk, blood and adipose tissue. Sci Total Environ, 1987. 64(3): p. 259-93.
208. Wassermann, M., et al., World PCBs map: storage and effects in man and his biologic environment in the 1970s. Ann N Y Acad Sci, 1979. 320: p. 69-124.
209. Altavilla, D., et al., IRFI 042, a novel dual vitamin E-like antioxidant, inhibits activation of nuclear factor-kappaB and reduces the inflammatory response in myocardial ischemia-reperfusion injury. Cardiovasc Res, 2000. 47(3): p. 515-28.
210. Price, D.T. and J. Loscalzo, Cellular adhesion molecules and atherogenesis. Am J Med, 1999. 107(1): p. 85-97.
211. Smith, W.L., D.L. DeWitt, and R.M. Garavito, Cyclooxygenases: structural, cellular, and molecular biology. Annu Rev Biochem, 2000. 69: p. 145-82.
212. Sidhu, K.S., Health benefits and potential risks related to consumption of fish or fish oil. Regul Toxicol Pharmacol, 2003. 38(3): p. 336-44.
124
213. Ikonomou, M.G., et al., Flesh quality of market-size farmed and wild British Columbia salmon. Environ Sci Technol, 2007. 41(2): p. 437-43.
214. Hennig, B., et al., Modification of environmental toxicity by nutrients: implications in atherosclerosis. Cardiovasc Toxicol, 2005. 5(2): p. 153-60.
215. Toborek, M., et al., Unsaturated fatty acids selectively induce an inflammatory environment in human endothelial cells. Am J Clin Nutr, 2002. 75(1): p. 119-25.
216. Boyle, E.M., Jr., et al., Inhibition of nuclear factor-kappa B nuclear localization reduces human E-selectin expression and the systemic inflammatory response. Circulation, 1998. 98(19 Suppl): p. II282-8.
217. Ungvari, Z.I., et al., Increased mitochondrial H2O2 production promotes endothelial NF-{kappa}B activation in aged rat arteries. Am J Physiol Heart Circ Physiol, 2007.
218. Forsell, P.K., et al., Polychlorinated biphenyls induce arachidonic acid release in human platelets in a tamoxifen sensitive manner via activation of group IVA cytosolic phospholipase A2-alpha. Biochem Pharmacol, 2005. 71(1-2): p. 144-55.
219. Brant, K.A. and R.L. Caruso, PCB 50 stimulates release of arachidonic acid and prostaglandins from late gestation rat amnion fibroblast cells. Reprod Toxicol, 2006. 22(4): p. 591-8.
220. Horia, E. and B.A. Watkins, Comparison of stearidonic acid and alpha-linolenic acid on PGE2 production and COX-2 protein levels in MDA-MB-231 breast cancer cell cultures. J Nutr Biochem, 2005. 16(3): p. 184-92.
221. Wan, M., et al., Eicosapentaenoic acid inhibits TNF-alpha-induced Lnk expression in human umbilical vein endothelial cells: involvement of the PI3K/Akt pathway. J Nutr Biochem, 2007. 18(1): p. 17-22.
222. Mishra, A., A. Chaudhary, and S. Sethi, Oxidized omega-3 fatty acids inhibit NF-kappaB activation via a PPARalpha-dependent pathway. Arterioscler Thromb Vasc Biol, 2004. 24(9): p. 1621-7.
223. Zhao, G., et al., Dietary {alpha}-linolenic acid inhibits proinflammatory cytokine production by peripheral blood mononuclear cells in hypercholesterolemic subjects. Am J Clin Nutr, 2007. 85(2): p. 385-91.
224. Perkins, D.J. and D.A. Kniss, Tumor necrosis factor-alpha promotes sustained cyclooxygenase-2 expression: attenuation by dexamethasone and NSAIDs. Prostaglandins, 1997. 54(4): p. 727-43.
225. Murakami, M. and I. Kudo, Recent advances in molecular biology and physiology of the prostaglandin E2-biosynthetic pathway. Prog Lipid Res, 2004. 43(1): p. 3-35.
226. Morita, I., et al., Different intracellular locations for prostaglandin endoperoxide H synthase-1 and -2. J Biol Chem, 1995. 270(18): p. 10902-8.
227. Razani, B., et al., Caveolin-1-deficient mice are lean, resistant to diet-induced obesity, and show hypertriglyceridemia with adipocyte abnormalities. J Biol Chem, 2002. 277(10): p. 8635-47.
228. Liou, J.Y., et al., Colocalization of prostacyclin synthase with prostaglandin H synthase-1 (PGHS-1) but not phorbol ester-induced PGHS-2 in cultured endothelial cells. J Biol Chem, 2000. 275(20): p. 15314-20.
229. Yang, B. and V. Rizzo, TNF-alpha potentiates protein-tyrosine nitration through activation of NADPH oxidase and eNOS localized in membrane rafts and
125
caveolae of bovine aortic endothelial cells. Am J Physiol Heart Circ Physiol, 2007. 292(2): p. H954-62.
230. Lim, E.J., et al., The role of caveolin-1 in PCB77-induced eNOS phosphorylation in human-derived endothelial cells. Am J Physiol Heart Circ Physiol, 2007.
231. Simons, K. and R. Ehehalt, Cholesterol, lipid rafts, and disease. J Clin Invest, 2002. 110(5): p. 597-603.
232. Rietveld, A. and K. Simons, The differential miscibility of lipids as the basis for the formation of functional membrane rafts. Biochim Biophys Acta, 1998. 1376(3): p. 467-79.
233. Sotgia, F., et al., Intracellular retention of glycosylphosphatidyl inositol-linked proteins in caveolin-deficient cells. Mol Cell Biol, 2002. 22(11): p. 3905-26.
234. Pettus, B.J., C.E. Chalfant, and Y.A. Hannun, Sphingolipids in inflammation: roles and implications. Curr Mol Med, 2004. 4(4): p. 405-18.
235. Johns, D.G., J.S. Jin, and R.C. Webb, The role of the endothelium in ceramide-induced vasodilation. Eur J Pharmacol, 1998. 349(2-3): p. R9-10.
236. Raines, M.A., R.N. Kolesnick, and D.W. Golde, Sphingomyelinase and ceramide activate mitogen-activated protein kinase in myeloid HL-60 cells. J Biol Chem, 1993. 268(20): p. 14572-5.
237. Saleem, A., et al., Activation of serine/threonine protein kinases and early growth response 1 gene expression by tumor necrosis factor in human myeloid leukemia cells. J Immunol, 1995. 154(8): p. 4150-6.
238. Manna, S.K., V. Haridas, and B.B. Aggarwal, Bcl-x(L) suppresses TNF-mediated apoptosis and activation of nuclear factor-kappaB, activation protein-1, and c-Jun N-terminal kinase. J Interferon Cytokine Res, 2000. 20(8): p. 725-35.
126
Vita
Personal Information
Name: Lei Wang
Date of Birth: 02/10/1978
Place of Birth: Shenyang City, Liaoning Province, P.R.China
Education
2003 MS of Food Science and Nutrition, China Agricultural University, Beijing China
2000 BS of Food Science and Technology, China Agricultural University, Beijing China
Professional Positions
2003-present Research Assistant
Graduate Center for Nutritional Sciences, University of Kentucky USA
2002-2003 Research Assistant
Space Nutritional Laboratory, China Research Institute of Space Medical Science
1999-2002 Research Assistant
Food Safety and Inspection Standard Laboratory, Chinese Ministry of Agriculture
Awards and Honors
2006-2007 Presidential Fellowship, University of Kentucky.
2006 Featured on NIEHS/NIH website as “Student Success Story”:
http://www.niehs.nih.gov/research/supported/sbrp/training/training2_s9.cfm
2004-2007 Annual Travel Grant Award, University of Kentucky
2001 Young Investigator Bursary, International Union of Nutritional Sciences Council.
The 17th International Conference of Nutrition, Austria Vienna (Only 25 students
worldwide received this award)
2001 Second-Prize of Excellent Dissertation, The 4th National Ocean Active Compounds
Conference, Shanghai China
1997-1998 Presidential Scholarship, China Agricultural University, Beijing China
127
Publications
Manuscripts already published:
1. Lei Wang, Gudrun Reiterer, Michal Toborek, Bernhard Hennig. Changing ratios of
omega-6 to omega-3 fatty acids can differentially modulate Polychlorinated
biphenyl toxicity in endothelial cells. Chemico-Biological Interactions (in press,
2007).
2. Bernhard Hennig, Lei Wang, Xabier Arzuaga, Debjani Das Ghosh, Viswanathan
Saraswathia,Michal Toborek. Linoleic acid induces proinflammatory events in
vascular endothelial cells via activation of the PI3K/Akt and ERK1/2 signaling.
Journal of Nutritional Biochemistry. 2006 17(11):766-72.
3. Wang, L, Bai Sh, Shi J. Effect of dietary n-3 polyunsaturated fatty acid on lipid
metabolism in the rat of micro-gravity model. Space Medicine & Medical
Engineering, Medline and Aerospace Database, 2002, 15(5):347-350.
4. Lei Wang, Shi Jieping. Dietary unsaturated fatty acid and lipid metabolism in mice
model. Nutrition and Functional Food Research and Progress, 2002, 1:44.
5. Shi J, Song L, Wang D, Wang, L. Advances on uses of biotechnology in modifying
plant lipids. (Review). Journal of Agricultural Biotechnology, 2001, 9(4):403-408.
6. Shi J, Zhao K, Song L, Wang, L, Wang D, Anti-Tumor functional factor in creamery,
Chinese Health Food, 2001, 4:5-7.
Manuscripts under review:
1. Lei Wang, Eun Jin Lim, Michal Toborek, Bernhard Hennig. The role of fatty acids and
caveolin-1 in TNF-α-induced endothelial cell activation. (submitted to Metabolism in
November 2007).
2. Yuanyuan Zheng, Eun Jin Lim, Lei Wang, Eric J. Smart, Michal Toborek, Bernhard
Hennig. Role of caveolin-1 in EGCG-mediated protection against linoleic acid-induced
endothelial cell activation. (submitted to Journal of Nutritional Biochemistry in October
2007).
128
129
Abstracts and Presentations
1. Lei Wang, Michal Toborek Bernhard Hennig. Omega-3 but not omega-6 fatty acids
protect against TNF-alpha induced endothelial cell activation. The FASEB
Journal, April, 2006: B240.
2. Lei Wang, M. Toborek and B. Hennig. Different ratios of omega-6 to omega-3 fatty
acids can modulate the pro-inflammatory response in endothelial cells mediated
by TNF-α. Journal of the American College of Nutrition, 24: 426, 2005.
3. Lei Wang, Reiterer Gudrun, Michal Toborek Bernhard Hennig. Different ratios of
omega-6 to omega-3 fatty acids can modulate proinflammatory events in
endothelial cells induced by TNF-alpha and environmental contaminants. The
FASEB Journal, April, 2005 19(4): A442.
4. Viswanathan Saraswathia, Lei Wang, Michal Toborek, Bernhard Hennig. Linoleic acid
activates vascular endothelial cells via peroxynitrite formation. The FASEB
Journal, April, 103.6 2004.
5. Wang, L, Bai Sh, Shi J. Effect of dietary n-3 polyunsaturated fatty acid on Ca
metabolism. Annals of Nutrition & Metabolism,Official Journal of the
Federation Of European Nutrition Societies, 2001,45: 92.
6. 2002 Essential Fatty Acids and Human Nutrition and Health International Conference,
Shanghai, China. Apr. 2002. “The prevention of osteoporosis by dietary fatty
acids.”
7. Superfund Annual meeting, Seattle, Washington. Nov. 2004. “Different ratios of
omega-6 and omega-3 fatty acids can mediate PCB toxicity in endothelial cells.”
8. PCB Workshop, University of Illinois at Urbana-Champaign. 2004. “Nutrient-
mediated protection of PCB toxicity in endothelial cells.”
9. The 17th International Congress of Nutrition, Vienna, Austria. Aug. 2001. “Effect of
dietary n-3 polyunsaturated fatty acid on lipid metabolism in the rat of micro-
gravity model.”