Characterization of Adipokine-Induced Responses for Inflammation and
Leukocyte Interaction in Endothelial Cells
A dissertation presented to
the faculty of
the College of Arts and Sciences of Ohio University
In partial fulfillment
of the requirements for the degree
Doctor of Philosophy
Manindra Singh
August 2017
© 2017 Manindra Singh. All Rights Reserved. 2
This dissertation titled
Characterization of Adipokine-Induced Responses for Inflammation and
Leukocyte Interaction in Endothelial Cells
by
MANINDRA SINGH
has been approved for
the Program of Molecular and Cellular Biology
and the College of Arts and Sciences by
Fabian Benencia
Associate Professor of Immunology
Robert Frank
Dean, College of Arts and Sciences 3
ABSTRACT
SINGH, MANINDRA, Ph.D., August 2017, Molecular and Cellular Biology
Characterization of Adipokine-Induced Responses for Inflammation and
Leukocyte Interaction in Endothelial Cells
Director of Dissertation: Fabian Benencia
Without a doubt, the epidemic of obesity is one of the most serious health problems that affects millions of people worldwide and increases the risk of premature death. Studies indicate that chronic, low-grade inflammation is a major contributor of the obesity-associated pathogenesis, and therapeutic measures that target this process have been envisioned to be promising anti-obesity therapies.
Adipokines – biologically active molecules secreted from adipose tissue – have been described as potential candidates for therapy given their capability to regulate numerous physiological responses, both at local and systemic levels. However, the mechanism by which adipokines regulate inflammatory processes, specifically in the vascular compartment of adipose tissue, is poorly understood. With a goal to provide evidence for their therapeutic value as well as to understand the mechanism by which adipokines regulate intercellular crosstalk between various signaling pathways, herewith we characterize in detail the regulatory role of an adipokine visfatin for inflammatory activation, leukocytosis (recruitment of leukocytes), and morphogenic responses in endothelial cells. It is hypothesized that visfatin treatment can activate endothelial cells to upregulate the secretion of chemokines and expression of inflammatory markers resulting in an enhanced 4 leukocyte interaction. Consistently, we report that visfatin-treated endothelial cells exhibit an increased pro-inflammatory expression profile, enhanced chemokine secretion, and a pronounced capacity for leukocyte migration and attachment. We also evaluated the angiogenic capability of visfatin and report visfatin-mediated upregulation of multiple angiogenesis-associated genes and endothelial cells capillary-like tube formation in vitro. Furthermore, we report that visfatin-stimulated pro-inflammatory expression and chemokine secretion is predominantly mediated by the components of MAP kinase and NFκB signaling pathways. Finally, we show that pharmacological inhibition of visfatin can suppress the secretion of inflammatory chemokines by human endothelial cells. Thus, this work provides a detailed characterization of visfatin mediated responses that lead to inflammation in endothelial cells and provide further evidence for its potential role as a therapeutic target to reduce disease progression. Future in vivo studies are required to further delineate the physiological effect of adipokines for their regulation of metabolic and inflammatory functions. Towards this, we provide a qualitative profile for adipokine-responsive cytokine secretion as preliminary data that, hopefully, will be extended to the murine system.
5
DEDICATION
To all teachers and advocates of science, especially my parents
6
ACKNOWLEDGMENTS
First and foremost, I am truly indebted and deeply thankful to my adviser,
Dr. Fabian Benencia, for his extensive guidance and support towards all the aspects of my research and career development. Thank you very much for being my mentor and allowing me to work in your laboratory, where I developed my scientific aptitude in immunology and fascination for the complexities of the immune system. I am thankful to the members of my dissertation advisory committee: Dr. John Kopchick, Dr. Robert Colvin, and Dr. Darlene Berryman, for their valuable comments and advice towards the design of my dissertation research project. Additionally, I express my gratitude to Dr. Ramiro Malgor, Dr.
Monica Burdick, Dr. Kelly McCall, Dr. Vishwajeet Puri, and Dr. Kevin Lee, for their helpful advice with my experiments and numerous scientific discussions on various occasions. Furthermore, I owe sincere and earnest thankfulness to Dr. Rita
Sharma and Dr. Vishwa Sharma for providing their continuous availability to help me troubleshoot my experiments and sharing reagents and experimental protocols that were integral to this research. I would also like to thank Michele Pate, Dr. Maria
Cecilia Courregges, and Dr. Harrison Muturi, for their support with molecular biology experiments. Sincere thanks are extended to my graduate and research advisers in India, Dr. Seemi Farhat Basir and Dr. Nasheeman Ashraf, to motivate me towards a career in scientific research. I also would like to thank Dr. Sarah
Wyatt for her amazing enthusiasm and guidance during the scientific writing course that immensely helped me improve my technical writing and grant preparation 7 skills. Additionally, I also thank Dr. Roxanne Male’-Brune for her unfailing support and assistance with various internal funding opportunities, and the Student
Enhancement Award and John J. Kopchick fellowship for funding this research.
Further, I am also grateful to all my colleagues, especially Dr. Maria Muccioli, Dr.
Grady Carlson, Dr. Reetobrata Basu, Dr. Amrita Basu, Dr. Sukanta Jash, Dr.
Sayani Banerjee, Ian Ackers, Quyen Luong, Gunjan Vidwans, and Samantha
Shaw, for the scientific discussions and friendships that made my Ph.D. journey cheerful. I am eternally thankful to my parents for their continuous support and extended involvement in my education. Most of all, I thank Margaret, my wife, critic, and my moral support, for the all her help and unparalleled sustenance that was instrumental throughout this research.
Thank you all for the encouragement!
8
TABLE OF CONTENTS
Page
Abstract…………………………………………………………………………………..3
Dedication………………………………………………………………………………..5
Acknowledgments…………………………………………………………..…………..6
List of Tables……………………………………………………………………………13
List of Figures…………………………………………………………………………..14
List of Abbreviations……………………………………………………………………19
Chapter 1: Introduction to Obesity-Associated Inflammation and Metabolic
Dysfunction .....……………………………………………………………..…...……..24
1.1 Obesity – A Global Epidemic and an Inflammatory Disease………….24
1.2 Adipose Tissue…………………………………………………………….25
1.2.1 Adipose tissue – Its Composition and Response in
Obesity……………………………………………………….…………25
1.2.2 Obesity-Induced Immunological Changes in Adipose
Tissue…………………………………………………………….……..28
1.2.3 Immunoinfiltration of the Adipose Tissue.…………….……..30
1.2.4 Inflammatory Signaling in Obesity...………………..………..34
1.3 Adipose Microenvironment and Endothelial Cells………………………39
1.3.1 Endothelial Inflammation and Angiogenic Responses……..40
1.4 Adipokines: The Signals from Fat and Role in Inflammation………….45
1.5 Hypotheses and Specific Aims………...…………………………………56 9
1.6 Summary of Project Significance………………..……………………….57
Chapter 2: Adipokine Treatment Upregulates Pro-Inflammatory Gene Expression,
Chemokine Secretion, and Enhanced Monocyte Interaction in Human Endothelial
Cells ……………………………………………………………..……………………...59
2.1 Abstract……………………………………………………………………..59
2.2 Introduction…………………………………………………………………60
2.3 Materials and Methods…………………………………………………….62
2.3.1 Culture and Maintenance of Endothelial Cells…...………….62
2.3.2 Adipokine Treatments..………………………………………..63
2.3.3 RNA Isolation…………………………………………………...64
2.3.4 Reverse-Transcriptase PCR (RT-PCR) …………………..…65
2.3.5 Real-Time Quantitative PCR (RT-qPCR)……………..……..65
2.3.6 Enzyme-Linked Immunosorbent Assay (ELISA)……………66
2.3.7. Antibody Microarray……………………………………………66
2.3.8 Leukocyte Transmigration Assay..…………………………...67
2.3.9 Leukocyte Adhesion Assay……………………………………68
2.3.10 Microscopy and Image Analysis………………………………69
2.3.11 Statistical Analysis …………………………………………….69
2.4 Results……………………………………………………………………...70
2.4.1 Adipokine Treatment Increases the Secretion of Pro-
Inflammatory Cytokines and Chemokines in Human Endothelial
Cells ..…………………………………………………………………...70 10
2.4.2 Visfatin Increases the Expression of Pro-Inflammatory Genes
and Chemokine Secretion in Human Endothelial Cells…………….76
2.4.3 Visfatin Treatment of Human Endothelial Cells Results in
Enhanced Chemoattractant Capacity for Monocyte Transmigration
…………………………………………………………………………...83
2.4.4 Visfatin Increases the Gene Expression of Adhesion Markers
in Human Endothelial Cells…………………….……………………..86
2.4.5 Treatment of Human Endothelial Cells with Visfatin Results in
an Enhanced Capacity for Monocyte Adhesion.……………………88
2.5 Discussion………………………………………………………………….90
Chapter 3: The Adipokine Visfatin Promotes Angiogenic Responses in Human
Endothelial Cells…………………..…………………………………………………...98
3.1 Abstract ……………………………...……………………………………..98
3.2 Introduction…………………………………………………………………98
3.3 Materials and Methods…………………………………………………..100
3.3.1 Human Endothelial Cell Culture and Treatments………….100
3.3.2 Real-Time Quantitative PCR (RT-qPCR)…………………..101
3.3.3 SDS-PAGE and Gelatin Zymography……..………………..102
3.3.4 Endothelial Cells Vascular Tube Formation Assay…….....103
3.3.5 Fluorescence Microscopy and Image Analysis…...……….104
3.3.6 Statistical Analysis ……...……………………………………104
3.4 Results……………………...……………………………………………..105 11
3.4.1 Visfatin Upregulates the Expression of Angiogenic Markers in
Human Endothelial Cells ..…………………………………………..105
3.4.2 Visfatin Enhances the Production of Functional Matrix
Metalloproteases in Human Endothelial Cells...…………………...107
3.4.3 Visfatin Shows an Angiogenic Effect and Causes an Early
Induction of Vascular Tube Formation in Human Endothelial
Cells…………………………………………………………………...109
3.5 Discussion……………….………………………………………………..113
Chapter 4: Identification of Signal Transduction Pathways Governing Visfatin-
Induced Pro-Inflammatory Expression in Human Endothelial Cells..……………118
4.1 Abstract …………………………………………………………………...118
4.2 Introduction ………………………………………………………….……118
4.3 Materials And Methods…………………………………………………..120
4.3.1 Cell Culture and Treatments.……………………….……….120
4.3.2 Pharmacological Inhibitors….....…………….………………121
4.3.3 Cell-Based Enzyme-Linked Immunosorbent Assay….…...121
4.3.4 Real-Time Quantitative PCR (RT-qPCR)…………………..122
4.3.5 Enzyme-Linked Immunosorbent Assay (ELISA)……...…..123
4.3.6 Statistical Analysis …………………………………………...124
4.4 Results…………………………………………………………………….124
4.4.1 Visfatin Treatment Activates ERK1/2 MAP-Kinase Pathway in
Human Endothelial Cells…………………………………………….124 12
4.4.2 Visfatin Treatment Upregulates Multiple Signaling Pathways
to Stimulate the Gene Expression of Pro-Inflammatory Chemokines,
Adhesion Markers, and Angiogenic Markers in Human Endothelial
Cells…………………………………………………………………...127
4.4.3 Pharmacologic Inhibition of Visfatin Suppresses Pro-
Inflammatory Chemokine Secretion in Human Endothelial Cells..146
4.5 Discussion………………………………………………………………...149
Chapter 5: Conclusion and Recommendation for Future Work….………………156
5.1 Summary of Results and Relevance of Study………….……………..156
5.2 Future Directions and Preliminary Data in Murine System.….………161
References ……………………………………………………………………………170
Appendix 1: Sequences of the Primers Used for RT-qPCR……………………...201
Appendix 2: Antibody Microarray…….……………………………….…………….203
Appendix 3: Luciferase Assay for Transcription Factor Activation…..…………..204
13
LIST OF TABLES
Page
Table 1.1 Key adipokines, their sources, and function in health and disease……47
Table 2.1 Endothelial and monocytic cells chosen for adipokine treatment and analyses...………………………………………………………………………………61
Table 4.1 Molecular pathway inhibitors used in the study and their mode of inhibition……………………………………………………………………………….120
14
LIST OF FIGURES
Page
Figure 1.1 Metabolic and immunological health complications of obesity………...25
Figure 1.2 Composition of white adipose tissue…………………………………….28
Figure 1.3 Phenotypic and functional changes in visceral white adipose tissue with respect to obesity………………………………………………………………………33
Figure 1.4 Inflammatory signaling in obesity………………………………………...36
Figure 1.5 The multistep process of leukocyte extravasation……………………...42
Figure 1.6 Steps involved in inflammation-driven angiogenesis…………………..44
Figure 1.7 An overview of physiological functions regulated by adipokines secreted from the adipose tissue……………………………………………….……………….46
Figure 1.8 Positive association between visfatin and medical conditions………...55
Figure 1.9 Diagrammatic hypothesis…………………………………………………57
Figure 2.1 Qualitative cytokine secretion profile of human endothelial cells
(HUVEC) in response to adipokine treatments...... ………………………………....72
Figure 2.2 Visfatin treatment promotes the secretion of multiple chemokines and cytokines in HUVEC in a dose-dependent manner………….…………...... ……….73
Figure 2.3 Leptin treatment promotes the secretion of multiple chemokines and cytokines in HUVEC...... ……………………………………….74
Figure 2.4 Secretion of pro-inflammatory cytokines and chemokines by HUVEC in response to vaspin treatment…………….………………………………………...…75 15
Figure 2.5 Pretreatment with vaspin suppresses tumor necrosis factor-alpha- induced cytokine secretion by HUVEC………………………………………………76
Figure 2.6 Visfatin increases the expression of chemokine ligand-2 (CCL2) in
HUVEC in a dose and time-sensitive manner.………………………………………78
Figure 2.7 Visfatin increases the expression of granulocyte-macrophage colony stimulating factor (GMCSF) in HUVEC in a dose and time-sensitive manner…..78
Figure 2.8 Visfatin increases the expression of interleukin-6 in HUVEC in a dose and time-sensitive manner………………………………………………….…………79
Figure 2.9 Visfatin increases the expression of CXC-chemokine ligand 2 (CXCL2) in HUVEC in a dose and time sensitive manner……………………………….……80
Figure 2.10 Visfatin increases the expression of CXC-chemokine ligand 8 (CXCL8) in HUVEC in a dose and time sensitive manner…………………………………….80
Figure 2.11 Visfatin stimulates enhanced secretion of the chemokine CXCL2 by
HUVEC in a dose and time-dependent manner…………...... 82
Figure 2.12 Visfatin stimulates enhanced secretion of the chemokine CXCL8 by
HUVEC in a dose and time-dependent manner…………………….…...... 82
Figure 2.13 Human monocytes (THP1) exhibit an enhanced migration capability towards visfatin-treated HUVEC culture supernatants……………………………..85
Figure 2.14 Visfatin treatment increases the expression of cell adhesion markers in HUVEC……………………………………………………………………………….87
Figure 2.15 Visfatin treated HUVEC exhibit an enhanced capacity for THP1 monocyte adherence…………………………………………………………………..89 16
Figure 3.1 Visfatin upregulates the expression of multiple angiogenic markers in
HUVEC ….…………………………………………………………………………….106
Figure 3.2 Visfatin treatment enhances the production of functional matrix metalloproteases (MMPs) in HUVEC in a time-dependent manner……………..108
Figure 3.3 Quantification of visfatin-stimulated gelatinolytic activity of MMPs….109
Figure 3.4 Vascular tube formation assay showing a capillary-like network in
HUVEC cultured on MatrigelTM…….………………………………………………...111
Figure 3.5 Analysis of vascular features in HUVEC after 6 hours visfatin treatment………………………………………………………………………………111
Figure 3.6 Visfatin treatment stimulates a rapid vascular tube formation in HUVEC cultured on MatrigelTM………………………………………………………………..112
Figure 3.7 Quantification of visfatin-induced vascular features in HUVEC…..….112
Figure 4.1 Visfatin stimulates the activation of ERK1/2 MAP-Kinase in HUVEC.126
Figure 4.2 Cell proliferation assay to determine the effect of pharmacological inhibitor treatment reveals no cytotoxicity within the tested dose range…………129
Figure 4.3 Visfatin upregulates the expression of CCL2 via NFκB, p38-MAPK, and
JNK signaling pathway in HUVEC…………………………………………………..131
Figure 4.4 Visfatin-stimulated upregulated expression of GMCSF in HUVEC is mediated by NFκB, p38-MAPK, and JNK signaling pathway…………………..…132
Figure 4.5 Visfatin-stimulated increase in the expression of IL6 in HUVEC is mediated by NFκB, p38-MAPK, and PI3K pathways……………………………...133 17
Figure 4.6 Visfatin increases the expression of CXCL2 in HUVEC via NFκB, and p38-MAPK pathways…………………………………………………………………134
Figure 4.7 Visfatin-stimulated increase in the expression of CXCL8 in HUVEC is mediated via p38-MAPK pathway…………………………………………………..135
Figure 4.8 Visfatin increases the secretion of chemokines CCL2, CXCL2, and
CXCL8 is regulated by p38-MAPK and NFκB pathways…………………………137
Figure 4.9 Visfatin upregulates the expression of E-selectin in HUVEC involves
NFκB, p38-MAPK, JNK, and ERK-MAPK pathways………………………………139
Figure 4.10 Visfatin-stimulated increase in the expression of intercellular cell adhesion molecule (ICAM1) in HUVEC involves NFκB and p38-MAPK pathways………………………………………………………………………………140
Figure 4.11 Visfatin-stimulated increase in the expression of vascular cell adhesion marker 1 (VCAM1) in HUVEC is mediated by NFκB and p38-MAPK pathways………………………………………………………………………………141
Figure 4.12 Visfatin upregulates the expression of platelet-endothelial cell adhesion molecule 1 (PECAM1) in HUVEC via p38-MAPK pathway……………142
Figure 4.13 Visfatin-induced expression in the levels of Angiopoietin 1 (ANGPT1) in HUVEC involves multiple signaling pathways…………………………………..144
Figure 4.14 Visfatin-stimulated upregulation for the expression of Angiopoietin 2
(ANGPT2) in HUVEC involves NFκB, p38-MAPK, JNK, and ERK-MAPK pathways………………………………………………………………………………145 18
Figure 4.15 Pathway regulation of visfatin-induced expression of inflammation- associated genes in HUVEC………………………………………………………...146
Figure 4.16 Cell proliferation assay to test the cytotoxic effect of pharmacological inhibitor FK866 reveals no adverse effect in HUVEC for the tested doses..……147
Figure 4.17 FK866-mediated inhibition of visfatin suppresses the chemokine secretion in HUVEC…………………………………………………………………..148
Figure 5.1 Diagrammatic model of visfatin-induced inflammatory responses in endothelial cells……………………………………………………………………….157
Figure 5.2 Summary of conclusions………………………………………………...161
Figure 5.3 Preliminary data in the murine system – differential regulation of cytokine and growth factor secretion in response to treatment with recombinant visfatin in murine endothelial cells (MS1) ……………………………………...... 166
Figure 5.4 Preliminary data in the murine system – differential regulation of cytokine and growth factor secretion in response to treatment with recombinant leptin in murine endothelial cells (MS1) ……………………………………...... 167
Figure 5.5 Preliminary data in the murine system – differential regulation of cytokine and growth factor secretion in response to treatment with recombinant vaspin in murine endothelial cells (MS1) ……………………………………...... 168
Figure A2.1 Layouts of the antibody arrays used for the simultaneous detection of cytokines and angiogenic molecules……………………………….………………203
Figure A3.1 Schematic representation of the Dual-Luciferase® Reporter Assay
System (Promega)……………………………………………………………………204 19
LIST OF ABBREVIATIONS
ABTS 2,2'-azino-bis (3-ethylbenzothiazoline-6-sulphonic acid)
Akt Protein Kinase B
AMPK 5' adenosine monophosphate-activated protein kinase
ANGPT Angiopoietin
ANGPTL Angiopoietin-Like Protein
ANOVA Analysis of Variance
AP1 Activator Protein 1
ATM Adipose Tissue Macrophages
ATP Adenosine triphosphate
BMAT Bone Marrow Adipose Tissue
BSA Bovine Serum Albumin
Casp-1 Caspase-1
CCL Chemokine CC-motif Ligand
CD Cluster of Differentiation
CDC Center for Disease Control
CRP C-Reactive Protein
CXCL CXC Chemokine-motif Ligand
DIO Diet-Induced Obesity
DMEM Dulbecco's Modified Eagle Medium
DMSO Dimethyl Sulfoxide 20
EBM Endothelial Basal Medium
EC Endothelial Cells
ECM Extracellular Matrix
EGM Endothelial Growth Medium eIF2α Eukaryotic-translation Initiation Factor 2α
ELISA Enzyme-Linked Immunosorbent Assay
ERK Extracellular signal-Regulated Kinases
FA Fatty Acids
FBS Fetal Bovine Serum
GAPDH Glyceraldehyde 3-phosphate dehydrogenase
GLUT Glucose Transporter
GMCSF Granulocyte-Macrophage Colony-Stimulating Factor
GRO Growth Related Cytokine
HFD High Fat Diet
HUVEC Human Umbilical Vein Endothelial Cells
ICAM Intercellular Cell Adhesion Molecule
IFNγ Interferon-gamma
IGF-1 Insulin-like Growth Factor-1 21
IκBα Inhibitor of NFκB
IKKα/β IκBα Kinase
IL Interleukin
IR Insulin Receptor
IRAK-1 Interleukin-1 Receptor-Associated Kinase-1
IRF Interferon Regulatory Factor
IRS Insulin Receptor Substrate
JAK-STAT Janus Kinase-Signal Transducer and Activator of Transcription
JAM Junctional Adhesion Molecule
JNK c-Jun N-terminal kinases
LPS Lipopolysaccharide
MAPK Mitogen-Activated Protein Kinases
MCP1 Monocyte Chemoattractant Protein-1
MEK Mitogen-Activated Protein Kinase Kinase
MMP Matrix Metalloproteinases
MS1 Mile Sven1 mTOR mammalian Target of Rapamycin
MyD88 Myeloid Differentiation Primary Response Gene 88
NAMPT Nicotinamide Phosphoribosyltransferase
NCHS National Center for Health Statistics
NFκB Nuclear Factor κB 22
NLRP3 NACHT, LRR and PYD domains-containing protein 3
NIH National Institutes of Health
NK Natural Killer Cells
NLR Nucleotide Oligomeric Domain (NOD)-like Receptor
NMN Nicotinamide Mononucleotide
OLETF Otsuka Long-Evans Tokishima Fatty rats
PAI-1 Plasminogen Activator Inhibitor-1
PBEF Pre-B-cell colony-Enhancing Factor
PBS Phosphate Buffered Saline
PECAM Platelet-Endothelial Cell Adhesion Molecule
PI3K Phosphatidylinositol-4,5-bisphosphate 3-Kinase
PKR Protein Kinase R
PV1 Plasmalemma Vesicle Protein 1
PRR Pattern Recognition Receptor
RBP4 Retinol Binding Protein 4
RNA Ribonucleic Acid
ROS Reactive Oxygen Species
RPMI Roswell Park Memorial Institute medium
RT-PCR Reverse Transcriptase-Polymerase Chain Reaction
RT-qPCR Real Time-quantitative Polymerase Chain Reaction
SDS-PAGE Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis 23
SFRP5 Secreted Frizzled-Related Protein 5
SVF Stromal Vascular Fraction
S6K S6 Kinase
TGFβ Transforming Growth Factor-beta
THP1 Human leukemic monocytic cell line (Peripheral Blood)
TIE2 Tyrosine kinase with Immunoglobulin-like and EGF-like domains2
TLR Toll-Like Receptor
TNFα Tumor Necrosis Factor-alpha
TRIF TIR-domain-containing adapter-inducing interferon-β
T2DM Type 2 Diabetes Mellitus uPA urokinase-type Plasminogen Activator uPAR uPA Receptor
Vaspin Visceral Adipose-tissue derived Serpin
VCAM1 Vascular Cell Adhesion Molecule 1
VE-Cadherin Vascular Endothelial-Cadherin
VEGF Vascular Endothelial Growth Factor
WAT White Adipose Tissue
WISP-1 WNT1-Inducible-Signaling Pathway protein 1
24
CHAPTER 1: INTRODUCTION TO OBESITY-ASSOCIATED INFLAMMATION
AND METABOLIC DYSFUNCTION
1.1 Obesity – A Global Epidemic and an Inflammatory Disease
Obesity is a global epidemic that affects individuals irrespective of their age, gender, and demographics, and leads to premature mortality. According to the
World Health Organization, over 300 million people were obese worldwide (1), and as per National Health and Nutrition Examination Survey (CDC/ NCHS), 35.7%
U.S. adults (78 million) and 16.9% (12.5 million) children and adolescents were obese in 2009-2010 (2). With a steep rise in the prevalence of obesity among both adults and children in the last decade, the healthcare sector is currently facing a big challenge to manage the financial and pathologic burden of obesity. Obesity is a strong, positive risk factor for diverse pathologies, including insulin resistance and type 2 diabetes, atherosclerosis, non-alcohol fatty liver disease, arthritis, immune system dysfunction, cardiovascular disorders, and some cancers (3).
Figure 1.1 illustrates a cluster of major immunological and metabolic complications associated with obesity. Owing to the association to these deadly diseases and their prevalence, it is not surprising that globally, more people die of obesity and overweight than hunger. It is therefore of utmost importance to devise anti-obesity therapies to reduce its deleterious effect on human health.
25
Figure 1.1 Metabolic and immunological health complications of obesity. Adapted from (4). Obesity is a leading risk factor for a plethora of medical complications. As shown in the diagram above, obesity affects a wide range of biological functions by its negative effect at the level of all major organs, bones, and cardiovascular system. Additionally, obese individuals are at a higher risk for several cancers.
1.2 Adipose Tissue
1.2.1 Adipose Tissue – Its Composition and Response in Obesity
Obesity is essentially a state of excessive fat accumulation caused by a disruption of energy balance due to the positive caloric intake. While some genetic factors may be involved, the environmental factors, such as nutrient-rich diets and sedentary lifestyle, are the major contributor for a predisposition to obesity- 26 associated metabolic complications (5). Dysfunction at the level of white adipose tissue (WAT) is one of the major determinants of the development of obesity- associated metabolic complications (6). As illustrated in Figure 1.2, WAT is a highly complex organ and based on its composition, can be divided in an adipocyte fraction, containing mature adipocytes, and a stromal vascular fraction (SVF) that is composed of pre-adipocytes, vascular cells, mesenchymal stem cells
(fibroblasts), and immune cells (7). WAT is distributed in various anatomical compartments referred to as ‘depots’; and WAT depots are classified into two major classes: subcutaneous (under the skin) and intra-abdominal (located within the abdominal cavity lining the internal organs) (8). The inguinal fat depot that is characterized by inguinal lymph node is the major representative of subcutaneous
WAT (sWAT). Another sWAT depot may include subscapular WAT. Intra- abdominal WAT is further classified into omental, mesenteric, perirenal, gonadal, and retroperitoneal depots, and together with epicardial fat depot, constitute visceral WAT (vWAT) (8, 9). In mice, mesenteric fat pad is the true vWAT that is present in the close association with the intestines (8). The perirenal and retroperitoneal fat pads are located below and around, and behind the kidneys, respectively. In male mice, epidydimal fat pads are typically the largest and most often studied owing to the ease of its isolation (8). The female equivalent of this fat pad is periovarian fat and surrounds the ovaries. Both epidydimal and periovarian fat pads are termed as gonadal (or perigonadal) fat depots. In addition to sWAT and vWAT, bone marrow is another site for fat accumulation and constitute bone 27 marrow adipose depot (BMAT). BMAT can constitute upto 70% of the bone marrow volume in adult humans (10). Interestingly, caloric restriction increases the BMAT expansion in animals, including humans (11). Recently, Cawthorn et al. described the function of BMAT as an essential endocrine organ capable of contributing to serum adiponectin levels under caloric restriction (12). Sufficient evidence suggests that individual WAT depots from different anatomical locations have different functions, including varying secretory profile, inflammatory status, and their contribution to disease. WAT performs several metabolic functions under the normal physiological state. Consequent to excess calorie intake, WAT undergoes extensive remodeling and vascular changes, and expands in biomass due to hypertrophy (adipocyte expansion) and hyperplasia (increase in adipocyte proliferation). Positive caloric imbalance and obesity leads to uncontrolled expansion of WAT and impairment of its normal architecture that eventually causes its ectopic expansion into other anatomical locations (e.g. liver, heart etc.) thereby affecting normal function (13). This phenomenon is called ‘lipotoxicity’ and is one of the initial features towards organ dysfunction in obesity. Over the course of time, obesity-associated changes (discussed in the following section) leads to an imbalance in metabolic and homeostatic functions leading to disease development. 28
Figure 1.2 Composition of white adipose tissue (WAT). Adapted from (7). WAT is highly heterogeneous and is composed of different cell populations. It can be divided into: (1) an adipose fraction containing mature adipocytes, which are the principal cellular component of WAT and are crucial for energy storage and secretory activity, and (2) stromal vascular fraction (SVF) containing pre- adipocytes, immune cells, vascular cells, and fibroblasts. ECM derived from fibroblasts is essential for mechanical strength and integrity of WAT. Intercellular communication between adipocytes and cells of SVF are critical to maintain homeostasis in adipose microenvironment and in the body.
1.2.2 Obesity-Induced Immunological Changes in Adipose Tissue
Increasing evidence suggests that reciprocal interactions between metabolic components and immune cells have an important role in obesity- associated disease development. Obesity is marked by the significantly enhanced serum levels of pro-inflammatory factors and infiltration of vWAT by immune cells 29
(1, 7). The initial, specific trigger for the recruitment of leukocytes to tissue is not known, albeit it is possible that metabolic imbalance coupled with pro-inflammatory factors in adipose tissue microenvironment can be responsible for this effect.
Progressive enlargement of adipose tissue results in the fibrosis (accumulation of excess connective tissue) and loss of its elasticity thereby impeding its ability to store lipids. Over time continued lipogenesis and fibrosis lead to a state of metabolic imbalance where cellular oxidative pathways for lipid metabolism become saturated resulting in a ‘lipid spillover’ (1). The resultant shuttling of lipids into non-oxidative metabolic pathways leads to the production of inflammatory precursors, such as ceramides and sphingosines, and their release in the bloodstream (6, 14). Continued expansion of WAT, increased fibrosis, and lack of oxygen are associated with endoplasmic reticulum stress, and considerable adipocyte death and release of the ‘danger signals’ such as ATP and uric acid, which are potent inflammatory signals and contribute to immunoinfiltration (1, 6,
14). Studies indicate that saturated fatty acids released from necrotic adipocytes and secretion of chemotactic molecules have a strong capacity to attract immune cells, which are initially recruited to WAT to suppress lipolytic signals and promote lipid uptake (15, 16). In this process, immune cells may become ‘activated’ and contribute to the production of inflammatory mediators. One of the first discoveries for obesity-associated inflammation was the identification of significantly higher levels of pro-inflammatory molecule tumor necrosis factor-alpha (TNFα) in WAT
(and adipocytes) of obese mice compared to lean controls (17). Since then, several 30 other reports confirmed that pro-inflammatory milieu in WAT can activate invading immune cells as well as other cells of SVF and lead to the secretion of various cytokines, C-reactive protein (CRP), IL6, IL1β, CCL2, and others, at both local as well as systemic levels, which are the predictive of insulin resistance and metabolic syndrome (14, 18–20). The intrinsic immune response that is induced by obesity and high-fat diet consumption (HFD) is markedly different than that elicited by infection with microbial pathogens and is characterized by chronic and low-grade inflammation, which lacks the characteristic signs of infection or injury-induced acute inflammation.
1.2.3 Immunoinfiltration of WAT
Immunoinfiltration is a process in which circulating leukocytes (i.e. immune cells) are recruited to WAT in response to the inflammatory signals. In the later stages of obesity, different types of immune cells can represent a predominant fraction of a dysfunctional adipose tissue. These may predominantly include macrophages, T-cells, and natural killer cells (NK) and are the major contributors of inflammation. CD8+ T-cells are important mediators of immunity and cytotoxicity and are among the immune cell population that is recruited to vWAT in the early stages of obesity. A three to four-fold increase in the number of CD8+ T-cells is found in the WAT of both obese humans and mice (21, 22). The increase in the number of CD8+ T-cells occurs at two weeks following 60% HFD that precedes the infiltration of macrophages to the AT, suggesting CD8+ T-cells might initiate the inflammatory response that predominate at the later stages of obesity (23). Upon 31 infiltration, CD8+ T-cells facilitate the recruitment of macrophages and secrete cytokines that cause macrophage activation. Mice deficient in CD8+ T-cells have fewer macrophages in their vWAT and lower serum levels of TNFα and IL6, suggesting the crucial role of CD8+ T-cells in adipose inflammation and insulin resistance.
+ CD4 Helper T-cells (TH) and Regulatory T-cells (Treg, a subset of CD4+ T cells with immunosuppressive functions) control several aspects of immune responses particularly through the secretion of specific cytokines. The specific functions mediated by the subsets of activated CD4+ T cells include activation of other immune cells (e.g. B-lymphocytes, cytotoxic T cells, and others), and non- immune cells, and suppression of immune reaction (24). The number of both anti- inflammatory TH and Treg is decreased in the vWAT of obese individuals. TH can be subdivided into three effector subtypes: (A) TH1 cells that secrete inflammatory cytokines, such as IFN-γ, (B) TH2 cells that secrete anti-inflammatory cytokines IL-
4 and IL-13, and (C) TH17 cells that produce IL-17. The ratio of TH1/ TH2 cells increases during HFD-induced obesity as IFN-γ secreting TH1 cells predominate the vWAT(22). Obesity is also associated with a reduction of Treg that regulate immuno-suppressive effects and secrete anti-inflammatory cytokines, such as IL4,
IL10, and IL13 (22, 25, 26). In wild-type lean mice, depletion of Treg cells causes the increase in the levels of insulin and pro-inflammatory cytokines in both the circulation and AT (26). On the other hand, upregulation of Treg proliferation causes an improvement of insulin sensitivity and secretion of the anti-inflammatory 32 cytokine IL-10 (26). So as animals become obese, the ratio of CD8+ T cells to CD4+
T cells increases and the depletion of immunosuppressive Treg favor the activation of pro-inflammatory responses and pathophysiology of the vWAT.
In the later stages of obesity, macrophages represent the principal immune cell population that infiltrates the vWAT, referred as adipose tissue macrophages
(ATM). The activation of ATM is dependent on the cytokine milieu in the WAT.
Stimulation with T-helper 1 (TH1) cytokines (e.g. interferon-γ (IFN-γ)) and lipopolysaccharide (LPS) leads to macrophage polarization into ‘classically activated’ M1 macrophage phenotype (27). M1 macrophages have capacity to mediate TH1 type immune response and produce pro-inflammatory cytokines, such as TNFα and IL6, whose elevated levels in obesity are positively correlated to insulin resistance (18). On the other hand, stimulation with TH2 cytokines, such as
IL-4 and IL-13, leads to the polarization of macrophages towards an ‘alternatively activated’ M2 phenotype, which suppresses the production of pro-inflammatory cytokine and promotes tissue repair and fibrotic responses (28). M2 macrophages produce anti-inflammatory cytokines, such as IL-10 that downregulate classical
NF-κB-dependent activation. ATM are likely to exist in various phenotypes along
M1/M2 axis depending on the fat-depot and the cytokine milieu, but the increase in adiposity tends to shift the macrophages towards M1 phenotype that contribute to the inflammatory status of vWAT in obesity (28). Interestingly, M1 macrophages have been shown to aggregate around necrotic adipocytes to form ‘crown-like structures’ [Figure 1.3], which is now considered a characteristic feature of 33 inflamed adipose tissue (29). Hence in obesity, the chronic inflammation is maintained by the pro-inflammatory output of vWAT and classically activated M1 macrophages. Other immune cells that infiltrate vWAT during the course of obesity include natural killer T-cells, mast cells, B-lymphocytes, and neutrophils, and they contribute to the inflammation suggesting a major role of immune system in the progression of metabolic syndrome and pathophysiology of obesity (26, 27).
Figure 1.3 Phenotypic and functional changes in vWAT with respect to obesity. Obesity leads to a progressive dysfunction at the level of vWAT characterized by loss of insulin sensitivity, increased immunoinfiltration, secretion of pro- inflammatory factors, and loss of metabolic control. Further, uncontrolled WAT expansion may result in hypoxia and impaired angiogenesis leading to adipocyte necrosis. Macrophages form aggregates around dying adipocytes forming ‘crown- like structures’ that is a characteristic feature of an inflamed vWAT. vWAT, visceral white adipose tissue; WAT, white adipose tissue. 34
Overall, obesity-associated dysregulation in metabolic responses promotes functional and compositional changes in adipose tissue marked by ectopic WAT expansion, adipocyte dysfunction, and immunoinfiltration, thereby causing a sustained metainflammation (a chronic, low-grade inflammatory response in obesity) in the obese individuals, thereby increasing their susceptibility to various pathologies. However, details regarding the interaction of immune cells and cellular components of the adipose tissue are still incomplete and may hold crucial information for the pathogenic processes. Dissecting the mechanism of this cellular crosstalk in adipose microenvironment is essential to determine therapeutically relevant information.
1.2.4 Inflammatory Signaling in Obesity
Contrary to the classical, acute immune response that is triggered primarily by the leukocytes in response to infection or tissue injury, the obesity-induced immune response is initiated and sustained by the metabolic cells e.g. adipocytes.
Nutrient overload, increasing metabolic demands at the level of adipose tissue, and lipotoxicity result in the production of factors that have inflammatory potential.
The major intracellular kinases that transduce nutrient signals into inflammation in metabolic tissues include inhibitor of κ kinase (IKK), c-Jun N-terminal kinase (JNK), and protein kinase-R (PKR) (32). These kinases have markedly increased activation and upregulation of their downstream pro-inflammatory targets in obese subjects compared to their lean counterparts (5, 32–34). In response to nutrient excess or inflammatory signaling – such as that induced by TNFα – JNK and IKK 35 are activated and inhibit insulin signaling by serine phosphorylation of insulin receptor substrate 1 (IRS1) thereby resulting in a reduced glucose uptake by target cells (35, 36). In addition to inhibiting insulin signaling, inflammatory kinases also activate transcriptional programs via the activation of their downstream effectors e.g. nuclear factor κB (NFκB), activator protein 1 (AP1), and interferon regulatory factor (IRF) resulting in an enhanced expression of pro-inflammatory cytokines. It is now well known that obesity increases the levels of several pro-inflammatory cytokines, (e.g. TNFα, IL6, CCL2/MCP1, IL1β, and others) not only in adipocytes but also in other metabolic tissues, such as liver (37), brain (38, 39), pancreas (40,
41), and skeletal muscle (42–44). These cytokines further induce the ongoing inflammatory signaling and further potentiate intracellular responses that interfere in insulin signaling. Figure 1.4 illustrates the schematic of obesity-induced inflammatory signaling.
36
Figure 1.4 Inflammatory signaling in obesity. The increased levels of metabolic signals (e.g. lipids) and pro-inflammatory cytokines (e.g. TNFα) can be sensed by PRRs and cytokine receptors resulting in the activation of multiple intracellular kinases and inflammasome. This leads to a downstream activation of factors that control inflammation, transcription, and organelle stress. Prolonged activation of inflammatory signaling interfere with proper insulin action by causing inhibitory modifications in IRS-1 or insulin receptor. Chronic activation of these pathways leads to inflammation and impairment of insulin sensitivity by multiple mechanisms. [Red arrows – negative regulation]. AP1, Activator protein-1; eIF2α, eukaryotic Initiation factor 2α; ER, Endoplasmic reticulum; ERK, Extracellular signal-regulated kinase; FA, Fatty acids; IκBα, Inhibitor of NFκB; IKK, Inhibitor of κB Kinase; IL, Interleukins; IRF, Interferon Regulatory Factor; IRS-1, Insulin Response Substrate-1; JNK, C-Jun N- terminal Kinase; mTORC, Mammalian target of Rapamycin; NFκB, Nuclear Factor κB; NLR, NOD-like receptor; PKR, Protein kinase R; S6K, S6 kinase; Ser-P, Serine phosphorylation; TLR, Toll-like receptor; Tyr-P, Tyrosine phosphorylation.
Studies indicate that pattern recognition receptors (PRRs) of the innate immune system have the capability to sense the metabolic signals (e.g. saturated fatty acids) to generate the activation of intracellular kinases that govern pro- 37 inflammatory responses. In this regard, different classes of toll-like receptors
(TLRs) have been extensively studied for their role in obesity-induced inflammation of metabolic tissues. Adipose tissue expresses nearly all TLRs; and TLR4 has gained the most attention because of its capability to sense free fatty acids (FA) and to generate inflammatory signals, and resulting in the downstream activation of inflammatory pathways (45). TLR4 deficient mice are protected from inflammatory activation and lipid-infused insulin resistance upon diet-induced obesity (DIO) (46). In a separate report, a TLR2 knockout in mice also results in a protection from insulin resistance and inflammatory disease progression in DIO
(47). Nod-like receptor (NLR) family of PRRs can also sense signals released from stressed and necrotic cells to facilitate leukocytosis to promote clearance and constrain tissue injury (48). In macrophages, NLR activation can stimulate NLRP3/ cryptopyrin inflammasome to induce caspase-1 mediated IL1β and IL18 production, contributing to pancreatic β-cell death and inflammatory disease progression (49, 50). The levels of caspase-1 and IL1β are increased in obesity, and consistently, mice deficient in Nlrp3 and casp-1 are resistant to DIO and inflammation (51), particularly due to a shift in the activation state of ATM (M2>M1) without an alteration in the total number. Pro-inflammatory cytokine receptor and
TLR-mediated downstream signaling may be mediated by the adaptor protein myeloid differentiation primary response gene 88 (MyD88) to activate multiple intracellular kinases that transduce metabolic signals into inflammatory functions.
IKKβ activation occurs downstream of MyD88 activation and is critical for the 38
NFκB-mediated inflammation of the liver, hypothalamus, and myeloid cells in obesity (34, 52).
Obesity-induced activation of stress-induced kinases (e.g. JNK) share common upstream mediators that also activate IKK/NFκB pathways in response to metabolic signals, cytokines, intracellular stress, and hyperglycemia (32). Of interest, JNK activation is well described in both immune cells and adipocytes in the context of obesity. Both JNK1 and JNK2 isoforms participate in metabolic regulation, energy expenditure, and body weight control, with JNK2 having a protective role in DIO (53). Hypothalamic inactivation of JNK1 results in the protection from DIO and weight loss in mice (54). Additionally, JNK1 inactivation in hematopoietic cells suppresses inflammation irrespective of adiposity and imparts improved metabolic profile under DIO (55). PKR, another stress kinase activated by metabolic and inflammatory signals, can negatively influence constitutive translation by inhibiting eukaryotic translation initiation factor 2α
(eIF2α) (56), thus further complicating intracellular responses to anabolic ligands and contributing to endoplasmic reticulum stress. In addition to JNK and IKK/NFκB, multiple kinases have also been described for their inhibitory role in insulin signaling and regulation of inflammatory expression in the context of obesity.
These include PKR, extracellular signal-regulated kinase (ERK), S6 kinase, and mammalian target of rapamycin (mTOR) that can contribute to insulin resistance by different mechanisms (57, 58). 39
Obesity induces a cluster of metabolic and inflammatory signals that may activate multiple signaling pathways in cells of both metabolic tissues and immune system. Thus, a single kinase system, one target, or a linear pathway that transduces metabolic signals to inflammatory responses is improbable, and a network of multiple interacting pathways likely operate to govern chronic inflammation in obesity.
1.3 Adipose Microenvironment and Endothelial Cells
WAT is a highly vascularized organ, and endothelial cells (EC) together with other vascular cells form a major portion of the vascular compartment. In WAT, EC support various important functions, including leukocyte distribution, nutrient circulation, lipogenesis, and adipose growth. EC from the omental WAT are reported to have increased expression of angiogenesis and inflammation associated genes and reduced expression of metabolic genes compared to those isolated from sWAT (59). EC are the first line of a barrier for the infiltrating leukocytes, and cross talk between different cells in the adipose microenvironment can influence the activation state of the EC leading to an enhanced interaction with leukocytes promoting their tissue infiltration (60). Indeed, obesity-induced pro- inflammatory responses in adipose microenvironment result in endothelial dysfunction, which further exacerbates the health of the adipose tissue. Details regarding the mechanism that regulate EC-leukocyte crosstalk and endothelial dysfunction in the adipose microenvironment are unclear and are under investigation. 40
1.3.1 Endothelial Inflammation and Angiogenic Responses
As mentioned earlier, inflammatory processes that are triggered due to metabolic dysfunction in WAT also contribute to the impairment of normal vascular function and activation of EC. Obesity causes vascular disturbances and reduction in capillary density in WAT, which results in a reduced nutrient supply and localized hypoxia in the adipose microenvironment (61, 62). More importantly, limited blood supply can trigger inflammation due to ischemia-induced adipocyte necrosis and lipolysis, which may facilitate the recruitment of macrophages to the WAT. In addition, obesity-induced downregulation of anti-inflammatory and vaso-protective molecules (e.g. adiponectin) with a simultaneous up-regulation of pro- inflammatory cytokines, like TNFα and IL6, further contributes to endothelial dysfunction (7). With inflammatory stimuli, activated EC exhibit a pronounced upregulation of cell adhesion markers that interact with immune cells and facilitate their ‘extravasation’ and immunoinfiltration into WAT (63–65).
The interaction between EC and leukocytes is a multi-step process, and can be divided into capture, rolling, arrest, crawling, and transendothelial migration of leukocytes into the tissue. Inflammatory mediators released at the site of injury can stimulate the upregulation and interaction of specific adhesion molecules on both
EC and leukocyte surface. The initial capture of circulating leukocytes is the result of leukocyte-endothelial interaction facilitated by selectins, a class of cell adhesion molecules expressed by both leukocytes and endothelial cells (Reviewed in (60,
66). L-selectin is expressed constitutively by leukocytes and is involved in the initial 41 capture and rolling, while P-selectin is overexpressed in response to pro- inflammatory stimuli by both activated platelets and inflamed EC and mediates early leukocyte recruitment (67). E-selectin is rapidly overexpressed (4 hours for peak expression) by EC in response to pro-inflammatory stimuli and potentiates the leukocyte recruitment under inflammatory conditions (68). Once the leukocyte is ‘captured’ over the endothelium, it begins to roll due to the force of blood flow.
Pro-inflammatory activation of EC also leads to an overexpression of cell adhesion molecules (e.g. ICAM1 and VCAM1) (69), which together with selectins and integrins lead to an arrest of leukocytes at the site of inflammation or injury.
Transendothelial migration of leukocytes (also called as extravasation or diapedesis) is a complex and least understood stage of the EC-leukocyte interaction. Although most transendothelial migration occurs through the endothelial junctions (paracellular), evidence indicates that leukocytes can cross the endothelial barrier through transcytosis (migration through the endothelial cells). Paracellular migration is predominantly mediated by molecules required to maintain endothelial integrity e.g. junctional adhesion molecules (JAM), platelet- endothelial cell adhesion molecule-1 (PECAM1), VE-Cadherins, and others (66,
67). Binding with surface ligands on leukocytes, intracellular signaling leads to VE- cadherin-mediated disruption in endothelial barrier facilitating the transmigration of the leukocyte in a paracellular fashion. Transcellular extravasation is mediated by a plasmalemma vesicle protein-1 (PV1, also PAL-E), which is a key element that form structures to cover fenestrae, stomata, and transendothelial channels (70). 42
Antibody-targeted blocking of PV1 was shown to delay the leukocyte transmigration through cultured EC (71). However, the exact mechanism for transcellular leukocyte migration and other participating molecules that contribute to this process are largely unknown. Figure 1.5 illustrates the steps involved in leukocyte extravasation.
Figure 1.5 The multistep process of leukocyte extravasation. Adapted from (66). Pro-inflammatory stimuli can result in the upregulation and overexpression of cell adhesion molecules that facilitate the interaction between circulating leukocytes and endothelium. Selectins mainly mediate the initial capture, followed by rolling and capture of the leukocyte because of multiple subsequent interactions facilitated by cell adhesion molecules (e.g. ICAM1 and VCAM1). Transendothelial migration (extravasation) is mediated by JAMs, mainly VE-Cadherin, which regulate paracellular leukocyte migration. PV1 is the key protein that mediate transcellular migration. ICAM1, Intercellular cell adhesion molecue-1; VCAM1, Vascular cell adhesion molecule-1; PECAM1, Platelet-endothelial cell adhesion molecule-1; JAM, Junctional adhesion molecules; PV1, Plasmalemma vesicle protein-1.
Angiogenesis or neovascularization is an essential physiological feature to enable an active tissue to cope with its increased metabolic and oxygen demand, 43 and factors that influence vascular growth are likely to play a crucial role in the homeostatic regulation of the corresponding tissue. Angiogenesis and adipogenesis are tightly coupled during embryonic development, and substantial evidence exists to suggest that these processes continue to reciprocally interact during adult life and are necessary to ensure proper maintenance of the adipose tissue (72). Interestingly, it has been reported that vasculature can regulate WAT expansion, and obese mice treated with anti-angiogenic reagents show reduced body weight with decreased adipose mass than the untreated controls (73).
Rapidly expanding WAT leads to inadequate nutrient supply to the tissue components and hypoxia, both of which are powerful triggers for angiogenic response, and lead to an upregulation of pro-angiogenic molecules in the adipose microenvironment (74). In response to inflammatory and pro-angiogenic factors
(e.g. VEGF), vessels nearest to the stimulus may undergo local destabilization and certain EC become morphologically specialized into highly polarized ‘tip cells’. Tip cells express VEGFR2 on their filopodia and have a strong capacity to sense
VEGF, and grow outward to form an angiogenic ‘sprout’ that is supported by the rapidly proliferating ‘stalk EC’ (75–77) [Figure 1.6]. If the inflammatory stimuli persist (as in chronic inflammation), the tip cells may invade the matrix to initiate a pro-angiogenic response and release proteolytic factors, e.g. matrix metalloproteinases (MMPs) for extracellular matrix degradation (78, 79). MMP- mediated ECM degradation further increases the availability of matrix-associated angiogenic molecules, such as VEGF, which together with cytokines can 44 synergistically act to result in a rapid growth of the ‘nascent sprout’ into neovessels
(75, 79, 80). However, in the absence of inflammatory stimuli, neovessels quickly regress and return to the original quiescent state.
Figure 1.6 Steps involved in inflammation-driven angiogenesis. (1) Local signals can lead to an activation and destabilization of the endothelium, resulting in the morphogenic changes, and (2) formation of specialized ‘endothelial tip cells’ that extend into the ECM leading to a nascent sprout. (3) Secretion of ECM modifying factors (e.g. MMPs) by the growing tip cell further increase the availability of cytokines and pro-angiogenic factors favoring growth of the neovessels. (4a) Vessels quickly regress to a quiescent state as soon as the signal disappears; (4b) persistence of inflammatory signals can further continue the angiogenic processes.
Overall, obesity-associated inflammatory changes at the level of adipose microenvironment leads to dysfunction of adipocytes and EC, and favor increased immunoinfiltration and a pro-inflammatory milieu, which further feed forward the 45 inflammatory responses in a ‘vicious cycle’ ultimately leading to a systemic release of pro-inflammatory mediators causing metabolic syndrome and insulin resistance.
1.4 Adipokines: The Signals from Fat and Role in Inflammation
WAT was long considered to be a storage organ with a few functions, including release of lipids under fasting conditions, thermoregulation, and mechanical protection. However, research in the last fifteen years has established its major role as an active endocrine organ. Mature adipocytes, preadipocytes, endothelial cells, and resident immune cells, together secrete a plethora of metabolites and bioactive proteins, called ‘adipokines’ that have a hormone-like signaling ability (7).
Although total human secretome of adipokines is not fully profiled, but it is considered that adipose tissue is a source of over 600 such proteins (81). At the systemic levels, adipokines can regulate a myriad physiological functions, such as hunger, blood clotting, insulin secretion, endothelial functions, inflammation, hypertension, and several others (7, 82, 83) [Figure 1.7]. A normal adipokine secretion ensures a fine balance between a ‘healthy’ crosstalk between WAT and other organs, including brain, liver, heart, lungs, muscles, vasculature, and immune system; however, abnormal adipokine production, as seen in obesity, can result in the disruption of homeostatic balance and pathological functions (82).
46
Figure 1.7 An overview of physiological functions regulated by adipokines secreted from the adipose tissue.
In context of immunity-associated functions, adipokines can be broadly categorized as pro-inflammatory or anti-inflammatory based on their function.
Obesity-induced vWAT dysfunction favors a dysregulation in the adipokine secretion, and favors an increased secretion of pro-inflammatory adipokines that can contribute to immunoinfiltration and insulin resistance. Indeed, obese individuals have higher levels of pro-inflammatory adipokines in their circulation
(20, 84, 85). HFD induces the blood levels of pro-inflammatory adipokines, such as TNFα, IL6, PAI-1, leptin and resistin, while simultaneously decreasing the anti- inflammatory adipokines e.g. adiponectin (13, 86). In addition to the classical adipokines – TNFα, IL6, leptin, several other adipokines have been identified that were associated with obesity-associated inflammation and insulin resistance.
These include resistin, retinol-binding protein 4, lipocalin 2, angiopoietin-like 47 protein 2, CC-chemokine ligand 2 (CCL2), CXC-chemokine ligand 5 (CXCL5),
NAMPT/visfatin, vaspin and others. Table 1.1 provides a list of key adipokines and their general role in health and disease. It is now well known that altered secretory profile of adipokines may link obesity to its inflammatory, metabolic, malignant, and cardiovascular pathologies, and their targeted ablation can be the basis of effective anti-obesity therapies.
Table 1.1
Key adipokines, their sources, and function in health and disease
Adipokine Source Receptor Function References
Adiponectin Adipocytes Adiponectin Insulin sensitizer, anti- (87) receptors 1 and 2, inflammatory T-cadherin, calreticulin–CD91
Adipsin Adipose tissue Unknown Activates the alternative (88) complement pathway
ANGPTL2 Adipocytes, Unknown Local and vascular inflammation (89, 90) other cells
CCL2/ SVF, CCR2 Monocyte recruitment (91, 92) MCP1 adipocytes
Chemerin Adipocytes Unknown Chemoattractant protein; (93–95) regulates adipogenesis
CXCL5 SVF, CXCR2 Antagonism of insulin signaling (96) macrophages through the JAK–STAT pathway
FGF21 Adipose tissue, FGF-Receptor Stimulates glucose uptake into (97–99) liver, skeletal adipocytes; increases muscle thermogenesis, energy expenditure, and fat utilization; improves glucose and lipid metabolism
IL18 SVF IL18 receptor, IL18 Broad-spectrum inflammation (100, 101) binding protein 48
Table 1.1: Continued
IL6 Adipocytes, IL6 Receptor Variable functions based on the (102–104) SVF, liver, tissue; pro-inflammatory muscle
Leptin Adipocytes Leptin Receptor Appetite control through the (105–108) central nervous system
Lipocalin-2 Adipocytes, Unknown Promotes insulin resistance and (109, 110) macrophages inflammation through TNF secretion from adipocytes
NAMPT/ Adipocytes, Unknown Monocyte chemotactic activity, (111, 112) Visfatin macrophages, angiogenesis other cells
Omentin Omental Unknown Anti-inflammatory; insulin (113, 114) adipose tissue sensitizing
PAI-1 Endothelial Unknown Inhibition of fibrinolysis (115, 116) cells, hepatocytes
RBP4 Liver, Retinol (Vitamin A), Implicated in systemic insulin (117–119) adipocytes, transthyretin resistance macrophages
Resistin PBMC (human), Unknown Promotes insulin resistance and (120–122) adipocytes inflammation through IL-6 and (rodent) TNF secretion from macrophages
SFRP5 Adipocytes WNT5a Suppression of pro-inflammatory (123) WNT signaling
TNFα SVF, TNFα Receptor Inflammation, antagonism of (36, 124, 125) adipocytes insulin signaling
Vaspin Adipose tissue Unknown Serine protease inhibitor; (126–128) decreases food intake; improves hyperglycemia
VEGF Adipose tissue, VEGF-Receptor Stimulates angiogenesis in (129–131) endothelial cells adipose tissue
WISP Adipocytes αvβ5 Integrin Regulates adipogenesis and (132–135) Receptor adipose tissue inflammation
49
In the present research, we have investigated the role of adipokines – visfatin, leptin, and vaspin – for their capacity to induce cytokine secretion in both human and mouse endothelial cells (HUVEC and MS1, respectively), and particularly focusing on defining the function of visfatin on the activation responses and monocyte interaction in human endothelial cells (HUVEC). We chose these adipokines owing to their strong biomedical relevance and association with inflammatory disorders (as mentioned below).
Leptin is a small protein product (16 kDa) of adipocyte Ob gene and is involved in the control of appetite and energy homeostasis through its affinity to specific hypothalamic receptors that signal for satiety (106). Mice with Ob gene disruption (ob/ob) are obese, and show hyperphagia, hyperlipidemia, metabolic abnormalities, insulin resistance and cardiovascular impairment. The administration of exogenous leptin reverses the cytotoxic effect of TNFα in obese mice (ob/ob) (136). Obese individuals have elevated leptin levels in the circulation owing to the increased accumulation of adipose mass and are insensitive to the anorexic and other beneficial effects of this adipokine due to receptor-saturation
(137–139). The average circulatory levels of leptin in humans have been reported in a wide range, with lean subjects (2.9 to 10.8 ng/ml) displaying significantly lower levels than obese individuals (12 to 37.8 ng/ml) (140, 141). In the prepubertal subjects, serum levels of leptin are significantly elevated (up to 4-fold) than their lean counterparts and with overall higher values in all female subjects, indicating an effect of gender on its secretion (141). Leptin belongs to helical-cytokine 50 superfamily, which also contains IL2 and growth hormone, and has been implicated in several pro-inflammatory responses associated with obesity. In monocytes, leptin upregulates the production of pro-inflammatory cytokines (such as TNFα and IL6) promotes ROS production, and cell proliferative and migratory responses (142). In macrophages, treatment with leptin stimulates JAK2-STAT3 signaling and upregulates chemokine secretion, including CCL3, 4 and 5 (143,
144) Further, pro-inflammatory stimuli, such as TNFα and LPS, increase the levels of leptin in both WAT and the circulation (145). Leptin also controls TH cell function and increases the production of TH1-type cytokines – IFNγ and IL2, and inhibits
TH2-type cytokines, such as IL4. Ob knockout mice (ob/ob) are resistant towards autoimmune brain disorders due to the polarization of TH cells into anti- inflammatory TH2 cells (146). Elevated leptin levels are associated with increased susceptibility to cardiovascular pathogenesis and directly contribute to peripheral vascular inflammation, coronary artery disease and atherosclerosis (147). At the level of vascular EC, treatment with leptin has been shown to activate NFκB and
JNK activation and stimulate the production of ROS (148), CRP (149), and nitric oxide (147). Furthermore, the increased oxidative stress promoted by leptin treatment in vitro also causes the enhanced production of MCP-1, which is a powerful inflammatory chemokine (150). However, the mechanism by which this adipokine exerts atherogenic and inflammatory effects on vasculature is controversial; although it seems to cause a pathogenic shift during hyperleptinemia, as healthy individuals with normal serum-leptin levels do not 51 exhibit abnormal EC-function. Additionally, hyperleptinemia has also been considered as an independent predictor of vascular inflammation and insulin resistance in several studies, but the cause-and-effect relationship has not been characterized. In context of EC-mediated immunoinfiltration of WAT, the role of leptin is yet to be established.
Visceral adipose-tissue derived Serpin (vaspin) was first described by Hida et al. as an insulin-sensitizing adipokine secreted from WAT of Otsuka Long-
Evans Tokishima Fatty (OLETF) rats –a model of obesity and type 2 diabetes
(151). Tissue expression and serum levels of vaspin peak with obesity, insulin levels and diabetes, and eventually decrease progressively with the worsening of diabetes and body weight loss in OLETF rats (152). Administration of vaspin into obese mice fed with high-fat high-sucrose diet improves both insulin sensitivity and glucose tolerance and normalizes the blood-glucose to normal levels (151).
Interestingly, insulin-sensitizing effects of vaspin appear to be limited to obese state only, as administration of this adipokine in lean mice does not influence glucose tolerance neither altered glucose uptake by cultured adipocytes (126). In addition to its protective effects against impaired glucose uptake, this adipokine reverses the expression of genes that are implicated in metabolic syndrome (151,
153). Vaspin administration results in the suppression of pro-inflammatory adipokines TNFα, resistin and leptin, while upregulated the levels of adiponectin and GLUT4 in the WAT of obese mice fed with high-fat high-sucrose diet (151). In vitro treatment of vascular-smooth muscle cells with recombinant vaspin abolishes 52 activation responses, such as ROS generation, cell proliferation, and chemokinesis, and blocks the NFκB, MAPK and PI3K-Akt pathway– describing the anti-atherogenic effects of this adipokine (154). In humans, the serum concentration ranges from 0.2 ng/ml to 2.5 ng/ml, which is positively correlated with the amount of body fat and type 2 diabetes (152, 155). In human aortic EC, vaspin is reported to oppose the effect of TNFα, and reduce the expression of adhesion markers – ICAM1, VCAM1, and E-selectin, and chemokine MCP-1 (156).
Additionally, vaspin treatment stimulates AMP kinase (AMPK) – an intracellular energy sensor pathway – with a simultaneous decrease in NFκB activity (157), suggesting a role of this adipokine in the regulation of both metabolic and inflammatory pathways. Due to its insulin-sensitizing functions during hyperglycemic state and protective role in vasculature and adipose, vaspin has been speculated as a beneficial adipokine and a therapeutic candidate. However, not much data are available on how this adipokine functions to influence the immunoinfiltration of WAT and EC-leukocyte interaction during obesity.
Visfatin is a small protein (55 kDa) with both intracellular and extracellular distribution. It was first identified in 1994 as pre-B-cell colony enhancing factor
(PBEF) from a bone marrow cDNA library and was described for its cytokine-like function in the B-cell maturation pathway (158). In 2002, a murine homologue of
PBEF was described as an enzyme Nicotinamide Phosphoribosyl Transferase
(NAMPT) that catalyzes the reaction between nicotinamide and 5-phosphoribosyl
1-pyrophosphate to yield nicotinamide mononucleotide (NMN) – a precursor of 53 nicotinamide adenine dinucleotide (159). In 2005, this molecule was called
‘visfatin’ because it was found to be secreted mainly by the visceral fat, and because it controlled glucose absorption in mice by signaling through the insulin receptors presumably by binding to different sites than insulin (160). Although the article was later retracted, because the insulin-mimetic effect of visfatin was batch- dependent, the study led other groups to investigate the role of visfatin in the control of glucose metabolism. Despite its mention as an ‘adipokine’, it has been shown that the predominant source for serum visfatin is leukocytes irrespective of obesity parameters (161). Nonetheless, studies indicate that extracellular visfatin may act as an immunomodulatory molecule and may provide an important link for paracrine signaling between adipocytes and stromal vascular cells to promote inflammatory responses (162, 163). However, the physiological relevance, and whether visfatin is a pro-inflammatory factor or an anti-inflammatory factor is a subject of debate (164, 165). On one hand, visfatin has been reported for its beneficial effects, such as β-cell function and insulin secretion (166), protection in neuronal injury (165), cardioprotective effects (167), and increased glucose uptake by kidney cells (168), while numerous other reports suggest its positive association with inflammatory disorders, including rheumatoid arthritis (169), myocardial infarction (170), atherosclerosis (171), and cardiovascular diseases (172).
Additionally, increased expression of visfatin correlates positively with several inflammatory disorders and some cancers (e.g. colorectal, breast, gastric, myeloma, melanoma, and others) (112). Figure 1.8 illustrates the positive 54 association of visfatin with several medical conditions. The fasting circulatory levels of this molecule are found to be in the ranges of 1.4 ng/ml to 29.3 ng/ml in lean subjects compared to 4.1 ng/ml to 32.7 ng/ml in the obese subjects (173–
175). As increased serum-concentration of visfatin correlates positively with visceral adiposity and certain cancers and because of several pro-inflammatory responses that it elicits in leukocytes, this adipokine has been considered a link between metabolic disturbances due to lipid accumulation and inflammatory and cardiovascular diseases. In human monocytes, visfatin up-regulates p38 mitogen- activated protein kinase (MAPK) and extracellular signal-regulated kinase (ERK) pathways and promotes the secretion of IL1β, TNFα and IL6 (176). In EC, visfatin treatment increases NFκB signaling to induce cytokine secretion and expression of adhesion markers that are implicated in the pathogenesis (177, 178). Although several research reports have indicated a pro-inflammatory role of visfatin in monocytes and vasculature (179–181)s, the mechanism by which this molecule initiates the signaling responses in the target cells is still not clear as a specific receptor has not been identified yet. As visfatin controls the activation responses in both leukocytes and EC, it is worth hypothesizing that it might also facilitate the interaction between leukocytes and vasculature to promote immunoinfiltration.
Therapeutic targeting of visfatin might reduce inflammation of both vasculature and
WAT, and thus may be a potential treatment to reduce atherogenic processes.
55
Figure 1.8 Positive association of visfatin with various medical conditions.
Research in the last decade emphasizes on the role of adipokines as a major contributing link between obesity and its comorbidities. Adipokines (e.g. visfatin), whose local expression and serum levels are positively associated with cancers and several inflammatory disorders, respectively, are likely to directly regulate functional responses. Another possibility is that they can target vasculature, specifically EC, to induce atherogenic changes that favor leukocyte chemotaxis and immunoinfiltration. It is worthwhile to speculate that a single adipokine can simultaneously target multiple cells in the microenvironment to regulate multiple cellular processes towards the development of a chronic, inflammatory state. Indeed, targeted interventions to block the adipokine function have been proposed as therapies. However, our current knowledge on adipokines 56 regarding their function and cell-specific response is still expanding and warrant more studies in this direction before specific therapies can be developed.
1.5 Hypotheses and Specific Aims
This project aims to characterize adipokine-induced pro-inflammatory responses in both human and mouse EC specifically aiming to determine a cytokine secretion profile, inflammation-associated gene expression, and interaction with leukocytes. Our long-term goal is to provide directions to develop novel anti-inflammatory therapies. Considering the evidence that a dysregulated adipokine profile and endothelial dysfunction are both positively correlated with obesity-associated inflammatory disorders, it is possible that that cellular crosstalk between adipocytes and EC, mediated through adipokine action, directly contributes to inflammatory responses. We hypothesize that EC will activate signal pathways resulting in pro-inflammatory gene expression and cytokine secretion in response to adipokines. Further, we propose that adipokine-induced endothelial activation will lead to an enhanced capability for their interaction with leukocytes and stimulate angiogenic responses [Figure 1.9].
57
Figure 1.9 Diagrammatic hypothesis.
To assess these hypotheses, the following specific aims were addressed.
1. Specific Aim 1: To Characterize Adipokine-Directed Cytokine Secretion
and Expression of Inflammation-Associated Genes in Human Endothelial
Cells, and Determine the Interaction with Leukocytes Specifically in
Response to Adipokine Visfatin
2. Specific Aim 2: To Characterize Angiogenic Gene Expression and
Determine the Capacity for Angiogenesis in Human Endothelial Cells in
Response to Adipokine Visfatin
3. Specific Aim 3: To Identify the Intracellular Signaling Pathways Underlying
Visfatin-Directed Gene Expression in Human Endothelial Cells
1.6 Summary of Project Significance
Adipokine-based anti-obesity therapies are considered promising candidates to ameliorate metabolic status and reverse inflammation. Before these 58 strategies can be translated into therapies, it is extremely important to understand the mechanism of adipokine action at the cellular and molecular levels. Adipokines can target multiple cells to evoke dysfunctional responses leading to inflammation and homeostatic disruption initially at the adipose microenvironment and subsequently at the whole-body level. It is therefore relevant to characterize the function of adipokines for their inflammatory and pathogenic potential. In this direction, several studies have been undertaken and the results present substantial evidence for the relevance of adipokines, including visfatin, in context of several metabolic, inflammatory, neurological, and ageing-related disorders.
However, anti-adipokine therapies are still far from being fully competent and only had mixed success in clinical trials, suggesting that details regarding the mechanism of adipokine action are still incomplete and more studies are warranted in this direction. EC provide a barrier between circulating leukocytes and a tissue microenvironment and endothelial dysfunction is often an initial step towards the exacerbation of tissue homeostasis and inflammation. The results presented here highlight the potential of targeting visfatin effects on EC for therapeutic purposes. 59
CHAPTER 2: ADIPOKINE TREATMENT UPREGULATES PRO-
INFLAMMATORY GENE EXPRESSION, CHEMOKINE SECRETION, AND
ENHANCED MONOCYTE INTERACTION IN HUMAN ENDOTHELIAL CELLS.
2.1 Abstract
Although the role of adipokines have been extensively characterized for the regulation of metabolic response and adipocyte function, details concerning their inflammatory role in the adipose microenvironment are largely unexplained. We tested the hypothesis that adipokine treatment results in a pro-inflammatory activation of endothelial cells to promote chemokine expression and enhanced interaction with monocytes. In the physiological context, similar processes have been shown to contribute to adipose inflammation and tumor growth (182–184).
Consistently, we show that treatment with adipokines – visfatin, leptin, and vaspin
– increased the secretion of pro-inflammatory cytokines in human endothelial cells.
Further, we investigated the role of visfatin in a greater detail and demonstrate that visfatin acts as a pro-inflammatory factor and upregulates several activation responses and enhanced monocyte interaction in human endothelial cells. These results emphasize the importance of visfatin as a therapeutic target and its relevance in pathogenic responses at the level of adipose and tumor microenvironment. We are in the process of revising a manuscript for publication in PLoS One.
60
2.2 Introduction
The epidemic of obesity and its comorbidities affects millions of people worldwide and imposes a difficult healthcare challenge to tackle. One of the major driving factor for obesity-associated disease progression is a ‘low-grade, chronic inflammation’ of visceral adipose tissue, correlating to poor clinical outcomes and extending links to diseases, such as atherosclerosis, myocardial infarction, and cancers. It has been postulated that adipose-derived bioactive molecules (called adipokines) may act in a paracrine manner to potentiate the inflammatory responses in the adipose microenvironment and promote immunoinfiltration and pathogenic activation at the cellular levels (84). However, limited evidence exists to substantiate this claim, particularly with respect to the vascular compartment and its role in leukocytosis under adipokine instruction. Details regarding adipokine-induced pro-inflammatory processes in endothelial activation and their interaction with leukocytes remain largely unexplored, though previous studies strongly indicate their therapeutic relevance in context of adipose dysfunction and pathogenesis (7, 185, 186). Thus, there is an urgent need to characterize these responses further to strengthen the understanding of obesity-associated inflammation and provide clues towards the development of effective interventions.
Here, we have investigated the function of primary human endothelial cells
(HUVEC), specifically for their capability to upregulate pro-inflammatory gene expression, cytokine secretion, and interaction with monocytes (THP1) under the effect of adipokine treatment in vitro. Some of the basic information regarding 61 these cellular models are listed in Table 2.1. We specifically chose this endothelial model (HUVEC) primarily because (i) it is an established endothelial model and a non-immortalized primary cell line that is directly derived from the host (187), (ii) it exhibits functional resemblance to native endothelial cells, and (iii) these cells have been extensively used as a choice-of-system to study immunological functions, cytokine secretion, proliferation, and vascular homeostasis (188). Further, we chose THP1 leukemic cell line as a model for human monocytes, because of their resemblance to primary monocytes-macrophages isolated from healthy or diabetic donors (189), and because of their capability to differentiate into native macrophage-like cells and functional resemblance (190). Additionally, for the treatments we used commercially available, human recombinant adipokines – visfatin, leptin, and vaspin. We described a comprehensive cytokine profile in human endothelial cells upon adipokine treatment, and extensively characterized pro-inflammatory responses and monocyte interaction in response to visfatin.
Table 2.1
Endothelial and monocytic cells chosen for adipokine treatment and analysis
Cultured Cells Origin (Organism) Site of Recovery Cell Type
HUVEC Human Umbilical Vein: Endothelium Primary
THP1 Human Peripheral blood: Monocytes Immortalized
62
2.3 Materials and Methods
2.3.1 Culture and Maintenance of Endothelial Cells
Primary Human Umbilical Vein Endothelial Cells, derived from umbilical vein from pooled donors (HUVEC, CC-2519), were purchased from Lonza
(Walkersville, MD) and cultured in a complete growth medium – Endothelial
Growth Medium-2 (EGM-2) as per manufacturer’s guidelines. To formulate EGM-
2, contents of the EGM™-2 SingleQuots™ Kit (Lonza Catalog No. CC-4176 containing human epidermal growth factor [hEGF], vascular endothelial growth factor [VEGF], R3-insulin-like growth factor-1 [R3-IGF-1], ascorbic acid, hydrocortisone, human fibroblast growth factor-beta [hFGF-β], heparin, fetal bovine serum [FBS], and gentamicin/amphotericin-B [GA]) were added to EBM™-
2 Basal Medium. Prior to seeding the cells, culture plates were coated with gelatin
(ScienCell, Carlsbad, CA) to allow the cells to efficiently adhere to the surface.
Specifically, gelatin solution was diluted to 0.1% with sterile water and applied to the culture dishes (1ml per 10 cm2) followed by an incubation of 30 minutes at
37oC. The solution was aspirated and surfaces were rinsed once with sterile PBS and plates were air-dried in a sterile hood for at least one hour prior to culture.
When initially culturing cells from the cryopreserved stocks, HUVEC were rapidly thawed in a 37oC water bath and added to culture vessels containing warm EGM-
o 2 followed by an incubation at 37 C and 5% CO2 in a humidified incubator. The media was replaced with fresh EGM-2 after 24 hours and cells were grown until about 80% confluent. Endothelial monolayers were subcultured by trypsinization 63 and split 1:3 – 1:5 in fresh EGM-2. HUVEC were never subcultured beyond passage 8, and stored by cryopreservation using a freezing medium (80% FBS,
10% EGM-2, and 10% DMSO) over slow cooling in an isopropanol container placed in a -80oC deep freezer. For long term storage, cryopreserved culture vials were transferred in liquid nitrogen. All the experiments were performed using
HUVEC between passages 2-5 as described previously (191)s.
THP1 monocytes (192) were cultured and maintained in a complete growth medium containing Roswell Park Memorial Institute basal medium (RPMI-1640;
Gibco, Grand Island, NY) supplemented with 10% FBS and 1X antibiotic mixture
(Antibiotic-Antimycotic, Gibco) at 37oC and 5% CO2. Upon reaching a density of approximately 1x106 cells/ml, the suspensions were subcultured in fresh complete media. The cells were stored and cryopreserved in a freezing medium (90% FBS and 10% DMSO) and stored as described above.
2.3.2 Adipokine Treatments
Recombinant human adipokines – leptin, vaspin, and visfatin – were purchased from Peprotech Inc. (Rocky Hill, NJ). The purity of these commercially obtained adipokines was marked greater than 98% as per the information supplied by the manufacturer. Specifically, HUVEC were treated with varying concentrations of adipokines (10 ng/ml to 1000 ng/ml) and for various durations (1 hour – 24 hours) in a growth factor deficient medium – EBM-2 – supplemented only with 0.5% FBS (treatment media). Depending on the experiment, samples 64 were collected following the treatments, which include culture conditioned supernatants, cell lysates, etc. for further analysis and target detection.
2.3.3 RNA Isolation
To determine the gene expression of chemokines and adhesion marker, adipokine treated HUVEC were processed for total RNA isolation using Trizol reagent (Invitrogen, Carlsbad, CA). Briefly, after the treatment, the culture medium was removed and endothelial monolayers were washed once with sterile PBS followed by the addition of 0.3-0.4 ml Trizol reagent per 1x106 cells to lyse the cells.
The lysates were prepared by homogenization with gentle pipetting followed by incubation at room temperature (RT) for 5 minutes for complete dissociation of the nucleoprotein complexes. To each sample, chloroform was added (0.2 ml per 1 ml
Trizol) and mixed vigorously and incubated for further 2-3 minutes. The mixture was centrifuged for 15 minutes at 12,000xg at 4oC to allow the separation of a bottom pink organic layer and a clear aqueous layer on the top. Aqueous layers
(containing RNA) were collected in fresh tubes and added to 100% isopropanol
(0.5 ml per 1 ml of Trizol) containing 10μg of RNase-free glycogen to allow the precipitation of RNA. The samples were incubated for 10 minutes at RT followed by centrifugation for 10 minutes at 12,000xg at 4oC. The supernatants were discarded and RNA pellets were washed once with 75% ethanol (0.5 ml) by centrifugation for 5 minutes at 7500xg at RT. The supernatants were discarded and RNA pellets were dissolved in 25μl of nuclease-free water, and samples were quantified for total RNA concentration in nanogram per microliter. 65
2.3.4 Reverse-Transcriptase PCR (RT-PCR)
RNA samples isolated from adipokine-treated HUVEC were reverse- transcribed to the corresponding cDNA by RT-PCR. Briefly, 1μg of total RNA was treated with DNase I (Invitrogen, Carlsbad, CA), followed by reverse transcription into cDNA in 20μl reactions by using a commercially available RT-PCR system
(High-Capacity cDNA Reverse Transcription Kit) as per manufacturer’s guidelines
(Applied Biosystems; Grand Island, NY). The cDNA samples were diluted 5 times, and a total of 2.5μl was used as template in the subsequent analysis.
2.3.5 Real-Time Quantitative PCR (RT-qPCR)
RT-qPCR analysis was performed to quantitatively measure the adipokine- responsive pro-inflammatory expression in HUVEC for the following targets: CCL2
(MCP1), GMCSF, IL6, CXCL2, CXCL8, E-selectin, ICAM1, VCAM1, and PECAM1, using Luminaris HiGreen qPCR Master Mix (Life Technologies; Grand Island, NY)
(193). GAPDH was used as a reference gene or housekeeping internal control gene, as was reported in the previously published reports in HUVEC (194–196).
Primer sequences for the target genes can be found in Appendix 1. The gene expression of chemokines and adhesion markers was normalized to GAPDH and expressed as fold change relative to the untreated control using 2-ddCt method
(197). The reactions were carried out in a CFX ConnectTM Real-Time PCR detection system (BioRad; Hercules, CA).
66
2.3.6 Enzyme-Linked Immunosorbent Assay (ELISA)
A sandwich-ELISA system (Peprotech; Rocky Hill, NJ) was used for the detection of secreted proteins produced by HUVEC following the guidelines in the manufacturer’s protocol. Briefly, high-binding 96 well plates were coated with 50μl of capture antibody diluted to 1μg in PBS and incubated overnight at RT. Wells were washed three times with 300μl of wash buffer (0.05% Tween-20 in PBS) using a multichannel pipette, and the surface was blocked with a blocking buffer
(1% BSA in PBS) for 1 hours at RT. Wells were washed as before and unknown samples or standards were applied and plates were incubated overnight at 4oC for antigen binding. Next day, wells were washed as before and incubated at RT for 2 hours in 50μl of biotinylated detection antibody solution (diluted to 0.5μg in wash buffer plus 0.1% BSA). The wells were washed as before and incubated at RT for
30 minutes in 50μl of a streptavidin-HRP (1:2000) solution. The washes were repeated as before and incubated with 100μl of a colorimetric substrate ABTS (Life
Technologies; Grand Island, NY) at RT for 25-30 minutes. The microplates were read using a microplate reader at 405 nM to measure the absorbance values due to color developed in proportion to the quantity of target antigen.
2.3.7 Antibody Microarray
For the semi-quantitative detection of secreted human proteins by HUVEC, a membrane-based Human Angiogenesis Antibody Array (Cat# AAH-ANG-1000-
4; Ray Biotech, Norcross, GA) was used as per the protocol provided by the manufacturer. First, the membranes containing antibody arrays were blocked with 67 a blocking buffer (supplied in the kit) for 30 minutes at RT. Culture-conditioned medium were collected from HUVEC after the adipokine treatments (visfatin, leptin or vaspin) or control (untreated sample), and applied to the blocked membrane and incubated overnight at 4oC for antigen binding. Membranes were washed sequentially in wash buffers I and II (supplied in the kit) for a total of three washes
(five minutes each) in each buffer. To each membrane, a cocktail of biotinylated antibody was added followed by an overnight incubation at 4oC. Next day, the solutions were removed and membranes were washed as above. A solution of 1X
HRP-Streptavidin was added to each membrane and incubated for two hours at
RT. The solutions were removed and a substrate solution was added for the chemiluminescence-based detection of target proteins using a ChemiDocTM XRS gel documentation system (BioRad; Hercules, CA). The information about the target molecules and the array layout can be found in Appendix 2.
2.3.8 Leukocyte Transmigration Assay
HUVEC were treated with visfatin and culture-conditioned medium was collected as a chemoattractant to test the capacity for monocyte migration in untreated (control) versus visfatin-treated (test) sample. Serum-free culture medium (EBM-2) supplemented with 50 ngml-1 MCP1 (a known chemoattractant for monocytes), was used as a positive control. A two-chamber, microplate-based transmigration system (ChemoTx® Neuroprobe; Gaithersburg, MD) was used to evaluate the transmigration of THP1 monocytes towards the culture medium through a filter of 8μM pore size. Briefly, microplate wells were filled with the 25 μl 68 of the conditioned medium collected from the untreated (control) or visfatin-treated
(test sample), and the positive control (MCP1 containing medium), and the plate was sealed with an 8μM filter (supplied in the kit). A suspension of THP1 human monocytes (50,000 cells in 25μl) was dispensed on the top of each well above the filter. The microplate was cover with a clear plastic lid, and the assembly was
o incubated at 37 C in humidified air with 5% CO2 for 4-8 hours to allow the chemotaxis of monocytes through the filter. Non-migrated cells on the origin side
(top) were washed away with PBS, and plates were centrifuged to collect the transmigrated cells bound to the inner side of the filter.
2.3.9 Leukocyte Adhesion Assay
To test the capacity of HUVEC to physically interact with leukocytes, a modified cell adhesion assay was performed (VybrantTM Cell Adhesion Assay Kit,
Molecular Probes; Eugene, OR). Endothelial monolayers were cultured on 0.1% gelatin-coated coverslips and treated with adipokines. Meanwhile, THP1 human monocytes were labeled with a fluorescent dye – Calcein AM (supplied in the kit).
Specifically, monocytes suspensions at 5 X 106 cells/ml were incubated at 37oC with a 1:200 dilution of Calcein AM for 30 minutes. Cells were washed twice with warm, serum-free RPMI. After the treatment with adipokines, endothelial monolayers (HUVEC) were subjected to Calcein AM-labeled monocyte suspension and the co-cultures were incubated at 37oC in humidified air with 5%
CO2 for 30 minutes. Unbound monocytes were washed away by gently rinsing the 69 coverslips with sterile PBS. Cells were fixed in 4% formaldehyde, and coverslips were mounted on a slide to be visualized by fluorescence microscopy.
2.3.10 Microscopy and Image Analysis
THP1 human monocytes that were transmigrated to the lower chamber were photographed with a camera equipped phase-contrast microscope (Nikon
Eclipse TS100) at 100X magnification, and the total number of transmigrated cells were counted using Cell Counter program of ImageJ (NIH). For the monocyte adhesion assay, fluorescence imaging was done using a confocal microscope
(Nikon Eclipse Ti), and photographs from random multiple fields were obtained.
Images were analyzed for the total number of adherent monocytes from each sample using an image analysis software (Nikon, NIS Elements).
2.3.11 Statistical Analysis
The graphing and statistical analyses of all the data and results were carried out by using GraphPad Prism software. The significance of difference amongst the treatment groups and control samples was tested by a two-tailed student’s t-test.
Multiple comparisons between the groups were performed using ANOVA with a posthoc analysis by the Tukey-Kramer multiple comparisons test. Results yielding a p-value <0.05 were considered significant for the difference between the compared groups. Each experiment was repeated two more times with each treatment group containing at least three independent replicates per experiment to obtain statistically relevant results.
70
2.4 Results
2.4.1 Adipokine Treatment Increases the Secretion of Pro-Inflammatory
Cytokines and Chemokines in Human Endothelial Cells
An integral, early component of the inflammatory response is the secretion of pro-inflammatory cytokines that act as endogenous mediators towards the leukocyte infiltration in a tissue. External stimuli, such as paracrine factors or stress, can induce the release of pro-inflammatory cytokines by endothelial cells
(198). To determine the cytokine secretion by human endothelial cells (HUVEC) in response to adipokines – visfatin, vaspin, and leptin – a membrane-based antibody microarray was used to profile multiple targets. Briefly, the culture supernatant was collected 24 hours after adipokine treatments and applied to a membrane spotted with specific antibodies for target cytokines. The antibody arrays were developed using a chemiluminescence-based detection method and spots were photographed to visually analyze the amount of cytokine secreted as proportional to the spot intensity. As presented in Figures 2.1, treatment with adipokines, visfatin, leptin, and vaspin, resulted in differential level of cytokines secretion as compared to the untreated control in each array. The pro-inflammatory cytokines
IL-1α, GMCSF, GRO, CCL2, IL6, and IL8 were chosen for further confirmation by
ELISA for a targeted analysis. HUVEC were cultured in the presence of varied doses of each adipokine and culture supernatants containing the secreted cytokines were collected for sandwich-ELISA analysis. As presented in Figure 2.2, treatment with visfatin increased the secretion of IL-1α, GMCSF, CXCL2 (GROβ), 71
CCL2, and CXCL8 (IL8) in a dose-dependent manner. Leptin treatment also resulted in an increased secretion of these cytokines [Figure 2.3]; however, vaspin treatment caused an increase only in the levels of IL-1α and CCL2 [Figure 2.4].
Previously published reports indicate that vaspin may have an anti-inflammatory effect (199), and administration of vaspin in obese mice improved glucose tolerance, and reversed the expression of genes associated with insulin resistance
(151). To test the anti-inflammatory function of vaspin, HUVEC were pre-treated for one hour with recombinant vaspin followed by stimulation with an inflammatory factor – TNFα – for 24 hours. Culture supernatant was collected for the detection of the pro-inflammatory cytokines released in the medium. Vaspin pre-treatment resulted in the suppression of TNFα-stimulated secretion of pro-inflammatory cytokines – CCL2 and IL6 [Figure 2.5], suggesting that vaspin may partially suppress endothelial activation under inflammation.
72
Figure 2.1 Qualitative cytokine secretion profile of human endothelial cells (HUVEC) in response to adipokine treatments. Protein microarray incubated with conditioned medium obtained after the 24 h treatment of HUVEC with visfatin, leptin, and vaspin (100 ngml-1 each) reveal differential levels of secreted cytokines in both Array 1 (upper panel) and Array 2 (lower panel). The molecules highlighted in red box (IL1α, IL6, IL8, GMCSF, GRO, and MCP1) were chosen for subsequent analyses based on their stronger intensity compared to control. The information about the array layout and target molecules can be found in Appendix 2. 73
Figure 2.2 Visfatin treatment promotes the secretion of multiple chemokines and cytokines in HUVEC in a dose-dependent manner. Pro-inflammatory factors secreted by HUVEC in the culture medium in response to 24 h treatment with recombinant visfatin were measured by ELISA. Levels are shown for (A) IL1α, (B) IL6, (C) GMCSF, (D) CCL2, (E) CXCL2, and (F) CXCL8. *p<0.05, **p<0.01, ***p<0.001 vs 0 (control, grey bar), (n=5). 74
Figure 2.3 Leptin treatment promotes the secretion of multiple chemokines and cytokines in HUVEC. Pro-inflammatory factors secreted in the culture medium by HUVEC in response to 24 h treatment with recombinant leptin were measured by ELISA. Levels are shown for (A) IL1α, (B) IL6, (C) GMCSF, (D) CCL2, (E) CXCL2, and (F) CXCL8. *p<0.05, **p<0.01 vs 0 (control, grey bar), (n=5). 75
Figure 2.4 Secretion of pro-inflammatory cytokines and cytokines by HUVEC in response to vaspin treatment. HUVEC were treated with recombinant vaspin for 24 h; cytokines released in the culture medium were measured by ELISA. Levels are shown for (A) IL1α, (B) IL6, (C) GMCSF, (D) CCL2, (E) CXCL2, and (F) CXCL8. *p<0.05, **p<0,01, ***p<0.001 vs 0 (control, grey bar), (n=5). 76
Figure 2.5 Pretreatment with vaspin suppresses tumor necrosis factor-alpha induced cytokine secretion by HUVEC. HUVEC were treated with TNFα (5 ngml-1) alone, or pre-treated for 2 h with the indicated concentrations of recombinant vaspin (50 and 100 ngml-1) followed by stimulation with TNFα (5 ngml-1) for 24 h, and pro-inflammatory factors secreted in the culture medium were quantified by ELISA. Levels are shown for (A) CCL2 (MCP1), and (B) IL6. ***p<0.001 vs Control; δδδp<0.001 vs TNFα, (n=3).
2.4.2 Visfatin Increases the Expression of Pro-Inflammatory Genes and
Chemokine Secretion in Human Endothelial Cells
Recently visfatin has been extensively reviewed for its possible contribution in a number of pathologies and inflammatory disorders, including chronic kidney disease, polycystic ovary syndrome, rheumatoid arthritis, and coronary artery disease. (163, 172), but the mechanism of action by which it contributes to the inflammatory processes is still a subject of debate (163). To test the direct actions 77 of visfatin in the context of inflammation, HUVEC were treated with recombinant visfatin for the indicated doses and duration, and samples were obtained for the gene expression analysis at both RNA and protein level. Briefly, endothelial monolayers were treated with visfatin and total RNA was isolated after 1h or 4h post-treatment. As presented in Figures 2.6-2.10, treatment with visfatin resulted in an upregulation of mRNA levels of several chemokine genes, including – CCL2
(MCP1), GMCSF, IL6, CXCL2 (GRO-β), and CXCL8 (IL8). Importantly, the induction in the expression of these genes was rapid, with peak levels attained at
1 hour followed by an eventual decline, perhaps due to a negative feedback regulation. Additionally, the gene expression response was also observed to be visfatin-dose dependent, with a working concentration of 100 ngml-1 as the most effective dose for the induction of most targets [Figures 2.6-2.10].
78
Figure 2.6 Visfatin increases the expression of chemokine ligand-2 (CCL2) in HUVEC in a dose and time-sensitive manner. Relative mRNA expression levels of CCL2 in HUVEC after visfatin treatment is shown. (A) Dose response at 1 h, (B) Time course over 4 h. **p<0.01, ***p<0.001 vs Control (grey bar), (n=3).
Figure 2.7 Visfatin increases the expression of granulocyte-macrophage colony stimulating factor (GMCSF) in HUVEC in a dose and time-sensitive manner. Relative mRNA expression levels of GMCSF in HUVEC after visfatin treatment is shown. (A) Dose response at 1 h, (B) Time course over 4 h. ***p<0.001 vs Control (grey bar), (n=3). 79
Figure 2.8 Visfatin increases the expression of interleukin-6 in HUVEC in a dose and time-sensitive manner. Relative quantification of the mRNA levels of IL6 in HUVEC in response to visfatin treatment, (A) Dose response at 1 h; (B) and (C) time course over 4 h. *p<0.05, ***p<0.001 vs Control (grey bar), (n=3). 80
Figure 2.9 Visfatin increases the expression of CXC-chemokine ligand 2 (CXCL2) in HUVEC in a dose and time sensitive manner. Relative quantification of the mRNA levels of CXCL2 in HUVEC in response to visfatin treatment, (A) Dose response at 1 h, (B) time course over 4 h. *p<0.05, **p<0.01, ***p<0.001 vs Control (grey bar), (n=3).
Figure 2.10 Visfatin increases the expression of CXC-chemokine ligand 8 (CXCL8) in HUVEC in a dose and time sensitive manner. Relative RNA expression levels of CXCL8 in HUVEC in response to visfatin treatment is shown. (A) Dose response at 1 h, (B) Time course over 4 h. **p<0.01, ***p<0.001 vs Control (grey bar), (n=3). 81
To determine the visfatin-induced expression of inflammatory mediators at the protein level, HUVEC were treated with recombinant visfatin and culture- conditioned supernatant was collected at different time points to measure the levels of secreted proteins by ELISA. We found that treatment with visfatin stimulated the production of CXC-chemokine ligands – CXCL2 (GROβ) [Figure
2.11], and CXCL8 (IL8) [Figure 2.12]. This effect was dependent on the dose of visfatin and duration of the treatment, with the maximum secretion observed at 2- hour post treatment. CXC-chemokine ligands are powerful mediators of inflammation and have been widely studied for their capacity to induce leukocyte chemotaxis and angiogenic potential (183, 200). Visfatin-stimulated increase in
CXC-chemokine secretion in human endothelial cells indicates the role of this adipokine as a possible contributor to inflammation in the adipose microenvironment.
82
Figure 2.11 Visfatin stimulates enhanced secretion of the chemokine CXCL2 by HUVEC in a dose and time dependent manner. Levels of CXCL2 secreted by visfatin-treated HUVEC in the culture supernatant were determined by ELISA. (A) Dose response at 2h, (B) Time course over 24 h. **p<0.01, ***p<0.001 vs Control (grey bar), (n=5).
Figure 2.12 Visfatin stimulates enhanced secretion of the chemokine CXCL8 by HUVEC in a dose and time dependent manner. Levels of CXCL2 secreted by visfatin-treated HUVEC in the culture supernatant were determined by ELISA. (A) Dose response at 2h, (B) Time course over 24 h. **p<0.01, ***p<0.001 vs Control (grey bar), (n=5). 83
2.4.3 Visfatin Treatment of Human Endothelial Cells Results in Enhanced
Chemoattractant Capacity for Monocyte Transmigration
Leukocyte chemotaxis and tissue-infiltration is an important component of an inflammatory response. Immune cells, such as monocytes and neutrophils, express receptors that sense chemokines, and follow the chemokine gradient to reach and transmigrate into the inflamed tissue (201). Besides adipocytes, endothelial cells can contribute to the chemokine milieu in the adipose microenvironment and aid in immune cell migration and contribute to ongoing inflammation. As demonstrated in the previous section, treatment with recombinant visfatin resulted in the upregulation of multiple cytokines and chemokines at both transcript and protein level in HUVEC. To test the effect of visfatin on endothelial cell-mediated leukocyte migration, a two-chamber transmigration system was used with culture-conditioned medium as attractant and human monocytes (THP1) as the representative immune cells. Briefly,
HUVEC were treated with visfatin and culture-conditioned medium representing the secreted chemokine milieu was collected and placed in the lower chamber of a transmigration assembly (ChemoTX, Neuroprobe). The plate was covered with a filter (8 μM) in a way that the liquid phase in the lower chamber makes a contact.
Above the filter, a suspension with equal number of THP1 monocytes was placed
o on the top of each well, and the assembly was incubated at 37 C, 5% CO2 in a humidified environment to allow the transmigration of immune cells through the filter to the lower chamber. THP1 monocytes that transmigrated to the lower 84 chamber were visualized over a phase-contrast microscope; photographs were obtained and the number of transmigrated monocytes were quantified. As shown in Figure 2.13(A), the number of transmigrated monocytes was visually higher in the visfatin-treated sample than the untreated control. The quantification of the images confirmed this observation, as shown in Figure 2.13(B). These data suggest, that in context of adipose microenvironment, visfatin may contribute to increased monocyte chemotaxis by acting on the vascular compartment, which further substantiate the role of this adipokine as a positive indicator of adipose tissue inflammation.
85
Figure 2.13 Human monocytes (THP1) exhibit an enhanced migration capability towards visfatin-treated endothelial culture supernatants. Culture conditioned medium was collected after 8 h treatment of HUVEC with visfatin and used as a chemoattractant for THP1 monocytes in a leukocyte transmigration assay. (A) Representative micrographs showing transmigrated monocytes in the lower chamber containing the culture medium (100X); (B) Quantification of transmigrated monocytes in six different fields. MCP1 (50 ngml-1) was used as a positive control. *p<0.05, ***p<0.001 vs 0 (grey bar); δp<0.05 vs MCP1; (n=6). MCP1, Monocyte Chemotactic Protein. 86
2.4.4 Visfatin Increases the Gene Expression of Adhesion Markers in Human
Endothelial Cells
To initiate an immune response, circulating leukocytes must interact and establish contact with vascular endothelium and transmigrate into the target tissue.
The leukocyte-endothelial interaction is a multi-step response and is effected by a variety of surface proteins commonly referred to as ‘cell adhesion molecules’, whose expression is usually increased during a pro-inflammatory response and inflammatory disorders (202–206). To determine the function of visfatin in context of leukocyte-endothelial interaction, HUVEC were cultured in the presence of recombinant visfatin and total RNA was isolated and reverse transcribed to measure the gene expression of classical adhesion molecules – E-selectin,
ICAM1, VCAM1, and PECAM1. As seen in Figure 2.14, treatment of HUVEC with visfatin for 4 hours resulted in an increase in the RNA expression of all the above- mentioned adhesion molecules. Importantly, visfatin treatment stimulated a 5-fold increase in the levels of E-selectin, and approximately 3-fold increase in VCAM1, suggesting the involvement of this adipokine in contributing to immunoinfiltration, specifically creating conditions for leukocyte rolling and attachment over the vascular endothelium.
87
Figure 2.14 Visfatin treatment increases the expression of cell adhesion markers in HUVEC. Relative quantification of the mRNA levels of adhesion markers in HUVEC in response to visfatin treatment after 4 h. (A) E-Selectin, (B) ICAM1, (C) VCAM1, and (D) PECAM1. *p<0.05, **p<0.01, ***p<0.001 vs Control (grey bar), (n=3). ICAM1, Intercellular cell adhesion molecule 1; VCAM1, Vascular cell adhesion molecule 1; PECAM1, Platelet endothelial cell adhesion molecule 1. 88
2.4.5 Treatment of Human Endothelial Cells with Visfatin Results in an
Enhanced Capacity for Monocyte Adhesion
Prior to immunoinfiltration into the target tissues, such as adipose tissue, leukocytes should cross the vascular-barrier posed by endothelial layer in the blood vessels. Under inflamed conditions the activated endothelial cells increase the expression of surface adhesion molecules that promote a physical interaction with the circulating immune cells (66, 206, 207). To test if visfatin has a contributing effect in this phenomenon in vitro a leukocyte adhesion assay was performed.
Briefly, coverslip cultures of HUVEC were treated with recombinant visfatin followed by a co-incubation with fluorescence-labeled THP1 monocytes. Unbound monocytes were washed away and those that remained bound were visualized and photographed using fluorescence microscopy. As seen in Figure 2.15(A), visfatin-treated endothelial monolayers exhibited a pronounced adhesion to THP1 monocytes compared to untreated sample. The adhered monocytes were counted using an image analysis software and the quantification is reported in Figure
2.15(B), which further confirms that visfatin-treatment resulted in an enhanced capacity for monocyte adhesion in human endothelial cells.
89
Figure 2.15 Visfatin treated HUVEC exhibit an enhanced capacity for THP1 monocyte adherence. Calcein-AM labeled THP1 monocytes were applied on 4 h visfatin-treated HUVEC monolayers cultured on a glass coverslip, (A) Representative fluorescence micrographs showing adherent monocytes, and (B) quantification of attached monocytes in the from six different fields. TNFα (10 ngml-1) was used as a positive control. ***p<0.001, p<0.059 vs 0 (control, grey bar), (n=6). TNFα, Tumor necrosis factor alpha. 90
2.5 Discussion
Obesity is present in epidemic proportions and affects populations worldwide irrespective of age and gender. Obesity represents a complex condition of metabolic and inflammatory imbalance at the whole-body level, and thus, obese individuals have a significantly higher risk of developing disease over time (13,
208). The pathogenic aspect of obesity has been well characterized due to its positive association with multiple life-threatening conditions, and together with diabetes, obesity is now seen as a ‘twin epidemic’ of the modern world (209) imposing a tremendous health burden. Hence, there is an immediate need to better understand the inflammatory mechanisms contributing to obesity-associated pathogenesis and develop targeted therapies to counter disease progression.
With a goal to elucidate the interaction between the components of adipose microenvironment in the context of inflammation, here we report, (i) a comprehensive cytokine/ chemokine profile representing 40 targets in response to three adipokines – visfatin, leptin, and vaspin; and (ii) an anti-inflammatory action of vaspin; and (iii) visfatin-induced pro-inflammatory gene expression and enhanced monocyte interaction, in human endothelial cells (HUVEC) in vitro.
Higher serum levels of these adipokines have been found to be positively associated in the individuals with obesity and metabolic complications, suggesting a possible involvement of these molecules in pathogenesis (210). However, the molecular mechanisms for their action and contribution to drive inflammatory processes, particularly visfatin and vaspin, are yet not fully understood. Using an 91 antibody array approach, we developed a detailed chemokine profile in human endothelial cells showing a differential pattern of expression under adipokine treatment [Figure 2.1]. A targeted, downstream analysis revealed a dose- dependent, significant upregulation in the levels of multiple pro-inflammatory cytokines – IL-1α, CCL2 (MCP1), GMCSF, IL6, CXCL2 (GRO-β), and CXCL8 (IL8) by HUVEC specifically in response to visfatin and leptin treatment [Figures 2.2-
2.3]. These cytokines control substantial immune responses and widespread cellular functions in a variety of inflammatory and infectious conditions (183, 211–
213). The significant upregulation of these factors in endothelial cells underscores the existence of an adipo-vascular axis and a paracrine loop for the contribution to the inflammatory milieu at the level of adipose microenvironment.
The adipokine vaspin was first described by Hida et al. as a visceral fat- derived factor with insulin-sensitizing and potentially anti-inflammatory properties in a murine model of diabetes (151). It was suggested that vaspin may function as a compensatory factor in obese rats to partially overcome poor metabolic control and may have an immunomodulatory effect (151). In our experiments, we found that treatment with vaspin resulted both stimulated (IL1α and CCL2), and suppressed (CXCL8) the basal secretion of pro-inflammatory molecules in human endothelial cells, suggesting a diverse mode of its action [Figure 2.4]. Several reports have confirmed a positive correlation between serum vaspin levels and metabolic and inflammatory disorders; however, its role in inflammation remains controversial (214–216). We show that vaspin pre-treatment suppressed TNFα- 92 stimulated secretion of CCL2, and IL6 in cultured HUVEC [Figure 2.5]. These data, together with other published reports, highlight that vaspin may have a protective role under inflammatory conditions, but more studies are needed to validate its biological relevance.
The adipokine visfatin has been in focus owing to its positive association with obesity parameters, insulin resistance, cardiovascular disorders, and multiple inflammatory diseases (162, 163). To gain an insight for the vascular function of visfatin at the cellular level, we investigated its putative pro-inflammatory function using HUVEC. We found that visfatin treatment caused a significant upregulation in the RNA expression of classical pro-inflammatory markers – CCL2, GMCSF,
IL6, CXCL2, and CXCL8 [Figures 2.6-2.10], and showed a significantly elevated secretion of both CXCL2 and CXCL8 over a period of 24 hours [Figures 2.11-2.12].
These pro-inflammatory mediators have been well studied for their substantial role in the regulation of immune responses under a diverse stimulus, including tissue injury, and infection (200). CCL2, also called as MCP1, is a powerful chemotactic factor for monocyte migration, and its levels are positively correlated with obesity parameters, atherosclerosis, cardiovascular disease, and cancer (217). We show that treatment of cultured human endothelial cells with adipokines significantly upregulated the levels of CCL2 (up to 2-fold) at both RNA and protein level [Figures
2.2-2.4, 2.6, respectively]. Previous studies using mouse models of atherosclerosis emphasize on a critical role of CCL2 for mediating monocyte infiltration into atherosclerotic lesions and disease progression (218, 219). In 93 addition, CCL2 has been implicated in the pathogenesis of a wide variety of cancers, including breast cancer (220), gastrointestinal cancers (221), pancreatic cancer (222), and lung cancer (223). Based on these findings and our results, it is noteworthy to speculate that increased CCL2 originated from adipokine-stimulated endothelial cells might be a potential factor for the dysfunction at the level of tissue microenvironment and possibly, disease progression. GMCSF is a pro- inflammatory cytokine and a colony stimulating factor, best characterized for its role in the production and differentiation of monocytes and macrophages from their bone marrow precursors. In context of disease, the increased levels of GM-CSF are found to be associated with a broad category of inflammatory disorders, notably, arthritis, multiple sclerosis, interstitial lung disease, colitis, aortic aneurism, asthma, and leukemia (213, 224, 225). We show that visfatin treatment caused a significant upregulation in the expression of GMCSF by cultured human endothelial cells [Figures 2.2 and 2.7]. Visfatin can cause a similar effect in the physiological context where enhanced release by GMCSF in the microenvironment (adipose or tumor) can result in exacerbated immune response and potentiate pathogenesis (224). We also show that visfatin treatment resulted in a significant upregulation (over 2-fold) in the RNA expression of IL6 [Figure 2.8].
IL-6 is a pleiotropic cytokine that controls widespread physiological responses, including lymphocyte development, insulin resistance, lipid metabolism, autoimmune responses, neural disorders, and tumor progression (212). Based on this finding and owing to a positive association for IL6 levels in obesity, it might be 94 possible that visfatin has an involvement for the regulation of metabolic and immune processes at the level of adipose microenvironment with IL6 as a mediator, and further experiments are needed to substantiate this claim. CXC- ligands, such as CXCL2 and CXCL8, belong to a chemokine family of potent chemotactic factors that mediate leukocyte migration to the inflamed tissues. We show that visfatin treatment resulted in a strong upregulation in the levels of
CXCL2 and CXCL8, both at mRNA and protein levels [Figures 2.9-2.12]. Both
CXCL2 and CXCL8 have been well-studied for their role in tumor survival and pro- angiogenic responses (226, 227), and play a crucial role in mediating leukocyte- endothelial interaction during an inflammation owing to the presence of shared receptors on these cell types. Our results are highly relevant in context of both adipose and tumor microenvironment, as it indicates a possible involvement of visfatin both for immunoinfiltration and pro-tumorigenic responses mediated by
CXC chemokines.
Our findings for the role of visfatin to control pro-inflammatory gene expression and chemokine secretion is consistent with the previous findings.
Apparently, extracellular visfatin can induce proinflammatory expression in diverse cellular targets. Adya et al. showed that extracellular visfatin treatment could upregulate MCP1 secretion in microvascular cells (228). It was also reported that both visfatin can regulate MCP1 levels in adipocytes, where visfatin treatment stimulated the mRNA expression and secretory levels of MCP1, respectively in both in vitro, and in vivo conditions (229). Moschen et al. reported that human 95 monocytes when treated with recombinant visfatin increased inflammatory cytokine secretion and T-cell stimulatory responses (176). Visfatin-mediated upregulation of these highly chemotactic and pro-inflammatory molecules (CCL2,
GMCSF, IL6, CXCL2 and CXL8) suggest a cellular cross-talk mechanism between endothelial cells, immune cells, and adipocytes where visfatin might play a causal link to drive inflammatory responses through the vascular compartment at the level of microenvironment. Consistent with these results, we also observed an enhanced migratory potential of THP1 human monocytes in response to conditioned medium obtained after visfatin treatment of endothelial cells [Figure
2.13], suggesting a strong relevance of visfatin for immune cell migration in the adipose and tumor microenvironment.
In response to pro-inflammatory signaling, activated endothelial cells increase the expression of cell adhesion molecules (CAM) that promote enhance leukocyte interaction ultimately resulting in their extravasation into the target tissue. In our experiments, we show that visfatin-treatment resulted in a significant upregulation in the mRNA expression of the adhesion markers – E-selectin,
ICAM1, VCAM1, and PECAM1 [Figure 2.14]. This increase in CAM expression was consistent with the corresponding biological function, where visfatin-treated endothelial cells showed a higher capacity for monocyte (THP1) adhesion [Figure
2.15]. Enhanced expression of adhesion markers, including ICAM1 and VCAM1 is crucial for monocyte infiltration to atherosclerotic lesions (230), and tumor progression and metastases (231). Based on our findings and given that obesity 96 increases the susceptibility to cardiovascular disease and cancer in the affected individuals (232), it is plausible that visfatin might play a direct role for disease progression through the abovementioned pro-inflammatory effects in endothelial cells in the stromal fraction, in context of both adipose, and tumor microenvironment. While visfatin can induce the expression of pro-inflammatory mediators in endothelial cells, it is possible that inflammation can lead to the production of visfatin itself by the vascular compartment. In support of this claim, a study by Romacho et al. revealed that in response to inflammatory cytokines, such as TNFα, IL1β, and angiotensin-II, HUVEC increase the expression of visfatin at both the RNA and protein levels, and promote its secretion in the extracellular space (233). Based on this report and our findings, it is possible that the pro- inflammatory action of visfatin is due to a paracrine loop, where visfatin first promotes the expression of inflammatory chemokines by endothelial cells, which in turn act on the endothelial cells to further promote visfatin secretion in a feed forward manner.
In conclusion, with a goal to understand the relevance of adipokines for inflammation we investigated cytokine gene expression and leukocyte interaction in human endothelial cells. We report, a comprehensive qualitative cytokine- secretory profile in human endothelial cells in response to adipokine treatment.
Our findings for visfatin-induced pro-inflammatory effects in endothelial cells are relevant from a pathogenic standpoint. Here we report that visfatin not only increased the pro-inflammatory gene expression in human endothelial cells, but 97 also enhanced the capacity for endothelial-monocyte interaction. Uncontrolled chemokine secretion and excessive immunoinfiltration has been associated with inflammation and poor outcome in context of both adipose tissue dysfunction and tumor progression. Over all, our results provide an evidence for visfatin as a pro- inflammatory factor for endothelial activation and its implications for adipose inflammation and possible link to carcinogenesis.
98
CHAPTER 3: THE ADIPOKINE VISFATIN PROMOTES ANGIOGENIC
RESPONSES IN HUMAN ENDOTHELIAL CELLS
3.1 Abstract
Angiogenesis is an essential physiological response by which an actively growing tissue attempts to overcome its increased nutrient and oxygen demand.
Evidence indicates the therapeutic relevance of anti-angiogenic approaches in context of obesity-associated pathologies and tumor progression. The pro- inflammatory milieu in the adipose microenvironment can positively contribute to dysfunctional angiogenesis leading to tissue dysfunction and disease progression.
It is therefore important to understand how these processes are regulated and what the potential mediators of a dysfunctional angiogenic response are. Here, we investigated the function of the adipokine visfatin for its angiogenic capabilities, and observed that visfatin stimulated morphogenic and functional responses and increased pro-angiogenic gene expression in human endothelial cells. These results support the growing evidence for the pro-angiogenic role of visfatin and underscore its therapeutic relevance.
3.2 Introduction
Angiogenesis is the formation of nascent vasculature in a series of events characterized by vascular and extracellular matrix (ECM) remodeling, cellular migration, and proliferation, and sprouting of microvessels, guided by the growth factors and inflammatory mediators to compensate the increased nutrient demand of a growing tissue (75). Under a state of chronic inflammation, as in obesity, 99 dysfunctional responses such as increased immunoinfiltration and secretion of pro-inflammatory cytokines, lipolysis, and hypoxia can lead to angiogenic signals in adipose microenvironment. In fact, studies indicate that angiogenesis and adipogenesis are functionally coupled processes (72). Interestingly, the angiogenic responses at the level of an inflamed adipose tissue share similar molecular characteristics to that in tumor microenvironment, and it has been speculated that antiangiogenic therapies that target adipogenesis will also prove to be potential anticancer measures.
Recently, several studies have focused on the role of adipokines and ECM components as regulators of vascular responses and angiogenesis. It is well characterized that growing adipocytes and adipose stromal cells secrete multiple angiogenic factors, including VEGF, FGF, TGFβ, HGF, resistin, leptin, adiponectin, and others which can induce functional responses, both directly, or in a synergistic manner to regulate angiogenic processes (72). Additionally, during inflammation, activated macrophages can also influence vascular responses by producing inflammatory cytokines with potent angiogenic ability, such as TNFα, CXCL8, IL6,
GMCSF, VEGF, etc. (72). Understanding the mechanism by which adipokines stimulate angiogenesis is crucial to better determine how crosstalk between different cells can support adipogenesis, and provide clues for the interventions to target these processes in inflammatory diseases. Several studies indicate the role of visfatin in dysregulated angiogenic processes and highlight its therapeutic importance (234, 235, 180, 236), but details regarding visfatin-induced pro- 100 angiogenic effectors are limited. Here we have investigated the capability of visfatin to stimulate angiogenic functions in human endothelial cells. We performed an in vitro angiogenesis assay and found that visfatin stimulated capillary-like tube formation in endothelial cells in vitro. In addition, we report that visfatin increased the activity of ECM modifying proteases. Finally, we show that visfatin-treatment upregulated the RNA expression of multiple regulators of angiogenic processes.
Our findings are relevant to both adipogenesis and tumor growth as both processes share several overlapping molecular mediators. Importantly, our results support the evidence that visfatin is a pro-angiogenic factor for human endothelial cells and may have a therapeutic relevance.
3.3 Materials and Methods
3.3.1 Human Endothelial Cell Culture and Treatments
Primary Human Umbilical Vein Endothelial Cells, derived from umbilical vein from pooled donors (HUVEC, CC-2519), were purchased from Lonza
(Walkersville, MD) and cultured in a complete growth medium – Endothelial
Growth Medium-2 (EGM-2) as per manufacturer’s guidelines. To formulate EGM-
2, contents of the EGM™-2 SingleQuots™ Kit (Lonza Catalog No. CC-4176 containing human epidermal growth factor [hEGF], vascular endothelial growth factor [VEGF], R3-insulin-like growth factor-1 [R3-IGF-1], ascorbic acid, hydrocortisone, human fibroblast growth factor-beta [hFGF-β], heparin, fetal bovine serum [FBS], and gentamicin/amphotericin-B [GA]) were added to EBM™- 101
2 Basal Medium. Prior to seeding the cells, culture plates were coated with gelatin
(ScienCell, Carlsbad, CA) to allow the cells to efficiently adhere to the surface.
For the experiments, HUVEC between passages 2-5 were used. Cells were seeded in 0.1% gelatin-coated, multi-well culture dishes and grown until the formation of a monolayer. Based on the experiments, the adipokine treatments were given in a low-serum medium (EBM-2, 0.5% FBS) for 1-24 hours, and samples were collected for further analysis.
3.3.2 Real-Time Quantitative PCR (RT-qPCR)
Real-time qPCR analyses were performed to measure the mRNA levels of inflammation and angiogenesis-associated genes – ANGPT1, ANGPT2, CXCL11, uPAR, Angiogenin, VEGFR1, VEGFR2, and MMP2 – expressed by HUVEC in response to visfatin treatment. These gees were chosen for analysis due to their relevance in both inflammatory and angiogenic processes. Primer sequences for these target genes can be found in Appendix 1. Reactions were carried out using
Luminaris HiGreen qPCR Master Mix (Life Technologies; Grand Island, NY) (193).
GAPDH was used as a reference gene or housekeeping internal control gene
(194–196). The gene expression of chemokines and adhesion markers was normalized to GAPDH and expressed as fold change relative to the untreated control using 2-ddCt method (197). The reactions were carried out in a CFX
ConnectTM Real-Time PCR detection system (BioRad; Hercules, CA).
102
3.3.3 SDS-PAGE and Gelatin Zymography
To determine the functional activity of matrix metalloproteases (MMPs) secreted by HUVEC in the culture-conditioned medium, gelatin zymography analysis was performed as described previously (237). Cells were treated with visfatin and culture-conditioned supernatants were collected at different time points. Total protein content in the samples was quantified using a BSA assay
(Pierce; Waltham, MA). A total of 5 μg protein from each sample was diluted to 10
μl in the gel-loading buffer, and samples were loaded in each lane of a 10% Novex®
Zymogram Gelatin Gel (Invitrogen; Carlsbad, CA). The protein samples were resolved by polyacrylamide gel electrophoresis (SDS-PAGE) in a Tris-Glycine-
SDS running buffer under non-denaturing conditions. After the electrophoresis, the gels were carefully removed from the assembly and in 1X Zymogram Renaturing
Buffer for 30 minutes at RT with gentle agitation. The solution was removed and gels were equilibrated in 1X Zymogram Developing Buffer for 30 minutes at RT with gentle agitation. The buffer was decanted and replaced with fresh 1X
Zymogram Developing Buffer and further incubated at 37oC overnight to allow the metalloprotease activity. Next day the gels were washed by rinsing three times (5 minutes per wash) with deionized water under gentle agitation at RT. The gels were stained by submerging in a solution of SimplyBlueTM SafeStain (Invitrogen) for 1-4 hours under gentle agitation at RT. The gels were destained by washing three times with deionized water as before to reveal the clear zones of gelatinase activity on the gels at the location corresponding to MMPs. Finally, the gels were 103 placed on a plastic sheet protector and scanned using a gel imaging instrument
(BioRad). Images were analyzed by ImageJ software (NIH) to quantify the bands
(zones of clearance) produced due to gelatinase activity of the metalloproteases.
3.3.4 Endothelial Cells Vascular Tube Formation Assay
To assess the capacity of capillary-like tubes by HUVEC in response to visfatin treatment, a tube formation assay was performed by culturing endothelial cells on basement matrix as described previously (238, 239). A basement matrix,
MatrigelTM (Corning; Corning, NY) was used to coat the culture surface.
Specifically, MatrigelTM was thawed overnight on an ice bath kept at 4oC. Using pre-chilled pipette tips, 50 μl of the fluid MatrigelTM was placed per well of 96-well plate kept on ice. The plate was incubated at 37oC for 30 minutes to allow the basement matrix to gel. Cells at a density of 15,000 in 100μl were added to each
o well and incubated at 37 C humidified air with 5% CO2 for 2-8 hours to allow the formation of the capillary network. The visfatin treatment was added simultaneously. After two hours, the plates were checked under a phase contrast microscope every hour to visually analyze the formation of tubes in response to treatment. The experiment was optimized for the time of tube formation and concluded after the vessel sprouting (tube formation) was observed. The assay was repeated at least two times with each sample (controls, and unknown) represented in triplicates.
104
3.3.5 Fluorescence Microscopy and Image Analysis
Fluorescence microscopy was used to visualize and analyze the angiogenic activity of HUVEC in response to visfatin treatments. To fluorescently label the cells, a 50 μl solution of 6 μM Calcein AM was added to each well and plates were
o incubated at 37 C and in 5% CO2 for 20 minutes. The cells were observed and photographed using a fluorescent inverted microscope (Leica, DMI 6000) with 485 nm excitation or 520 nm emission filter. The images for brightfield and green channel fluorescence were acquired from 3 random fields from each well and analyzed for the vascular features by ImageJ software (NIH).
3.3.6 Statistical Analysis
For all the experiments, the plotting of data and statistical analysis was performed by using GraphPad Prism software. Data are presented as Mean ±
SEM. The significance of difference amongst differentially treated groups (control and unknown) was tested by a two-tailed student’s t-test. Results yielding a p- value <0.05 were considered significant for the difference between the compared groups. Each experiment was repeated at least two times with each treatment group containing at least three independent replicates per experiment to obtain statistically relevant results.
105
3.4 Results
3.4.1 Visfatin Upregulates the Expression of Angiogenic Markers in Human
Endothelial Cells
Obesity-induced disruption of metabolic homeostasis and inflammation is partially due to inadequate vascular supply to the enlarged adipose tissue (240). It has been speculated that adipose tissue secretions (e.g. adipokines) may act in a local and systemic fashion to influence angiogenesis (129). To verify the contribution of the adipokine visfatin in context of angiogenesis, we measured the expression of multiple angiogenesis-associated genes in HUVEC. As seen in
Figure 3.1, visfatin treatment resulted in an upregulation in the RNA expression of multiple angiogenic markers, including ANGPT1, ANGPT2, CXCL11, uPA,
Angiogenin, VEGFR1, VEGFR2, and MMP2. These genes regulate different aspects of angiogenesis and may have effect in both central and peripheral to the adipose microenvironment. Notably, visfatin treatment increased both ANGPT1 and ANGPT2 in a time-dependent manner. These findings are relevant from the pathogenic standpoint as both of these genes are heavily implicated in regulating tumor angiogenesis and thus provide a crucial link between angiogenic and inflammatory pathways (241, 242). 106
Figure 3.1 Visfatin upregulates the expression of multiple angiogenic markers in HUVEC. Time course of relative mRNA quantification of multiple angiogenesis-associated genes expressed in HUVEC after 1 h and 4 h post-visfatin treatment (100 ngml-1). Levels are shown for (A) ANGPT1, (B) ANGPT2, (C) MMP2, (D) Angiogenin, (E) uPA, (F) CXCL11, (G) VEGFR1, and (H) VEGFR2. *p<0.05, **p<0.001, ***p<0.001 vs Control (black bar), (n=3). ANGPT1, Angiopoietin 1; ANGPT2, Angiopoietin 2; MMP2, Matrix metalloproteinase 2; uPA, urokinase plasminogen activator; CXCL11, CXC-chemokine ligand 11; VEGFR, Vascular endothelial growth factor receptor. 107
3.4.2 Visfatin Enhances the Production of Functional Matrix Metalloproteases in
Human Endothelial Cells
The formation of vascular network is a function of angiogenesis and extracellular matrix (ECM) modification that is dependent on the reciprocal interaction between vascular cells and the components of the ECM. Matrix metalloproteases (MMPs) are a class of zinc-dependent proteases that regulate several functions, including ECM modification, cytokine availability, and tissue remodeling in various physiological and pathological conditions (243, 244). It has been reported that the expression of MMPs is increased in inflammatory conditions, such as obesity, atherosclerosis, and some cancers. In line with these findings and in context of adipose microenvironment, we sought to determine if visfatin has an influence on MMP action. To test the effect of visfatin to influence the MMP activity, gelatin zymography was used. Briefly, HUVEC were treated with recombinant visfatin and culture supernatants (containing secreted MMPs) were collected, and loaded on a 10% zymogram gelatin gel (Novex) and electrophoresed to resolve protein bands. The gels were renatured, and incubated at 37oC to reactivate MMPs and allow proteolysis of gelatin in the gel matrix. The
Zymographs were stained and developed, and photographed for the band density quantification using an image analysis software. As seen in Figure 3.2, visfatin treatment resulted in an enhanced gelatinolytic activity of MMP2 (65 kDa) and
MMP9 (88 kDa). The band density, or ‘zone of gelatin clearance’ was quantified using ImageJ, and data are shown in Figure 3.3. Notably, the visfatin-stimulated 108 increase in MMP9 activity was time dependent. Visfatin-mediated stimulation of
MMPs in endothelial cells highlight the relevance of this adipokine in abnormal angiogenesis and tissue remodeling and further consolidate its role in the regulation of inflammatory responses.
Figure 3.2 Visfatin treatment enhances the production of functional matrix metalloproteases (MMPs) in HUVEC in a time-dependent manner. HUVEC were treated with visfatin (100 ngml-1) for 6h, 12h, and 24 h to obtain the culture conditioned medium for the detection of functional metalloproteinases by gelatin zymography. Representative zymogram shows the gelatinase activity (clear bands) by MMP2 (65 kDa) and MMP9 (88 kDa).
109
Figure 3.3 Quantification of visfatin-stimulated gelatinolytic activity of MMPs. Zymogram gels were developed and photographed, and gelatinase activity of functional metalloproteinases was quantified using the ‘gel analysis’ feature of ImageJ. The mean area of gelatin clearance (band density) is shown above for (A) MMP2, and (B) MMP9. *p<0.05 vs Untreated (control, black bar), (n=6).
3.4.3 Visfatin Shows an Angiogenic Effect and Causes an Early Induction of
Vascular Tube Formation in Human Endothelial Cells
When plated on the reconstituted basement membrane extracellular matrix, such as Matrigel, endothelial cells readily form capillary-like tubular networks. The tube formation assay has long been used as an established measure to study angiogenesis for both in vitro and ex vivo applications (245), and various standard protocols are available to validate this effect (238, 239, 246). We investigated the role of visfatin in its capacity to stimulate tube formation in vitro using a modified protocol as described by Arnautova and Kleinman (238). Briefly, HUVEC were 110 cultured on surfaces coated with growth factor-reduced Matrigel and treated with recombinant visfatin for varied durations ranging from 4-16 hours. After the incubation, cells were carefully labeled with a fluorescent dye – Calcein AM and visualized using fluorescence microscopy. The photographs were obtained from 2 random fields per well and 3 wells were used for each treatment condition per experiment to obtain statistically significant observations. Upon treatment with visfatin for longer durations (over 8 hours), no difference in the degree of tube formation was observed between the various treatment groups (data not shown).
For a comparison, Figures 3.4 and 3.5 indicate that even at 6-hours post- treatment, there was no distinguishable difference observed between the groups.
Interestingly, the angiogenic effect of visfatin was rapid and clearly observable at an early time point of 4 hours [Figure 3.6], where a visual difference was observed for vascular sprouting, branching, and tube formation in the visfatin-treated group.
The vascular features – number of tubes and number of branch points – were quantified using ImageJ, and these parameters were found to be significantly higher under visfatin treatment in proportion to the dose [Figure 3.7]. These data suggest that visfatin can act a potent angiogenic factor and can stimulate morphological changes in endothelial cells.
111
Figure 3.4 Vascular tube formation assay showing a capillary-like network in HUVEC cultured on MatrigelTM. Representative micrographs of endothelial capillary-tube formation on basement matrix – MatrigelTM after 6 h with, or without visfatin supplementation (100 ngml-1). Scale bar - 200μM. Upper panel – phase contrast; lower panel – fluorescence (calcein-AM).
Figure 3.5 Analysis of vascular features in HUVEC after 6 hours visfatin treatment. Vascular features in HUVEC after 6 h visfatin treatment (100 ngml-1) were counted from multiple fields and quantified using ImageJ. Data shows the average number of (A) Number of tubes (branches), and (B) No. of nodes (branch points), (n=6). 112
Figure 3.6 Visfatin treatment stimulates a rapid vascular tube formation in HUVEC cultured on MatrigelTM. Representative micrographs of endothelial-tube formation on MatrigelTM after 4 hours with, or without visfatin supplementation (100 ngml-1). Scale bar – 100uM. Upper panel – phase contrast; lower panel – fluorescence (calcein-AM).
Figure 3.7 Quantification of visfatin-induced vascular features in HUVEC. The vascular features formed in HUVEC after 4 h visfatin treatment were quantified from multiple fields using ImageJ. Data shows the average number of (A) Number of tubes (branches), and (B) No. of nodes (branch points). **p<0.01 vs 0 (control, grey bar), (n=6). 113
3.5 Discussion
Substantial evidence exists to suggest that angiogenesis and adipogenesis are coupled processes, and both adipose and vascular compartments interact reciprocally in the tissue microenvironment to regulate physiological function and homeostasis. Obesity-induced hypertrophy of the adipose tissue results in a metabolic and oxygen deficit leading to hypoxia and concomitant production of pro- inflammatory and pro-angiogenic factors to cope with the stress response (74). It has been proposed that therapeutic targeting of angiogenesis can be an effective strategy to treat obesity and restore the metabolic balance at the level of adipose tissue. Interestingly, similar anti-angiogenic strategies have been extensively researched in the context of tumor progression and have shown promise in clinical trials (247, 248). In fact, hypoxia-triggered angiogenesis and immunoinfiltration is a common feature in both adipose and tumor microenvironment, and it is possible that common molecular mediators may be involved. The molecular processes that guide the specialized morphological and functional responses in the endothelial cell during microvessel formation have several commonalities between adipogenesis and tumor growth. Hence, strategies to target pro-angiogenic processes may be effective towards the treatment of both obesity and cancer.
Studies indicate that adipose tissue-derived factors can influence angiogenic response, and indeed several inflammatory adipokines, including TNFα, IL6, leptin, and resistin, have been reported to play an important role in angiogenesis
(186). However, studies describing their direct vascular effects and pro-angiogenic 114 gene expression profile are limited. Here we investigated the role of the pro- inflammatory adipokine visfatin for its capacity to induce angiogenic responses in human endothelial cells (HUVEC). We show that treatment of HUVEC with recombinant human visfatin – (i) significantly upregulated the mRNA expression of multiple angiogenic markers, (ii) increased the functional activity of matrix metalloproteases, and (iii) stimulated the vascular tube formation in vitro.
We determined visfatin-induced angiogenic marker expression and found an increase in the relative expression of multiple genes. Visfatin increased the
RNA levels of angiopoietins – ANGPT1 and ANGPT2 – in a time-dependent manner, with a greater increase in ANGPT2 levels proportional to ANGPT1 [Figure
3.1(A-B)]. Angiopoietins are the ligands for the TIE2 receptor (ANGPT-TIE2) and are extensively studied in the context of their therapeutic relevance to cancer (241,
242). ANGPT1 binding to TIE2 leads to downstream signaling regulating vascular homeostatic responses, such as endothelial cell survival, endothelial barrier maintenance, and quiescent vasculature. While on the other hand, ANGPT2 acts as an antagonist and promotes vascular leakage, endothelial activation, and disruption of endothelial barrier towards angiogenesis. Studies have shown that higher ANGPT2/ANGPT1 ratio is correlated with inflammation and poor outcome in several malignancies, including acute myeloid leukemia, breast cancer, multiple myeloma, hepatocellular carcinoma, colorectal cancer, and prostate cancer (241).
Therapeutic interventions that target ANGPT-TIE2 signaling for anti-tumorigenic responses are already in preclinical and clinical trial stages. The visfatin-mediated 115 higher increase in ANGPT2 relative to ANGPT1 underscores its capability to promote pro-inflammatory action and its implication in cancer development via the
ANGPT-TIE2 system. Other angiogenic markers that we found to be upregulated in response to visfatin were – Urokinase-type plasminogen activator (uPA),
Angiogenin, and CXCL11 [Figure 3.1 (C-E)]. Our results are relevant from a carcinogenesis-standpoint, as signaling by uPA-uPAR, angiogenin, and CXCL11 has been characterized in the context of tumor angiogenesis, where these molecules can synergistically influence tumorigenesis by exerting pro-angiogenic effects (249–251). For instance, in vitro uPA/uPAR downregulation was shown to suppress TIE2 phosphorylation and angiogenin-mediated angiogenesis in human microvascular cells (252). In a separate study, uPA/uPAR downregulation in a cellular model of pancreatic carcinoma was shown to suppress angiogenin with a corresponding increase in the chemokine secretion (249). CXCL11 has been shown to exert a tumorigenic and pro-angiogenic effect in epithelial ovarian cancer cell line overexpressing mesenchymal factors (251). We also found that visfatin increased the mRNA expression of VEGFR1, and VEGFR2 (p>0.05) [Figure 3.1
(F-G)]. Both receptors can transduce a variety of downstream effects in endothelial cells; including survival, inflammation, and angiogenesis in response to vascular endothelial growth factor (VEGF). Interestingly, it was recently shown that
VEGFR1 can be stimulated by a VEGF isoform to promote angiogenesis and improve the metabolic profile and insulin sensitivity in obese mice (253). These findings are consistent with the previous studies, where visfatin was reported to 116 induce production of soluble angiogenic regulators, VEGF and FGF, and regulate angiogenic responses in microvascular endothelial cells (254, 255). Further, we found that visfatin treatment resulted in enhanced gelatinolytic activity of matrix metalloproteinases, MMP2 and MMP9 [Figures 3.2-3.3]. The release of these enzymes is stimulated by inflammatory signaling, and MMP-mediated degradation of ECM components has been described as a crucial process in the neovessels growth (256, 257).
Previously, we confirmed the role of visfatin as a pro-inflammatory factor
(Chapter 2). Inflammation-driven angiogenesis is a hallmark of the progression of diseases, including cancer (182). To test the angiogenic capability of visfatin, we performed an in vitro endothelial tube formation assay (Matrigel assay) and found that visfatin treatment promoted a rapid tube formation (4.5 hours) compared to untreated control [Figures 3.6]. However, upon longer incubation, this angiogenic response could not be distinguished [Figures 3.4]. Recently, Park et al. reported a regulatory role of visfatin in breast carcinoma through notch-1 upregulation (258).
It is possible that oncogenic effect of visfatin is partially due to an upregulation of pro-angiogenic responses as mentioned above.
Our results on visfatin-mediated overexpression of multiple angiogenesis- associated genes and its capability to induce morphogenic changes in endothelial cells are consistent with the previous findings and indicate that visfatin may be a potent contributor for pathologies, such as tumor progression, where abnormal angiogenesis is involved. Given the positive association of visfatin in inflammatory 117 disorders, and that inflammation and angiogenesis are closely linked, we provide evidence for the possible role of visfatin as a molecular connection for the cellular crosstalk mediating disease progression. Therapeutic targeting of visfatin may be beneficial to reduce adipogenesis and tumor progression.
118
CHAPTER 4: IDENTIFICATION OF SIGNAL TRANSDUCTION PATHWAYS
GOVERNING VISFATIN-INDUCED PRO-INFLAMMATORY EXPRESSION IN
HUMAN ENDOTHELIAL CELLS
4.1 Abstract
It has been previously shown that extracellular visfatin can stimulate pro- inflammatory expression in human endothelial cells. However, not enough information is available for signaling pathways responsible for the regulation of visfatin-mediated pro-inflammatory effect. We found that treatment with visfatin activated ERK-MAPK pathway in human endothelial cells. Using co-incubation with signal pathway specific inhibitors, we identified that visfatin-stimulated pro- inflammatory mRNA expression is predominantly mediated by p38-MAPK and
NFκB signaling pathways. We also demonstrate that pharmacological inhibition of visfatin with a specific inhibitor FK866 suppressed pro-inflammatory the secretion of pro-inflammatory chemokines by human endothelial cells. These findings highlight the relevance of visfatin in pathogenesis and its importance as a therapeutic target.
4.2 Introduction
In Chapter 2, we provided detailed information about the visfatin-induced pro-inflammatory expression of multiple cytokine-chemokine targets in human endothelial cells towards their enhanced capacity to attract and physically adhere to THP1 monocytes. But details regarding visfatin-induced signaling in endothelial cells are limited, and it is possible that visfatin treatment can activate multiple 119 signaling pathway regulators to influence inflammatory functions in the tissue microenvironment. It is therefore important to investigate the visfatin-associated signal transduction to understand its regulatory effect and therapeutic value. To evaluate the effect of visfatin in MAPK pathway activation, we used a cell-based
ELISA to quantify the intracellular levels of activated (phosphorylated) ERK1/2 relative to its total fraction in response to visfatin treatment. Pathway-specific inhibitors are potent molecules that have been extensively studied for their antitumor capacity and have been used in numerous studies for the identification of signaling pathways stimulated in response to a particular stimulus (259–264).
We used pathway-specific pharmacological inhibitors to identify the molecular pathways responsible for extracellular visfatin-induced expression of pro- inflammatory and pro-angiogenic genes in human endothelial cells (HUVEC). The information of specific inhibitors, their target pathway, and corresponding reference are listed in Table 4.1. We observed that visfatin-stimulated upregulation of its downstream effectors involved multiple major signaling pathways, including
NFκB, p38-MAPK, JNK, ERK-MAPK, and PI3K. In addition, we also investigated if pharmacological inhibition of visfatin using its specific inhibitor, FK866, can influence visfatin-induced chemokine secretion. We report that visfatin can stimulate multiple pathways to regulate gene expression in human endothelial cells, and its pharmacological inhibition can be an effective strategy to suppress its pro-inflammatory effect.
120
Table 4.1
Molecular pathway inhibitors used in the study and their mode of inhibition
Inhibitor Target Pathway Step of Inhibition Reference(s)
Bay11-7085 NFκB IκB-α Phosphorylation (69, 228, 259, 265–267)
SB202190 p38-MAPK p38 Kinase (260, 268–274)
SP600125 JNK-MAPK c-Jun phosphorylation (261, 275–282)
U0126 ERK-MAPK MEK1/2 Kinase (41, 58, 61–69)
Wortmannin PI3K PI3 Kinase (262, 264, 291–295)
4.3 Materials and Methods
4.3.1 Cell Culture and Treatments
Primary Human Umbilical Vein Endothelial Cells, derived from umbilical vein from pooled donors (HUVEC, CC-2519), were purchased from Lonza
(Walkersville, MD) and cultured in a complete growth medium – Endothelial
Growth Medium-2 (EGM-2) as per manufacturer’s guidelines. To formulate EGM-
2, contents of the EGM™-2 SingleQuots™ Kit (Lonza Catalog No. CC-4176 containing human epidermal growth factor [hEGF], vascular endothelial growth factor [VEGF], R3-insulin-like growth factor-1 [R3-IGF-1], ascorbic acid, hydrocortisone, human fibroblast growth factor-beta [hFGF-β], heparin, fetal bovine serum [FBS], and gentamicin/amphotericin-B [GA]) were added to EBM™-
2 Basal Medium. Prior to seeding the cells, culture plates were coated with gelatin
(ScienCell, Carlsbad, CA) to allow the cells to efficiently adhere to the surface. For the experiments, HUVEC between passages 2-5 were used. Cells were seeded in 121
0.1% gelatin-coated, multi-well culture dishes and grown until the formation of a monolayer. Based on the experiments, the adipokine treatments were given in a low-serum medium (EBM-2, 0.5% FBS) for the indicated durations, and samples were collected for further analysis.
4.3.2 Pharmacological Inhibitors
To identify the signal transduction pathways involved in visfatin-stimulated gene expression, HUVEC were pretreated for 60 minutes with pathway-specific inhibitors, followed by treatment with visfatin (100 ngml-1) for the indicated duration.
Unless otherwise indicated, the inhibitors and their working concentration used were as follows: NFκB inhibitor (BAY11-7085, 10 μM), p38-MAPK inhibitor
(SB202190, 10 μM), JNK inhibitor (SP600125, 25 μM), ERK-MAPK inhibitor
(U0126, 10 μM), and PI3K inhibitor (Wortmannin, 0.1 μM). FK866, a chemical compound known to block the activity of visfatin as described previously (296–
298), was used with varying concentrations (0-100nM) to assess its effect on cell viability, and pharmacological inhibition of extracellular visfatin for the subsequent assay of chemokine secretion. For all these experiments, the solvent DMSO (10
μl) was mixed in the experimental groups (control and test samples) to maintain the proper treatment conditions.
4.3.3 Cell-Based Enzyme-Linked Immunosorbent Assay
An ELISA-based assay using fluorogenic substrates to measure phosphorylated ERK1/ERK2 in whole cells (R&D Systems; Minneapolis, MN) was used to determine the activation of MAPK pathway in HUVEC in response to 122 visfatin treatment. HUVEC were cultured on 0.1% gelatin-coated surface in a 96- well plate; upon monolayer formation, cells were treated with visfatin (100 ngml-1), or complete medium EGM-2, for 5-30 minutes. After the experiment, the media was removed and cells were fixed with 4% formaldehyde in 1X PBS (100μl per well) for 20 minutes at room temperature (RT), and permeabilized in a quenching buffer (0.6% H2O2; 100μl per well) for 20 minutes at RT. The wells were washed three times with a wash buffer (5 minutes per wash), and a primary antibody mixture was added to each well and plates were incubated overnight at 4oC. Next day, the solution was removed, wells were washed as before, and a secondary antibody mixture was applied to each well followed by incubation at RT for two hours. The solution was removed and wells were washed as before and a final wash with 1X PBS was given. A solution of fluorogenic substrates was added to each well, and the plate was incubated at RT protected from light for 30-45 minutes. Data were acquired using a fluorescence plate reader to read the plate first with excitation at 540 nm and emission at 600 nm (represent phosphorylated
ERK1/2), and then reading with excitation at 360 nm and emission at 450 nm
(represent total ERK1/2).
4.3.4 Real-Time Quantitative PCR (RT-qPCR)
RT-qPCR was used to measure the RNA expression of chemokines (MCP1,
GMCSF, IL6, CXCL2, and CXCL8), adhesion markers (PECAM1, ICAM1, VCAM1, and E-Selectin), and angiogenesis-associated genes (ANGPT1, and ANGPT2) in
HUVEC treated with of visfatin alone, or in combination with signal pathway 123 inhibitors. Primer information for the target genes can be found in Appendix 1.
Reactions were carried out using Luminaris HiGreen qPCR Master Mix (Life
Technologies; Grand Island, NY) (193). GAPDH was used as a reference gene or housekeeping internal control gene (194–196). The gene expression of chemokines and adhesion markers was normalized to GAPDH and expressed as fold change relative to the untreated control using 2-ddCt method (197). The reactions were carried out in a CFX ConnectTM Real-Time PCR detection system
(BioRad; Hercules, CA).
4.3.5 Enzyme-Linked Immunosorbent Assay (ELISA)
A sandwich-ELISA system (Peprotech; Rocky Hill, NJ) was used for the detection of secreted proteins produced by HUVEC. High-binding 96 well plates were coated with 50μl of capture antibody diluted to 1μg in PBS and incubated overnight at RT. Wells were washed three times with 300μl of wash buffer (0.05%
Tween-20 in PBS) using a multichannel pipette, and the surface was blocked with a blocking buffer (1% BSA in PBS) for 1 hours at RT. Wells were washed as before and unknown samples or standards were applied and plates were incubated overnight at 4oC for antigen binding. Next day, wells were washed as before and incubated at RT for 2 hours in 50μl of biotinylated detection antibody solution
(diluted to 0.5μg in wash buffer plus 0.1% BSA). The wells were washed as before and incubated at RT for 30 minutes in 50μl of a streptavidin-HRP (1:2000) solution.
The washes were repeated as before and incubated with 100μl of a colorimetric substrate ABTS (Life Technologies; Grand Island, NY) at RT for 25-30 minutes. 124
The microplates were read using a plate reader at 405 nM to measure the absorbance values due to color developed in proportion to the quantity of target antigen.
4.3.6 Statistical Analysis
For all the experiments, the plotting of data (Mean ± SEM) and statistical analysis was performed by using GraphPad Prism software. The data values for differentially treated groups (control and unknown) were tested by a two-tailed student’s t-test. Multiple comparisons between the groups were performed using
ANOVA with a posthoc analysis by the Tukey-Kramer multiple comparisons test.
Results-producing a p-value <0.05 were considered significant for the difference between the compared groups. Each experiment was repeated at least twice with each treatment group containing at least three independent replicates per experiment to obtain statistically relevant results.
4.4 Results
4.4.1. Visfatin Treatment Activates ERK1/2 MAP-Kinase Pathway in
Human Endothelial Cells
Mitogen-activated protein kinases (MAPK) consists of a family of evolutionarily conserved kinases that transduce extracellular signals to regulate several fundamental cellular processes, including cell growth, differentiation, migration, angiogenesis, and inflammation. Dysregulated or aberrant activation of
MAPK pathways are linked to pathologies, such as cancer, neurodegenerative disorders, cardiovascular diseases and obesity and have been extensively 125 reviewed (154, 299–303). The activation of extracellular signal-regulated kinase
1/2 (ERK1/2) arm of MAPK pathway has been shown to be crucial for adipocyte differentiation and adipogenesis, and have been implicated in inflammatory conditions, such as asthma (284), rheumatoid arthritis (304), and cancers (301,
303). Based on these findings, it is possible that ERK1/2 pathway may also be involved in the regulation of pro-inflammatory and angiogenic gene expression, and cytokine secretion induced by the adipokine visfatin in human endothelial cells, as shown previously [Chapter 2, Figures 2.2, 2.6-2.10, 2.11, 2.13; Chapter 3,
Figure 3.1]. To determine if treatment with visfatin stimulates the activation of the
MAPK-ERK1/2 pathway, we used a cell-based ELISA (R&D Systems) to quantify the relative proportion of phosphorylated ERK1/2 to total ERK1/2 in a chemiluminescent detection method. To accomplish this, HUVEC were cultured in a 96-well plate in triplicates for each condition per experiment and treated with recombinant visfatin (100 ngml-1) for different durations (5-30 minutes). After treatment, cells were immediately fixed with 4% formaldehyde to preserve the instantaneous phosphorylation state, and samples were processed according to the manufacturer’s guidelines. As shown in Figure 4.1, treatment with visfatin increased the activation (phosphorylation) of ERK 1/2 in human endothelial cells.
The amount of phosphorylated ERK1/2 was highest at 5-minutes post-treatment with visfatin compared to the untreated sample, while a number of total ERK1/2 remained unchanged across all time points [Figure 4.1]. The above results show 126 the relevance of MAPK-ERK pathway in visfatin-stimulated pro-inflammatory and pro-angiogenic gene expression in human endothelial cells.
Figure 4.1 Visfatin stimulates the activation of ERK1/2-MAP-Kinase in HUVEC. HUVEC were treated with visfatin (100 ngml-1) and activation of intracellular ERK1/2 MAPK was determined using a cell-based ELISA. Data shows the fluorescence values proportional to the levels of (A) Phosphorylated ERK1/2, (B) total ERK1/2, and (C) ratio of phosphorylated to total ERK1/2. The complete growth medium EGM2 (containing multiple growth factors and 2% FBS) was used as a positive control. **p<0.01, ***p<0.001 vs 0 (untreated control), (n=3). EGM2, Endothelial growth medium; ERK1/2, Extracellular signal-regulated kinase. 127
4.4.2. Visfatin Treatment Upregulates Multiple Signaling Pathways to Stimulate the Gene Expression of Pro-Inflammatory Chemokines, Adhesion Markers, and
Angiogenic Markers in Human Endothelial Cells
The chronic, low-grade inflammation in obesity has been described as a major link to several pathologies, including cancer (305). Plasma proteins and soluble mediators such as adipose tissue-derived cytokines may act on components of adipose microenvironment and elicit signaling pathways that further potentiate the inflammatory response (306). In response to inflammatory stimuli, endothelial cells are typically restored to an ‘activated’ state and exhibit increased expression of chemokines and adhesion makers, leukocyte adhesion and extravasation, and aberrant angiogenesis (198, 230, 307). Several published reports suggest that the inflammatory phenotype in endothelial cells can be a result of activation of multiple intracellular signaling pathways, including IKKα/β-NFκB pathway (195, 198, 307), p38-MAPK pathway (272–274, 276), ERK-MAPK pathway (289, 308, 309), JNK-MAPK pathway (195, 276, 280–282), and PI3K-Akt pathway (198, 295). Molecular crosstalk can take place between the inflammatory signaling pathways to generate the downstream effector molecules that translate the function, such as leukocyte-endothelial interaction (5, 198, 288, 310). One reliable way for the general identification of the signaling pathways in the stimulated cells is pathway inhibitor approach. The pathway inhibitor approach has been widely used in the context of inflammation to identify the signaling mechanism, or to study the effect of a drug, both in vitro and in vivo (195, 259– 128
261, 266, 277, 284, 286, 288, 290–292, 311). A list of chemical inhibitors used, their target pathways, and corresponding references is presented in Table 4.1. To assess for any cytotoxic effect and a suitable dose, CellTiter non-radioactive proliferation (viability) assay (Promega) was performed with a varied concentration of inhibitor treatment of HUVEC. As shown in Figure 4.2, no cytotoxicity was observed in response to inhibitor treatments. For each pathway inhibitor, the highest concentration was chosen for pathway analysis in the subsequent experiments.
129
Figure 4.2 Cell proliferation assay to determine the effect of pharmacological inhibitor treatment reveals no cytotoxicity within the tested dose range. HUVEC were incubated with the increasing dose of pathway inhibitors, or solvent (DMSO) alone for 8 h and assayed for cytotoxicity using MTS assay (Promega). Levels are shown for (A) BAY 11-7085, (B) SB202190, (C) SP600125, (D) U0126, and (E) Wortmannin. DMSO, Dimethyl sulfoxide. 130
We aimed to identify the signal pathways responsible for visfatin-induced pro-inflammatory gene expression in human endothelial cells. To accomplish this, we used commercially available chemical inhibitors of the target pathway to abrogate a specific signal loop and measured the resultant effect on the target gene expression. Specifically, HUVEC were incubated with solvent alone
(untreated control, DMSO), or with pathway inhibitors for 60 minutes (pre- treatment) followed by the addition of visfatin and further incubation for 1-4 hours.
After the treatment, total RNA was isolated, reversed transcribed, and mRNA expression was measured by real-time qPCR for chemokines, CCL2 (MCP1),
GMCSF, IL6, CXCL2 (GROβ), and CXCL8 (IL8); adhesion markers, PECAM1,
ICAM1, VCAM1, and E-selectin; and angiogenic markers, ANGPT1 and ANGPT2.
Consistent with the previous results [Chapter 2, Figures 2.6-2.10, 2.14; Chapter 3,
Figure 3.1A&B], treatment with visfatin resulted in an increase in inflammatory gene expression profile. Notably, a visfatin-mediated increase in RNA expression of chemokines was observed to be predominantly mediated by NFκB and p38-
MAPK pathway [Figures 4.3-4.7]. Specifically, increase in CCL2 (MCP1) appeared to be regulated by NFκB, p38-MAPK, and JNK, with some contribution from ERK-
MAPK pathway [Figure 4.3]; increase in GMCSF was regulated by NFκB, p38-
MAPK, and JNK [Figure 4.4]; increase in IL6 was regulated by NFκB, p38-MAPK, and PI3K [Figure 4.5]; increase in CXCL2 (GROβ) was regulated by NFκB and p38-MAPK [Figure 4.6], and increase in CXCL8 (IL8) was regulated by p38-MAPK
[Figure 4.7]. Interestingly, it was also observed that PI3K was predominantly 131 involved in negative regulation of CCL2 (MCP1), GMCSF, CXCL2 (GROβ), and
CXCL8 (IL8) [Figures 4.3-4.4, 4.6-4.7]. This was consistent with the multiple previously published findings highlighting the role of the PI3K pathway in the negative regulation of inflammatory signaling in innate immune responses (293,
312–314).
Figure 4.3 Visfatin upregulates the expression of CCL2 via NFκB, p38-MAPK, and JNK signaling pathway in HUVEC. HUVEC were pretreated for 60 mins with the pathway inhibitors, or solvent (DMSO) followed by the treatment with visfatin (100 ngml-1) for further 1 h. Data show the relative quantification levels for the mRNA expression of CCL2 (MCP1). Levels are shown relative to untreated control (Unt., first bar). *p<0.05 vs Unt.; δp<0.05, δδp<0.01, δδδp<0.001 vs visfatin; (n=3). CCL2 (MCP1), CC-chemokine ligand 2 (Monocyte chemotactic protein 1). 132
Figure 4.4 Visfatin-stimulated upregulated expression of GMCSF in HUVEC is mediated by NFκB, p38-MAPK, and JNK signaling pathway. HUVEC were pretreated for 60 mins with the pathway inhibitors, or solvent (DMSO) followed by the treatment with visfatin (100 ngml-1) for further 1 h. Data show the relative quantification levels for the mRNA expression of GMCSF. Levels are shown relative to untreated control (Unt., first bar). ***p<0.001 vs Unt.; δp<0.05, δδδp<0.001 vs visfatin; (n=3). GMCSF, Granulocyte-macrophage colony stimulating factor.
133
Figure 4.5 Visfatin-stimulated increase in the expression of IL6 in HUVEC is mediated by NFκB, p38-MAPK, and PI3K pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors, or solvent (DMSO) followed by the treatment with visfatin (100 ngml-1) for further 1 h. Data show the relative quantification levels for the mRNA expression of IL6. Levels are shown relative to untreated control (Unt., first bar). ***p<0.001 vs Unt.; δp<0.05, δδp<0.01, δδδp<0.001 vs visfatin; (n=3). IL6, Interleukin 6.
134
Figure 4.6 Visfatin increases the expression of CXCL2 in HUVEC via NFκB and p38-MAPK pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors, or solvent (DMSO) followed by the treatment with visfatin (100 ngml-1) for further 1 h. Data show the relative quantification levels for the mRNA expression of CXCL2 (GROβ). Levels are shown relative to untreated control (Unt., first bar). **p<0.01 vs Unt.; δδp<0.01, δδδp<0.001 vs visfatin; (n=3). CXCL2 (GROβ), CXC-chemokine ligand 2 (Growth-related oncogene β).
135
Figure 4.7 Visfatin-stimulated increase in the expression of CXCL8 in HUVEC is mediated via p38-MAPK pathway. HUVEC were pretreated for 60 mins with the pathway inhibitors, or solvent (DMSO) followed by the treatment with visfatin (100 ngml-1) for further 1 h. Data show the relative quantification levels for the mRNA expression of CXCL8 (IL8). Levels are shown relative to untreated control (Unt., first bar). **p<0.01 vs Unt.; δδp<0.01, δδδp<0.001 vs visfatin; (n=3). CXCL8 (IL8), CXC-chemokine ligand 8 (Interleukin 8).
136
In addition to determining the regulation at the level of RNA, we also determined the expression at the protein level for the chemokines CCL2 (MCP1),
CCXL2 (GROβ), and CXCL8 (IL8). Briefly, HUVEC were pretreated with pathway inhibitors, or the solvent (DMSO) for 30 minutes as above, followed by the addition of visfatin and further incubation for 8 hours. After the treatment, culture supernatants were collected and secretory levels of chemokines were determined by ELISA. As presented in Figure 4.8, we observed a reduced secretion of all three chemokines CCL2, CXCL2, and CXCL8, by visfatin-stimulated HUVEC in the presence of NFκB and p38-MAPK inhibitor. Together with the results of RNA expression above, it can be inferred that both NFκB and p38-MAPK pathways are predominantly involved in the transduction of visfatin-stimulated chemokine expression and secretion in human endothelial cells.
137
Figure 4.8 Visfatin increases the secretion of chemokines CCL2, CXCL2, and CXCL8 is regulated by p38-MAPK and NFκB pathways. HUVEC were pretreated with the solvent (DMSO), or pathway inhibitors for 30 min followed by the treatment with visfatin (100 ngml-1) for 8 h. Culture supernatants were assayed by ELISA to determine the levels of chemokines released in the medium. Levels are shown for (A) CCL2 (MCP1), (B) CXCL2 (GRO-β), and (C) CXCL8 (IL8). **p<0.01 vs Unt.; δp<0.05, δδp<0.01, δδδp<0.001 vs visfatin; (n=5). CCL2 (MCP1), CC-chemokine ligand 2 (Monocyte chemotactic protein 1); CXCL2 (GROβ), CXC-chemokine ligand 2 (Growth-related oncogene β); CXCL8 (IL8), CXC-chemokine ligand 8 (Interleukin 8). 138
It is well established that inflammation or stress can activate NFκB pathway, and stimulation of NFκB can lead to an enhanced expression of adhesion markers in endothelial cells (69, 315–318). Additionally, MAPK pathway, specifically p38-
MAPK activation can also be involved in the positive regulation of adhesion molecules, both independently (273, 317, 319) or in a concerted manner with NFκB
(317, 320, 321). In our study, pathway inhibitor analysis revealed that visfatin- induced transcription of adhesion molecules was a function of both NFκB and p38-
MAPK activation. Notably, both NFκB and p38 were implicated in visfatin-induced mRNA expression of E-selectin [Figure 4.9], ICAM1 [Figure 4.10], and VCAM1
[Figure 4.11], while only p-38-MAPK was implicated in PECAM1 [Figure 4.12].
139
Figure 4.9 Visfatin upregulates the expression of E-selectin in HUVEC involves NFκB, p38-MAPK, JNK, and ERK-MAPK pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors or solvent (DMSO) alone, followed by the treatment with visfatin (100 ngml-1) for further 4 h. Data show the relative quantification levels for the mRNA expression of E-selectin. Levels are shown relative to untreated control (Unt., first bar). ***p<0.001 vs Unt.; δp<0.05, δδδp<0.001 vs visfatin; (n=3).
140
Figure 4.10 Visfatin-stimulated increase in the expression of intercellular cell adhesion molecule-1 (ICAM1) in HUVEC involves NFκB and p38-MAPK pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors or solvent (DMSO) alone, followed by the treatment with visfatin (100 ngml-1) for further 4 h. Data show the relative quantification levels for the mRNA expression of ICAM1. Levels are shown relative to untreated control (Unt., first bar). **p<0.01 vs Unt.; δp<0.05, δδp<0.01, δδδp<0.001 vs visfatin; (n=3). ICAM1, Intercellular cell adhesion molecule1.
141
Figure 4.11 Visfatin-stimulated increase in the expression of vascular cell adhesion marker 1 (VCAM1) in HUVEC is mediated by NFκB and p38-MAPK pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors or solvent (DMSO) alone, followed by the treatment with visfatin (100 ngml-1) for further 4 h. Data show the relative quantification levels for the mRNA expression of VCAM1. Levels are shown relative to untreated control (Unt., first bar). ***p<0.001 vs Unt.; δδδp<0.001 vs visfatin; (n=3). VCAM1, Vascular cell adhesion molecule 1.
142
Figure 4.12 Visfatin upregulates the expression of platelet-endothelial cell adhesion molecule 1 (PECAM1) in HUVEC via p38-MAPK pathway. HUVEC were pretreated for 60 mins with the pathway inhibitors or solvent (DMSO) alone, followed by the treatment with visfatin (100 ngml-1) for further 4 h. Data show the relative quantification levels for the mRNA expression of PECAM1. Levels are shown relative to untreated control (Unt., first bar). **p<0.01 vs Unt.; δp<0.05, δδp<0.01, p0.07 vs visfatin; (n=3). PECAM1, Platelet endothelial cell adhesion molecule 1.
143
In the recent years, there has been an increasing focus on the overlapping mechanisms and mediators that integrate the obesity-associated pathogenesis and cancer. Angiopoietin family (ANGPT) of proteins have been shown to mediate a pro-angiogenic and pro-inflammatory effect, and elevated serum levels of
ANGPT2 have been described as a predictive marker for colorectal cancer in obesity (322). We evaluated the gene expression of angiopoietins (ANGPT1 and
ANGPT2) in response to visfatin treatment, alone or in combination with pathway inhibitors, in human endothelial cells. We observed that visfatin significantly increased the expression of ANGPT1 with the involvement of NFκB, p38-MAPK,
JNK, ERK-MAPK, and PI3K [Figure 4.13]; while the visfatin-induced increase in transcription of ANGPT2 involved NFκB, p38-MAPK, JNK, and ERK-MAPK [Figure
4.14].
144
Figure 4.13 Visfatin-induced expression in the levels of Angiopoietin 1 (ANGPT1) in HUVEC involves multiple signaling pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors or solvent (DMSO) alone, followed by the treatment with visfatin (100 ngml-1) for 1 h. Data show the relative quantification levels for the mRNA expression of ANGPT1. Levels are shown relative to untreated control (Unt., first bar). ***p<0.001 vs Unt.; δp<0.05, δδp<0.01, δδδp<0.001 vs visfatin; (n=3). ANGPT1, Angiopoietin 1.
145
Figure 4.14 Visfatin-stimulated upregulation for the expression of Angiopoietin 2 (ANGPT2) in HUVEC involves NFκB, p38-MAPK, JNK, and ERK-MAPK pathways. HUVEC were pretreated for 60 mins with the pathway inhibitors or solvent (DMSO) alone, followed by the treatment with visfatin (100 ngml-1) for 1 h. Data show the relative quantification levels for the mRNA expression of ANGPT2. Levels are shown relative to untreated control (Unt., first bar). ***p<0.001 vs Unt.; δp<0.05, δδp<0.01, δδδp<0.001 vs visfatin; (n=3). ANGPT2, Angiopoietin 2.
146
The involvement of multiple molecular pathways for visfatin-stimulated angiopoietin expression suggests a complex regulation with a possible crosstalk and overlapping signaling mediators. Overall, these data indicate that visfatin- stimulated inflammatory gene expression is predominantly mediated by p38-
MAPK and NFκB pathways in endothelial cells [Figure 4.15], while visfatin has a strong capacity to activate multiple signaling pathways to regulate the expression of diverse gene targets related to inflammatory response.
Figure 4.15 Pathway regulation of visfatin-induced expression of inflammation- associated genes in HUVEC. The genes are grouped into three different functional categories: Cytokines/Chemokines, Adhesion Markers, and Angiogenic Markers. Green color denotes upregulation; red color denotes downregulation; no color denotes no effect.
4.4.3. Pharmacologic Inhibition of Visfatin Suppresses Pro-Inflammatory
Chemokine Secretion in Human Endothelial Cells
A low molecular weight compound FK866 was first described by Hasmann et al. and was shown to have a specific inhibitory effect on intracellular visfatin
(296). FK866 has been extensively used in the pharmacological studies targeting the intracellular enzymatic form of visfatin (NAMPT) mainly in the context of 147 cancer, both in vivo (323, 324) and in vitro (298, 325–327). We aimed to determine if FK866-mediated pharmacological inhibition can suppress visfatin-induced pro- inflammatory chemokine secretion in human endothelial cells. Cell proliferation
(viability) assay revealed no cytotoxic effect of this inhibitor in the human endothelial cells for the indicated dose and treatment duration [Figure 4.16]. Based on this data, the highest concentration of FK866 (100 nM) was chosen for subsequent experiments.
Figure 4.16 Cell proliferation assay to test the cytotoxic effect of pharmacological inhibitor FK866 reveals no adverse effect in HUVEC for the tested doses. HUVEC were treated with solvent alone (DMSO), or increasing concentrations of FK866 for 8 h and cell toxicity was determined by MTS assay.
HUVEC were incubated with varying concentrations of FK866 or solvent alone (DMSO, control) for 1 hour (pretreatment) followed by addition of visfatin, and further incubation for 8 hours. Culture supernatants were collected and ELISA 148 was used to measure the secretory levels of CCL2 (MCP1), CXCL2 (GROβ), and
CXCL8 (IL8). As illustrated in Figure 4.17, FK866 suppressed the visfatin-induced chemokine secretion in dose-dependent manner for all three targets. These data indicate that therapeutic targeting of visfatin can be used as an effective strategy to reduce the inflammatory milieu of chemokines and slow down the endothelial cell-mediated inflammatory responses.
Figure 4.17 FK866-mediated inhibition of visfatin suppresses the chemokine secretion in HUVEC. HUVEC were pretreated with FK866 for 1 h followed by the addition of visfatin (100 ngml-1) and further treatment for 8 h, after which the culture medium was assayed to quantify the secreted chemokines by ELISA. Levels are shown for (A) CCL2 (MCP1), (B) CXCL2 (GROβ), and (C) CXCL8 (IL8). *p<0.05, **p<0.01, ***p<0.001 vs Unt.; δp<0.05, δδp<0.01 vs visfatin; (n=5). CCL2 (MCP1), CC-chemokine ligand 2 (Monocyte chemotactic protein 1); CXCL2 (GROβ), CXC-chemokine ligand 2 (Growth-related oncogene β); CXCL8 (IL8), CXC-chemokine ligand 8 (Interleukin 8).
149
4.5. Discussion
Intracellular signaling pathways convert extracellular stimulus into physiological responses, such as gene expression, cytoskeleton changes, cell differentiation etc. Studies show that obesity leads to an upregulation of multiple intracellular signaling pathways in adipocytes and endothelial cells leading to pro- inflammatory gene expression, cytokine secretion, and morphological changes that facilitate leukocyte extravasation. Visfatin is an adipokine whose serum levels are increased in obesity and inflammatory disorders. We previously showed that treatment with visfatin stimulated the expression of several targets that regulate inflammatory and angiogenic functions (Chapters 2 and 3). Here, we have investigated the role of visfatin to for the intracellular signaling in human endothelial cells (HUVEC) and show that visfatin treatment resulted in the activation of ERK1/2
MAPK pathway [Figure 4.1]. In addition, using pathway-specific inhibitors, we identified multiple signaling pathways for visfatin-induced pro-inflammatory gene expression and cytokine secretion [Figures 4.3-4.14]. Finally, we show that pretreatment with a specific visfatin inhibitor, FK866, blocked the visfatin- stimulated inflammatory cytokines release [Figure 4.16].
NFκB activation is one of the most well-characterized central mechanism for immunity-associated responses and can be activated in response to inflammatory stimuli, including cytokines, tissue injury, or infection. NFκB activation is characterized in a multitude of inflammatory disorders, including rheumatoid arthritis, atherosclerosis, coronary artery disease, and cancers (265, 307, 328). 150
Given the discovery that NFκB pathway activation is also observed in obesity and insulin resistance, it was proposed that inflammation could be a mechanistic link between obesity and metabolic disorders. Indeed, several obesity-induced pro- inflammatory factors, such as TNFα, IL6 etc. and adipokines, such as visfatin can result in an activation of NFκB signaling in diverse cellular targets to exacerbate dysfunctional responses leading to increased immunoinfiltration and pathogenesis
(7, 329). In line with this notion, we observed that specific inhibition of NFκB by
Bay11-7085 suppressed visfatin-stimulated increase in the mRNA levels of multiple pro-inflammatory markers, including (i) cytokines – CCL2, GMCSF, IL6, and CXCL2 [Figures 4.3-4.6], (ii) adhesion markers – E-selectin, ICAM1, and
VCAM [Figures 4.9-4.11], and (iii) angiogenic molecules, ANGPT1, and ANGPT2
[Figures 4.13-4.14] in human endothelial cells. Since NFκB is involved in the inflammatory control of diverse pathologies, it can be speculated that visfatin may be a potential contributing factor for this response.
MAPK pathways control a variety of cellular responses and constitute three major intracellular components – (i) ERK1/2, that is predominantly activated by growth factors, and (ii) p38 and (iii) JNK, both of which can be activated in response to inflammatory cytokines and stress (thus are called a stress-activated protein kinases). Activation of all three MAPK pathways has been observed in inflammatory disorders, such as rheumatoid arthritis (330) and inflammatory bowel disease (331), where they controlled multiple responses ranging from inflammatory cytokine secretion, leukocyte adhesion and chemotaxis, and 151 angiogenesis. In addition, MAPK pathways are well characterized in the context of tumor biology and are often deregulated in multiple cancers (303). We observed that treatment with visfatin resulted in increased phosphorylation (activation) of
ERK1/2 MAPK pathway [Figure 4.1], indicating its possible involvement in the above-mentioned pathologies. Previously, Moschen et al. reported that visfatin can stimulate pro-inflammatory cytokine expression and immunoactivation of human
PBMC-derived monocytes predominantly by the activation of p38 MAPK pathway
(179). Consistently, we found that specific inhibition of p38 pathway by SB202190 abrogated the mRNA expression of (i) cytokines – CCL2, GMCSF, IL6, CXCL2, and CXCL8 [Figures 4.3-4.7], (ii) adhesion markers – E-selectin, ICAM1, VCAM1, and PECAM1 [Figures 4.9-4.12], and (iii) angiogenic molecules – ANGPT1 and
ANGPT2 [Figures 4.13-4.14] in human endothelial cells. Additionally, we observed that JNK inhibition by its specific inhibitor (SP600125) also suppressed visfatin- stimulated mRNA expression of CCL2, GMCSF, E-selectin, ANGPT1, and
ANGPT2 [Figures 4.3, 4.4, 4.9, 4.13, and 4.14, respectively]. It was previously reported inflammatory cytokine TNFα and free fatty acids can increase JNK activity
(33). In addition, several studies indicate JNK-mediated regulation in chronic inflammatory disorders, e.g. obesity was shown to increase widespread JNK activity in mice (33), and JNK deficient mice showed resistance to rheumatoid arthritis and atherosclerosis (332), and it was proposed that inhibition of JNK can be a potential strategy for the treatment of diabetic cardiomyopathy (277). With respect to these findings, it can be speculated that visfatin-induced JNK activity 152 and resultant pro-inflammatory expression may contribute to pathogenesis.
ERK1/2 MAPK has been well studied in the context of several chronic inflammatory conditions and cancer (299–301). We tested if ERK1/2 is also involved in visfatin- induced mRNA expression, and found that inhibition of ERK by U0126 resulted in the suppression of CCL2, E-selectin, ANGPT1, and ANGPT2 [Figures 4.3, 4.9,
4.13, and 4.14, respectively], however, it also resulted in the increased expression of GMCSF, IL6, CXCL2, ICAM1, VCAM1, and PECAM1 [Figures 4.4, 4.5, 4.6,
4.10, 4.11, and 4.12, respectively], suggesting a possibility of negative regulation, or a compensatory activation of other pathways to contribute to this effect.
PI3K pathways are another class of protein kinases that control a diversity of cellular responses, including growth, survival, migration, insulin sensitivity, and angiogenesis. In endothelial cells, the PI3K pathway is activated in response to growth factors (VEGF) and activates downstream regulators of survival and proliferation to influence neovascularization and other angiogenic responses
(333). In the context of inflammation, PI3K pathways have been implicated in both pro-inflammatory and anti-inflammatory responses through their effect on signaling mediators downstream of receptors of innate immunity (314). In our experiments, we observed a predominant negative role of PI3K for visfatin-induced pro- inflammatory expression as evidenced by the increased mRNA levels of these molecules under specific inhibition of PI3K pathway. Particularly, PI3K-specific inhibition leads to an increase in the mRNA expression of (i) cytokines – CCL2,
GMCSF, CXCL2, and CXCL8 [Figures 4.3-4.4, and 4.6-4.7], (ii) adhesion markers 153
– E-selectin, ICAM1, VCAM1, and PECAM1 [Figures 4.9-4.12], and (iii) angiogenic molecule, ANGPT2 [Figure 4.14]. These results for the negative regulation of PI3K for inflammatory expression was consistent with the previous findings. Guha et al. reported that blockade of PI3K with specific inhibitors significantly enhanced LPS- induced pro-inflammatory signaling by MAPK pathways (ERK1/2, p38, and JNK) and upregulated the expression of downstream inflammatory effectors in human monocytes (264). Another study by Schabbauer et al. suggested that in vivo blockade of the PI3K pathway by Wortmannin resulted in LPS-induced inflammation, coagulation, and pro-inflammatory cytokine secretion, and poor survivability in a mouse model of endotoxemia, suggesting a protective role of the
PI3K pathway (295). Based on these findings and our results, it can be suggested that visfatin-stimulated expression of pro-inflammatory mediators can be negatively influenced by sustained activation of PI3K, and thus, PI3K agonists may be used to partially limit endothelial inflammation in the adipose microenvironment.
Our results for visfatin-stimulated multiple signaling pathways are relevant, as visfatin can induce similar pro-inflammatory responses by acting on other cells of the microenvironment. Indeed, it has been shown that visfatin has diverse cellular targets, and by the activation of pathways, including NFκB, MAPK, and
PI3K in these cellular targets, visfatin can be a mediator of dysfunctional response leading to disease. However, a visfatin-specific receptor has not be identified thus far (112), hence future studies are needed in this direction to delineate the signaling scheme of visfatin before specific therapies can be devised. 154
FK866 (also called as Apo866) was first described by Hasman et al. (296) as a specific inhibitor of the intracellular enzymatic form of visfatin (NAMPT) and was shown to have a pro-apoptotic effect on hepatic carcinoma cells by the gradual depletion of intracellular NAD+. Since, FK866 has been used in several reports to assess the function of visfatin (both extracellular and intracellular) in the context of inflammatory disorders and cancers (168, 236, 298, 323, 325, 326, 334, 335). In our experiments, pre-treatment with FK866 (1 hour) resulted in the suppression of extracellular visfatin-stimulated release of pro-inflammatory chemokines CCL2,
CXCL2, and CXCL8 [Figure 4.16]. These molecules have been implicated in inflammatory disorders and tumor progression (336), and their reduction by pharmacological inhibition of visfatin further emphasizes the relevance of this adipokine in disease.
Overall, we conclude that extracellular visfatin has a capacity to induce signaling pathways that regulate widespread cellular responses, including inflammation and cell proliferation. This is the first report for the simultaneous identification of multiple signaling pathways for visfatin-induced pro-inflammatory gene expression in human endothelial cells. We found that p38 MAPK and NFκB pathways are the predominant regulators of visfatin-stimulated mRNA expression of chemokines, adhesion, and angiogenic markers [Figure 4.15]. Our results also highlight the use of pathway inhibitors as a potential strategy to suppress adipokine-mediated inflammatory processes. It is highly possible that visfatin- induced activation of multiple pathways results in signal integration and 155 convergence upon common regulators. Future studies to identify those common mediators will be helpful to provide clues for targeted therapies to impair visfatin- mediated inflammation. In conclusion, these results suggest that visfatin is a multifaceted molecule with a capability to activate multiple signaling pathways for the regulation of diverse cellular responses, and pharmacological inhibition of visfatin can be an effective strategy to reduce chemokine secretion and inflammatory disease progression.
156
CHAPTER 5: CONCLUSION AND RECOMMENDATION FOR FUTURE WORK
5.1 Summary of Results and Relevance of Study
The major findings of this research project are diagrammatically illustrated in Figure 5.1. In the present research, we performed a set of gene expression studies and functional analyses to characterize the role of adipokines, specifically visfatin, for inflammation using primary human umbilical vein endothelial cells.
Abnormal adipokine (visfatin) levels have been associated with a wide spectrum of diseases, including type 2 diabetes, coronary artery disease, atherosclerosis, neurological disorders, gallbladder dysfunction, rheumatoid arthritis, fatty liver disease, dyslipidemia, some cancers, and even aging (7, 112). Inflammatory processes, such as increased immunoinfiltration and cytokine secretion, are the major determinant of adipokine-mediated adverse effects. Indeed, conditions like obesity, or inflammatory disorders e.g. arthritis, have been found to be associated with increased circulatory levels of adipokines that support pro-inflammatory effect
(e.g. leptin) with a simultaneous downregulation of anti-inflammatory adipokines
(e.g. adiponectin) (7). Given that adipokines target multiple tissues and can be potential regulators of pathophysiology, it is therefore important to understand their mechanisms of action by which these effects are governed. Although, recent studies have emphasized their clinical relevance, the clinical trials with adipokine inhibitors have not been particularly promising and achieved mixed results (112,
337). 157
Figure 5.1 Diagrammatic model of visfatin-induced inflammatory responses in endothelial cells. Based on our results, we propose that recombinant visfatin promotes multiple inflammation-associated responses in endothelial cells. Extracellular visfatin (receptor unknown) can stimulate the phosphorylation of ERK1/2 as well as the activation of other kinases of MAPK pathway (e.g. JNK and p38) and NFκB pathway. Visfatin-mediated expression of multiple inflammation-associated genes, particularly cytokines and chemokines (blue labels), adhesion molecules (green labels), and angiopoietins (red labels), are predominantly mediated by p38 and NFκB pathways. On the other hand, PI3K pathway may be involved in the negative regulation of these genes. Additionally, visfatin treatment can also cause enhanced interaction with monocytes (migration and adhesion) and angiogenesis in endothelial cells.
Endothelial activation is often the first step towards aberrant leukocyte extravasation and immunoinfiltration, and subsequent inflammatory responses in the corresponding tissue. It is established that adipokines can target endothelial cells and modulate their activation state. In addition, adipokines can stimulate classical pro-inflammatory signaling, including NFκB pathway to govern 158 inflammatory processes – cytokine secretion – in which other cells of the microenvironment, including adipocytes, immune cells, and stromal cells all can participate in paracrine responses collectively leading to exacerbation of the metabolic status of the tissue (7, 210). Studies attempting to describe adipokine- mediated vascular functions are warranted to not only understand their function in disease progression but also to provide clues for the development of effective and targeted combination therapies.
Here, we developed a comprehensive adipokine responsive-cytokine profile in human endothelial cells – treated with leptin, vaspin, and visfatin – that showed several differentially expressed molecules (Chapter 2). We found that this treatment resulted in a significantly increased secretion of multiple cytokines with established functions in the context of inflammatory disorders and cancers. The adipokine visfatin has been a subject of interest due to its strong positive association with obesity and inflammatory disorders. Thus, we proposed that visfatin may have a capacity to exert a similar pro-inflammatory effect for endothelial activation. Consistently, we found that visfatin-treated primary human endothelial cells exhibited a significant upregulation in the gene expression of multiple chemokines – CCL2, GMCSF, IL6, CXCL2, and CXCL8 – and adhesion markers – E-Selection, ICAM1, VCAM1, and PECAM1. Importantly, this increase in the expression of several inflammatory markers was consistent with increased chemotactic ability of THP1 monocytes and their significantly enhanced adherent capacity to endothelial monolayers under the effect of visfatin. These data indicate 159 that visfatin has a strong capacity to exert a direct inflammatory effect on endothelial cells to promote their interaction with leukocytes, and hence, further underscores its importance as an inflammatory link and therapeutic relevance for targeted abrogation.
Dysfunctional angiogenesis has been implicated in the context of both adipose tissue expansion and tumorigenesis. Higher levels of visfatin have been associated with both responses and proposed to be a potential link between obesity and cancer (338). Our results with visfatin and human endothelial cells indicate its pro-angiogenic role, where visfatin treatment caused a significantly increased angiogenic marker expression, metalloprotease activity, and in vitro tube formation in human endothelial cells (Chapter 3). Further, we conducted pathway inhibitor analysis (Chapter 4) to broadly identify the contribution of the intracellular signaling pathway for visfatin-stimulated expression of pro- inflammatory markers. We found that p38-MAPK and NFκB were the predominant molecular pathways for visfatin-mediated upregulation of inflammation-associated genes, with a possible negative regulation by the PI3K pathway. Finally, we show that visfatin-specific pharmacological inhibition by an anti-tumor agent FK866 successfully suppressed the chemokine secretion by visfatin. We propose that
FK866-mediated targeted inhibition of visfatin may be an effective method to antagonize its function for inflammatory activation and leukocyte interaction n endothelial cells. Altogether, this research project provides crucial insights into the relevance of adipokines for their contribution to endothelial inflammation and 160 leukocyte interaction. We further propose that visfatin can be a potential regulator of inflammation and a link between obesity and its associated comorbidities, and our results from detailed in vitro analyses warrant the need for further specific studies involving the administration of visfatin in the experimental in vivo models to study it potential inflammatory role in physiological context. As such, Section 5.2 discusses future experimentation to gain a comprehensive insight into adipokine function using mice models.
In summary, the data presented in Chapter 2 support our hypothesis that treatment of primary human endothelial cells with the adipokine visfatin results in an upregulated pro-inflammatory expression profile and enhanced leukocyte interaction. Furthermore, the data presented in Chapter 3 support the evidence for a pro-angiogenic role of visfatin. Finally, the data presented in Chapter 4 further supports our hypothesis that the adipokine visfatin can stimulate classical inflammatory pathways to promote the expression of markers of endothelial activation. Figure 5.2 provides a concise summary of major conclusions derived from these research findings.
161
Figure 5.2 Summary of conclusions.
5.2 Future Directions and Preliminary Data in Murine System
In our studies, we demonstrated that treatment with recombinant adipokines, particularly visfatin, resulted in the overexpression of pro-inflammatory chemokines, adhesion markers, and angiogenic markers in primary human endothelial cells. Additionally, we also reported that visfatin-treatment induced several functional responses in the endothelial cells, such as enhanced capability for monocyte migration, monocyte adherence, and capillary-like vessel formation.
Finally, using pathway inhibitor analyses we demonstrated that visfatin-stimulated pro-inflammatory expression is predominantly mediated by p38-MAPK and NFκB 162 signal pathway. Overall, these results support the hypothesis for a pro- inflammatory function of visfatin and emphasize on its role as a potential link between obesity and disease. As such, some details remain to be investigated.
First, a specific receptor for the cytokine-like activity of visfatin has not been identified thus far, so the whole intracellular pathway for visfatin signaling is still unknown. We showed that treatment of visfatin resulted in the activation of multiple pathways, including NFκB, p38-MAPK, JNK, ERK-MAPK, and PI3K. Further studies are warranted to identify the upstream elements for visfatin signaling.
Previously, in a controversial report, visfatin was proposed as an insulin mimetic with a capacity to bind to the insulin receptor (IR) and promote glucose uptake in both the cultured cells and mice model of diabetes (160). Subsequent reports indicated that visfatin can activate IR in pancreatic β-cells (339), and may regulate insulin signaling and pro-inflammatory effect in chondrocytes (340). Additionally, it has been proposed that adipokines can regulate pro-inflammatory signaling in their target cells by activating Toll-like receptor (TLR)-mediated pathways of the innate immune system (341). Notably, in a recent analysis using computational modeling,
Camp et. al reported a structural homology between visfatin and a TLR4 ligand and showed that binding of visfatin to TLR4 may activate downstream NFκB activation (342). The activation of TLRs can be experimentally determined by the accumulation of downstream effectors (IRAK-1, MyD88, TRIF, TECAM-2, IRF-3) and subsequent NFκB activation (343). Overall, these studies indicate that extracellular visfatin may be a potential ligand for both IR and the TLRs for its 163 metabolic and inflammatory functions. To validate this claim for the visfatin- induced activation responses in cultured endothelial cells (in vitro), the activation of putative receptors can be determined by a Western blot-based detection. Briefly, cells will be cultured in the presence of visfatin and for various durations (5-120 minutes) followed by lysis and homogenization in the presence of phosphatase and protease inhibitors to obtain total protein samples. An equal amount (μg) of the protein samples will be loaded on a polyacrylamide gel and electrophoresed to resolve the protein bands. Total protein bands will be blotted on a nitrocellulose membrane, followed by an incubation with appropriate dilutions of primary
(phospho-IR, or TLR signal components) and secondary antibodies, and subsequent detection by antibody-conjugated enzymatic assays. The band density of the phosphorylated form of the target receptors (proportional to the activation) can be quantified using a densitometry software, or gel analysis feature of ImageJ
(NIH) and compared among treatment groups.
We showed that visfatin-induced chemokine expression involved NFκB and p38-MAPK pathway (Chapter 4). NFκB activation is marked by the phosphorylation of IKKα/β, and the subsequent degradation of its inhibitor (IκBα), resulting in the nuclear translocation of NFκB dimers where it can directly bind to its promoter elements. The experiments detailing these events are not documented herewith; thus, it would be of interest to perform these experiments to obtain relevant information. One way to determine endogenous activation of NFκB is through a luciferase-reporter assay [Appendix 3]. Briefly, plasmids containing a reporter 164 gene (luciferase) under the control of NFκB promoter are transfected in the endothelial cells using a suitable career (e.g. lipofectamine), followed by the culture of cells in an opaque, 96-well plate in the presence of visfatin for various durations
(5-60 minutes) to study the kinetics of NFκB transactivation. Cells can be lysed after the treatments and luciferase activity (proportional to NFκB activation) can be determined using a plate reader. The activation of other intracellular signaling factors, including IKKα/β, IκBα, and p38-MAPK can be determined by western blotting (as before) using phospho-specific antibodies.
Preliminary data in the murine system. To better understand the adipokine action in the physiological context, it is important to initiate the in vivo studies in mice. As a preliminary step towards this direction, we developed a comprehensive cytokine/growth factor profile in a murine pancreatic microvascular cell line, MS1 upon treatment with adipokines – visfatin, vaspin, and leptin [Figure 5.3]. MS1 is a well-established endothelial model of murine microvascular cells and has been extensively used in immunological, cancers, and metabolic studies (344–349).
MS1 cells were maintained in Dulbecco Modified Eagle Medium (DMEM) supplemented with 10% FBS. For the adipokine treatments, cells were grown in multi-well culture dishes and cultured in the presence of recombinant adipokines visfatin, vaspin, and leptin (100 ng/ml each). After 24 hours, the culture supernatant was collected and used for the detection of secreted proteins by a protein microarray (RayBio® Mouse Cytokine Antibody Array G-Series 3) following manufacturer’s protocol. The microarray slides were processed and sent to the 165 company for data acquisition. As demonstrated in Figure 5.3, treatment with each adipokine resulted in differential secretory levels of multiple proteins, which are known for their role in metabolic regulation, immunomodulation, and growth responses. This early-stage preliminary data provides initial direction towards the investigation of the various functional processes that can be regulated by the above-mentioned adipokines. For instance, we observed a marked upregulation in the levels of IGFBP6 – which has been reported for its role in cancer cell migration and angiogenesis – in response to all three adipokines. Also, VEGF, a master regulator of endothelial proliferation and angiogenesis, whose levels are positively associated with tumor progression, was found to be strongly upregulated in all treatments. This data provides clues for the potential involvement of these adipokines in a wide variety of cellular processes and disease development. Future studies are needed to enhance the understanding of adipokine biology to extend our knowledge for their potential role in obesity-associated pathogenesis.
166
Figure 5.3 Preliminary data in the murine system – differential regulation of cytokine and growth factor secretion in response to treatment with recombinant visfatin in murine endothelial cells (MS1). (A) Semi-quantitative cytokine and growth factor profile obtained upon assay of conditioned medium via protein microarray shows relative levels of secretory factors produced by MS1 endothelial cells in response to recombinant visfatin treatment (100 ngml-1) for 24 h. 167
Figure 5.4 Preliminary data in the murine system – differential regulation of cytokine and growth factor secretion in response to treatment with recombinant leptin in murine endothelial cells (MS1). (B) Semi-quantitative cytokine and growth factor profile obtained upon assay of conditioned medium via protein microarray shows relative levels of secretory factors produced by MS1 endothelial cells in response to recombinant leptin treatment (100 ngml-1) for 24 h. 168
Figure 5.5 Preliminary data in the murine system – differential regulation of cytokine and growth factor secretion in response to treatment with recombinant vaspin in murine endothelial cells (MS1). (C) Semi-quantitative cytokine and growth factor profile obtained upon assay of conditioned medium via protein microarray shows relative levels of secretory factors produced by MS1 endothelial cells in response to recombinant vaspin treatment (100 ngml-1) for 24 h. 169
The adipokine research has advanced exponentially in the recent years.
Recently, a WAT secretome was described that revealed over 600 proteins, including previously known and novel molecules. Given their involvement in the regulation of almost all physiological aspects, more studies like ours are needed to describe their targeted function in a context of disease and inflammation.
Visfatin, whose function we thoroughly characterized in this project for its inflammatory role, is now considered to be one of the most important adipokine with a strong therapeutic value and a potential marker for inflammatory disorders.
Still, there are caveats about its clinical efficacy due to mixed results (112), which further underscores the need for more investigation for its physiological function.
Similarly, other recently discovered adipokines may have a therapeutic value and a potential role in disease-related processes. Thus, more studies like this research are needed to constantly provide data to strengthen our existing understanding of the pathophysiology of obesity and help discover uncover novel therapies.
170
REFERENCES
1. M. F. Gregor, G. S. Hotamisligil, Inflammatory Mechanisms in Obesity, Annu. Rev. Immunol. 29, 415–445 (2011). 2. K. M. Flegal, M. D. Carroll, B. K. Kit, C. L. Ogden, Prevalence of obesity and trends in the distribution of body mass index among US adults, 1999-2010., JAMA 307, 491–7 (2012). 3. T.-D. Kanneganti, V. D. Dixit, Immunological complications of obesity, Nat. Immunol. 13, 707–712 (2012). 4. N. Ouchi, H. Kobayashi, S. Kihara, M. Kumada, K. Sato, T. Inoue, T. Funahashi, K. Walsh, Adiponectin Stimulates Angiogenesis by Promoting Cross-talk between AMP-activated Protein Kinase and Akt Signaling in Endothelial Cells, J. Biol. Chem. 279, 1304–1309 (2004). 5. M. F. Gregor, G. S. Hotamisligil, Inflammatory mechanisms in obesity., Annu. Rev. Immunol. 29, 415–45 (2011). 6. C. M. Kusminski, P. E. Bickel, P. E. Scherer, Targeting adipose tissue in the treatment of obesity-associated diabetes, Nat. Rev. Drug Discov. 15, 639–660 (2016). 7. N. Ouchi, J. L. Parker, J. J. Lugus, K. Walsh, Adipokines in inflammation and metabolic disease., Nat. Rev. Immunol. 11, 85–97 (2011). 8. D. E. Berryman, E. O. List, L. Sackmann-Sala, E. Lubbers, R. Munn, J. J. Kopchick, Growth hormone and adipose tissue: Beyond the adipocyteGrowth Horm. IGF Res. 21, 113–123 (2011). 9. Y.-Y. Chau, R. Bandiera, A. Serrels, O. M. Martínez-Estrada, W. Qing, M. Lee, J. Slight, A. Thornburn, R. Berry, S. McHaffie, R. H. Stimson, B. R. Walker, R. M. Chapuli, A. Schedl, N. Hastie, Visceral and subcutaneous fat have different origins and evidence supports a mesothelial source., Nat. Cell Biol. 16, 367–75 (2014). 10. P. K. Fazeli, M. C. Horowitz, O. A. MacDougald, E. L. Scheller, M. S. Rodeheffer, C. J. Rosen, A. Klibanski, Marrow fat and bone-new perspectivesJ. Clin. Endocrinol. Metab. 98, 935–945 (2013). 11. M. J. Devlin, A. M. Cloutier, N. A. Thomas, D. A. Panus, S. Lotinun, I. Pinz, R. Baron, C. J. Rosen, M. L. Bouxsein, Caloric restriction leads to high marrow adiposity and low bone mass in growing mice, J. Bone Miner. Res. 25, 2078–2088 (2010). 12. W. P. Cawthorn, E. L. Scheller, B. S. Learman, S. D. Parlee, B. R. Simon, H. Mori, X. Ning, A. J. Bree, B. Schell, D. T. Broome, S. S. Soliman, J. L. Delproposto, C. N. Lumeng, A. Mitra, S. V. Pandit, K. A. Gallagher, J. D. Miller, V. Krishnan, S. K. Hui, M. A. Bredella, P. K. Fazeli, A. Klibanski, M. C. Horowitz, C. J. Rosen, O. 171
A. Macdougald, Bone marrow adipose tissue is an endocrine organ that contributes to increased circulating adiponectin during caloric restriction, Cell Metab. 20, 368–375 (2014). 13. S. Sun, Y. Ji, S. Kersten, L. Qi, Mechanisms of inflammatory responses in obese adipose tissue., Annu. Rev. Nutr. 32, 261–86 (2012). 14. S. Sun, Y. Ji, S. Kersten, L. Qi, Mechanisms of Inflammatory Responses in Obese Adipose Tissue, Annu. Rev. Nutr. 32, 261–286 (2012). 15. A. Kosteli, E. Sugaru, G. Haemmerle, J. F. Martin, J. Lei, R. Zechner, A. W. Ferrante, Weight loss and lipolysis promote a dynamic immune response in murine adipose tissue, J. Clin. Invest. 120, 3466–3479 (2010). 16. X. Prieur, C. Y. L. Mok, V. R. Velagapudi, V. Núñez, L. Fuentes, D. Montaner, K. Ishikawa, A. Camacho, N. Barbarroja, S. O’Rahilly, J. K. Sethi, J. Dopazo, M. Orešič, M. Ricote, A. Vidal-Puig, Differential lipid partitioning between adipocytes and tissue macrophages modulates macrophage lipotoxicity and M2/M1 polarization in obese mice, Diabetes 60, 797–809 (2011). 17. P. A. Kern, M. Saghizadeh, J. M. Ong, R. J. Bosch, R. Deem, R. B. Simsolo, The expression of tumor necrosis factor in human adipose tissue. Regulation by obesity, weight loss, and relationship to lipoprotein lipase, J Clin Invest 95, 2111– 2119 (1995). 18. H. Xu, G. T. Barnes, Q. Yang, G. Tan, D. Yang, C. J. Chou, J. Sole, A. Nichols, J. S. Ross, L. A. Tartaglia, H. Chen, Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance, 112, 1821–1830 (2003). 19. L. Fontana, J. C. Eagon, M. E. Trujillo, P. E. Scherer, S. Klein, Visceral Fat Adipokine Secretion Is Associated With Systemic Inflammation in Obese Humans, 56, 4–7 (2007). 20. V. Cataln, J. Gomez-Ambrosi, B. Ramirez, F. Rotellar, C. Pastor, C. Silva, A. Rodriguez, M. J. Gil, J. A. Cienfuegos, G. Frhbeck, Proinflammatory cytokines in obesity: Impact of type 2 diabetes mellitus and gastric bypass, Obes. Surg. 17, 1464–1474 (2007). 21. M. E. Rausch, S. Weisberg, P. Vardhana, D. V Tortoriello, Obesity in C57BL/6J mice is characterized by adipose tissue hypoxia and cytotoxic T-cell infiltration., Int. J. Obes. (Lond). 32, 451–463 (2008). 22. S. Winer, Y. Chan, G. Paltser, D. Truong, H. Tsui, J. Bahrami, R. Dorfman, Y. Wang, J. Zielenski, F. Mastronardi, Y. Maezawa, D. J. Drucker, E. Engleman, D. Winer, H.-M. Dosch, Normalization of obesity-associated insulin resistance through immunotherapy., Nat. Med. 15, 921–9 (2009). 23. S. Nishimura, I. Manabe, M. Nagasaki, K. Eto, H. Yamashita, M. Ohsugi, M. Otsu, K. Hara, K. Ueki, S. Sugiura, K. Yoshimura, T. Kadowaki, R. Nagai, CD8+ 172 effector T cells contribute to macrophage recruitment and adipose tissue inflammation in obesity., Nat. Med. 15, 914–20 (2009). 24. R. V. Luckheeram, R. Zhou, A. D. Verma, B. Xia, CD4 +T cells: Differentiation and functionsClin. Dev. Immunol. 2012 (2012), doi:10.1155/2012/925135. 25. M. M. Tiemessen, A. L. Jagger, H. G. Evans, M. J. C. van Herwijnen, S. John, L. S. Taams, CD4+CD25+Foxp3+ regulatory T cells induce alternative activation of human monocytes/macrophages., Proc. Natl. Acad. Sci. U. S. A. 104, 19446– 19451 (2007). 26. M. Feuerer, L. Herrero, D. Cipolletta, A. Naaz, J. Wong, A. Nayer, J. Lee, A. B. Goldfine, C. Benoist, S. Shoelson, D. Mathis, Lean, but not obese, fat is enriched for a unique population of regulatory T cells that affect metabolic parameters., Nat. Med. 15, 930–939 (2009). 27. M. J. Davis, T. M. Tsang, Y. Qiu, J. K. Dayrit, J. B. Freij, G. B. Huffnagle, M. A. Olszewski, Macrophage M1/M2 polarization dynamically adapts to changes in cytokine microenvironments in Cryptococcus neoformans infection, MBio 4 (2013). 28. C. N. Lumeng, J. L. Bodzin, A. R. Saltiel, Obesity induces a phenotypic switch in adipose tissue macrophage polarization, 117, 175–184 (2007). 29. I. Murano, G. Barbatelli, V. Parisani, C. Latini, G. Muzzonigro, M. Castellucci, S. Cinti, Dead adipocytes, detected as crown-like structures, are prevalent in visceral fat depots of genetically obese mice, J. Lipid Res. 49, 1562–1568 (2008). 30. S. Sun, Y. Ji, S. Kersten, L. Qi, Mechanisms of inflammatory responses in obese adipose tissue., Annu. Rev. Nutr. 32, 261–86 (2012). 31. G. Cildir, S. C. Akıncılar, V. Tergaonkar, Chronic adipose tissue inflammation: all immune cells on the stage., Trends Mol. Med. 19, 487–500 (2013). 32. G. Solinas, M. Karin, JNK1 and IKKbeta: molecular links between obesity and metabolic dysfunction, Faseb J 24, 2596–2611 (2010). 33. J. Hirosumi, G. Tuncman, L. Chang, C. Z. Gorgun, K. T. Uysal, K. Maeda, M. Karin, G. S. Hotamisligil, A central role for JNK in obesity and insulin resistance, Nature 420, 333–336 (2002). 34. M. C. Arkan, A. L. Hevener, F. R. Greten, S. Maeda, Z.-W. Li, J. M. Long, A. Wynshaw-Boris, G. Poli, J. Olefsky, M. Karin, IKK-β links inflammation to obesity- induced insulin resistance, Nat. Med. 11, 191–198 (2005). 35. P. Peraldi, G. S. Hotamisligil, W. a Buurman, M. F. White, B. M. Spiegelman, Tumor necrosis factor (TNF)-alpha inhibits insulin signaling through stimulation of the p55 TNF receptor and activation of sphingomyelinase., J. Biol. Chem. 271, 13018–13022 (1996). 36. G. S. Hotamisligil, D. L. Murray, L. N. Choy, B. M. Spiegelman, Tumor necrosis 173 factor alpha inhibits signaling from the insulin receptor, Proc.Natl.Acad.Sci.U.S A 91, 4854–4858 (1994). 37. E. J. Park, J. H. Lee, G. Y. Yu, G. He, S. R. Ali, R. G. Holzer, C. H. Österreicher, H. Takahashi, M. Karin, Dietary and Genetic Obesity Promote Liver Inflammation and Tumorigenesis by Enhancing IL-6 and TNF Expression, Cell 140, 197–208 (2010). 38. A.-L. Dinel, C. André, A. Aubert, G. Ferreira, S. Layé, N. Castanon, Cognitive and emotional alterations are related to hippocampal inflammation in a mouse model of metabolic syndrome., PLoS One 6, e24325 (2011). 39. B. T. Jeon, E. A. Jeong, H. J. Shin, Y. Lee, D. H. Lee, H. J. Kim, S. S. Kang, G. J. Cho, W. S. Choi, G. S. Roh, Resveratrol attenuates obesity-associated peripheral and central inflammation and improves memory deficit in mice fed a high-fat diet, Diabetes 61, 1444–1454 (2012). 40. a D. Dobrian, Q. Ma, J. W. Lindsay, K. a Leone, K. Ma, J. Coben, E. V Galkina, J. L. Nadler, Dipeptidyl peptidase IV inhibitor sitagliptin reduces local inflammation in adipose tissue and in pancreatic islets of obese mice., Am. J. Physiol. Endocrinol. Metab. 300, E410-21 (2011). 41. L. E. Nicol, W. F. Grant, S. M. Comstock, M. L. Nguyen, M. S. Smith, K. L. Grove, D. L. Marks, Pancreatic inflammation and increased islet macrophages in insulin-resistant juvenile primates, J. Endocrinol. 217, 207–213 (2013). 42. I. M. Khan, X.-Y. Perrard, G. Brunner, H. Lui, L. M. Sparks, S. R. Smith, X. Wang, Z.-Z. Shi, D. E. Lewis, H. Wu, C. M. Ballantyne, Intermuscular and perimuscular fat expansion in obesity correlates with skeletal muscle T cell and macrophage infiltration and insulin resistance., Int. J. Obes. (Lond). 39, 1607–18 (2015). 43. M. Perreault, a Marette, Targeted disruption of inducible nitric oxide synthase protects against obesity-linked insulin resistance in muscle., Nat. Med. 7, 1138– 1143 (2001). 44. C. P. Lambert, N. R. Wright, B. N. Finck, D. T. Villareal, Exercise but not diet- induced weight loss decreases skeletal muscle inflammatory gene expression in frail obese elderly persons, J. Appl. Physiol. (Bethesda, Md. 1985) 105, 473–8 (2008). 45. M. B. Fessler, L. L. Rudel, J. M. Brown, Toll-like receptor signaling links dietary fatty acids to the metabolic syndrome, Curr. Opin. Lipidol. 20, 379–385 (2009). 46. H. Shi, M. V. Kokoeva, K. Inouye, I. Tzameli, H. Yin, J. S. Flier, TLR4 links innate immunity and fatty acid-induced insulin resistance, J. Clin. Invest. 116, 3015–3025 (2006). 47. R. W. Himes, C. W. Smith, Tlr2 is critical for diet-induced metabolic syndrome 174 in a murine model, FASEB J. 24, 731–739 (2010). 48. G. Chen, M. H. Shaw, Y.-G. Kim, G. Nuñez, NOD-like receptors: role in innate immunity and inflammatory disease, Annu. Rev. Pathol. 4, 365–398 (2009). 49. K. Schroder, R. Zhou, J. Tschopp, The NLRP3 Inflammasome: A Sensor for Metabolic Danger?, Science (80-. ). 327, 296–300 (2010). 50. D. De Nardo, E. Latz, NLRP3 inflammasomes link inflammation and metabolic diseaseTrends Immunol. 32, 373–379 (2011). 51. B. Vandanmagsar, Y.-H. Youm, A. Ravussin, J. E. Galgani, K. Stadler, R. L. Mynatt, E. Ravussin, J. M. Stephens, V. D. Dixit, The NLRP3 inflammasome instigates obesity-induced inflammation and insulin resistance, Nat. Med. 17, 179– 188 (2011). 52. D. Cai, M. Yuan, D. F. Frantz, P. A. Melendez, L. Hansen, J. Lee, S. E. Shoelson, Local and systemic insulin resistance resulting from hepatic activation of IKK-β and NF-κB, Nat. Med. 11, 183–190 (2005). 53. G. Tuncman, J. Hirosumi, G. Solinas, L. Chang, M. Karin, G. S. Hotamisligil, Functional in vivo interactions between JNK1 and JNK2 isoforms in obesity and insulin resistance, Proc. Natl. Acad. Sci. 103, 10741–10746 (2006). 54. B. F. Belgardt, J. Mauer, F. T. Wunderlich, M. B. Ernst, M. Pal, G. Spohn, H. S. Bronneke, S. Brodesser, B. Hampel, A. C. Schauss, J. C. Bruning, Hypothalamic and pituitary c-Jun N-terminal kinase 1 signaling coordinately regulates glucose metabolism, Proc. Natl. Acad. Sci. 107, 6028–6033 (2010). 55. G. Solinas, C. Vilcu, J. G. Neels, G. K. Bandyopadhyay, J. L. Luo, W. Naugler, S. Grivennikov, A. Wynshaw-Boris, M. Scadeng, J. M. Olefsky, M. Karin, JNK1 in Hematopoietically Derived Cells Contributes to Diet-Induced Inflammation and Insulin Resistance without Affecting Obesity, Cell Metab. 6, 386–397 (2007). 56. S. Dabo, E. F. Meurs, dsRNA-dependent protein kinase PKR and its role in stress, signaling and HCV infection, Viruses 4, 2598–2635 (2012). 57. S. Boura-Halfon, Y. Zick, Phosphorylation of IRS proteins, insulin action, and insulin resistance., Am. J. Physiol. Endocrinol. Metab. 296, E581--91 (2009). 58. J. F. Tanti, F. Ceppo, J. Jager, F. Berthou, Implication of inflammatory signaling pathways in obesity-induced insulin resistanceFront. Endocrinol. (Lausanne). 3 (2013), doi:10.3389/fendo.2012.00181. 59. A. Villaret, P. D. J. Galitzky, D. Este`ve, C. Marie-Adeline Marques, Sengene`s, M. L. Patrick Chiotasso, Tamara Tchkonia, J. L. Kirkland, And, A. Bouloumie, Adipose Tissue Endothelial Cells From Obese Human Subjects: Differences Among Depots in Angiogenic, Metabolic, and Inflammatory Gene Expression and Cellular Senescence, Diabetes 59, 2755–2763 (2010). 175
60. W. A. Muller, How endothelial cells regulate transmigration of leukocytes in the inflammatory response, Am. J. Pathol. 184, 886–896 (2014). 61. M. Pasarica, O. R. Sereda, L. M. Redman, D. C. Albarado, D. T. Hymel, L. E. Roan, J. C. Rood, D. H. Burk, S. R. Smith, Reduced adipose tissue oxygenation in human obesity evidence for rarefaction, macrophage chemotaxis, and inflammation without an angiogenic response, Diabetes 58, 718–725 (2009). 62. M. Blüher, Adipose tissue dysfunction in obesity, Exp. Clin. Endocrinol. Diabetes 117, 241–250 (2009). 63. K. M. Nieman, I. L. Romero, B. Van Houten, E. Lengyel, Adipose tissue and adipocytes support tumorigenesis and metastasis, Biochim. Biophys. Acta - Mol. Cell Biol. Lipids 1831, 1533–1541 (2013). 64. K. Landgraf, D. Friebe, T. Ullrich, J. Kratzsch, K. Dittrich, G. Herberth, V. Adams, W. Kiess, S. Erbs, A. Kor̈ner, Chemerin as a mediator between obesity and vascular inflammation in children, J. Clin. Endocrinol. Metab. 97 (2012). 65. E. Porreca, C. Di Febbo, L. Fusco, V. Moretta, M. Di Nisio, F. Cuccurullo, Soluble thrombomodulin and vascular adhesion molecule-1 are associated to leptin plasma levels in obese women, Atherosclerosis 172, 175–180 (2004). 66. D. Vestweber, How leukocytes cross the vascular endothelium, Nat. Rev. Immunol. 15, 692–704 (2015). 67. T. Garrood, L. Lee, C. Pitzalis, Molecular mechanisms of cell recruitment to inflammatory sites: General and tissue-specific pathways, Rheumatology 45, 250– 260 (2006). 68. M. P. Bevilacqua, J. S. Pober, D. L. Mendrick, R. S. Cotran, M. A. Gimbrone, Identification of an inducible endothelial-leukocyte adhesion molecule, Med. Sci. 84, 9238–9242 (1987). 69. I. Kim, S.-O. Moon, S. Hoon Kim, H. Jin Kim, Y. Soon Koh, G. Young Koh, Vascular Endothelial Growth Factor Expression of Intercellular Adhesion Molecule 1 (ICAM-1), Vascular Cell Adhesion Molecule 1 (VCAM-1), and E-selectin through Nuclear Factor- B Activation in Endothelial Cells, J. Biol. Chem. 276, 7614–7620 (2001). 70. R. V Stan, M. Kubitza, G. E. Palade, PV-1 is a component of the fenestral and stomatal diaphragms in fenestrated endothelia., Proc Natl Acad Sci U S A 96, 13203–13207 (1999). 71. J. Keuschnigg, T. Henttinen, K. Auvinen, M. Karikoski, M. Salmi, S. Jalkanen, The prototype endothelial marker PAL-E is a leukocyte trafficking molecule, Blood 114, 478–484 (2009). 72. Y. Cao, Angiogenesis modulates adipogenesis and obesityJ. Clin. Invest. 117, 2362–2368 (2007). 176
73. M. A. Rupnick, D. Panigrahy, C.-Y. Zhang, S. M. Dallabrida, B. B. Lowell, R. Langer, M. J. Folkman, Adipose tissue mass can be regulated through the vasculature., Proc. Natl. Acad. Sci. U. S. A. 99, 10730–10735 (2002). 74. D.-H. Kim, R. Gutierrez-Aguilar, H.-J. Kim, S. C. Woods, R. J. Seeley, Increased adipose tissue hypoxia and capacity for angiogenesis and inflammation in young diet-sensitive C57 mice compared with diet-resistant FVB miceInt. J. Obes. (2012), doi:10.1038/ijo.2012.141. 75. A. G. Arroyo, M. L. Iruela-Arispe, Extracellular matrix, inflammation, and the angiogenic response, Cardiovasc. Res. 86, 226–235 (2010). 76. R. Blanco, H. Gerhardt, VEGF and Notch in tip and stalk cell selection, Cold Spring Harb. Perspect. Med. 3 (2013), doi:10.1101/cshperspect.a006569. 77. D. Ribatti, E. Crivellato, “Sprouting angiogenesis”, a reappraisalDev. Biol. 372, 157–165 (2012). 78. I. Yana, H. Sagara, S. Takaki, K. Takatsu, K. Nakamura, K. Nakao, M. Katsuki, S. Taniguchi, T. Aoki, H. Sato, S. J. Weiss, M. Seiki, Crosstalk between neovessels and mural cells directs the site-specific expression of MT1-MMP to endothelial tip cells, J. Cell Sci. 120, 1607–1614 (2007). 79. F. De Smet, I. Segura, K. De Bock, P. J. Hohensinner, P. Carmeliet, Mechanisms of vessel branching: Filopodia on endothelial tip cells lead the way, Arterioscler. Thromb. Vasc. Biol. 29, 639–649 (2009). 80. J. R. Carmeliet P, Molecular mechanisms and clinical applications of angiogenesis., Nature 473, 298–307 (2011). 81. S. Lehr, S. Hartwig, H. Sell, Adipokines: A treasure trove for the discovery of biomarkers for metabolic disorders, Proteomics - Clin. Appl. 6, 91–101 (2012). 82. M. Fasshauer, M. Bluher Department, Adipokines in health and disease, Trends Pharmacol. Sci. 36, 461–470 (2015). 83. M. Khan, F. Joseph, Adipose Tissue and Adipokines: The Association with and Application of Adipokines in Obesity, Scientifica (Cairo). 2014, 1–7 (2014). 84. D. C. W. Lau, B. Dhillon, H. Yan, P. E. Szmitko, S. Verma, Adipokines: molecular links between obesity and atheroslcerosis., Am. J. Physiol. Heart Circ. Physiol. 288, H2031–H2041 (2005). 85. J. Vendrell, M. Broch, N. Vilarrasa, A. Molina, J. M. Gómez, C. Gutiérrez, I. Simón, J. Soler, C. Richart, Resistin, adiponectin, ghrelin, leptin, and proinflammatory cytokines: relationships in obesity., Obes. Res. 12, 962–71 (2004). 86. H. Xu, G. T. Barnes, Q. Yang, G. Tan, D. Yang, C. J. Chou, J. Sole, A. Nichols, J. S. Ross, L. A. Tartaglia, H. Chen, Chronic inflammation in fat plays a crucial role 177 in the development of obesity-related insulin resistance., J. Clin. Invest. 112, 1821– 1830 (2003). 87. T. Kadowaki, T. Yamauchi, Adiponectin and adiponectin receptorsEndocr. Rev. 26, 439–451 (2005). 88. B. S. Rosen, K. S. Cook, J. Yaglom, D. L. Groves, J. E. Volanakis, D. Damm, T. White, B. M. Spiegelman, Adipsin and complement factor D activity: an immune- related defect in obesity., Science 244, 1483–7 (1989). 89. M. Tabata, T. Kadomatsu, S. Fukuhara, K. Miyata, Y. Ito, M. Endo, T. Urano, H. J. Zhu, H. Tsukano, H. Tazume, K. Kaikita, K. Miyashita, T. Iwawaki, M. Shimabukuro, K. Sakaguchi, T. Ito, N. Nakagata, T. Yamada, H. Katagiri, M. Kasuga, Y. Ando, H. Ogawa, N. Mochizuki, H. Itoh, T. Suda, Y. Oike, Angiopoietin- like Protein 2 Promotes Chronic Adipose Tissue Inflammation and Obesity-Related Systemic Insulin Resistance, Cell Metab. 10, 178–188 (2009). 90. M. Hirasawa, K. Takubo, H. Osada, S. Miyake, E. Toda, M. Endo, K. Umezawa, K. Tsubota, Y. Oike, Y. Ozawa, Angiopoietin-like protein 2 is a multistep regulator of inflammatory neovascularization in a murine model of age-related macular degeneration, J. Biol. Chem. 291, 7373–7385 (2016). 91. H. Kanda, S. Tateya, Y. Tamori, K. Kotani, K. I. Hiasa, R. Kitazawa, S. Kitazawa, H. Miyachi, S. Maeda, K. Egashira, M. Kasuga, MCP-1 contributes to macrophage infiltration into adipose tissue, insulin resistance, and hepatic steatosis in obesity, J. Clin. Invest. 116, 1494–1505 (2006). 92. H. R. Zhou, E.-K. Kim, H. Kim, K. J. Claycombe, Obesity-associated mouse adipose stem cell secretion of monocyte chemotactic protein-1., Am. J. Physiol. Endocrinol. Metab. 293, E1153-8 (2007). 93. H. Sell, J. Laurencikiene, A. Taube, K. Eckardt, A. Cramer, A. Horrighs, P. Arner, J. Eckel, Chemerin is a novel adipocyte-derived factor inducing insulin resistance in primary human skeletal muscle cells, Diabetes 58, 2731–2740 (2009). 94. K. B. Goralski, T. C. McCarthy, E. A. Hanniman, B. A. Zabel, E. C. Butcher, S. D. Parlee, S. Muruganandan, C. J. Sinal, Chemerin, a novel adipokine that regulates adipogenesis and adipocyte metabolism, J. Biol. Chem. 282, 28175– 28188 (2007). 95. M. C. Ernst, C. J. Sinal, Chemerin: At the crossroads of inflammation and obesityTrends Endocrinol. Metab. 21, 660–667 (2010). 96. C. Chavey, G. Lazennec, S. Lagarrigue, C. Clapé, I. Iankova, J. Teyssier, J. S. Annicotte, J. Schmidt, C. Mataki, H. Yamamoto, R. Sanches, A. Guma, V. Stich, M. Vitkova, B. Jardin-Watelet, E. Renard, R. Strieter, A. Tuthill, G. S. Hotamisligil, A. Vidal-Puig, A. Zorzano, D. Langin, L. Fajas, CXC Ligand 5 Is an Adipose-Tissue Derived Factor that Links Obesity to Insulin Resistance, Cell Metab. 9, 339–349 178
(2009). 97. G. Gaich, J. Y. Chien, H. Fu, L. C. Glass, M. A. Deeg, W. L. Holland, A. Kharitonenkov, T. Bumol, H. K. Schilske, D. E. Moller, The effects of LY2405319, an FGF21 Analog, in obese human subjects with type 2 diabetes, Cell Metab. 18, 333–340 (2013). 98. F. F. Fisher, S. Kleiner, N. Douris, E. C. Fox, R. J. Mepani, F. Verdeguer, J. Wu, A. Kharitonenkov, J. S. Flier, E. Maratos-Flier, B. M. Spiegelman, FGF21 regulates PGC-1α and browning of white adipose tissues in adaptive thermogenesis, Genes Dev. 26, 271–281 (2012). 99. J. Xu, D. J. Lloyd, C. Hale, S. Stanislaus, M. Chen, G. Sivits, S. Vonderfecht, R. Hecht, Y. S. Li, R. A. Lindberg, J. L. Chen, D. Y. Jung, Z. Zhang, H. J. Ko, J. K. Kim, M. M. V??niant, Fibroblast growth factor 21 reverses hepatic steatosis, increases energy expenditure, and improves insulin sensitivity in diet-induced obese mice, Diabetes 58, 250–259 (2009). 100. I. S. Wood, B. Wang, J. R. Jenkins, P. Trayhurn, The pro-inflammatory cytokine IL-18 is expressed in human adipose tissue and strongly upregulated by TNFalpha in human adipocytes., Biochem. Biophys. Res. Commun. 337, 422–9 (2005). 101. C. Darimont, O. Avanti, F. Blancher, S. Wagniere, R. Mansourian, I. Zbinden, P. Leone-Vautravers, a Fuerholz, V. Giusti, K. Macé, Contribution of mesothelial cells in the expression of inflammatory-related factors in omental adipose tissue of obese subjects., Int. J. Obes. (Lond). 32, 112–20 (2008). 102. J. M. Harkins, N. Moustaid-Moussa, Y.-J. Chung, K. M. Penner, J. J. Pestka, C. M. North, K. J. Claycombe, Expression of interleukin-6 is greater in preadipocytes than in adipocytes of 3T3-L1 cells and C57BL/6J and ob/ob mice., J. Nutr. 134, 2673–2677 (2004). 103. B. K. Pedersen, M. Febbraio, Muscle-derived interleukin-6--a possible link between skeletal muscle, adipose tissue, liver, and brain., Brain. Behav. Immun. 19, 371–376 (2005). 104. V. Rotter, I. Nagaev, U. Smith, Interleukin-6 (IL-6) Induces Insulin Resistance in 3T3-L1 Adipocytes and Is, Like IL-8 and Tumor Necrosis Factor-α, Overexpressed in Human Fat Cells from Insulin-resistant Subjects, J. Biol. Chem. 278, 45777–45784 (2003). 105. K. Chen, F. Li, J. Li, H. Cai, S. Strom, A. Bisello, D. E. Kelley, M. Friedman- Einat, G. A. Skibinski, M. A. McCrory, A. J. Szalai, A. Z. Zhao, Induction of leptin resistance through direct interaction of C-reactive protein with leptin., Nat. Med. 12, 425–432 (2006). 106. Y. Zhang, R. Proenca, M. Maffei, M. Barone, L. Leopold, J. M. Friedman, Positional cloning of the mouse obese gene and its human homologue., Nature 179
372, 425–432 (1994). 107. V. Abella, M. Scotece, J. Conde, J. Pino, M. A. Gonzalez-Gay, J. J. Gómez- Reino, A. Mera, F. Lago, R. Gómez, O. Gualillo, Leptin in the interplay of inflammation, metabolism and immune system disorders, Nat. Rev. Rheumatol. 13, 100–109 (2017). 108. H. Cui, M. López, K. Rahmouni, The cellular and molecular bases of leptin and ghrelin resistance in obesity., Nat. Rev. Endocrinol. 13, 338–351 (2017). 109. Q. W. Yan, Q. Yang, N. Mody, T. E. Graham, C. H. Hsu, Z. Xu, N. E. Houstis, B. B. Kahn, E. D. Rosen, The adipokine lipocalin 2 is regulated by obesity and promotes insulin resistance, Diabetes 56, 2533–2540 (2007). 110. I. K. M. Law, A. Xu, K. S. L. Lam, T. Berger, T. W. Mak, P. M. Vanhoutte, J. T. C. Liu, G. Sweeney, M. Zhou, B. Yang, Y. Wang, Lipocalin-2 deficiency attenuates insulin resistance associated with aging and obesity, Diabetes 59, 872– 882 (2010). 111. T. B. Dahl, S. Holm, P. Aukrust, B. Halvorsen, Visfatin/NAMPT: a multifaceted molecule with diverse roles in physiology and pathophysiology., Annu. Rev. Nutr. 32, 229–43 (2012). 112. A. Garten, S. Schuster, M. Penke, T. Gorski, T. de Giorgis, W. Kiess, Physiological and pathophysiological roles of NAMPT and NAD metabolism, Nat. Rev. Endocrinol. , 1–12 (2015). 113. JN Fain, HS Sacks, B. Buehrer, SWBahouth, E. Garrett, RYWolf, R. Carter, D. Tichansky, A. Madan, Identification of omentin mRNA in human epicardial adipose tissue: comparison to omentin in subcutaneous, internal mammary artery periadventitial and visceral abdominal depots., Int. J. Obes. (Lond). 32, 810–815 (2008). 114. H. Yamawaki, J. Kuramoto, S. Kameshima, T. Usui, M. Okada, Y. Hara, Omentin, a novel adipocytokine inhibits TNF-induced vascular inflammation in human endothelial cells, Biochem. Biophys. Res. Commun. 408, 339–343 (2011). 115. M. C. Alessi, D. Bastelica, A. Mavri, P. Morange, B. Berthet, M. Grino, I. Juhan-Vague, Plasma PAI-1 levels are more strongly related to liver steatosis than to adipose tissue accumulation, Arterioscler. Thromb. Vasc. Biol. 23, 1262–1268 (2003). 116. Y. Zhu, P. Carmeliet, W. P. Fay, Plasminogen activator inhibitor-1 is a major determinant of arterial thrombolysis resistance., Circulation 99, 3050–3055 (1999). 117. T. E. Graham, Q. Yang, M. Blüher, A. Hammarstedt, T. P. Ciaraldi, R. R. Henry, C. J. Wason, A. Oberbach, P.-A. Jansson, U. Smith, B. B. Kahn, Retinol- binding protein 4 and insulin resistance in lean, obese, and diabetic subjects., N. Engl. J. Med. 354, 2552–2563 (2006). 180
118. Q. Yang, T. E. Graham, N. Mody, F. Preitner, O. D. Peroni, J. M. Zabolotny, K. Kotani, L. Quadro, B. B. Kahn, Serum retinol binding protein 4 contributes to insulin resistance in obesity and type 2 diabetes, Nature 436, 356–362 (2005). 119. P. M. Moraes-Vieira, M. M. Yore, P. M. Dwyer, I. Syed, P. Aryal, B. B. Kahn, RBP4 activates antigen-presenting cells, leading to adipose tissue inflammation and systemic insulin resistance, Cell Metab. 19, 512–526 (2014). 120. M. Bokarewa, I. Nagaev, L. Dahlberg, U. Smith, A. Tarkowski, Resistin, an Adipokine with Potent Proinflammatory Properties, J. Immunol. 174, 5789–5795 (2005). 121. C. M. Steppan, S. T. Bailey, S. Bhat, E. J. Brown, R. R. Banerjee, C. M. Wright, H. R. Patel, R. S. Ahima, M. A. Lazar, The hormone resistin links obesity to diabetes, Nature 409, 307–312 (2001). 122. I. Nagaev, M. Bokarewa, A. Tarkowski, U. Smith, Human resistin is a systemic immune-derived proinflammatory cytokine targeting both leukocytes and adipocytes, PLoS One 1 (2006), doi:10.1371/journal.pone.0000031. 123. N. Ouchi, A. Higuchi, K. Ohashi, Y. Oshima, N. Gokce, R. Shibata, Y. Akasaki, A. Shimono, K. Walsh, Sfrp5 is an anti-inflammatory adipokine that modulates metabolic dysfunction in obesity., Science 329, 454–7 (2010). 124. R. W. O’Rourke, A. E. White, M. D. Metcalf, A. S. Olivas, P. Mitra, W. G. Larison, E. C. Cheang, O. Varlamov, C. L. Corless, C. T. Roberts, D. L. Marks, Hypoxia-induced inflammatory cytokine secretion in human adipose tissue stromovascular cells, Diabetologia 54, 1480–1490 (2011). 125. G. S. Hotamisligil, N. S. Shargill, B. M. Spiegelman, Adipose expression of tumor necrosis factor-alpha: direct role in obesity-linked insulin resistance, Science (80-. ). 259, 87–91 (1993). 126. K. Hida, J. Wada, J. Eguchi, H. Zhang, M. Baba, A. Seida, I. Hashimoto, T. Okada, A. Yasuhara, A. Nakatsuka, K. Shikata, S. Hourai, J. Futami, E. Watanabe, Y. Matsuki, R. Hiramatsu, S. Akagi, H. Makino, Y. S. Kanwar, Visceral adipose tissue-derived serine protease inhibitor: A unique insulin-sensitizing adipocytokine in obesity, Proc. Natl. Acad. Sci. 102, 10610–10615 (2005). 127. H. Li, W. Peng, J. Zhuang, Y. Lu, W. Jian, Y. Wei, W. Li, Y. Xu, Vaspin attenuates high glucose-induced vascular smooth muscle cells proliferation and chemokinesis by inhibiting the MAPK, PI3K/Akt, and NF-κB signaling pathways, Atherosclerosis 228, 61–68 (2013). 128. N. Klöting, P. Kovacs, M. Kern, J. T. Heiker, M. Fasshauer, M. R. Schön, M. Stumvoll, A. G. Beck-Sickinger, M. Blüher, Central vaspin administration acutely reduces food intake and has sustained blood glucose-lowering effects, Diabetologia 54, 1819–1823 (2011). 181
129. H. K. Sung, K. O. Doh, J. E. Son, J. G. Park, Y. Bae, S. Choi, S. M. L. Nelson, R. Cowling, K. Nagy, I. P. Michael, G. Y. Koh, S. L. Adamson, T. Pawson, A. Nagy, Adipose vascular endothelial growth factor regulates metabolic homeostasis through angiogenesis, Cell Metab. 17, 61–72 (2013). 130. J. I. Jung, H. J. Cho, Y. J. Jung, S. H. Kwon, S. Her, S. S. Choi, S. H. Shin, K. W. Lee, J. H. Y. Park, High-fat diet-induced obesity increases lymphangiogenesis and lymph node metastasis in the B16F10 melanoma allograft model: Roles of adipocytes and M2-macrophagesInt. J. Cancer (2014). 131. S. Nishimura, I. Manabe, M. Nagasaki, Y. Hosoya, H. Yamashita, H. Fujita, M. Ohsugi, K. Tobe, T. Kadowaki, R. Nagai, S. Sugiura, Adipogenesis in obesity requires close interplay between differentiating adipocytes, stromal cells, and blood vessels, Diabetes 56, 1517–1526 (2007). 132. V. Murahovschi, O. Pivovarova, I. Ilkavets, R. M. Dmitrieva, S. Döcke, F. Keyhani-Nejad, Ö. Gögebakan, M. Osterhoff, M. Kemper, S. Hornemann, M. Markova, N. Klöting, M. Stockmann, M. O. Weickert, V. Lamounier-Zepter, P. Neuhaus, A. Konradi, S. Dooley, C. Von Loeffelholz, M. Blüher, A. F. H. Pfeiffer, N. Rudovich, WISP1 Is a novel adipokine linked to inflammation in obesity, Diabetes 64, 856–866 (2015). 133. J. R. Grünberg, A. Hammarstedt, S. Hedjazifar, U. Smith, The Novel Secreted Adipokine WNT1-Inducible-Signaling Pathway Protein2/WISP2 is a Mesenchymal Cell Activator of Canonical WNT, J. Biol. Chem. (2014), doi:10.1074/jbc.M113.511964. 134. A. Hammarstedt, S. Hedjazifar, L. Jenndahl, S. Gogg, J. Grünberg, B. Gustafson, E. Klimcakova, V. Stich, D. Langin, M. Laakso, U. Smith, WISP2 regulates preadipocyte commitment and PPARγ activation by BMP4., Proc. Natl. Acad. Sci. U. S. A. 110, 2563–8 (2013). 135. Z. Chen, X. Ding, S. Jin, B. Pitt, L. Zhang, T. Billiar, Q. Li, WISP1-αvβ3 integrin signaling positively regulates TLR-triggered inflammation response in sepsis induced lung injury, Sci Rep 6, 28841 (2016). 136. N. Takahashi, W. Waelput, Y. Guisez, Leptin is an endogenous protective protein against the toxicity exerted by tumor necrosis factor., J. Exp. Med. 189, 207–212 (1999). 137. W. A. Banks, A. J. Kastin, W. Huang, J. B. Jaspan, L. M. Maness, Leptin enters the brain by a saturable system independent of insulin, Peptides 17, 305– 311 (1996). 138. C. Bjørbaek, Central leptin receptor action and resistance in obesity., J. Investig. Med. 57, 789–94 (2009). 139. J. Hebebrand, T. D. Muller, K. Holtkamp, B. Herpertz-Dahlmann, The role of leptin in anorexia nervosa: clinical implications., Mol. Psychiatry 12, 23–35 (2007). 182
140. F. Krempler, E. Hell, C. Winkler, D. Breban, W. Patsch, Plasma Leptin Levels. Interaction of Obesity With a Common Variant of Insulin Receptor Substrate-1, , 1686–1690 (1998). 141. A. Falorni, V. Bini, D. Molinari, F. Papi, F. Celi, G. Di Stefano, M. G. Berioli, M. L. Bacosi, G. Contessa, Leptin serum levels in normal weight and obese children and adolescents: relationship with age, sex, pubertal development, body mass index and insulin 415, Int.J Obes Relat Metab Disord. 21, 881–890 (1997). 142. S. Loffreda, S. Q. Yang, H. Z. Lin, C. L. Karp, M. L. Brengman, D. J. Wang, A. S. Klein, G. B. Bulkley, C. Bao, P. W. Noble, M. D. Lane, A. M. Diehl, Leptin regulates proinflammatory immune responses., FASEB J. 12, 57–65 (1998). 143. J. Santos-Alvarez, R. Goberna, V. Sánchez-Margalet, Human leptin stimulates proliferation and activation of human circulating monocytes., Cell. Immunol. 194, 6–11 (1999). 144. N. Kiguchi, T. Maeda, Y. Kobayashi, Y. Fukazawa, S. Kishioka, Leptin enhances CC-chemokine ligand expression in cultured murine macrophage, Biochem. Biophys. Res. Commun. 384, 311–315 (2009). 145. C. Grunfeld, C. Zhao, J. Fuller, A. Pollock, A. Moser, J. Friedman, K. R. Feingold, Endotoxin and cytokines induce expression of leptin, the ob gene product, in hamsters: A role for leptin in the anorexia of infection, J. Clin. Invest. 97, 2152–2157 (1996). 146. G. Matarese, A. Di Giacomo, V. Sanna, G. M. Lord, J. K. Howard, A. Di Tuoro, S. R. Bloom, R. I. Lechler, S. Zappacosta, S. Fontana, Requirement for leptin in the induction and progression of autoimmune encephalomyelitis., J. Immunol. 166, 5909–5916 (2001). 147. A. U. Momin, N. Melikian, A. M. Shah, D. J. Grieve, S. B. Wheatcroft, L. John, A. El Gamel, J. B. Desai, T. Nelson, C. Driver, R. A. Sherwood, M. T. Kearney, Leptin is an endothelial-independent vasodilator in humans with coronary artery disease: Evidence for tissue specificity of leptin resistance., Eur. Heart J. 27, 2294–2299 (2006). 148. a Bouloumie, T. Marumo, M. Lafontan, R. Busse, Leptin induces oxidative stress in human endothelial cells., FASEB J. 13, 1231–8 (1999). 149. P. Singh, M. Hoffmann, R. Wolk, A. S. M. Shamsuzzaman, V. K. Somers, Leptin induces C-reactive protein expression in vascular endothelial cells., Arterioscler. Thromb. Vasc. Biol. 27, e302–e307 (2007). 150. S. Aleffi, I. Petrai, C. Bertolani, M. Parola, S. Colombatto, E. Novo, F. Vizzutti, F. A. Anania, S. Milani, K. Rombouts, G. Laffi, M. Pinzani, F. Marra, Upregulation of proinflammatory and proangiogenic cytokines by leptin in human hepatic stellate cells., Hepatology 42, 1339–1348 (2005). 183
151. K. Hida, J. Wada, J. Eguchi, H. Zhang, M. Baba, A. Seida, I. Hashimoto, T. Okada, A. Yasuhara, A. Nakatsuka, K. Shikata, S. Hourai, J. Futami, E. Watanabe, Y. Matsuki, R. Hiramatsu, S. Akagi, H. Makino, Y. S. Kanwar, Visceral adipose tissue-derived serine protease inhibitor: a unique insulin-sensitizing adipocytokine in obesity., Proc. Natl. Acad. Sci. U. S. A. 102, 10610–10615 (2005). 152. S. Teshigawara, J. Wada, K. Hida, A. Nakatsuka, J. Eguchi, K. Murakami, M. Kanzaki, K. Inoue, T. Terami, A. Katayama, I. Iseda, Y. Matsushita, N. Miyatake, J. F. McDonald, K. Hotta, H. Makino, Serum vaspin concentrations are closely related to insulin resistance, and rs77060950 at SERPINA12 genetically defines distinct group with higher serum levels in Japanese population., J. Clin. Endocrinol. Metab. 97, E1202-7 (2012). 153. M. Blüher, Vaspin in obesity and diabetes: Pathophysiological and clinical significanceEndocrine 41, 176–182 (2012). 154. H. Li, W. Peng, J. Zhuang, Y. Lu, W. Jian, Y. Wei, W. Li, Y. Xu, Vaspin attenuates high glucose-induced vascular smooth muscle cells proliferation and chemokinesis by inhibiting the MAPK, PI3K/Akt, and NF-κB signaling pathways., Atherosclerosis 228, 61–8 (2013). 155. B. Youn, N. Klo, N. Lee, J. W. Park, E. Song, K. Ruschke, A. Oberbach, M. Fasshauer, M. Stumvoll, M. Blu, Serum Vaspin Concentrations in Human Obesity and Type 2 Diabetes, 57, 372–377 (2008). 156. S. Phalitakul, M. Okada, Y. Hara, H. Yamawaki, Vaspin prevents TNF-α- induced intracellular adhesion molecule-1 via inhibiting reactive oxygen species- dependent NF-κB and PKCθ activation in cultured rat vascular smooth muscle cells., Pharmacol. Res. 64, 493–500 (2011). 157. C. H. Jung, M. J. Lee, Y. M. Kang, Y. La Lee, H. K. Yoon, S.-W. Kang, W. J. Lee, J.-Y. Park, Vaspin inhibits cytokine-induced nuclear factor-kappa B activation and adhesion molecule expression via AMP-activated protein kinase activation in vascular endothelial cells., Cardiovasc. Diabetol. 13, 41 (2014). 158. B. Samal, Y. Sun, G. Stearns, S. Suggs, I. A. N. Mcniece, a novel human pre- B-cell colony-enhancing factor . Cloning and Characterization of the cDNA Encoding a Novel Human Pre-B-Cell Colony-Enhancing Factor, 14 (1994), doi:10.1128/MCB.14.2.1431.Updated. 159. A. Rongvaux, R. J. She, M. H. Mulks, D. Gigot, J. Urbain, O. Leo, F. Andris, Pre-B-cell colony-enhancing factor, whose expression is up-regulated in activated lymphocytes, is a nicotinamide phosphoribosyltransferase, a cytosolic enzyme involved in NAF biosynthesisEur. J. Immunol. 32, 3225–3234 (2002). 160. A. Fukuhara, M. Matsuda, M. Nishizawa, K. Segawa, M. Tanaka, K. Kishimoto, Y. Matsuki, M. Murakami, T. Ichisaka, H. Murakami, E. Watanabe, T. Takagi, M. Akiyoshi, T. Ohtsubo, S. Kihara, S. Yamashita, M. Makishima, T. 184
Funahashi, S. Yamanaka, R. Hiramatsu, Y. Matsuzawa, I. Shimomura, Visfatin: a protein secreted by visceral fat that mimics the effects of insulin., Science 307, 426–30 (2005). 161. D. Friebe, M. Neef, J. Kratzsch, S. Erbs, K. Dittrich, A. Garten, S. Petzold- Quinque, S. Blüher, T. Reinehr, M. Stumvoll, M. Blüher, W. Kiess, A. Körner, Leucocytes are a major source of circulating nicotinamide phosphoribosyltransferase (NAMPT)/pre-B cell colony (PBEF)/visfatin linking obesity and inflammation in humans, Diabetologia 54, 1200–1211 (2011). 162. T. B. Dahl, S. Holm, P. Aukrust, B. Halvorsen, Visfatin/NAMPT: a multifaceted molecule with diverse roles in physiology and pathophysiology., Annu. Rev. Nutr. 32, 229–43 (2012). 163. A. Garten, S. Schuster, M. Penke, T. Gorski, T. de Giorgis, W. Kiess, Physiological and pathophysiological roles of NAMPT and NAD metabolism., Nat. Rev. Endocrinol. 11, 1–12 (2015). 164. B. Zhao, M. Zhang, X. Han, X. Y. Zhang, Q. Xing, X. Dong, Q. J. Shi, P. Huang, Y. B. Lu, E. Q. Wei, Q. Xia, W. P. Zhang, C. Tang, Cerebral ischemia is exacerbated by extracellular nicotinamide phosphoribosyltransferase via a non- enzymatic mechanism, PLoS One 8, 1–14 (2013). 165. Z. Jing, J. Xing, X. Chen, R. A. Stetler, Z. Weng, Y. Gan, F. Zhang, Y. Gao, J. Chen, R. K. Leak, G. Cao, Neuronal NAMPT is Released after Cerebral Ischemia and Protects against White Matter Injury, J. Cereb. Blood Flow Metab. 34, 1613–1621 (2014). 166. J. R. Revollo, A. Körner, K. F. Mills, A. Satoh, T. Wang, A. Garten, B. Dasgupta, Y. Sasaki, C. Wolberger, R. R. Townsend, J. Milbrandt, W. Kiess, S. ichiro Imai, Nampt/PBEF/Visfatin Regulates Insulin Secretion in β Cells as a Systemic NAD Biosynthetic Enzyme, Cell Metab. 6, 363–375 (2007). 167. S. Y. Lim, S. M. Davidson, A. J. Paramanathan, C. C. T. Smith, D. M. Yellon, D. J. Hausenloy, The novel adipocytokine visfatin exerts direct cardioprotective effects., J. Cell. Mol. Med. 12, 1395–1403 (2008). 168. H. K. Song, M. H. Lee, B. K. Kim, Y. G. Park, G. J. Ko, Y. S. Kang, J. Y. Han, S. Y. Han, K. H. Han, H. K. Kim, D. R. Cha, Visfatin: a new player in mesangial cell physiology and diabetic nephropathy, AJP Ren. Physiol. 295, F1485–F1494 (2008). 169. F. Brentano, O. Schorr, C. Ospelt, J. Stanczyk, R. E. Gay, S. Gay, D. Kyburz, Pre-B cell colony-enhancing factor/visfatin, a new marker of inflammation in rheumatoid arthritis with proinflammatory and matrix-degrading activities, Arthritis Rheum. 56, 2829–2839 (2007). 170. F. Montecucco, I. Bauer, V. Braunersreuther, S. Bruzzone, A. Akhmedov, T. F. Luscher, T. Speer, A. Poggi, E. Mannino, G. Pelli, K. Galan, M. Bertolotto, S. 185
Lenglet, A. Garuti, C. Montessuit, R. Lerch, C. Pellieux, N. Vuilleumier, F. Dallegri, J. Mage, C. Sebastian, R. Mostoslavsky, A. Gayet-Ageron, F. Patrone, F. Mach, A. Nencioni, Inhibition of nicotinamide phosphoribosyltransferase reduces neutrophil-mediated injury in myocardial infarction., Antioxid. Redox Signal. 18, 630–641 (2013). 171. T. B. Dahl, A. Yndestad, M. Skjelland, E. Øie, A. Dahl, A. Michelsen, J. K. Damås, S. H. Tunheim, T. Ueland, C. Smith, B. Bendz, S. Tonstad, L. Gullestad, S. S. Frøland, K. Krohg-Sørensen, D. Russell, P. Aukrust, B. Halvorsen, Increased expression of visfatin in macrophages of human unstable carotid and coronary atherosclerosis: possible role in inflammation and plaque destabilization., Circulation 115, 972–980 (2007). 172. T. Romacho, C. F. Sánchez-Ferrer, C. Peiró, Visfatin/Nampt: An adipokine with cardiovascular impactMediators Inflamm. 2013, 1–15 (2013). 173. M. Jurdana, A. Petelin, M. černelič Bizjak, M. Bizjak, G. Biolo, Z. Jenko- PraŽnikar, Increased serum visfatin levels in obesity and its association with anthropometric/biochemical parameters, physical inactivity and nutrition, ESPEN. J. 8 (2013), doi:10.1016/j.clnme.2013.02.001. 174. D. Taşkesen, B. Kirel, T. Us, Serum visfatin levels, adiposity and glucose metabolism in obese adolescents, JCRPE J. Clin. Res. Pediatr. Endocrinol. 4, 76– 81 (2012). 175. G. Sun, J. Bishop, S. Khalili, S. Vasdev, V. Gill, D. Pace, D. Fitzpatrick, E. Randell, Y. G. Xie, H. Zhang, Serum visfatin concentrations are positively correlated with serum triacylglycerols and down-regulated by overfeeding in healthy young men, Am. J. Clin. Nutr. 85, 399–404 (2007). 176. a. R. Moschen, a. Kaser, B. Enrich, B. Mosheimer, M. Theurl, H. Niederegger, H. Tilg, Visfatin, an Adipocytokine with Proinflammatory and Immunomodulating Properties, J. Immunol. 178, 1748–1758 (2007). 177. W.-J. Lee, C.-S. Wu, H. Lin, I.-T. Lee, C.-M. Wu, J.-J. Tseng, M.-M. Chou, W. H.-H. Sheu, Visfatin-induced expression of inflammatory mediators in human endothelial cells through the NF-kappaB pathway., Int. J. Obes. (Lond). 33, 465– 72 (2009). 178. R. Adya, B. K. Tan, J. Chen, H. S. Randeva, Nuclear Factor-κB Induction by Visfatin in Human Vascular Endothelial Cells, Diabetes Care 31, 758–760 (2008). 179. A. R. Moschen, A. Kaser, B. Enrich, B. Mosheimer, M. Theurl, H. Niederegger, H. Tilg, Visfatin, an Adipocytokine with Proinflammatory and Immunomodulating Properties, J. Immunol. 178, 1748–1758 (2007). 180. J. Y. Kim, Y. H. Bae, M. K. Bae, S. R. Kim, H. J. Park, H. J. Wee, S. K. Bae, Visfatin through STAT3 activation enhances IL-6 expression that promotes endothelial angiogenesis, Biochim. Biophys. Acta - Mol. Cell Res. 1793, 1759– 186
1767 (2009). 181. P. Cirillo, V. Di Palma, F. Maresca, F. Pacifico, F. Ziviello, M. Bevilacqua, B. Trimarco, A. Leonardi, M. Chiariello, The adipokine visfatin induces tissue factor expression in human coronary artery endothelial cells: Another piece in the adipokines puzzle, Thromb. Res. 130, 403–408 (2012). 182. D. Raman, P. J. Baugher, Y. M. Thu, A. Richmond, Role of chemokines in tumor growthCancer Lett. 256, 137–165 (2007). 183. P. Romagnani, L. Lasagni, F. Annunziato, M. Serio, S. Romagnani, CXC chemokines: The regulatory link between inflammation and angiogenesisTrends Immunol. 25, 201–209 (2004). 184. H. Sell, C. Habich, J. Eckel, Adaptive immunity in obesity and insulin resistance., Nat. Rev. Endocrinol. 8, 709–16 (2012). 185. R. C. M. van Kruijsdijk, E. van der Wall, F. L. J. Visseren, Obesity and cancer: the role of dysfunctional adipose tissue., Cancer Epidemiol. Biomarkers Prev. 18, 2569–78 (2009). 186. J. M. Rutkowski, K. E. Davis, P. E. Scherer, Mechanisms of obesity and related pathologies: The macro- and microcirculation of adipose tissueFEBS J. 276, 5738–5746 (2009). 187. E. A. Jaffe, R. L. Nachman, C. G. Becker, C. R. Minick, Culture of human endothelial cells derived from umbilical veins. Identification by morphologic and immunologic criteria, J. Clin. Invest. 52, 2745–2756 (1973). 188. D. Onat, D. Brillon, P. C. Colombo, A. M. Schmidt, Human vascular endothelial cells: A model system for studying vascular inflammation in diabetes and atherosclerosisCurr. Diab. Rep. 11, 193–202 (2011). 189. Z. Qin, The use of THP-1 cells as a model for mimicking the function and regulation of monocytes and macrophages in the vasculatureAtherosclerosis 221, 2–11 (2012). 190. J. Auwerx, The human leukemia cell line, THP-1: A multifacetted model for the study of monocyte-macrophage differentiation, Experientia 47, 22–31 (1991). 191. A. L. Cheung, Isolation and culture of human umbilical vein endothelial cells (HUVEC)., Curr. Protoc. Microbiol. Appendix 4, Appendix 4B (2007). 192. S. Tsuchiya, M. Yamabe, Y. Yamaguchi, Y. Kobayashi, T. Konno, K. Tada, Establishment and characterization of a human acute monocytic leukemia cell line (THP-1)., Int. J. Cancer 26, 171–6 (1980). 193. M. C. Longo, M. S. Berninger, J. L. Hartley, Use of uracil DNA glycosylase to control carry-over contamination in polymerase chain reactions, Gene 93, 125– 128 (1990). 187
194. E. F. Grabowski, C. A. Carter, J. R. Ingelfinger, O. Tsukurov, N. Conroy, W. M. Abbott, R. W. Orkin, Comparison of tissue factor pathway in human umbilical vein and adult saphenous vein endothelial cells: implications for newborn hemostasis and for laboratory models of endothelial cell function, Pediatr Res 46, 742–747 (1999). 195. S. Gerhardt, V. König, M. Doll, T. Hailemariam-Jahn, I. Hrgovic, N. Zöller, R. Kaufmann, S. Kippenberger, M. Meissner, Dimethylfumarate protects against TNF-α-induced secretion of inflammatory cytokines in human endothelial cells, J. Inflamm. 12, 49 (2015). 196. S. H. Mangan, A. Van Campenhout, C. Rush, J. Golledge, Osteoprotegerin upregulates endothelial cell adhesion molecule response to tumor necrosis factor- alpha associated with induction of angiopoietin-2., Cardiovasc. Res. 76, 494–505 (2007). 197. K. J. Livak, T. D. Schmittgen, Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method., Methods 25, 402–408 (2001). 198. J. S. Pober, W. C. Sessa, Evolving functions of endothelial cells in inflammation., Nat. Rev. Immunol. 7, 803–15 (2007). 199. A. Saalbach, J. Tremel, D. Herbert, K. Schwede, E. Wandel, C. Schirmer, U. Anderegg, A. G. Beck-Sickinger, J. T. Heiker, S. Schultz, T. Magin, J. C. Simon, Anti-Inflammatory Action of Keratinocyte-Derived Vaspin, Am. J. Pathol. , 1–13 (2016). 200. M. D. Turner, B. Nedjai, T. Hurst, D. J. Pennington, Cytokines and chemokines: At the crossroads of cell signalling and inflammatory diseaseBiochim. Biophys. Acta - Mol. Cell Res. 1843, 2563–2582 (2014). 201. A. D. Luster, R. Alon, U. H. von Andrian, Immune cell migration in inflammation: present and future therapeutic targets., Nat. Immunol. 6, 1182–90 (2005). 202. B. Devaux, D. Scholz, a Hirche, W. P. Klövekorn, J. Schaper, Upregulation of cell adhesion molecules and the presence of low grade inflammation in human chronic heart failure., Eur. Heart J. 18, 470–9 (1997). 203. B. Rossi, S. Angiari, E. Zenaro, S. L. Budui, G. Constantin, Vascular inflammation in central nervous system diseases: adhesion receptors controlling leukocyte-endothelial interactions., J. Leukoc. Biol. 89, 539–56 (2011). 204. J.-K. Min, Y.-M. Kim, S. W. Kim, M.-C. Kwon, Y.-Y. Kong, I. K. Hwang, M. H. Won, J. Rho, Y.-G. Kwon, TNF-Related Activation-Induced Cytokine Enhances Leukocyte Adhesiveness: Induction of ICAM-1 and VCAM-1 via TNF Receptor- Associated Factor and Protein Kinase C-Dependent NF- B Activation in Endothelial Cells, J. Immunol. 175, 531–540 (2005). 188
205. U. Maus, S. Henning, H. Wenschuh, K. Mayer, W. Seeger, J. Lohmeyer, Role of endothelial MCP-1 in monocyte adhesion to inflamed human endothelium under physiological flow., Am. J. Physiol. Heart Circ. Physiol. 283, H2584–H2591 (2002). 206. C. W. Smith, Endothelial adhesion molecules and their role in inflammation., Can. J. Physiol. Pharmacol. 71, 76–87 (1993). 207. K. Ley, C. Laudanna, M. I. Cybulsky, S. Nourshargh, Getting to the site of inflammation: the leukocyte adhesion cascade updated, Nat. Rev. Immunol. 7, 678–689 (2007). 208. S. E. Kahn, R. L. Hull, K. M. Utzschneider, Mechanisms linking obesity to insulin resistance and type 2 diabetes., Nature 444, 840–846 (2006). 209. S. Smyth, A. Heron, Diabetes and obesity: the twin epidemics, Nat. Med. 12, 75–80 (2006). 210. C. R. Balistreri, C. Caruso, G. Candore, The role of adipose tissue and adipokines in obesity-related inflammatory diseases., Mediators Inflamm. 2010, 802078 (2010). 211. J. E. Sims, D. E. Smith, The IL-1 family: regulators of immunity, Nat. Rev. Immunol. 10, 117 (2010). 212. C. A. Hunter, S. A. Jones, IL-6 as a keystone cytokine in health and disease, Nat. Immunol. 16, 448–457 (2015). 213. I. P. Wicks, A. W. Roberts, Targeting GM-CSF in inflammatory diseases, Nat. Rev. Rheumatol. 12, 37–48 (2015). 214. A. Saalbach, J. Tremel, D. Herbert, K. Schwede, E. Wandel, C. Schirmer, U. Anderegg, A. G. Beck-Sickinger, J. T. Heiker, S. Schultz, T. Magin, J. C. Simon, Anti-inflammatory action of keratinocyte-derived vaspin relevance for the pathogenesis of psoriasis, Am. J. Pathol. 186, 639–651 (2016). 215. E. Esaki, H. Adachi, Y. Hirai, S. ichi Yamagishi, T. Kakuma, M. Enomoto, A. Fukami, E. Kumagai, K. Ohbu, A. Obuchi, A. Yoshimura, S. Nakamura, Y. Nohara, T. Fujiyama, Y. Fukumoto, T. Imaizumi, Serum vaspin levels are positively associated with carotid atherosclerosis in a general population, Atherosclerosis 233, 248–252 (2014). 216. B.-D. Fu, H. Yamawaki, M. Okada, Y. Hara, Vaspin can not inhibit TNF-alpha- induced inflammation of human umbilical vein endothelial cells., J. Vet. Med. Sci. 71, 1201–7 (2009). 217. S. L. Deshmane, S. Kremlev, S. Amini, B. E. Sawaya, Monocyte Chemoattractant Protein-1 (MCP-1): An Overview, J. Interf. Cytokine Res. 29, 313–326 (2009). 218. C. Combadière, S. Potteaux, M. Rodero, T. Simon, A. Pezard, B. Esposito, 189
R. Merval, A. Proudfoot, A. Tedgui, Z. Mallat, Combined inhibition of CCL2, CX3CR1, and CCR5 abrogates Ly6Chi and Ly6Clo monocytosis and almost abolishes atherosclerosis in hypercholesterolemic mice, Circulation 117, 1649– 1657 (2008). 219. L. Gu, Y. Okada, S. K. Clinton, C. Gerard, G. K. Sukhova, P. Libby, B. J. Rollins, Absence of monocyte chemoattractant protein-1 reduces atherosclerosis in low density lipoprotein receptor-deficient mice., Mol. Cell 2, 275–281 (1998). 220. H. Fujimoto, T. Sangai, G. Ishii, A. Ikehara, T. Nagashima, M. Miyazaki, A. Ochiai, Stromal MCP-1 in mammary tumors induces tumor-associated macrophage infiltration and contributes to tumor progression, Int. J. Cancer 125, 1276–1284 (2009). 221. J. O. Alem??n, L. H. Eusebi, L. Ricciardiello, K. Patidar, A. J. Sanyal, P. R. Holt, Mechanisms of obesity-induced gastrointestinal neoplasiaGastroenterology 146, 357–373 (2014). 222. P. Monti, B. E. Leone, F. Marchesi, G. Balzano, A. Zerbi, F. Scaltrini, C. Pasquali, G. Calori, F. Pessi, C. Sperti, V. Di Carlo, P. Allavena, L. Piemonti, The CC chemokine MCP-1/CCL2 in pancreatic cancer progression: regulation of expression and potential mechanisms of antimalignant activity, Cancer Res 63, 7451–7461 (2003). 223. Z. Cai, Q. Chen, J. Chen, Y. Lu, G. Xiao, Z. Wu, Q. Zhou, J. Zhang, Monocyte chemotactic protein 1 promotes lung cancer-induced bone resorptive lesions in vivo., Neoplasia 11, 228–36 (2009). 224. J. A. Hamilton, Colony-stimulating factors in inflammation and autoimmunity, Nat. Rev. Immunol. 8, 533–544 (2008). 225. F. O. Nestle, S. Alijagic, M. Gilliet, Y. Sun, S. Grabbe, R. Dummer, G. Burg, D. Schadendorf, Vaccination of melanoma patients with peptide- or tumor lysate- pulsed dendritic cells. (1998). 226. R. M. Strieter, J. A. Belperio, M. D. Burdick, S. Sharma, S. M. Dubinett, M. P. Keane, in Annals of the New York Academy of Sciences, (2004), vol. 1028, pp. 351–360. 227. E. C. Keeley, B. Mehrad, R. M. Strieter, CXC chemokines in cancer angiogenesis and metastases.Adv. Cancer Res. 106, 91–111 (2010). 228. R. Adya, B. K. Tan, J. Chen, H. S. Randeva, Pre-B cell colony enhancing factor (PBEF)/visfatin induces secretion of MCP-1 in human endothelial cells: Role in visfatin-induced angiogenesis, Atherosclerosis 205, 113–119 (2009). 229. G. Sommer, S. Kralisch, N. Kloting, M. Kamprad, K. Schrock, J. Kratzsch, A. Tonjes, U. Lossner, M. Bluher, M. Stumvoll, M. Fasshauer, Visfatin Is a Positive Regulator of MCP-1 in Human Adipocytes In Vitro and in Mice In Vivo, Obesity 18, 190
1486–1492 (2009). 230. H. Ulbrich, E. E. Eriksson, L. Lindbom, Leukocyte and endothelial cell adhesion molecules as targets for therapeutic interventions in inflammatory diseaseTrends Pharmacol. Sci. 24, 640–647 (2003). 231. K. A. Paschos, D. Canovas, N. C. Bird, The role of cell adhesion molecules in the progression of colorectal cancer and the development of liver metastasisCell. Signal. 21, 665–674 (2009). 232. M. J. Khandekar, P. Cohen, B. M. Spiegelman, Molecular mechanisms of cancer development in obesityNat. Rev. Cancer 11, 886–895 (2011). 233. T. Romacho, L. A. Villalobos, E. Cercas, R. Carraro, C. F. Sánchez-Ferrer, C. Peiró, Visfatin as a Novel Mediator Released by Inflamed Human Endothelial Cells, PLoS One 8 (2013). 234. C. Peiró, T. Romacho, R. Carraro, C. F. Sánchez-Ferrer, Visfatin/PBEF/Nampt: A New Cardiovascular Target?, Front. Pharmacol. 1, 135 (2010). 235. J.-W. Park, W.-H. Kim, S.-H. Shin, J. Y. Kim, M. R. Yun, K. J. Park, H.-Y. Park, Visfatin exerts angiogenic effects on human umbilical vein endothelial cells through the mTOR signaling pathway, Biochim. Biophys. Acta - Mol. Cell Res. 1813, 763–771 (2011). 236. T. Q. Bi, X. M. Che, Nampt/PBEF/visfatin and cancer, Cancer Biol. Ther. 10, 119–125 (2010). 237. X. Hu, C. Beeton, Detection of functional matrix metalloproteinases by zymography., J. Vis. Exp. , 1–4 (2010). 238. I. Arnaoutova, H. K. Kleinman, In vitro angiogenesis: endothelial cell tube formation on gelled basement membrane extract, Nat. Protoc. 5, 628–635 (2010). 239. K. L. DeCicco-Skinner, G. H. Henry, C. Cataisson, T. Tabib, J. C. Gwilliam, N. J. Watson, E. M. Bullwinkle, L. Falkenburg, R. C. O’Neill, A. Morin, J. S. Wiest, Endothelial Cell Tube Formation Assay for the In Vitro Study of Angiogenesis, J. Vis. Exp. 10, 1–8 (2014). 240. Y. Cao, Angiogenesis and vascular functions in modulation of obesity, adipose metabolism, and insulin sensitivity., Cell Metab. 18, 478–89 (2013). 241. H. Huang, A. Bhat, G. Woodnutt, R. Lappe, Targeting the ANGPT–TIE2 pathway in malignancy, Nat. Rev. Cancer 10, 575–585 (2010). 242. R. Mazzieri, F. Pucci, D. Moi, E. Zonari, A. Ranghetti, A. Berti, L. S. Politi, B. Gentner, J. L. Brown, L. Naldini, M. De Palma, Targeting the ANG2/TIE2 Axis Inhibits Tumor Growth and Metastasis by Impairing Angiogenesis and Disabling Rebounds of Proangiogenic Myeloid Cells, Cancer Cell 19, 512–526 (2011). 191
243. Q. Chen, M. Jin, F. Yang, J. Zhu, Q. Xiao, L. Zhang, Matrix metalloproteinases: Inflammatory regulators of cell behaviors in vascular formation and remodelingMediators Inflamm. 2013 (2013), doi:10.1155/2013/928315. 244. K. Kessenbrock, V. Plaks, Z. Werb, Matrix Metalloproteinases: Regulators of the Tumor MicroenvironmentCell 141, 52–67 (2010). 245. I. Arnaoutova, J. George, H. K. Kleinman, G. Benton, The endothelial cell tube formation assay on basement membrane turns 20: State of the science and the artAngiogenesis 12, 267–274 (2009). 246. M. L. Ponce, Tube formation: an in vitro matrigel angiogenesis assay., Methods Mol. Biol. 467, 183–188 (2009). 247. S. W. Tas, C. X. Maracle, E. Balogh, Z. Szekanecz, Targeting of proangiogenic signalling pathways in chronic inflammation, Nat. Rev. Rheumatol. 12, 111–122 (2015). 248. A. E. El-Kenawi, A. B. El-Remessy, Angiogenesis inhibitors in cancer therapy: Mechanistic perspective on classification and treatment rationalesBr. J. Pharmacol. 170, 712–729 (2013). 249. B. Gorantla, S. Asuthkar, J. S. Rao, J. Patel, C. S. Gondi, Suppression of the uPAR-uPA System Retards Angiogenesis, Invasion, and In Vivo Tumor Development in Pancreatic Cancer Cells, Mol. Cancer Res. 9, 377–389 (2011). 250. N. Montuori, P. Ragno, Role of uPA/uPAR in the modulation of angiogenesis, Chem. Immunol. Allergy 99, 105–122 (2014). 251. Y.-J. Koo, T.-J. Kim, K.-J. Min, K.-A. So, U.-S. Jung, J.-H. Hong, CXCL11 mediates TWIST1-induced angiogenesis in epithelial ovarian cancer, Tumor Biol. 39, 101042831770622 (2017). 252. H. Raghu, S. S. Lakka, C. S. Gondi, S. Mohanam, D. H. Dinh, M. Gujrati, J. S. Rao, Suppression of uPA and uPAR attenuates angiogenin mediated angiogenesis in endothelial and glioblastoma cell lines, PLoS One 5 (2010), doi:10.1371/journal.pone.0012458. 253. M. R. Robciuc, R. Kivelä, I. M. Williams, J. F. De Boer, T. H. Van Dijk, H. Elamaa, F. Tigistu-Sahle, D. Molotkov, V. M. Leppänen, R. Käkelä, L. Eklund, D. H. Wasserman, A. K. Groen, K. Alitalo, VEGFB/VEGFR1-Induced Expansion of Adipose Vasculature Counteracts Obesity and Related Metabolic Complications, Cell Metab. 23, 712–724 (2016). 254. R. Adya, B. K. Tan, A. Punn, J. Chen, H. S. Randeva, Visfatin induces human endothelial VEGF and MMP-2/9 production via MAPK and PI3K/Akt signalling pathways: Novel insights into visfatin-induced angiogenesis, Cardiovasc. Res. 78, 356–365 (2008). 255. Y. H. Bae, M. K. Bae, S. R. Kim, J. H. Lee, H. J. Wee, S. K. Bae, Upregulation 192 of fibroblast growth factor-2 by visfatin that promotes endothelial angiogenesis, Biochem. Biophys. Res. Commun. 379, 206–211 (2009). 256. L. Nissinen, V. M. Kähäri, Matrix metalloproteinases in inflammation, Biochim. Biophys. Acta - Gen. Subj. 1840, 2571–2580 (2014). 257. E. I. Deryugina, J. P. Quigley, Tumor angiogenesis: MMP-mediated induction of intravasation- and metastasis-sustaining neovasculatureMatrix Biol. 44–46, 94– 112 (2015). 258. H.-J. Park, S.-R. Kim, S. S. Kim, H.-J. Wee, M.-K. Bae, M. H. Ryu, S.-K. Bae, Visfatin promotes cell and tumor growth by upregulating Notch1 in breast cancer., Oncotarget 5, 5087–99 (2014). 259. K. Nasu, M. Nishida, T. Ueda, A. Yuge, N. Takai, H. Narahara, Application of the nuclear factor-kappaB inhibitor BAY 11-7085 for the treatment of endometriosis: an in vitro study., Am. J. Physiol. Endocrinol. Metab. 293, E16–E23 (2007). 260. C. L. Manthey, S. W. Wang, S. D. Kinney, Z. Yao, SB202190, a selective inhibitor of p38 mitogen-activated protein kinase, is a powerful regulator of LPS- induced mRNAs in monocytes., J. Leukoc. Biol. 64, 409–17 (1998). 261. K. Assi, R. Pillai, A. Gómez-Muñoz, D. Owen, B. Salh, The specific JNK inhibitor SP600125 targets tumour necrosis factor-α production and epithelial cell apoptosis in acute murine colitis, Immunology 118, 112–121 (2006). 262. a Arcaro, M. P. Wymann, Wortmannin is a potent phosphatidylinositol 3- kinase inhibitor: the role of phosphatidylinositol 3,4,5-trisphosphate in neutrophil responses., Biochem. J. 296, 297–301 (1993). 263. J. H. Kim, R. K. Studer, N. V. Vo, G. A. Sowa, J. D. Kang, p38 MAPK inhibition selectively mitigates inflammatory mediators and VEGF production in AF cells co- cultured with activated macrophage-like THP-1 cells, Osteoarthr. Cartil. 17, 1662– 1669 (2009). 264. M. Guha, N. Mackman, The phosphatidylinositol 3-kinase-Akt pathway limits lipopolysaccharide activation of signaling pathways and expression of inflammatory mediators in human monocytic cells, J. Biol. Chem. 277, 32124– 32132 (2002). 265. J. W. Pierce, R. Schoenleber, G. Jesmok, J. Best, S. a Moore, T. Collins, M. E. Gerritsen, Novel inhibitors of cytokine-induced IkappaBalpha phosphorylation and endothelial cell adhesion molecule expression show anti-inflammatory effects in vivo., J. Biol. Chem. 272, 21096–103 (1997). 266. S. Strickson, D. G. Campbell, C. H. Emmerich, A. Knebel, L. Plater, M. S. Ritorto, N. Shpiro, P. Cohen, The anti-inflammatory drug BAY 11-7082 suppresses the MyD88-dependent signalling network by targeting the ubiquitin system, 193
Biochem. J. 451, 427–437 (2013). 267. S. Y. Park, J. H. Lee, Y. K. Kim, C. D. Kim, B. Y. Rhim, W. S. Lee, K. W. Hong, Cilostazol prevents remnant lipoprotein particle-induced monocyte adhesion to endothelial cells by suppression of adhesion molecules and monocyte chemoattractant protein-1 expression via lectin-like receptor for oxidized low- density lipoprotein receptor a, J. Pharmacol. Exp. Ther. 312, 1241–8 (2005). 268. J. C. Lee, J. T. Laydon, P. C. McDonnell, T. F. Gallagher, S. Kumar, D. Green, D. McNulty, M. J. Blumenthal, J. R. Heys, S. W. Landvatter, A protein kinase involved in the regulation of inflammatory cytokine biosynthesis.Nature 372, 739– 46 (1994). 269. S. W. Wang, J. Pawlowski, S. T. Wathen, S. D. Kinney, H. S. Lichenstein, C. L. Manthey, Cytokine mRNA decay is accelerated by an inhibitor of p38-mitogen- activated protein kinase, Inflamm. Res. 48, 533–538 (1999). 270. Y. Takanami-Ohnishi, S. Amano, S. Kimura, S. Asada, A. Utani, M. Maruyama, H. Osada, H. Tsunoda, Y. Irukayama-Tomobe, K. Goto, M. Karin, T. Sudo, Y. Kasuya, Essential role of p38 mitogen-activated protein kinase in contact hypersensitivity, J Biol Chem 277, 37896–37903 (2002). 271. K. Ipaktchi, A. Mattar, A. D. Niederbichler, L. M. Hoesel, S. Vollmannshauser, M. R. Hemmila, G. L. Su, D. G. Remick, S. C. Wang, S. Arbabi, Attenuating burn wound inflammatory signaling reduces systemic inflammation and acute lung injury, J. Immunol. 177 (2006). 272. J. H. Kim, R. K. Studer, N. V. Vo, G. A. Sowa, J. D. Kang, p38 MAPK inhibition selectively mitigates inflammatory mediators and VEGF production in AF cells co- cultured with activated macrophage-like THP-1 cells, Osteoarthr. Cartil. 17, 1662– 1669 (2009). 273. L. Roussel, F. Houle, C. Chan, Y. Yao, J. Berube, R. Olivenstein, J. G. Martin, J. Huot, Q. Hamid, L. Ferri, S. Rousseau, IL-17 Promotes p38 MAPK-Dependent Endothelial Activation Enhancing Neutrophil Recruitment to Sites of Inflammation, J. Immunol. 184, 4531–4537 (2010). 274. S. Dragoni, N. Hudson, B.-A. Kenny, T. Burgoyne, J. A. McKenzie, Y. Gill, R. Blaber, C. E. Futter, P. Adamson, J. Greenwood, P. Turowski, Endothelial MAPKs Direct ICAM-1 Signaling to Divergent Inflammatory Functions, J. Immunol. , 1600823 (2017). 275. B. L. Bennett, D. T. Sasaki, B. W. Murray, E. C. O’Leary, S. T. Sakata, W. Xu, J. C. Leisten, A. Motiwala, S. Pierce, Y. Satoh, S. S. Bhagwat, A. M. Manning, D. W. Anderson, SP600125, an anthrapyrazolone inhibitor of Jun N-terminal kinase, Proc. Natl. Acad. Sci. 98, 13681–13686 (2001). 276. H. Yamawaki, K. Saito, M. Okada, Y. Hara, Methylglyoxal mediates vascular inflammation via JNK and p38 in human endothelial cells, AJP Cell Physiol. 295, 194
C1510–C1517 (2008). 277. Y. Pan, Y. Wang, Y. Zhao, K. Peng, W. Li, Y. Wang, J. Zhang, S. Zhou, Q. Liu, X. Li, L. Cai, G. Liang, Inhibition of JNK phosphorylation by a novel curcumin analog prevents high glucose-induced inflammation and apoptosis in cardiomyocytes and the development of diabetic cardiomyopathy, Diabetes 63, 3497–3511 (2014). 278. Z. Han, D. L. Boyle, L. Chang, B. Bennett, M. Karin, L. Yang, A. M. Manning, G. S. Firestein, c-Jun N-terminal kinase is required for metalloproteinase expression and joint destruction in inflammatory arthritis, J. Clin. Invest. 108, 73– 81 (2001). 279. J. Hirosumi, G. Tuncman, L. Chang, C. Z. Görgün, K. T. Uysal, K. Maeda, M. Karin, G. S. Hotamisligil, A central role for JNK in obesity and insulin resistance., Nature 420, 333–336 (2002). 280. H. Huang, G. Jing, J. J. Wang, N. Sheibani, S. X. Zhang, ATF4 is a novel regulator of MCP-1 in microvascular endothelial cells, J. Inflamm. 12, 31 (2015). 281. C.-H. Zhou, J. Pan, H. Huang, Y. Zhu, M. Zhang, L. Liu, Y. Wu, Salusin-β, but Not Salusin-α, Promotes Human Umbilical Vein Endothelial Cell Inflammation via the p38 MAPK/JNK-NF-κB Pathway, PLoS One 9, e107555 (2014). 282. M. Du, A. Martin, F. Hays, J. Johnson, R. A. Farjo, K. M. Farjo, Serum retinol- binding protein-induced endothelial inflammation is mediated through the activation of toll-like receptor 4, 6, 185–197 (2017). 283. M. F. Favata, K. Y. Horiuchi, E. J. Manos, a J. Daulerio, D. a Stradley, W. S. Feeser, D. E. Van Dyk, W. J. Pitts, R. a Earl, F. Hobbs, R. a Copeland, R. L. Magolda, P. a Scherle, J. M. Trzaskos, Identification of a Novel Inhibitor of Mitogen-activated Protein Kinase Kinase, J. Biol. Chem. 273, 18623–18632 (1998). 284. W. Duan, J. H. P. Chan, C. H. Wong, B. P. Leung, W. S. F. Wong, Anti- inflammatory effects of mitogen-activated protein kinase kinase inhibitor U0126 in an asthma mouse model., J. Immunol. 172, 7053–7059 (2004). 285. S. K. Jo, W. Y. Cho, S. A. Sung, H. K. Kim, N. H. Won, MEK inhibitor, U0126, attenuates cisplatin-induced renal injury by decreasing inflammation and apoptosis, Kidney Int. 67, 458–466 (2005). 286. G. Mohammad, M. Mairaj Siddiquei, M. Imtiaz Nawaz, A. M. Abu El-Asrar, The ERK1/2 inhibitor U0126 attenuates diabetes-induced upregulation of MMP-9 and biomarkers of inflammation in the retina, J. Diabetes Res. 2013 (2013), doi:10.1155/2013/658548. 287. D. Prusty, Activation of MEK/ERK Signaling Promotes Adipogenesis by Enhancing Peroxisome Proliferator-activated Receptor gamma (PPARgamma ) 195 and C/EBPalpha Gene Expression during the Differentiation of 3T3-L1 Preadipocytes, J. Biol. Chem. 277, 46226–46232 (2002). 288. F. Syeda, J. Grosjean, R. A. Houliston, R. J. Keogh, T. D. Carter, E. Paleolog, C. P. D. Wheeler-Jones, Cyclooxygenase-2 induction and prostacyclin release by protease-activated receptors in endothelial cells require cooperation between mitogen-activated protein kinase and NF-κB pathways, J. Biol. Chem. 281, 11792– 11804 (2006). 289. M. Kawaguchi, L. F. Onuchic, S. K. Huang, Activation of extracellular signal- regulated kinase (ERK)1/2, but not p38 and c-Jun N-terminal kinase, is involved in signaling of a novel cytokine, ML-1, J. Biol. Chem. 277, 15229–15232 (2002). 290. A. Maddahi, L. Edvinsson, Cerebral ischemia induces microvascular pro- inflammatory cytokine expression via the MEK/ERK pathway., J. Neuroinflammation 7, 14 (2010). 291. H. Yano, S. Nakanishi, K. Kimura, N. Hanai, Y. Saitoh, Y. Fukui, Y. Nonomura, Y. Matsuda, Inhibition of histamine secretion by wortmannin through the blockade of phosphatidylinositol 3-kinase in RBL-2H3 cells, J. Biol. Chem. 268, 25846– 25856 (1993). 292. T. Okada, L. Sakuma, Y. Fukui, O. Hazeki, M. Ui, Blockage of chemotactic peptide-induced stimulation of neutrophils by wortmannin as a result of selective inhibition of phosphatidylinositol 3-kinase, J. Biol. Chem. 269, 3563–3567 (1994). 293. B. Chaurasia, J. Mauer, L. Koch, J. Goldau, A.-S. Kock, J. C. Brüning, Phosphoinositide-dependent kinase 1 provides negative feedback inhibition to Toll-like receptor-mediated NF-kappaB activation in macrophages., Mol. Cell. Biol. 30, 4354–66 (2010). 294. A. Abliz, W. Deng, R. Sun, W. Guo, L. Zhao, W. Wang, Attenuates Thyroid Injury Associated With Severe Acute Pancreatitis in Rats, 8, 13821–13833 (2015). 295. G. Schabbauer, M. Tencati, B. Pedersen, R. Pawlinski, N. Mackman, PI3K- Akt pathway suppresses coagulation and inflammation in endotoxemic mice, Arterioscler. Thromb. Vasc. Biol. 24, 1963–1969 (2004). 296. M. Hasmann, I. Schemainda, FK866, a Highly Specific Noncompetitive Inhibitor of Nicotinamide Phosphoribosyltransferase, Represents a Novel Mechanism for Induction of Tumor Cell Apoptosis, Cancer Res. 63, 7436–7442 (2003). 297. A. Matsuda, W.-L. Yang, A. Jacob, M. Aziz, S. Matsuo, T. Matsutani, E. Uchida, P. Wang, FK866, a Visfatin Inhibitor, Protects Against Acute Lung Injury After Intestinal Ischemia–Reperfusion in Mice via NF-κB Pathway, Ann. Surg. 259, 1007–1017 (2014). 298. I. Gehrke, E. D. J. Bouchard, S. Beiggi, A. G. Poeppl, J. B. Johnston, S. B. 196
Gibson, V. Banerji, On-target effect of FK866, a nicotinamide phosphoribosyl transferase inhibitor, by apoptosis-mediated death in chronic lymphocytic leukemia cells, Clin. Cancer Res. 20, 4861–4872 (2014). 299. M. C. Lawrence, A. Jivan, C. Shao, L. Duan, D. Goad, E. Zaganjor, J. Osborne, K. McGlynn, S. Stippec, S. Earnest, W. Chen, M. H. Cobb, The roles of MAPKs in disease, Cell Res. 18, 436–442 (2008). 300. A. J. Muslin, MAPK signalling in cardiovascular health and disease: molecular mechanisms and therapeutic targets, Clin. Sci. 115, 203–218 (2008). 301. E. K. Kim, E.-J. Choi, Pathological roles of MAPK signaling pathways in human diseases, Biochim. Biophys. Acta - Mol. Basis Dis. 1802, 396–405 (2010). 302. F. Bost, M. Aouadi, L. Caron, B. Binétruy, in Biochimie, (2005), vol. 87, pp. 51–56. 303. A. S. Dhillon, S. Hagan, O. Rath, W. Kolch, MAP kinase signalling pathways in cancer, Oncogene 26, 3279–3290 (2007). 304. T. Thalhamer, M. A. McGrath, M. M. Harnett, MAPKs and their relevance to arthritis and inflammationRheumatology 47, 409–414 (2008). 305. S. M. Louie, L. S. Roberts, D. K. Nomura, Mechanisms linking obesity and cancerBiochim. Biophys. Acta - Mol. Cell Biol. Lipids 1831, 1499–1508 (2013). 306. T. J. Guzik, D. Mangalat, R. Korbut, Adipocytokines - Novel link between inflammation and vascular function?, J. Physiol. Pharmacol. 57, 505–528 (2006). 307. L. Xiao, Y. Liu, N. Wang, New paradigms in inflammatory signaling in vascular endothelial cells., Am. J. Physiol. Heart Circ. Physiol. 306, H317-25 (2014). 308. X. Xing, J. Yang, X. Yang, Y. Wei, L. Zhu, D. Gao, M. Li, IL-17A induces endothelial inflammation in systemic sclerosis via the ERK signaling pathway, PLoS One 8, 1–10 (2013). 309. W. Chen, J. G. N. Garcia, J. R. Jacobson, Integrin ??4 attenuates SHP-2 and MAPK signaling and reduces human lung endothelial inflammatory responses, J. Cell. Biochem. 110, 718–724 (2010). 310. P. Dent, D. T. Curiel, P. B. Fisher, S. Grant, Synergistic combinations of signaling pathway inhibitors: Mechanisms for improved cancer therapy, Drug Resist. Updat. 12, 65–73 (2009). 311. J. S. Logue, D. K. Morrison, Complexity in the signaling network: Insights from the use of targeted inhibitors in cancer therapy, Genes Dev. 26, 641–650 (2012). 312. F. Y. Liew, D. Xu, E. K. Brint, L. A. J. O’Neill, Negative regulation of Toll-like receptor-mediated immune responses, Nat. Rev. Immunol. 5, 446–458 (2005). 313. T. Lang, A. Mansell, The negative regulation of Toll-like receptor and 197 associated pathways, Immunol. Cell Biol. 85, 425–434 (2007). 314. T. Fukao, S. Koyasu, PI3K and negative regulation of TLR signalingTrends Immunol. 24, 358–363 (2003). 315. A. Denk, Activation of NF-kappa B via the Ikappa B Kinase Complex Is Both Essential and Sufficient for Proinflammatory Gene Expression in Primary Endothelial Cells, J. Biol. Chem. 276, 28451–28458 (2001). 316. N. Marui, M. K. Offermann, R. Swerlick, C. Kunsch, C. A. Rosen, M. Ahmad, R. W. Alexander, R. M. Medford, Vascular cell adhesion molecule-1 (VCAM-1) gene transcription and expression are regulated through an antioxidanct-sensitive mechanism in human vascular endothelial cells., J.Clin.Invest. 92, 1866–1874 (1993). 317. R. Kacimi, J. S. Karliner, F. Koudssi, C. S. Long, R. Kacimi, J. S. Karliner, F. Koudssi, C. S. Long, Expression and Regulation of Adhesion Molecules in Cardiac Cells by Cytokines Response to Acute Hypoxia, , 576–586 (1998). 318. T. Collins, M. A. Read, A. S. Neish, M. Z. Whitley, D. Thanos, T. Maniatis, Transcriptional regulation of endothelial cell adhesion molecules: NF-κB and cytokine-inducible enhancers, Faseb J 9, 899–909 (1995). 319. Q. Cheng, S. J. McKeown, L. Santos, F. S. Santiago, L. M. Khachigian, E. F. Morand, M. J. Hickey, Macrophage Migration Inhibitory Factor Increases Leukocyte-Endothelial Interactions in Human Endothelial Cells via Promotion of Expression of Adhesion Molecules, J. Immunol. 185, 1238–1247 (2010). 320. M. Bawadekar, M. De Andrea, I. Lo Cigno, G. Baldanzi, V. Caneparo, A. Graziani, S. Landolfo, M. Gariglio, The Extracellular IFI16 Protein Propagates Inflammation in Endothelial Cells Via p38 MAPK and NF-κB p65 Activation, J. Interf. Cytokine Res. 35, 441–453 (2015). 321. J. Kaur, R. C. Woodman, P. Kubes, P38 MAPK: critical molecule in thrombin- induced NF-kappa B-dependent leukocyte recruitment., Am. J. Physiol. Heart Circ. Physiol. 284, H1095-103 (2003). 322. E. Volkova, J. A. Willis, J. E. Wells, B. A. Robinson, G. U. Dachs, M. J. Currie, Association of angiopoietin-2, C-reactive protein and markers of obesity and insulin resistance with survival outcome in colorectal cancer, Br. J. Cancer 104, 51–59 (2011). 323. A. Matsuda, W.-L. Yang, A. Jacob, M. Aziz, S. Matsuo, T. Matsutani, E. Uchida, P. Wang, FK866, a visfatin inhibitor, protects against acute lung injury after intestinal ischemia-reperfusion in mice via NF-κB pathway., Ann. Surg. 259, 1007– 17 (2014). 324. E. Esposito, D. Impellizzeri, E. Mazzon, G. Fakhfouri, R. Rahimian, C. Travelli, G. C. Tron, A. A. Genazzani, S. Cuzzocrea, The NAMPT inhibitor FK866 198 reverts the damage in spinal cord injury, J. Neuroinflammation 9, 554 (2012). 325. S. Schuster, M. Penke, T. Gorski, R. Gebhardt, T. S. Weiss, W. Kiess, A. Garten, FK866-induced NAMPT inhibition activates AMPK and downregulates mTOR signaling in hepatocarcinoma cells, Biochem. Biophys. Res. Commun. 458, 334–340 (2015). 326. Z. Moore, G. Chakrabarti, X. Luo, A. Ali, Z. Hu, F. J. Fattah, R. Vemireddy, R. J. DeBerardinis, R. A. Brekken, D. A. Boothman, NAMPT inhibition sensitizes pancreatic adenocarcinoma cells to tumor-selective, PAR-independent metabolic catastrophe and cell death induced by β-lapachone, Cell Death Dis. 6, e1599 (2015). 327. M. Pittelli, L. Formentini, G. Faraco, A. Lapucci, E. Rapizzi, F. Cialdai, G. Romano, G. Moneti, F. Moroni, A. Chiarugi, Inhibition of nicotinamide phosphoribosyltransferase: Cellular bioenergetics reveals a mitochondrial insensitive NAD pool, J. Biol. Chem. 285, 34106–34114 (2010). 328. E. Pikarsky, R. M. Porat, I. Stein, R. Abramovitch, S. Amit, S. Kasem, E. Gutkovich-Pyest, S. Urieli-Shoval, E. Galun, Y. Ben-Neriah, NF-kappaB functions as a tumour promoter in inflammation-associated cancer., Nature 431, 461–466 (2004). 329. M. Karin, F. R. Greten, NF-kappaB: linking inflammation and immunity to cancer development and progression., Nat. Rev. Immunol. 5, 749–759 (2005). 330. M. A. Amin, P. J. Mansfield, A. Pakozdi, P. L. Campbell, S. Ahmed, R. J. Martinez, A. E. Koch, Interleukin-18 induces angiogenic factors in rheumatoid arthritis synovial tissue fibroblasts via distinct signaling pathways, Arthritis Rheum. 56, 1787–1797 (2007). 331. F. Scaldaferri, M. Sans, S. Vetrano, C. Correale, V. Arena, N. Pagano, G. Rando, F. Romeo, A. E. Potenza, A. Repici, A. Malesci, S. Danese, The role of MAPK in governing lymphocyte adhesion to and migration across the microvasculature in inflammatory bowel disease, Eur. J. Immunol. 39, 290–300 (2009). 332. G. L. Johnson, K. Nakamura, The c-jun kinase/stress-activated pathway: regulation, function and role in human disease., Biochim. Biophys. Acta 1773, 1341–8 (2007). 333. G. Q. Li, Y. Zhang, D. Liu, Y. Y. Qian, H. Zhang, S. Y. Guo, M. Sunagawa, T. Hisamitsu, Y. Q. Liu, PI3 kinase/Akt/HIF-1alpha pathway is associated with hypoxia-induced epithelial-mesenchymal transition in fibroblast-like synoviocytes of rheumatoid arthritis, Mol Cell Biochem 372, 221–231 (2013). 334. N. Busso, M. Karababa, M. Nobile, A. Rolaz, F. Van Gool, M. Galli, O. Leo, A. So, T. De Smedt, Pharmacological inhibition of nicotinamide phosphoribosyltransferase/visfatin enzymatic activity identifies a new inflammatory 199 pathway linked to NAD, PLoS One 3 (2008), doi:10.1371/journal.pone.0002267. 335. Y. Li, Y. Zhang, B. Dorweiler, D. Cui, T. Wang, C. W. Woo, C. S. Brunkan, C. Wolberger, S. I. Imai, I. Tabas, Extracellular nampt promotes macrophage survival via a nonenzymatic interleukin-6/STAT3 signaling mechanism, J. Biol. Chem. 283, 34833–34843 (2008). 336. F. Balkwill, Cancer and the chemokine network, Nat. Rev. Cancer 4, 540–550 (2004). 337. M. Blüher, Adipokines - removing road blocks to obesity and diabetes therapyMol. Metab. 3, 230–240 (2014). 338. A. Garten, S. Petzold, A. Körner, S. ichiro Imai, W. Kiess, Nampt: linking NAD biology, metabolism and cancerTrends Endocrinol. Metab. 20, 130–138 (2009). 339. J. E. P. Brown, D. J. Onyango, M. Ramanjaneya, A. C. Conner, S. T. Patel, S. J. Dunmore, H. S. Randeva, Visfatin regulates insulin secretion, insulin receptor signalling and mRNA expression of diabetes-related genes in mouse pancreatic beta-cells., J. Mol. Endocrinol. 44, 171–8 (2010). 340. C. Jacques, M. Holzenberger, Z. Mladenovic, C. Salvat, E. Pecchi, F. Berenbaum, M. Gosset, Proinflammatory actions of visfatin/nicotinamide phosphoribosyltransferase (Nampt) involve regulation of insulin signaling pathway and Nampt enzymatic activity, J. Biol. Chem. 287, 15100–15108 (2012). 341. S. N. Wang, S. Te Wang, K. T. Lee, The potential interplay of adipokines with toll-like receptors in the development of hepatocellular carcinoma, Gastroenterol. Res. Pract. 2011 (2011), doi:10.1155/2011/215986. 342. S. M. Camp, E. Ceco, C. L. Evenoski, S. M. Danilov, T. Zhou, E. T. Chiang, L. Moreno-Vinasco, B. Mapes, J. Zhao, G. Gursoy, M. E. Brown, D. M. Adyshev, S. S. Siddiqui, H. Quijada, S. Sammani, E. Letsiou, L. Saadat, M. Yousef, T. Wang, J. Liang, J. G. N. Garcia, Unique Toll-Like Receptor 4 Activation by NAMPT/PBEF Induces NFκB Signaling and Inflammatory Lung Injury, Sci. Rep. 5, 13135 (2015). 343. M. R. Dasu, S. Devaraj, S. Park, I. Jialal, Increased Toll-Like Receptor (TLR) activation and TLR ligands in recently diagnosed type 2 diabetic subjects, Diabetes Care 33, 861–868 (2010). 344. L. Qiao, Q. Gu, Y. Dai, Z. Shen, X. Liu, R. Qi, J. Ma, B. Zou, Z. Li, H. Y. Lan, B. C. Y. Wong, XIAP-associated factor 1 (XAF1) suppresses angiogenesis in mouse endothelial cells, Tumor Biol. 29, 122–129 (2008). 345. J. L. Arbiser, H. Larsson, L. Claesson-Welsh, X. Bai, K. LaMontagne, S. W. Weiss, S. Soker, E. Flynn, L. F. Brown, Overexpression of VEGF 121 in immortalized endothelial cells causes conversion to slowly growing angiosarcoma and high level expression of the VEGF receptors VEGFR-1 and VEGFR-2 in vivo., Am. J. Pathol. 156, 1469–1476 (2000). 200
346. K.-C. Lan, C.-Y. Chiu, C.-W. Kao, K.-H. Huang, C.-C. Wang, K.-T. Huang, K.- S. Tsai, M.-L. Sheu, S. H. Liu, Advanced Glycation End-Products Induce Apoptosis in Pancreatic Islet Endothelial Cells via NF-κB-Activated Cyclooxygenase- 2/Prostaglandin E2 Up-Regulation., PLoS One 10, e0124418 (2015). 347. S. Cherqui, K. M. Kingdon, C. Thorpe, S. M. Kurian, D. R. Salomon, Lentiviral gene delivery of vMIP-II to transplanted endothelial cells and endothelial progenitors is proangiogenic in vivo., Mol. Ther. 15, 1264–72 (2007). 348. K. Ghosh, C. K. Thodeti, A. C. Dudley, A. Mammoto, M. Klagsbrun, D. E. Ingber, Tumor-derived endothelial cells exhibit aberrant Rho-mediated mechanosensing and abnormal angiogenesis in vitro, Proc. Natl. Acad. Sci. 105, 11305–11310 (2008). 349. M. Bode, Y. Wu, X. Pi, P. Lockyer, W. Dechyapirom, A. Portbury, C. Patterson, Regulation of ASB4 expression in the immortalized murine endothelial cell lines MS1 and SVR: a role for TNF-α and oxygen, Cell Biochem Funct. 29, 334–341 (2011). 201
APPENDIX1: SEQUENCES OF THE PRIMERS USED FOR RT-qPCR
Target gene Primer sequence (5' 3') Product size (bp)
Forward – TGAAGCTCGCACTCTCGCCTC h-CCL2 (MCP1) 114 Reverse - AATCGATGACAGCGCCGTAGCC
Forward - CGCCCAAACCGAAGTCATAGCC h-CXCL2 (GROβ) 80 Reverse - TTTCTTAACCATGGGCGATGCGG Forward - TGACTTCCAAGCTGGCCGTG h-CXCL8 (IL8) 81 Reverse - CCTTGGCAAAACTGCACCTTCAC
Forward – GCCGCGCCCCCGGTTTCTAT h-GAPDH 114 Reverse - AGCGATGTGGCTCGGCTGGC
Forward - TGACCTCCAGGAGCCGACCTG h-GMCSF 87 Reverse - GGGCCCTTGAGCTTGGTGAGG
Forward – TCCACAAGCGCCTTCGGTCC h-IL6 89 Reverse - TCTCCTGGGGGTACTGGGGC
Forward – CCCTATGCTACACAGCTGCC h-E-Selectin 106 Reverse - CCACTGAAGCCAGGGTCACAC
Forward – CAGACCTTTGTCCTGCCAGCG h-ICAM1 87 Reverse - GGAACAGACCACGGTCCCCT Forward - GTCAATGTTGCCCCCAGAGATAC h-VCAM1 112 Reverse - TTTCGGAGCAGGAAAGCCCTG
Forward – AAAGCTGTCCCTGATGCCGTG h-PECAM1 85 Reverse - GTGCATCTGGCCTTGCTGTC
Forward - GCAGCCTGATCTTACACGGTGCTG h-ANGPT1 108 Reverse - AAGCATCAAACCACCATCCTCCTGT
Forward - AGCAGCATCAGCCAACCAGGAAATG h-ANGPT2 109 Reverse - ATGCATCAAACCACCAGCCTCCTGT
Forward - TGCTACAGTTGTTCAAGGCTTCCCC h-CXCL11 85 Reverse - TGCCACTTTCACTGCTTTTACCCCA
Forward - GGGCCCAAAACCGAAAAACCAAAGC h-UPAR 85 Reverse - GGCGTCACCCAGGTGGGCAT Forward - ACTTGTTCTGAGGCCGAGGAGCCTG h-Angiogenin 111 Reverse - CTGAGCCAGGGTCGGTGGGGT
Forward - GGCCACGGCCAGCGAGTACA h-VEGFR1 117 Reverse - AGAGGCCCTCCTTGCTTGGTGC 202
Forward – TTTCGCCCGGCTCGAGGTGC h-VEGFR2 99 Reverse - TAGGCAAACCCACAGAGGCGGC
Forward - TGCCCTGAGACCGCCATGTCCA h-MMP2 107 Reverse - CCGGCGCTGGTGCAGCTCTC
203
APPENDIX 2: ANTIBODY MICROARRAY
For the simultaneous, qualitative detection of multiple pro-inflammatory cytokines (Chapter 2, Figure 2.1), a commercial antibody array system was used
(Cat#AAH-ANG-1000-4, RayBiotech, Norcross, GA). The array layouts with the target molecules are depicted as follows:
Figure A2.1 Layouts of the antibody arrays used for the simultaneous detection of cytokines and angiogenic molecules.
204
APPENDIX 3: LUCIFERASE ASSAY FOR TRANSCRIPTION FACTOR ACTIVATION The following schematic is adapted from Dual-Luciferase® Reporter Assay
System by Promega. The system can be efficiently used to detect the activation of a transcription factor based on its specific promoter that has been cloned upstream of the firefly luciferase.
Figure A3.1. Schematic representation of the Dual-Luciferase® Reporter Assay System (Promega) to determine NFκB activation.
! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! !
! ! Thesis and Dissertation Services