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Regulation of the Cd36 Scavanger Receptor by the Antioxidant 7,8-dihydroneopterin

TABLE OF CONTENTS

TABLE OF CONTENTS

LIST OF FIGURES

ABBREVIATIONS

ABSTRACT

1.  INTRODUCTION

1.1 Overview

1.2 Atherosclerosis

1.2.1 Atherosclerosis and cardiovascular disease

1.2.2 The stages of plaque progression

1.2.3 Lipoproteins and oxidised LDL

1.2.4 Macrophages, foam cell formation and cell death

1.3 Plaque initiation theories

1.3.1 Response to injury theory

1.3.2 Response to retention theory

1.3.3 Oxidative modification hypothesis

1.4 Scavenger receptors

1.4.1 Scavenger receptors

1.4.2 CD36

1.4.3 CD36 structure

1.4.4 CD36 in atherosclerosis

1.5 7,8-Dihydroneopterin

1.5.1 GTP, 7,8-Dihydydroneopterin and products

1.5.3 Antioxidant down regulation of CD36

1.5.4 Other modes of CD36 down regulation

1.6 Peroxisome proliferator activated receptor-ɣ

1.6.1 PPARγ structure and function

1.6.2 Modulation of PPARγ activity

1.7. MAP kinases

1.7.1 ERK cascade

1.7.2 JNK cascade

1.7.3 P38 cascade

1.7.4 MAP kinases & atherosclerosis

1.8 Objective of research

2. MATERIALS & METHODS

2.1 Reagents, media and buffers

2.1.1 Reagents

2.1.2 Media

2.1.3 General solutions and buffers

A. Roswell Park Memorial Institute (RPMI)-1640 media (with or without phenol red)

B. Phosphate buffered saline (PBS)

C. 7,8-dihydroneopterin solution

D. D-neopterin solution

E. Triton lysis solution

F. MOPS buffer for SDS-PAGE

G. Transfer buffer for western blotting

H. Tris-buffered saline (TBS)

I. TBSM

J. Cracker buffer for SDS-PAGE

K. Restore stripping buffer

L. Ponceau S stain

M. Propidium iodide solution

N. Phorbol 12-myristate 12-acetate (PMA)

2.2 Methods

2.2.1 Cell culture based experiments

2.2.2 Cell viability assays

2.2.3 Determination of protein concentration

2.2.4 SDS-PAGE

2.2.5 Western blot analysis

2.2.6 Flow cytometry determination of U937 CD36 cell surface expression

2.2.7 Statistical analysis

3. RESULTS

3.1 Characterising the expression of CD36

3.1.1 The U937 cell line produces detectable basal levels of CD36

3.1.2 Triton X-100 as a lysing agent

3.1.3 Effect of PMA of U937 expression

3.2 Characterization of CD36 down regulation via 7,8-NP

3.2.1 7,8-NP reduces the expression of the CD36 scavenger receptor in the U937 cell Line

3.2.2 7,8-NP induced down regulation occurs in both adherent and suspension well plates

3.2.3 7,8-NP down regulation can be observed using both wet and semi-dry protein transfer protocols

3.2.4 Time course of 7,8-NP induced CD36 down regulation

3.2.5 Recovery of CD36 expression after 7,8-NP removal

3.2.6 Cellular viability of the recovery experiment

3.2.7 Effect of neopterin on CD36 expression in the U937 cell line

3.3 Transcriptional control of CD36

3.3.1 Effect of PPARγ inhibitor GW9662 on CD36 expression

3.3.2 Effect of GW9662 on cellular viability

3.3.3 Effect of GW9662 in conjunction with 7,8-NP on CD36 expression

3.3.4 Effect of increasing concentrations of cycloheximide on cellular viability

3.3.5 Effect of cycloheximide on CD36 expression

3.3.6 Cellular viability of the cycloheximide experiment

3.4 Effect of MAP kinase inhibitors on CD36 down regulation

3.4.1 Effect of JNK inhibition via SP600125 on 7,8-NP induced down regulation in the U937 cell line

3.4.2 Effect of JNK inhibition on cellular viability

3.4.3 Effect of MEK inhibition via PD98059 on 7,8-NP induced down regulation in the U937 cell line

3.4.4 Effect of MEK inhibition on cellular viability

3.4.5 Effect of p38 inhibition via SB202190 on 7,8-NP induced down regulation in the U937 cell line

3.4.6 Effect of p38 inhibition on cellular viability

3.4.7 Effect of NF-kB inhibition via BAY 11-7082 on 7,8-NP induced down regulation in the U937 cell line

3.4.8 Effect of NF-kB Inhibition on cellular viability

3.5 CD36 flow cytometry

3.5.1 CD36 flow cytometry protocol design

3.5.2 Down regulation of CD36 detected via flow cytometry

4. DISCUSSION

4.1 Characterising CD36 down regulation

    4.2 CD36 Down Regulation Measured via Flow Cytometry………………………………..92

4.3 Transcriptional Control of CD36

4.4 The Effect of MAP Kinase Inhibitors

4.4.1 JNK Inhibition

4.4.2 ERK1/2 Inhibition

4.4.3 p38 Inhibition

4.4.4 NF-κB Inhibition

4.5 A different mechanism of effect

4.6 Incomplete Down regulation

4.7 Future Research

4.8 Summary

BIBLIOGRAPHY

LIST OF FIGURES

INTRODUCTION

Figure 1.1      The development of an atherosclerotic plaque

Figure 1.2      The formation of a lipid laden foam

Figure 1.3      The structure of CD36

Figure 1.4      The biosynthetic pathway of 7,8-NP and its reduced products

Figure 1.5       Protective mechanisms of 7,8-NP

Figure 1.6       Regulation mechanics of PPARγ

RESULTS

Figure 3.1.1    The U937 cell line produces detectable basal levels of CD36

Figure 3.1.2    0.5% TritonX-100 effectively lysis cells

Figure 3.1.3    PMA has a limited effect on U937 expression

Figure 3.2.1    7,8-NP reduces the expression of the CD36 scavenger receptor in the U937 cell line

Figure 3.2.2    7,8-NP down regulation can be observed using both adherent and suspension plates

Figure 3.2.3    7,8-NP down regulation can be observed using both wet and semi-dry transfer protocols

Figure 3.2.4    7,8-NP down regulates CD36 over 24 hours

Figure 3.2.5    CD36 expression recovers 24 hours after 7,8-NP removal

Figure 3.2.6    CD36 expression recovers 24 hours after 7,8-NP removal

Figure 3.2.7    Neopterin has a limited effect on CD36 expression

Figure 3.2.8    Images of U937 cells after treatment with increasing concentrations of neopterin

Figure 3.3.1    PPARγ inhibitor GW9662 decreases CD36 expression

Figure 3.3.2    Effect of GW9662 on cellular viability using propidium iodide

Figure 3.3.3    GW9662 and 7,8-NP do not have an additive effect on 7,8-NP expression

Figure 3.3.4    Effect of increasing concentrations of cycloheximide on cellular viability

Figure 3.3.5    Effect of cycloheximide on CD36 expression

Figure 3.3.6    Effect of cycloheximide and cycloheximide+7,8-NP on U937 cell viability using trypan blue

Figure 3.4.1    JNK inhibition via SP600125 does not alleviate CD36 down regulation

Figure 3.4.2    Effect of SP600125 on U937 cell viability using propidium iodide

Figure 3.4.3    MEK inhibition via PD98059 does not alleviate CD36 down regulation

Figure 3.4.4    Effect of PD98059 on U937 cell viability using propidium iodide

Figure 3.4.5    p38 inhibition via SB202190 does not alleviate CD36 down regulation

Figure 3.4.6    Effect of SB202190 on U937 cell viability using propidium iodide

Figure 3.4.7    NF-kB inhibition via BAY 11-7082 does not alleviate CD36 down regulation

Figure 3.4.8    Effect of BAY 11-7082 on U937 cell viability using propidium iodide

Figure 3.5.1.1 Flow cytometer traces for CD36 antibody on U937 cells

Figure 3.5.1.2 Use of the flow cytometry CD36 antibody on the U937 cell line………..86

Figure 3.5.1.2 Flow cytometer traces for increasing concentrations of 7,8-NP  on U937    cells………………………………………………………………………..88

Figure 3.5.2.2 Effect of 7,8-NP on U937 cells detected using flow cytometry………….89

ABBREVIATIONS

7,8-NP 7,8-Dihydroneopterin
AAPH 2,2′-Azobis(2-amidinopropane) dihydrochloride
ApoB100 Apoplipoprotein B100
BCA Bicinchoninc acid
BMM Bone marrow-derived macrophage
BSA Bovine serum albumin
CD36 Cluster of differentiation 36
CE Cholesterol Esters
DMSO Dimethyl sulphoxide
DNA Deoxyribonucleic acid
EDTA Ethylenediaminetetraacetic acid
ERK extracellular signal-regulated kinase
FL Fluorescence filter
FRB Free Radical Biochemsitry
FSC Forward Scatter
GAG Glycosaminoglycan
GCH-1 GTP cyclohydrolase-I
GTP Guanosine triphosphate
H2O2 Hydrogen peroxide
HCl Hydrochloric acid
HDL High density lipoprotein
HETE Hydroxyeicosatetraenoic acid
HMDM Human monocyte derived macrophage
HOCL Hypochlorous acid
HODE Hydroxyoctadecadienoic acid
HRP Hydrogen peroxidase
IL Interleukin
INF-γ Interferon-γ
JNk c-Jun N-terminal kianse
LDL Low-density lipoprotein
LOX-1 Lectin-like oxidized low-density lipoprotein receptor-1
LPS Lipopolysaccharide
MAP Mitogen activated protein
MAPKAPK MAPK- activated protein kianse
MAPKK Map kinase kinase
MAPKKK Map kinase kinase kinase
MOPS 4-Morpholine-propanesulfonic acid
MPM Peritoneal macrophage
NaCl Sodium chloride
NaCO3 Sodium carbonate
NADPH Reduced nicotinamide adenine dinucleotide phosphate
NaHCO3 Sodium bicarbonate
NaOH Sodium hydroxide
oxLDL Oxidatievly modified low-density lipoprotein
PBS Phosphate buffered saline
PI Propidium iodide
PKA Protein Kianse A
PKB Protein kinase b
PKC Protein kinase c
PMA Phorbol 12-myristate 12-acetate
PPARγ Peroxisome proliferator-activated receptor-γ
PTPS 6-pyruvonyltetrahydropterin synthase
ROS Reactive oxygen species
RPMI-1640 Roswell Park Memorial Institute 1640
RXR Retinoid X Receptor
SDS Sodium dodecyl sulphate
SR-A Scavanger Receptor A
STAT Janus kinase-signal transducers and activators of transcription
TBS Membrane blocking solution
TBS Tris-buffered saline
TBSM Membrane blocking solution + milk
TGF Trasforming growth factor
TLR Toll like receptor
TNFα Tumor necrosis factor α
UV Ultraviolte
XO Xanthine Oxidase

ABSTRACT

OxLDL uptake via the CD36 scavenger receptor leads to foam cell formation and is at the core of the development of atherosclerosis. One potential protective mechanism against this process involves the human macrophage derived anti-oxidant 7,8-dihydroneopterin (7,8-NP) down regulating CD36. This study characterised CD36 down regulation in the U937 monocyte-like cell line, and examined a mechanism of action involving MAP kinase mediated control of the PPARYγ transcription factor.

Western blot analysis showed that it in U937 cells 7,8-NP concentrations up to 150µM down regulated CD36 to ~40% of basal levels over 24 hours. The effect seen here was stronger than that previously observed with human monocyte derived macrophages (HMDM). The oxidised product of 7,8-NP, neopterin, had no significant effect. CD36 levels were able to recover after down regulation and neared control levels 24 hours after 7,8-NP removal.

CD36 protein levels were found to be under control of PPARγ and it was shown that 7,8-NP likely only has its effect at the transcriptional level, and did not enhance the proteolytic removal of the CD36. PPARγ contains a MAP kinase binding site which when phosphorylated prevents the transcription of CD36. Co-incubation of selective MAP kinase inhibitors SP600125 (JNK) and PD98059 (ERK1/2) with 7,8-NP failed to block CD36 down regulation. The effect of p38 and NF-κB signalling in CD36 down regulation was additionally explored using their inhibitors (SB202190 and BAY 11-7082 respectively), but likewise did not block 7,8-NP’s effect.

The results confirm that in the U937 cell line 7,8-NP can decrease the levels of CD36, and that a regulatory pathway involving PPARγ is likely. It is also shown that 7,8-NPs mechanism of action does not involve the activation of a MAP kinase cascade phosphorylating PPARγ. This points towards a different mechanism of action, possibly involving PPARγ’s lipid ligand binding site.

  1. INTRODUCTION
  1.         Overview

Cardiovascular disease is a leading cause of death accounting for 16.7 million fatalities globally each year (Dahlöf, 2010). A major pathology of cardiovascular disease is atherosclerosis, otherwise known as the hardening of the artery wall (Hansson, 2005).  This process is characterised by the recruitment of immune cells and the deposition of lipids alongside cellular debris within the arterial wall. Over time this leads to the formation of atherosclerotic plaques and results in the occlusion of blood flow (Lusis, 2000). Cardiovascular disease may manifest as angina, stroke or myocardial infarction (Roger et al., 2011).

Oxidative stress and inflammatory cell death are at the core of the development of atherosclerosis. One of the fundamental interactions during this process is the death of macrophage cells after exposure to oxidatively modified low density lipoprotein (oxLDL) (Moore & Tabas, 2011). Low density lipoprotein (LDL) is a key component of the lipoprotein pool that functions in the transportation of lipids around the body. Oxidative modification of this particle results in oxLDL which is pro-inflammatory and cytotoxic (Berliner & Heinecke, 1996). OxLDL will interact with and be taken up by macrophage cells leading to the formation of foam cells (Henriksen, Mahoney, & Steinberg, 1983). These foam cells remain within the artery wall and make up a significant part of the atherosclerotic plaque (Stocker & Keaney, 2004). The inflammatory properties of oxLDL will also contribute to higher levels of oxidative stress within the arterial wall and in doing so recruit more macrophages to this location (Park, Febbraio, & Silverstein, 2009).

The initial step that results in the uptake of oxLDL is mediated by a class of cell surface receptors termed scavenger receptors. Cluster of differentiation 36 (CD36) is the scavenger receptor most associated with oxLDL triggered cytotoxicity and oxLDL uptake as CD36 contains motifs on its extracellular portion that allows for recognition and binding of the oxidised components of oxLDL (Ohidar, 2010). Mechanisms that are able to decrease levels of CD36 are of increasing interest in order to slow the progression of atherosclerosis.

One potential mechanism for CD36 down regulation relates to the human macrophage derived anti-oxidant 7,8-dihydroneopterin (7,8-NP). The protective role of 7,8-NP against macrophage foam cell formation is believed to be two-fold; 1) Its antioxidant activity which scavenges any reactive oxygen species (ROS) that may be generated in response to oxLDL (Steven Gieseg, Crone, Flavall, & Amit, 2008; Steven Gieseg, Reibnegger, Wachter, & Esterbauer, 1995). 2) The down regulation of CD36 resulting in a decrease in the amount of oxLDL that is taken up by macrophages in the first place (S. Gieseg, Z. Amit, Y.-T. Yang, A. Shchepetkina, & H. Katouah, 2010).

This research focuses on the second mode of action and explores how 7,8-NP is able to down regulate CD36 expression. By testing the effects of various inhibitors of intracellular signalling, we explore the mechanism of CD36 down regulation by 7,8-NP, with the aim of increasing our understanding its protective effect in atherosclerosis. Specifically 7,8-NP’s ability to down regulate the CD36 scavenger receptor and thus prevent foam cell formation.

  1.         Atherosclerosis
    1.   Atherosclerosis and cardiovascular disease

Atherosclerosis is the underlying process in cardiovascular disease which includes coronary heart disease, strokes and peripheral vascular disease. Altogether cardiovascular disease accounts for 30% of deaths globally (Santulli, 2013). The progression of atherosclerosis is a chronic and complex process. Its development involves endothelial dysfunction, arterial inflammation, immune cell recruitment and the thickening of the arterial intima through a build-up of cellular debris, lipids, cholesterol and calcium deposition (Bobryshev, 2006). Remodelling of the arterial wall results in large occlusions known as atherosclerotic plaques that restricts blood flow and reduces oxygen supply. As an atherosclerotic plaque progresses the lesion can become unstable and may rupture. Once ruptured the plaque will release its contents into the blood stream triggering thrombus formation, which if lodged in the small capillary beds, can further restrict or completely block blood supply (Falk, 1983). The end result of this blockage may lead to myocardial or cerebral infarction depending on the location of the plaque (Hansson & Hermansson, 2011; Libby, Ridker, & Maseri, 2002). Many risk factors will influence the likelihood of developing atherosclerosis and its severity. These include an individual’s genetic predisposition, along with obesity, age, gender, cigarette and alcohol use, sedentary lifestyle, hypertension and the presence of type II diabetes and hyperlipidaemia (Libby et al., 2002; Meyer & Schmitt, 2000).

1.2.2 The stages of plaque progression

The formation of an atherosclerotic plaque develops over a lifetime and goes through several stages (Figure 1.1). Six stages of plaque progression have been identified as determined by plaque complexity (Ross, 1995). Plaques found in adolescents are often identified as being in stage 1 or 2 of development. Here monocyte infiltration into the intimal layer of the arterial wall, along with the presence of plasma-derived lipid, initiates the thickening of the arterial wall (Ross, 1986; Stary et al., 1995; D Steinberg, 1989). As this thickening expands the addition of further lipids and cells, along with cell death leads to the development of Type III plaques. Necrotic core formation may then occur in which a lipid laden area devoid of cells develops deep within the arterial wall. These plaques are classified as being Type IV and can also be characterized by a thin layer of tissue that separates the necrotic core from the lumen. This layer will develop into a thick fibrous cap in Type V plaques. The final stage of plaque formation involves a more complex structure in which calcium deposits and ulceration develop, along with atrial vascular remodelling (Ross, 1986; Stary et al., 1995; D Steinberg, 1989).

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Figure 1.1 The development of an atherosclerotic plaque

The various stages of atherosclerotic plaque development, depicting the normal artery (1), lipid deposition (2), fatty streak formation (3), necrotic core and fibrous cap (4) and plaque rupture forming a thrombus (5). Adapted from Cross, 2016.

1.2.3 Lipoproteins and oxidised LDL

Several theories have been proposed to explain the initial process that leads to the beginning of plaque formation including the “Response to injury”, “Response to retention” and the “Oxidative modification” hypothesis. A common theme in all these theories is the role that LDL and immune cells play in atherosclerotic progression.

The body’s lipoprotein pool is an important source of lipids, both in normal circulation and throughout atherosclerotic plaques (Lusis, 2000). Lipoproteins are not homogenous but rather are split into many sub-fractions based on size, density and apolipoprotein content. Different lipoproteins have different metabolisms and functional properties (Lewis, 1973). The major lipoprotein fraction linked to atherosclerosis is LDL, the dominant cholesterol ester carrier in the body (Moore & Tabas, 2011). The clearance of LDL from the circulatory system is reliant on the LDL receptor which facilitates its endocytosis into hepatic and macrophage cells. Recognition of the LDL by its receptor is based upon several specific amino acid residues located on the apolipoprotein B100 (ApoB100), the primary lipid recognition protein on LDL. These residues include lysine, arginine and histidine. Oxidative modification of LDL to oxLDL will disrupt this tightly regulated process. Oxidative induced degradation of ApoB100 results in its fragmentation forming smaller peptides and altering the positive lysine residues. This results in the failure of oxLDL to be recognised by the LDL receptor (Esterbauer, Dieber-Rotheneder, Waeg, Striegl, & Juergens, 1990; Levitan, Volkov, & Subbaiah, 2010; Obama et al., 2007). Recognition of oxLDL instead switches to macrophage scavenger receptors. Here rapid and relatively unregulated uptake can occur, resulting in the over accumulation of cholesterol within the cell and subsequent differentiation into lipid-laden foam cells (Figure 1.2) (Bobryshev, 2006; Steinbrecher, 1991). OxLDL will further facilitate extensive cellular damage triggering a cellular apoptotic or necrotic response (Katouah, Chen, Othman, & Gieseg, 2015). Cellular contents (including cellular debris, oxidants, chemotactins and cytokines) can then be released into the surrounding environment increasing inflammation. This leads to the further recruitment of macrophages and production of oxLDL, culminating in a cyclic process. (Libby, Ridker, & Hansson, 2009; Madamanchi, Vendrov, & Runge, 2005).

1.2.4 Macrophages, foam cell formation and cell death

Macrophages are immune cells derived from monocytes and are involved in both innate and adaptive immunity. Macrophage recruitment to a site of infection enables the body to trigger an appropriate response for host defence (Mosser & Edwards, 2008). This may include the removal of pathogens and other large molecules, the clearance of damaged cells and the production of other chemicals used in immune response. These benefits do not always outweigh the consequences in terms of atherosclerosis. Macrophage infiltration has been shown to play an important part in all stages of atherosclerotic plaque development from the initial thickening of the artery wall to the advanced plaque stage (Nakashima, Wight, & Sueishi, 2008). Murine models lacking monocyte-macrophage recruitment mechanisms show reduced lipid deposition into atherosclerotic lesions while histological studies indicate the presence of macrophages within lipid-rich areas of the plaque core (Potteaux et al., 2011).

As aforementioned the role of macrophages in atherosclerotic plaque development relies on their internalization of oxLDL to form lipid laden foam cells, ultimately resulting in macrophage cell death and deposition into a growing atherosclerotic plaque (Daniel Steinberg, 2009). Foam cells are characterised by their accumulation of cholesterol esters (CE) along with oxidation products similar to those seen in oxidatively modified lipoproteins (Jessup & Kritharides, 2000).  Macrophages will accumulate CE and oxysterols when exposed to oxLDL (Asmis & Jelk, 2000; Boström et al., 2006; Brown, Dean, & Jessup, 1996). OxLDL has been shown to facilitate macrophage death in vitro and is suggested to be able to do this also within atherosclerotic plaques (Baird, Reid, Hampton, & Gieseg, 2005; Katouah et al., 2015). While the exact mechanisms of this process is disputed, the binding, uptake and processing of oxLDL plays a key role in the fate of a macrophage (Daniel Steinberg, 2009). When modified lipoproteins are taken up by a macrophage they are first transported to endosomes and then to lysosomes (Brown et al., 2000; Maor & Aviram, 1994). CE content is then hydrolysed into free cholesterol and fatty acids at an acidic pH. As oxLDL uptake by scavenger receptors is relatively un-regulated, its uptake leads to the accumulation of free cholesterol which can be re-esterified via ACAT leading to a large pool of esterified cholesterol within the cytosol. Experimental inhibition of ACAT also causes cholesterol accumulation in oxLDL treated cells, resulting in Endoplasmic Reticulum stress, protease activation, and apoptosis (Ozcan & Tabas, 2016; Wang et al., 1996). Under standard conditions excess cholesterol can be efluxed out of the macrophage through reverse cholesterol transport. This process is characterised by CE hydrolysis mediated by a family of enzymes with carboxyl ester hydrolase/esterase activity. Cholesterol and also cholesterol products are then efluxed out of the macrophage onto high density lipoprotein (HDL) particles (Johnson, Mahlberg, Rothblat, & Phillips, 1991) via ATP-binding cassette transporters ABCA1 and ABCG1 (Yvan-Charvet, Wang, & Tall, 2010). Yet modification of LDL has been shown to inhibit this process resulting in macrophages internalizing higher amounts of oxLDL along with reducing efflux culminating in foam cell formation.

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Figure 1.2 The formation of a lipid laden foam

Native LDL enters the atrial intima space and undergoes oxidation by cellular oxidants produced by endothelial, macrophage and smooth muscle cells. OxLDL then recruits macrophage differentiated monocytes to the site of inflammation. OxLDL uptake via scavenger receptors triggers the formation of lipid loaded foam cells. Foam cell death then results in the macrophage cellular content spilling into the environment, further propagating inflammation within the intima space.

  1.         Plaque initiation theories

1.3.1 Response to injury theory

“Response to injury” is an early theory proposed by Ross (1986) to explain the initial process of plaque formation. In this model disturbance of vascular homeostasis mediated via injury is proposed to initiate atherosclerosis. Suggested triggers include disruption of laminar flow and oscillatory shear stress at the arterial bifurcation, infection of the vascular wall, along with phagocyte associated myeloperoxidase activity (Malek, Alper, & Izumo, 1999). Disturbed endothelium loses its ability to vaso-regulate making the affected area more susceptible to circulating pro-atherogenic lipoproteins (Malek et al., 1999). Endothelial injury will also recruit immune cells which will release pro-inflammatory cytokines leading to the promotion and propagation of the inflammatory response (Wick, Knoflach, & Xu, 2004).  Part of this response results in the recruitment of leukocytes to the site of inflammation with the aim of resolving stress, promoting healing and can lead to the migration and adherence of monocytes to the site of endothelial inflammation. Damage caused to the endothelial layer also allows for the penetration of LDL into the arterial intima layer where oxidative modification may occur due to the inflammatory environment along with the presence of immune cells. These factors can result in foam cell formation and the growth of an atherosclerotic plaque.

1.3.2 Response to retention theory

(Williams & Tabas, 1995)  suggested a second “response-to-retention” theory in which plaque development is triggered by the diffusion and retention of lipoprotein within the sub-endothelial layer of the arterial wall where it binds with high affinity to muscle associated proteoglycan chains. A major feature of proteoglycans are long carbohydrate side chains of glycosaminoglycan’s (GAGs) covalently attached to a core protein via glyosidic linkages. This theory is reliant upon the repeating disaccharide units of the GAGs which contain negatively charged sulphate or carbohydrate groups. As the LDL moves through the sub-endothelial layer the positively charged lysine groups of ApoB100 are attracted to the negatively charged sulphate proteoglycans resulting in the adherence of LDL to the GAG matrix (Evanko, Raines, Ross, Gold, & Wight, 1998; Libby et al., 2009). Retention and aggregation of lipoprotein within the vascular intima allows for its oxidative modification by oxidants formed by vascular cells and macrophages. Modification then promotes its uptake into macrophages via scavenger receptors leading to foam cell formation.

1.3.3 Oxidative modification hypothesis

The oxidative modification hypothesis is also commonly accepted in order to explain the initial stages of plaque development. This theory states that LDL particles enter the arterial intima layer where they become trapped and oxidatively modified by cellular oxidants produced either by endothelial or smooth muscle cells. These cells produce pro-inflammatory cytokines, adhesion molecules and chemotactic proteins to promote the recruitment and migration of monocytes to the inflammatory area within the intima (Hansson, 2001).  Once activated monocytes can roll via cell adhesion proteins along the surface of the lumen endothelial layer triggered by a response to endothelial cell released pro-inflammatory signals. As the monocytes move through the area they undergo differentiation into macrophages in response to inflammatory cytokines. Macrophages in turn will produce oxidants such as superoxide, hydrogen peroxide and hypochlorite via the NADPH oxidase and myeloperoxidase complexes respectively, in order to oxidise any pathogenic threat. This will additionally cause further lipid and protein oxidation of LDL forming oxLDL (Bobryshev, 2006; Libby et al., 2009; Steinbrecher, 1991; Stocker & Keaney, 2004). OxLDL can then be taken up by macrophage scavenger receptors forming lipid laden foam cells which can then be added to a growing plaque (Lusis, 2000).

Inflammatory cytokines released during this process also play an important role in the destabilization of the plaques fibrous cap increasing the likelihood of plaque rupture. Released inflammatory cytokines inrerleukin-1, tumour necrosis factor, CD140 and CD-40 can stimulate an up-regulation of matrix metalloproteinases within macrophages and smooth muscle cells (Libby & Aikawa, 1998; Mach, Schönbeck, Bonnefoy, Pober, & Libby, 1997). It is these enzymes that are in part responsible for the remodelling of the extracellular matrix and the degradation of the plaques fibrous cap increasing the chances of rupture (Galis, Sukhova, Lark, & Libby, 1994; Geng et al., 1995), Once ruptured debris will form an acute thrombosis which may get lodged within the artery blocking or restricting blood supply resulting in nutrient deprivation and tissue death. This may often manifest as either a heart attack or stroke (Bobryshev, 2006; Libby et al., 2002).

  1.         Scavenger receptors

1.4.1 Scavenger receptors

As previously described the oxidative modification of LDL causing foam cell formation and cell death lead to the development of the oxidative theory of atherosclerosis. Part of the basis of this theory is the observation that cultured monocytes/macrophages will internalise oxLDL much more rapidly than unmodified LDL (Ho, Brown, Bilheimer, & Goldstein, 1976). The uptake of oxLDL is mediated via macrophage scavenger receptors. Scavenger receptors were first identified on activated macrophages and subsequently found to function in cholesterol and lipoprotein metabolism, among other roles, by binding and endocytosing a range of modified lipoproteins including oxLDL (Endemann et al., 1993). After binding, scavenger receptor-ligand interaction facilities a signalling cascade regulating macrophage activation, lipid metabolism, and inflammatory pathways (Ashraf & Gupta, 2011). Importantly in terms of atherosclerosis, and unlike the normal LDL receptor, scavenger receptors will not be down regulated by intracellular cholesterol levels allowing cholesteryl ester accumulation to the point of foam cell formation. OxLDL binding has even been shown to result in an upregulation event (D’Archivio et al., 2008). As such scavenger receptors are considered to be a major factor in the development of atherosclerotic plaques.

To date many classes of scavenger receptor have been identified and range from class A to class G. Some function as multi-ligand receptors while others only recognise specific structural motifs (Ashraf & Gupta, 2011). OxLDL is initially recognised by several scavenger receptors including the class A scavenger receptor A (SR-A) (Kodama, Reddy, Kishimoto, & Krieger, 1988), class B scavenger receptors SR-BI and CD36 (Endemann et al., 1993), class D scavenger receptor CD68  (Ramprasad, Terpstra, Kondratenko, Quehenberger, & Steinberg, 1996), and the class E scavenger receptor, lectin-like oxidised low density lipoprotein receptor-1 (LOX-1) (Yoshida, Kondratenko, Green, Steinberg, & Quehenberger, 1998). While in vivo evidence is still lacking for the particular role each plays in atherosclerosis, in vitro experimental evidence points towards an important role for CD36.

1.4.2 CD36

CD36 is an integral membrane bound multi-ligand scavenger receptor found in a number of cells including monocytes/macrophages, platelets, endothelial and smooth muscle cells (Asch, Barnwell, Silverstein, & Nachman, 1987; Collot-Teixeira, Martin, McDermott-Roe, Poston, & McGregor, 2007; Matsumoto et al., 2000; McGregor et al., 1989; Swerlick, Lee, Wick, & Lawley, 1992). CD36 present on the cell surface of macrophages plays a crucial role in the progression of atherosclerotic legions from the fatty streak to the necrotic plaque due to its ability to recognize, bind and endocytose oxLDL.

CD36 was first identified as a 88 kDa membrane glycoprotein on monocytes using the monoclonal antibody OKM5 (Talle et al., 1983). Later on it was discovered that the 88 kDa band was the incompletely glycosylated CD36 located in the Golgi apparatus, whereas the fully glycosylated 100kDa band is mature CD36 residing on the plasma membrane (Alessio et al., 1996). Further studies examining functional and structural characteristics led to CD36 being classified as a member of the class B scavenger receptor family. The gene encoding CD36 has been identified as being more than 46kb in size and located on the q11.2 band of chromosome 7 (Armesilla & Vega, 1994; Fernández-Ruiz, Armesilla, Sánchez-Madrid, & Vega, 1993). The gene consists of 15 exons, of which exons 4-13 and parts of exons 3 and 15 contribute to the protein with exons 1,2 and 15 making up the 3’ and 5’ untranslated regions.

1.4.3 CD36 structure

The structure of CD36 consists of a ditopic configuration in which an extracellular domain is flanked by two transmembrane and two cytoplasmic domains (Figure 1.3). The extra-cellular region consists of many N-linked glycosylation sites along with containing a hydrophobic domain capable of potentially interacting with the plasma membrane, a proline rich region between amino acids 242-333 and several functional domains (Greenwalt et al., 1992).  One domain located between amino acids 155 to 183 and is able to bind to oxLDL, along with growth hormone releasing peptides, hexarelin, and advanced glycated products (Navazo, Daviet, Ninio, & McGregor, 1996; Ohgami et al., 2001; Pearce et al., 1998).

The binding of oxLDL to macrophage CD36 occurs via a lipid moiety (Nicholson, Frieda, Pearce, & Silverstein, 1995) differing to other scavenger receptors in which binding occurs via a apoprotein moiety. Here high affinity binding of oxLDL to CD36 appears to be related to the recognition of oxidized phospholipids covalently bound to apoB in oxLDL (Boullier et al., 2000). The main structural characteristic related here is a phospholipid with a sn-2 acyl group incorporating a terminal γ-hydroxy(or oxo)-α,β-unsaturated carbonyl which is formed during LDL oxidation (Podrez et al., 2002). A computational modelling study undertaken by Doi et al. (1993) showed that the CD36 oxLDL binding domain contains a positively charged groove made by a lysine cluster which enables specific interactions with negatively charged ligands including oxLDL. Bound oxLDL is then able to be endocytosed utilizing a raft mediated pathway seemingly separate from caveolae, caveolin-1 and clathrin mediated uptake (Zeng, Tao, Chung, Heuser, & Lublin, 2003)

Once endocytosed oxLDL brings about important transcriptional changes including the up-regulation of CD36 expression (Shiffman et al., 2000). Upon stimulation via oxLDL Peroxisome proliferator-activated receptor-γ (PPARγ) is activated via the p38 kinase pathway (M. Zhao et al., 2002) forming a heterodimer with the retinoid X receptor (RXR). This PPARγ:RXRα complex can then bind to elements in the CD36 promoter leading to increased CD36 expression (Tontonoz, Nagy, Alvarez, Thomazy, & Evans, 1998). Various kinases have also been shown to be involved in CD36 upregulation. Protein kinase C was shown to be part of oxLDL-mediated CD36 up-regulation and PPARγ activation (Feng et al., 2000) along with protein kinase B (PKB),  in which PKB over expression in macrophages increased the CD36 promoter along with a PPARγ element-driven reporter gene (Munteanu et al., 2006).

STAT1 signalling also plays a role CD36 transcription. The ligand 15-Hydroxyeicosatetraenoic acid (15(S)-HETE) acts via this pathway and with inhibition of STAT1 activity significantly decreasing the expression of CD36 and subsequent foam cell formation (Agrawal et al., 2007). STAT1 activation is mediated by IFN-α, IFN-γ, and several other cytokines (Gao, 2005). IFN-γ binding to its receptor triggers a cascade of events resulting in the phosphorylation of the receptor and receptor-janus kinase proteins. Activated STAT1 can then form homodimers which when translocated to the nucleus will bind consensus DNA γ-activated sites and promote or repress various target genes including CD36 (O’Shea, Gadina, & Schreiber, 2002; Ramana, Gil, Schreiber, & Stark, 2002).

Figure 1.3 The structure of CD36

Structure of CD36 showing the oxLDL binding site located between amino acids 155-183 and the various glycosylation sites required for maturation and transportation to the cell membrane.

1.4.4 CD36 in atherosclerosis

Initial work exploring the role of CD36 in atherosclerosis came from Endemann et al. (1993) in which the HEK293 cell line transfected with CD36 subsequently acquired the ability to specifically bind oxLDL. Later it was demonstrated that CD36, along with class A scavenger receptors, accounted for the uptake of between 75-90% of all acetylated and oxidised LDL (Kunjathoor et al., 2002). Looking at a human population Nozaki et al. demonstrated that a small subset of the Japanese population whose monocytes lack CD36  showed a 40-50% reduction in the levels of oxLDL binding to HMDMs compared to a normal population (Nozaki et al., 1995). The same levels of reduced binding were also seen when CD36 binding sites were blocked by specific monoclonal antibodies reducing oxLDL binding to HMDMS by as much as 50% (Nozaki et al., 1995), with similar results seen in a separate study usings THP-1’s (Endemann et al., 1993).

The in vivo role of CD36 is human atherosclerotic development has yet to be fully explored with most evidence coming from animal models. Mice containing a double knockout of ApoE and CD36 are a common model used. Lack of ApoE triggers very high circulating levels of cholesterol rich lipoproteins and enables the development of atherosclerotic lesions at a much faster rate than wild type populations. After 12 weeks ApoE/CD36 knockout mice exhibit a 76.5% decrease in aortic lesion size compared to mice only lacking AopE, while 60% less oxLDL was internalized in the double knockouts compared to ApoE alone  (Febbraio et al., 2000). Further evidence for role of CD36 in plaque development came when CD36 was reintroduced to knockout mice via a stem-cell transfer, resulting in an increased size of the atherosclerotic lesion area (Febbraio et al., 2000). The sum total of these and similar publications demonstrates the key role that this scavenger receptor plays via oxLDL uptake and foam cell formation in atherosclerotic development.

  1.         7,8-Dihydroneopterin

1.5.1 GTP, 7,8-Dihydydroneopterin and products

Interferon-γ (INF-γ) is a dimerized & soluble cytokine critical in both innate and adaptive immunity. INF-γ has been reported to be involved in the regulation of over 500 genes (Leon & Zuckerman, 2005) and has been associated with both pro- and anti-atherogenic properties. One of INF-γ’s role in atherosclerosis relates to the stimulation of the production of 7,8-diydrneopterin and neopterin (Huber et al., 1984) which have both been detected within atherosclerotic plaques and associated with cardiovascular events (Janmale et al., 2015). Neopterin is considered to be a viable marker of plaque stability and atherosclerotic prognosis (Liu & Li, 2013), while 7,8-NP has been demonstrated in vitro to have a protective role against oxLDL mediated cell death (Steven Gieseg, Leake, et al., 2008). 7,8-NP and neopterin belong to the pteridine class of compounds which as a common characteristic have a pyrazino-2,3-pyrimidine bicyclic nitrogen containing ring system, and depending on the size of substituent groups can further be described and conjugated or un-conjugated (Wachter et al., 1992). Further sub-classification for unconjugated pteridines is determined on the corresponding oxidation state composing of the fully reduced tetrahydropterins, to the partially reduced dihydroneopterins and aromatic pterins (Oettl & Reibnegger, 2002).

All members of the pteridine class of compounds are synthesized from a guanosine triphosphate (GTP) precursor (Figure 1.4). INF-γ stimulates the first step via the enzyme GTP cyclohydrolase 1 which metabolises GTP into 7,8-NP  triphosphate (Werner et al., 1990). Phosphatase enzymes them hydrolyse 7,8-NP triphosphate  to generate 7,8-NP (Wachter et al., 1992). Co-stimulation with tumour necrosis factor-α (TNFα), dexamethasone or LPS has also been demonstrated to enhance synthesis (Werner-Felmayer et al., 1995; Werner-Felmayer et al., 1990) although not as strongly as INF-γ. Breakdown of 7,8-NP after reaction with oxidant species will form neopterin, 7,8-dihydroxanthopterin and xanthopterin.

\fileUsersA$ajy25HomeDesktopchemcial pathway.jpg

Figure 1.4 The biosynthetic pathway of 7,8-NP and its reduced products

The biosynthetic pathway of 7,8-NP involves the conversion of GTP to 7,8-dihydroneopterin triphosphate (7,8-DNT) catalysed by GTP cyclohydrolase-I (GCH-I). Accumulation of 7,8-DNT in cells of a myelocytic origin drives the dephosphorisation of 7,8-DNT to 7,8-NO due to insufficient 6-pyruvonyltetrahydropterin synthase (PTPS). Alternative cells with functional PTPS will drive tetrahydrobiopterin formation. Breakdown of 7,8-NP after reaction with oxidant species will form neopterin, 7,8-dihydroxanthopterin and xanthopterin. Adapted (Dántola, Vignoni, Capparelli, Lorente, & Thomas, 2008; Fuchs et al., 2009)

1.5.2 7,8-NP as an antioxidant

The atherosclerotic plaque is a site of chronic inflammation as indicated by the high number of immune cells such as macrophages, and the presence of various inflammatory markers. Macrophage cells can release a range of oxidizing agents including superoxide, hydrogen peroxide, lipid peroxides, lipoxygenases and possibly hypochlorite, which are all known to contribute to the production of oxLDL (Steven Gieseg, Leake, et al., 2008). While the oxidative theory of atherosclerosis has come under some criticism due to the inability of various antioxidant intervention trials to exhibit protection, these have focused on ascorbate and tocopherol which are tightly controlled in vivo and at high levels can act as pro-oxidants. Reduced pterins, such as 7,8-NP, have also been shown to act as antioxidants and are produced by macrophage cells in the atherosclerotic plaque. 7,8-NP has been demonstrated to have a protective effect with in vitro studies showing its ability to interfere with LDL oxidation (Steven Gieseg et al., 1995), reactive oxygen species (ROS) mediated reactions (Oettl, Greilberger, Dikalov, & Reibnegger, 2004) and free radical damage including from hydroxyl and peroxyl radicals (Duggan, Rait, Platt, & Gieseg, 2002)

7,8-NP is a scavenger of superoxide and peroxyl radicals generated via 2,2’-azobis-2-methyl-propanimidamide dihydrochloride (AAPH) (Oettl et al., 2004) while also reacting with hydrogen peroxide and chloramine-T (Weiss et al., 1993). The reaction rate of 7,8-NP with AAPH derived peroxyl radicals which nears the rate of diffusion makes it a very good inhibitor of this oxidation pathway (Duggan, Rait, Gebicki, & Gieseg, 2001). 7,8-NP has also been shown to inhibit oxidation of the polyunsaturated omega-6 fatty acid linoleate via AAPH and the formation of diene on LDL particles during both AAPH- and copper-mediated oxidation of LDL (Steven Gieseg et al., 1995). Another biologically relevant experimental system showed a 7,8-NP antioxidant effect in protecting bovine serum albumin from AAPH–mediated oxidation (Duggan et al., 2001).

7,8-NP causes a reduction in erythrocyte haemolysis induced by HOCl, H2O2 and AAPH treatments (Steven Gieseg, Maghzal, & Glubb, 2001; Y.-t. T. Yang, Whiteman, & Gieseg, 2012). Complete protection was observed in the presence of AAPH and partial protection (40%) in the presence of H2O2 (Steven Gieseg et al., 2001). 7,8-NP has also been reported to protect against lipid hydroperoxide formation and protein oxidation (Steven Gieseg et al., 2001). Additional studies confirmed this effect of 7,8-NP in which protein hydroxide and lipid peroxide formation on LDL was slowed in HMDM and THP-1 cells (Firth, Crone, Flavall, Roake, & Gieseg, 2008).

Further studies into protective roles of 7,8-NP have shown its ability to protects cells of myelocytic origin from the cytotoxic effects of oxLDL. It has been found that in monocyte like THP-1 and HMDM cells oxLDL formation was fully inhibited with only micro molar concentrations of 7,8-NP (Steven  Gieseg & Cato, 2003), while a separate study showed that 7,8-NP protected U937 cells from the cytotoxicity of oxLDL (Baird et al., 2005). This protective role is thought to occur via the use of 7,8-NP’s antioxidant properties to scavenge any ROS that may be generated in response to oxLDL.

1.5.3 Antioxidant down regulation of CD36

Besides 7,8-NP’s antioxidant properties a second mode of protection is hypothesised to occur via the down regulation of CD36 itself, preventing oxLDL uptake and foam cell formation (Figure 1.5). In HMDM cells the addition of 7,8-NP triggered the down regulation of the 100-kDa form of CD36 and partial reduction of the immature intracellular 88-kDA form, while also reducing the levels of reactive oxygen species (S. Gieseg et al., 2010). 7,8-NP is not the only antioxidant known to down regulate CD36 expression. α-tocopherol (Vitamin E) is well known as a lipophilic antioxidant and has been shown to inhibit oxLDL uptake by the down regulation of CD36 mRNA and protein expression (Ricciarelli, Zingg, & Azzi, 2000). Yet the same effect was not seen when cells are given β-tocopherol or probucol thus ruling out a general anti-oxidant regulated mechanism of down regulation (Ricciarelli et al., 2000). One alternative mechanism is that α-tocopherol inhibited protein kinase C (PKC) activity and it is the phosphorylation/dephosphorylation of the CD36 PPARγ transcription factor that may be involved in the down regulation of CD36. A second proposed mechanism was that α-tocopherol could modulate gene expression indirectly via inhibiting the activated oxidised form of PPARγ ligands. Evidence of α-tocopherol acting by this pathway came from an experiment showing that it reduced the level of PPARγ activators 9-Hydroxyoctadecadienoic acid (9-HODE) and 13-HODE by inhibiting their enzymatic production (Belkner, Stender, & Kühn, 1998).

1.5.4 Other modes of CD36 down regulation

Transforming growth factor-β1 (TGF-β1) and TGF-β2 have been shown to down- regulate CD36. Han et al. showed that TGF-β1 and TGF-β2 decreased expression of CD36 in THP-1 cells by a mechanism involving phosphorylation of mitogen activated protein (MAP) kinase which in turn initiated phosphorylation of PPARγ leading to a decrease in CD36 gene transcription (J. Han et al., 2000).

Alongside the transforming growth factors Boyer et al. showed a mechanism for CD36 down regulation in the form of the cytokine TNF-α which is known to play a major role in the development of atherosclerosis (Boyer et al., 2007). Using human monocytes Boyer et al. showed that TNF-α inhibited both CD36 membrane expression and mRNA expression in a process involving a reduction in PPARγ activation (Boyer et al., 2007). These results corresponded well to studies in which a reduction in PPARγ activation by TNF-α was reported in human adipocytes and hepatocytes (Sung, She, Xiong, & Tsukamoto, 2004). Further evidence for the role that this cytokine plays in CD36 down regulation came from a study by Zamora et al. who showed TNF-α trigged CD36 down regulation in toll like receptor (TLR) 2 (Pam3CSK4 and FSL1) and TLR4 ligand lipopolysaccharide (LPS) stimulated human peripheral blood mononuclear cells (PMBC) (Zamora et al., 2012). Additional anti-inflammatory cytokines have also been suggested to play a role in CD36 down regulation,. Rubic et al. using THP-1 cells and peripheral monocytes showed that anti-inflammatory cytokine interleukin-10 (IL-10) suppressed basal and PPARγ stimulated transcription of CD36 due to reduced PPARγ protein expression (Rubic & Lorenz, 2006). However this is contrasted in the study by Zamora et al. (2012) who although agreed that IL-10 along with IL-6 were produced during TLR induced down regulation, they themselves were not able to explain this effect.

Figure 1.5 Protective mechanisms of 7,8-NP

7,8-NP produced by GTP stimulated macrophages is thought to protect against atherosclerosis by scavenging radicals and in so preventing oxLDL and subsequent foam cell formation, along with down-regulating CD36, the main scavenger receptor of oxLDL.

 

  1.         Peroxisome proliferator activated receptor-ɣ

CD36 transcription is known to be controlled by the heterodimer Peroxisome Proliferator Activated Receptor-γ/Retinoid X Receptor (PPARγ/RXR) (S. Han & Sidell, 2002) with PPARγ ligands 15-deoxyD12 and 14-prostaglandin J2 increasing CD36 expression (Feng et al., 2000). PPARγ is a member of the PPAR subfamily of nuclear hormone receptors which act as ligand-activated transcription factor, regulating gene expression by interacting with specific DNA sequences upstream of its target genes. The PPAR subfamily names relates to the ability for the original member PPARα, to induce peroxisome proliferation in response to a variety of compounds (S. Lee et al., 1995). Other PPAR members do not necessarily share this function, instead acting as major regulators of the numerous aspects of lipid metabolism and metabolic control. Using PPARα as a base PPARγ was first identified using homology cloning in Xenopus (Dreyer et al., 1992) and subsequently in mice (Zhu, Alvares, Huang, Rao, & Reddy, 1993). Afterwards it was discovered to act as the main regulator of adipogenic differentiation (Tontonoz & Spiegelman, 2008).

1.6.1 PPARγ structure and function

Common to all members of the PPAR subfamily, PPARγ forms an obligate heterodimer with RXR in order to carry out its function (Kliewer et al. 1997), with lack of dimerization compromising PPARγ’s ability to bind to DNA with high affinity. The C-terminal region of PPARγ is required for RXR dimerization and also contains AF2, the major transcriptional activation domain, along with forming a ligand-binding pocket (Tontonoz 2008). The 120 amino acids that make up the N-terminal region exhibits transcriptional activity when interacting with a heterologous DNA-binding domain. Contradictory evidence using deletion studies have shown that when the N-terminal domain is removed PPARγ exhibits increased greater transcriptional activity (Tontonoz, Hu, & Spiegelman, 1994) suggesting that this domain may also contribute a level of inhibitory function.

1.6.2 Modulation of PPARγ activity

PPARγ is a phosphoprotein in which it can be post-translationally modified by the attachment of a phosphate group by MAP Kinases (Figure 1.6). In contrast to the likes of insulin this modification decreases transcriptional activity (Adams, Reginato, Shao, Lazar, & Chatterjee, 1997; Hu, Kim, Sarraf, & Spiegelman, 1996). In vitro assays have shown that two MAP Kinases ERK2, and JNK are able to phosphorylate the PPARγ leading to a decrease in transcriptional activity, including that of CD36 (Adams et al., 1997; Camp & Tafuri, 1997). Phosphorylation mediate by both these MAP kinases has been shown to occur at the same site at serine 82 of mouse PPARγ1 (Shao et al., 1998). The substitution by alanine here causes the loss of platelet-derived growth factor mediated repression of PPARγ activity (Camp & Tafuri, 1997). In humans phosphorylation at the corresponding S84 site was shown to inhibit ligand dependent as well and independent trans-activating function.  A separate study also showed that mutation at the MAP kinase binding site lead to a decrease in ligand binding affinity (J. Han et al., 2000) which may suggest communication between phosphorylation and ligand binding sites, similar to that seen in PPARα. In murine models PPARγ 2 is able to be phosphorylated at serine 112 culminating in decreased PPARγ activity (Adams et al., 1997; Hu et al., 1996). In macrophages TGF-β increased PPARγ phosphorylation and subsequently decreased thiazolidinedione (TZD) (a PPAR activator) induced CD36 expression (S. Han & Sidell, 2002).

PPARγ function can also be regulated by ligand binding. Bound ligands induce a conformational change which allows for the release of transcriptional inhibitors and the recruitment of transcriptional co-activators. One class of ligands involves redox-regulation. The oxidised lipid 15-deoxy-Δ12,14-PGJ2 has been shown to act as a ligand (Forman et al., 1995), along with 9-(HODE), 13-(HODE) (Lenz et al., 1990),  and 15(S)-HETE (Nagy, Tontonoz, Alvarez, Chen, & Evans, 1998).

One pathway of ligand activation involves the STAT1 signalling. Activated STAT1 forms a homodimer which when translocated to the nucleus will bind consensus DNA γ-activated sites and activate transcription (O’Shea, Gadina, & Schreiber, 2002; Ramana, Gil, Schreiber, & Stark, 2002). Interaction of STAT1 with PPARγ requires it to become actytelated, but not phosphorylated. Acetylation requires p300 activation which in turn requires Syk2 and Pyk2 stimulation and XO and NADPH oxidise mediated ROS production. Although the exact mechanism of how 15(S)-HETE activates XO/NADPH activity is not yet known

 

 

 

 

 

 

 

Figure 1.6 Regulation mechanics of PPARγ

The regulation mechanics of PPARγ induced down regulation shows a activated MAP Kinase cascade within the macrophage resulting in the phosphorylation of PPARγ’s kinase binding site subsequently decreasing the transcriptional activity for CD36. Oxidised ligand and STAT1 activation of PPARγ is also shown.

1.7. MAP kinases

Mitogen-activated protein kinases (MAP kinase) are a highly conserved family of protein kinases specific to the amino acids serine, threonine, and tyrosine. MAP kinase signalling pathways are involved in a wide variety of cellular functions including proliferation, differentiation, motility, stress response, apoptosis, and survival. Classic MAP kinases include the extracellular signal-regulated kinase 1 and 2 (Erk1/2), the c-Jun N-terminal kinases 1-3 (JNK1-3)/ the p38 isoforms (p38α, β, γ, and δ) and Erk5. These share number of characteristics including similar substrate sites, phosphorylation dependent activation and a “three-tiered” pathway structure in which phosphorylation cascades transmit movement of the signal from one “tier” to the next.

A diverse range of extracellular stimuli trigger MAP Kinase activation. Such stimuli include mitogens, cytokines and growth factors. A stimuli will initially activate one or more MAP kinase kinase kinases (MAPKKKs) either by a receptor dependent or independent mechanism in the initial “tier”. MAPKKKs can then phosphorylate and activate a downstream MAP kinase kinase (MAPKK), the second “tier” which in turn phosphorylates and activates MAP kinase. MAP kinase activation can then trigger the phosphorylation and activation of MAP kinase-activated protein kinases (MAPKAPKs) which function to amplify and mediate the wide range of biological process required.

1.7.1 ERK cascade

ERK MAP kinases (otherwise known as the classical MAP kinase) cascades are involved in a wide variety of cellular functions including in adhesion, progression, migration, survival, differentiation, metabolism, proliferation, and transcription (Roskoski, 2012). ERK1/2 is activated by dual phosphorylation on tyrosine and threonine by the MAPKK MEK1 and MEK2 (Cobb, Boulton, & Robbins, 1991). These in turn are activated by the MAPKKK’s Raf-1, B-af and c-MOS. Termination of ERK1/2 activation is dependent on regulatory mechanisms that remove one ore both phosphates by tyrosine or serine/threonine phosphatases (Todd, Tanner, & Denu, 1999).

1.7.2 JNK cascade

JNK kinase cascades are involved in mediating the cellular apoptotic response to pro-inflammatory genotoxins along with having a role in the regulation of cell proliferation, survival and differentiation (C. Davies & Tournier, 2012). JNK is selectively phosphorylated and activated by JNKK’s MKK4 and SEK, which in turn are activated by the MAPKKK’s MEKK1/2 and Tpl-2.

1.7.3 P38 cascade

The p38 MAP kinase and its signalling cascades are activated by stress and plays a role in immune response, cellular survival and differentiation (Cuadrado & Nebreda, 2010). Four p38 MAP kinase isoforms have been identified with the p38α and p38β isoforms being ubiquitously expressed (Ono & Han, 2000). The p38 pathway is activated by the MAPKK’s MKK3, JNKK and MEK6 and MAPKKK’s Rac-1, Cdc42 and PAK. In most inflammatory cells p38α is the predominately activated isoform with extracellular stimuli coming from a variety of cytokines and a number of pathogens that activate though different toll-like receptors. Several growth factors as well as environmental factors such as heat shock, osmotic regulation, UV light, hypoxic stress and oxygen radicals and also capable of inducing activation.

1.7.4 MAP kinases & atherosclerosis

Reactive oxygen species such as superoxide and hydrogen peroxide have been shown to be produced in vascular smooth muscle cells, endothelial cells and cardiomyocytes (Abe & Berk, 1998; Griendling, Sorescu, & Ushio-Fukai, 2000; Yoshizumi, Tsuchiya, & Tamaki, 2001). There is evidence that MAP kinases play a role in vascular remodelling alongside these reactive oxygen species.  MAP kinases have been shown to be impacted by oxidative stress with hydrogen peroxide significantly stimulating ERK1/2, JNK and p38 is rat aortic smooth muscle cells (SMC) (Yoshizumi, Abe, Haendeler, Huang, & Berk, 2000) Angiotensin II and endotheilan I also activate MAP kinases in VSMC and have been suggested to be a cause of vascular remodelling (Ishida, Ishida, Thomas, & Berk, 1998) (Yoshizumi et al., 1998).

1.8 Objective of research

OxLDL uptake by macrophage CD36 leading to foam cell formation is one process at the core of atherosclerotic plaque development. Mechanisms that are able to decrease levels of CD36 are of increasing interest in order to slow the progression of atherosclerosis. One potential mechanism for CD36 down regulation involves the human macrophage derived anti-oxidant 7,8-NP which has been shown to decrease basal CD36 levels in HMDM. In this study it is hypothesised the 7,8-NP is exerting its effect via interaction with the PPARγ transcription factor. Specifically it is thought that 7,8-NP is activating a MAP Kinase cascade with the end result being the phosphorylation of the PPARγ MAP kinase binding site and a subsequent decrease in CD36 transcription.

The first objective of this research will be to confirm that the 7,8-NP induced down-regulation seen in HMDM occurs in the U937 monocyte like cell line. This will include looking at the effects of time and concentration on CD36 levels, and also whether recovery after down regulation is possible. 7,8-NP induced down-regulation will be explored mainly using western blotting with cell lysate after experimental treatment being immunoblotted for the CD36 antibody. The effectiveness of flow-cytometry will additionally be explored using CD36 antibody binding to the cell surface of whole cells. The next part of this thesis will confirm that CD36 transcription is under the control of PPARγ in the U937 cell line using a selective PPARγ inhibitor in order to confirm our choice of target. It will also be investigated whether 7,8-NP alternatively has its effect via enhanced CD36 degradation using the protein synthesis inhibitor cycloheximide. The research will then explore the effects of MAP Kinases on 7,8-NP induced CD36 down-regulation. This will be achieved using selective MAP Kinase inhibitors including those to ERK1/2, JNK and p38. The effect of NF-κB inhibition will also be explored.

This research will help determined the mechanism via which 7,8-NP induces CD36 down regulation and help to improve our understanding on the role the 7,8-NP plays in the development of atherosclerosis.

  1. MATERIALS & METHODS

2.1 Reagents, media and buffers

All reagents were of analytical grade. All solutions were prepared with deionized water purified with a Milli-Q ultrafiltration system (Millipore, Massachusetts, USA). This water is referred to as Nano-pure water in this thesis.

2.1.1 Reagents

β-Mercaptoethanol      Sigma Chemical Co., Missouri, USA

4-Morpholine-propanesulfonic acid (MOPS)   Sigma Chemical Co., Missouri, USA

7,8-Dihydroneopterin (7,8-NP)     Schircks Laboratory, Switzerland

Acetic acid (glacial)      BDH Lab Supplies, Poole, England

Anchor non-fat milk powder  Fonterra Ltd, New Zealand

BAY 11-7082 (NF-κB Inhibitor)    Sigma-Aldrich Co. LLC, New Zealand

Bicinchoninic acid (BCA) protein determination kit  Pierce Biotechnolgy Inc., Illionois, USA

Bovine serum albumin (BSA)     Sigma-Aldrich Co. LLC, New Zealand

Bromophenol blue      Sigma-Aldrich Co. LLC, New Zealand

cOmplete™, Mini Protease Inhibitor Cocktail   Sigma-Aldrich Co. LLC, New Zealand

Coumassie blue      Bio-Rad Laboratories, California, USA

Dimethyl sufloxide (DMSO)     BDH Lab Supplies Ltd, Poole, England

Ethanol       BDH Lab Supplies Ltd, Poole, England

Ethylene-diamine-tetra-acetic acid (EDTA)  Sigma-Aldrich Co. LLC, New Zealand

Glycerol       Sigma-Aldrich Co. LLC, New Zealand

GW9662 (PPAR-ɣ inhibitor)     Sigma-Aldrich Co. LLC, New Zealand

Hydrochloric acid (HCl)     Merck, Germnay

Methanol       Merck, Fischer Scientific, Belgium

Molecular weight marker     Bio-Rad Laboratories, California, USA

Neopterin      Schircks Laboratory, Switzerland

NuPAGE 4-12% Bis-tris Gels, 1.0mm x10 well   Invitrogen, California, USA

NuPAGE 4-12% Bis-tris Gels, 1.5mm x15 well   Invitrogen, California, USA

PD98059 (MEK Inhibitor)     Sigma-Aldrich Co. LLC, New Zealand

Phorbol 12-myristate 13-acetate (PMA)    Sigma Chemical Co., Missouri, USA

Ponceau S       Sigma Chemical Co., Missouri, USA

SB202190 (p38 inhibitor)     Sigma-Aldrich Co. LLC, New Zealand

Sodium bicarbonate (NaHCO3)    Sigma-Aldrich Co. LLC, New Zealand

Sodium chloride (NaCl)     BDH Lab Supplies Ltd, Poole, England

Sodium dodecyl sulphate (SDS)    Sigma-Aldrich Co. LLC, New Zealand

Sodium hydroxide (NaOH)     BDH Lab Supplies Ltd, Poole, England

SP600125 (JNK Inhibitor)     Sigma-Aldrich Co. LLC, New Zealand

Supersignal West Dura Chemiluminescence   Pierce Biotechnolgy Inc., Illionois, USA

Thimerosal       Sigma Chemical Co., Missouri, USA

Tris(hydroxymethyl)aminomethane    Sigma-Aldrich Co. LLC, New Zealand

Triton X100      Sigma-Aldrich Co. LLC, New Zealand

Trypan blue solution (0.4%)     Sigma-Aldrich Co. LLC, New Zealand

Tween 20®       Sigma-Aldrich Co. LLC, New Zealand

2.1.2 Media

Penicillin/Streptomycin solution:  Invitrogen, Life Technologies, New Zealand 1000 U of Penicillin G &                                                                                                 1000 µg of Streptomycin/mL

RPMI 1640 media, with phenol red   Sigma-Aldrich Co. LLC, New Zealand

RPMI 1640 media, without phenol red  Sigma-Aldrich Co. LLC, New Zealand

2.1.3 General solutions and buffers

  1. Roswell Park Memorial Institute (RPMI)-1640 media (with or without    phenol red)

Powered RPMI-1640 (Sigma-Aldrich Co. LLC, New Zealand) was prepared in accordance to the manufacturer’s instructions. RPMI-1640 was dissolved in nano-pure water before addition of sodium bicarbonate (NaHCO3) and adjustment of pH to 7.4 by the addition of 11.4M hydrochloric acid (HCl). The RPMI-1640 medium was next sterilized by filtration through a 0.22 µm Millex® filter (Sartorius AG, Goettingen, Germany) and a peristaltic pump (CP-600, Life Technologies, Maryland, USA) into sterile 500 mL bottles previously washed with nitric acid and autoclaved. Prepared media was stored at 4oC.

Premade sterilized RPMI-1640 was also used (HycloneTM, GE Healthcare Life Sciences, Utah, USA) pre-supplemented with sodium bicarbonate, pH 7.4

  1. Phosphate buffered saline (PBS)

PBS buffer consisted of 150mM sodium chloride (NaCl) and 10mM sodium dihydrogen orthophosphate (NaH2PO4) at pH 7.4. The solution was prepared by mixing 50 mL of 3M NaCl, 40 mL of 250mM NaH2PO4, pH 7.4 and 910 mLof nano-pure water. PBS required for sterile cell culture experiments was sterilized by autoclaving (15 minutes, 121oC, 15 PSI).

  1. 7,8-dihydroneopterin solution

A 1mM 7,8-Dihydroneopterin (7,8-NP) (Schircks Laboratories, Switzerland) stock solution was prepared in RPMI 1640 media, in a 15 mL screw top centrifuge tube (Greiner, Greiner Bio-one, Neuburg, Germany). The solution was sonicated for 10 minutes or until dissolved before being sterilized by passage through a 0.22µM PES syringe filter (Membrane Solution, USA). The 7,8-NP solution was kept on ice at all times and used immediately after preparation.

  1. D-neopterin solution

Neopterin stock solution (1mM) was prepared by dissolving D-neopterin powder (Schircks Laboratories, Switzerland) in RPMI-1640 via sonication in a 15 mL screw top centrifuge tube (Greiner, Greiner Bio-one, Neuburg, Germany). The solution was sonicated until dissolved before being sterilized by passage through a 0.22µM PES syringe filter (Membrane Solution, USA). The neopterin solution was kept on ice at all times and used immediately after preparation.

  1. Triton lysis solution

Lysis buffer consisted of 0.5% Triton-X100 (diluted from 1% stock solution in nano-pure water). Protease inhibitor (ProteaseMini, Roche Diagnostics) prepared and frozen to the manufacturers recommendations (ProteaseMini, Roche Diagnostics), was defrosted and added to the lysis solution on the day of use.

  1. MOPS buffer for SDS-PAGE

A 10x stock solution of MOPS was made with 500mM MOPS, 500mM Tris base, 1% SDS (w/v), and 10mM EDTA in water with pH adjusted to 7.7 (adjusted with concentrated HCl). Stock solution was diluted with nano-pure water to 1x concentration before use.

  1. Transfer buffer for western blotting

Transfer buffer consisted of 25mM Tris, 200mM glycine and 20% methanol (v/v) in water and was stored at 4oC. Transfer buffer was able to be reused once.

  1. Tris-buffered saline (TBS)

TBS used for nitrocellulose membrane washing, consisted of 40mM Tris-HCl, 150mM NaCl, 0.05% Tween20 (w/v) and 0.01% thimerosal (w/v) dissolved in nano-pure water. Final pH was adjusted to 7.5 using concentrated HCl.

  1. TBSM

The membrane blocking solution (TBS-milk, TBSM) consisted of 5% (w/v) of nonfat milk powder (Anchor, New Zealand) dissolved in TBS. It was stored at 4C for up to one week.

  1. Cracker buffer for SDS-PAGE

To make the cracker buffer, 125mM Tris-HCl, pH 6.8 (adjusted with concentrated HCl), 1% SDS (w/v), 20% glycerol (w/v) and 0.1% bromophenol blue (w/v) were mixed in nano-pure water. Immediately prior to use, 2% (v/v) of β–mercaptoethanol and 0.5mM EDTA was added to the amount required of the above solution.

  1. Restore stripping buffer

Restore stripping buffer solution used to remove antibodies from the nitrocellulose membrane in preparation for reprobing consisted of 2 % SDS, 50mM Tris pH 6.8 , and 100µM β–mercaptoethanol and made up to volume in nano-pure water.

  1. Ponceau S stain

Ponceau S stain consisted of 0.01% Ponceau S (w/v) in 5% acetic acid (v/v).

  1. Propidium iodide solution

A 1 mg/mL stock solution of propidium iodide dye (Sigma-Aldrich Co. LLC, New Zealand) was prepared in nano-pure water and was kept in a sterilized schott bottle, wrapped in tin foil at 4oC

  1. Phorbol 12-myristate 12-acetate (PMA)

A 5mM PMA stock solution was prepared by dissolving PMA powder in DMSO. 20 µL aliquots were pipetted into 1.7 mL centrifuge tubes and stored at -20oC

2.2 Methods

2.2.1 Cell culture based experiments

2.2.1.1 Aseptic technique

All work involving cell culture was performed under aseptic conditions using a Class II biological safety cabinet (Clyde-Apex BH 200). All cell culture equipment used was sterile-brought plastic ware (Falcon, Terumo, Unomedical, Greiner Bio-one) or had been sterilized by autoclaving (15 miniutes, 121oC, 15 PSI). All media/solutions added to cells was sterilized by autoclaving or by filtration using a 0.22 µm filter (Membrane Solutions, USA). All equipment was sprayed with 70% ethanol (diluted from stock solution in distilled water) before use in the Class II safety cabinet. Cells were kept incubated in a humidified atmosphere at 37°C containing 95% air w/ 5% CO2 (Sunnyo Electric Cp. LTD, Japan).

2.2.1.2 Standard RPMI-1640 media for U937 cell culture

RPMI-1640 with phenol red enriched with 100 U/mL penicillin G, 100 µg/mL streptomycin and 5% (v/v) foetal bovine serum (FBS) was used as standard in U937 cell culture maintenance.

2.2.1.3 Preparation and culture of the U937 cell line

The U937 cell line is a type of immortal cell line derived from the malignant cells of a pleural effusion from a patient with diffuse histiocytic lymphoma, and are one of the few human cell lines expressing monocyte like characteristics exhibited by cells of histiocytic origin. The U937 cells used in this work were from a line gifted by the Haematology Resesearch Laboratory of the Christchurch School of Medicine, University of Otago.

Prior to revival stock U937 cells were stored in liquid nitrogen using 1 mL storage vials containing 10×106 cells/mL in dimethyl sulfoxide (DMSO) freezing medium. To prepare stock U937 for use one vial was defrosted in a 37oC water bath until almost fully thawed. This suspension was next poured into 20 mL of RPMI-1640 and centrifuged at 500g for 5 minutes in order to separate the DMSO from the cells. The cell pellet was re-suspended in 10 mL of RPMI-1640 before being cultured in 25 cm3 tissue culture flasks (Flacon, BD, USA) until the cells were deemed fully recovered from their storage state and transferred into a standard 75 cmtissue culture flask (Greiner Bio-one, Germany).

Cells were deemed suitable for experimental work once they had reached an approximate density of ~1×106 cells/mL and division rate had normalized to a doubling time of 92-96 hours. These cells showed a spherical morphology with small grain like protrusions on the membrane from cytoplasmic organelles, and with little clustering or blebbing. Subsequent cell density was maintained between 0.3-1.5×10cells/mL by passaging every 2-3 days.

2.2.1.4 Cell culture experimental conditions and procedures

Experiments with the U937 cell line were performed using either 12 or 24-well suspension culture plates (Cellstar ®, Greiner Bio-one). The total volume used during experimental work was 1 mL per well. Cells were counted using a haemocytometer in conjunction with a light microscope after dilution with trypan blue (ratio of 1:1). The required amount of cells was centrifuged at 500 g for 5 minutes at room temperature then re-suspended in the required experimental medium pre-heated to 37oC using a water bath. The cells were then aliquoted into the plate wells to a final concentration of 0.5×106 cells/mL.

2.2.2 Cell viability assays

2.2.2.1 Propidium iodide cell viability assay

U937 cell viability was measured after experimental treatment via the use of propidium iodide (PI) staining and flow cytometery (BD AccuriTM C6 flow cytometer, BD Biosciences, California, USA). PI is able to intercalate between DNA bases when the cell membrane becomes compromised during cell death. Intercalation can be measured via fluorescence at an excitation wavelength of 535 nM and emission wavelength of 617 nM. Combined with the use of a flow cytometer PI can be used to examine cell viability along with identifying necrotic, apoptotic and healthy cells.

After experimental treatment 250 µL of cell suspension was incubated with 4 µL of 1mg/mL PI and mixed via inversion. Incubation was allowed to occur in the dark for 10 minutes. Viability was measured via using a set volume (30 µL) analyzed on the flow cytometer using forward scatter (FSC) and the fluorescence filter FL-3 (FL-3). Cellular debris were identified and excluded from analysis and the cellular viability ratio (PI+ve:PI-ve) was compared to control samples.

2.2.2.2 Trypan blue viability test

The trypan blue viability tests is a dye exclusion test which is used to determine the number of viable cells present in a cell suspension. Live cells contain intact cell membranes that exclude the Trypan Blue dye, whereas non-viable cells have compromised cellular membranes that allow for the dye to enter the cellular cytoplasm staining it blue.

After experimental treatment the cellular suspension was diluted in a 1:1 ratio with 0.4% Trypan Blue solution and incubated for two minutes. The suspension was then transferred to a hemocytometer and the number of viable and non-viable cells counted. Viable cells were identified as having a white or clear cytoplasm, while non-viable cells were identified as having a blue cytoplasm. The ratio of viable:non-viable cells was then calculated and cellular viability determined.

2.2.3 Determination of protein concentration

Protein concentration of cell lysate was quantified using the bicinchoninic acid (BCA) protein determination kit assay (PierceTM, Rockford, USA) in which the reduction of Cu2+ to Cu+ by protein in an alkaline medium can be measured via a BCA reagent. The reaction product of this assay exhibits strong absorbance at 562 nM with increasing protein concentration in the range of 25–250µg/mL.

Fresh working reagent was prepared prior to each use by combining Reagent A (NaCO3, NaHCO3, BCA and sodium tartrate in 0.1M sodium hydroxide) and reagent B (4% CuS04.5H2O) at a ratio of 50:1 (5 mL Reagent A to 100 µL Reagent B). The assay was run by mixing 50 µL of cell lysate (diluted in distilled water as required) with 1 mL of working reagent prior to a 30 minute incubation on a shaking heating block at 60o C to start the reaction. When complete the reaction was stopped by placing the samples on ice until cool and the absorbance read at 562 nM against a water blank. Protein concentration was determined using a standard curve by measuring the absorbance of known concentrations of BSA (0-250 µg/mL) under the same conditions.

2.2.4 SDS-PAGE

2.2.4.1 Cell lysate preparation.

U937 samples for western blot were prepared in the following manner. All medium within the wells were pippeted up and down to dislodge the cells into suspension before transfer into 1.7 mL centrifuge tubes. Cells were centrifuged for 15,000g, 0C for 5 minutes in order to pellet cells. The supernatant was discarded and cells re-suspended in sterilized ice cold PBS before being centrifuged at 15,000 g, at 0oC for 5 miniutes in order to wash the cells. The supernatant was discarded and 150 µL of ice-cold lysis buffer (containing protease inhibitor) was added to each centrifuge tube and left on ice for 25 miniutes in order to ensure lysation. Cell lysate was stored at -80oC degrees until analyzed.

2.2.4.2 SDS-polyacrylamide gel electrophoresis

The required amount of sample as determined by protein concentration (generally 50 µg at volumes ~30-50 µL) was aliquoted into 1.7 mL centrifuge tubes. 400 µL of ice-cold acetone was added to each tube and centrifuged for 15,000, 0oC for 10 minutes to pellet the cells. The supernatant was discarded and the tubes were left in a fume hood for any residual acetone to evaporate. The pellet was re-suspended in fresh cracker buffer containing 2% β-mercaptoethanol to a final protein concentration of 2µg protein per µL. The samples were then heated in a heating block at 90oC for 2 minutes to denature the proteins. The samples were centrifuged for 5 minutes at 20,000 g. 5 µL of pre-stained molecular weight marker mix (Fermentas International Inc, Ontario, Canada) and 25 µL/well of sample were loaded into the gel wells. The gel (4-12% Bis-Tris Gel, Invitrogen, California, USA) was run at 100V at 300mA in 1x MOPS buffer until the dye had run past the wells, and 200V at 300mA thereafter until the dye had reached the bottom of the gel.

2.2.5 Western blot analysis

2.2.5.1 Protein transfer

Proteins separated on the SDS-PAGE gel were transferred onto a nitrocellulose membrane using either wet or semi-dry transfer protocols. For the wet method transfer occurred over 800 minutes at 70V in a water cooled tank transfer electrophoresis unit (TE22, Hoefer, USA) containing the transfer buffer. For the semi-dry method transfer occurred at 1.3A, 25V for 10 minutes using the Trans-Blot® Turbo™ Transfer System (Bio-Rad Laboratories, California, USA) following the manufactures instructions.

The following steps were all undertaken on a rocking platform mixer (Ratex Instruments, Australia). Once transfer had been completed the nitrocellulose membrane was rinsed with nano-pure water and incubated with 0.01% Ponceau S stain for 1 minute to check transfer efficiency. Efficient transfer showed distinct pink columns forming at the location of each well. The membrane was then blocked using 5% TBSM for 1.5-2 hours with the TBSM being replaced every 30 minutes. During this time the Ponceau S stain completely disappeared.

2.2.5.2 CD36 immunostaining

Following blocking the nitrocellulose membrane was incubated with rabbit polyclonal affinity purified antibody raised against human CD36 (NB400-145, Novus Biologicals Inc., USA) for 1.5 hours diluted to 1:1,000-2000 in 1% TBSM. The nitrocellulose membrane was then washed 5 times for 5 minutes in TBS before undergoing incubation for 1 hour with the secondary antibody; goat polyclonal anti-rabbit IgG, conjugated to hydrogen peroxidase (HRP) (NB730-H, Novus Biologicals Inc., USA) diluted to 1:2,000. Incubation was followed by 5 lots of 5 minute washes in TBS before being briefly rinsed with nano-water prior to visualization.

2.2.5.3 Visualization

Visualization of the antibody staining was achieved via chemiluminescence signaling corresponding to the location of the HRP-conjugated secondary antibody. The signal was produced by incubating the nitrocellulose membrane with Supersignal West Dura Chemiluminescence (TerumoScientific, USA) substrates A & B, mixed at a ratio of 1:1. Images containing the signal were obtained at high resolution, without filter or light, using the time-lapse auto-exposure setting on a Syngene Chemigenius-2 bioimaging system (USA). The images were then analyzed for band intensity using GeneSnap (Syngene, USA) and ImageJ64 1.42q.

2.2.5.4 Nitrocellulose membrane stripping and β-Actin probing

Re-probing of the nitrocellulose membrane for β-actin was always subsequent to visualization of CD36 during which time the nitrocellulose membrane was kept in TBS at 4C. Prior to re-probing for β-actin the nitrocellulose membrane was stripped of all previous antibodies via incubation with stripping buffer (2% SDS, 50mM Tris, 100µM β–MEtOH, pH 6.8, heated to 50C on a water bath) for 20 minutes, followed by 5 lots of 5 minute washes in TBS. Reprobing for β-actin was done directly after stripping without blocking.

Once stripping was completed the nitrocellulose membrane was incubated with mouse monoclonal antibody raised against β-actin (A5316, Sigma-Aldrich Chemical Co., USA) for 1.5 hours diluted to 1:7,500 in 1% TBSM. The nitrocellulose membrane was then washed 5 times for 5 minutes in TBS before undergoing incubation for 1 hour with the secondary antibody; anti-mouse IgG Peroxidase-linked NA931 (GE Healthcare, UK) diluted to 1:7,500 in 2% TBSM. Incubation was followed by 5 lots of 10 minute washes in TBS before being briefly rinsed with nano-pure water prior to visualization.

2.2.6 Flow cytometry determination of U937 CD36 cell surface expression

Flow cytometry is a powerful technique that is able to analyze the fluorescence of individual cells passing through a laser light source. The fluidic systems within the flow cytometer uses hydrodynamic focusing to arrange the particles in a sample into a straight stream that passes through the light source. The flow cytometer then collects light scattering and fluorescent emission information about the particle to reveal information about its properties. The flow cytometer contains two separate detectors, the Forward Scatter Channel (FSC) which is offset 20 degrees from the beam axis, and the Side Scatter Channel which is offset 90 degrees. The patterns generated from these two detectors are unique for every particle and provides information of particle size, granular content, while also allowing identification of live cells and cellular debris.

Cells were extracted from experimental wells and centrifuged in a pre-cooled centrifuge tube (4oC) for 5 minutes at 2000 rpm. The supernatant was then discarded and the cells re-suspended in 245 µL PBS + 0.5% BSA. 5 µL of Purified Mouse Anti-Human CD36 primary antibody (BD Biosciences) was added and incubated on ice for 20 minutes (final concentration: 10µg/mL mouse anti-human CD36). The cells were then centrifuged for 5 minutes at 2000 rpm before being re-suspended in 190 µL PBS +0.5% BSA and 10 µL of secondary antibody Polyclonal Goat Anti-Mouse Immunoglobulins/RPE Goat F(ab’)2 added. Incubation on ice was allowed to occur for 20 minutes before the cells where centrifuged 5 minutes at 2000 rpm and brought back up in 1mL PBS +0.5% BSA. 10,000 events were then analysed on the FL2 filter using the BD Biosciences Accuri C6 Flow Cytometer. A primary antibody control (IgMk) was used also to ensure that all labeling detected was seen purely due to CD36 antibody binding.

2.2.7 Statistical analysis

The results presented in this thesis were graphed and statistically analyzed by GraphPad Prisim version 6.01 for Windows (GraphPad Software, San Diego, California, USA). Significance was confirmed via either, one-way or two-way analysis of variance (ANOVA). Significance levels are indicated as: (*) p≤  0.05, (**) p≤ 0.01 and (***) p≤ 0.001. Unless stated otherwise results presented within this thesis are shown in triplicate from at least three representative experiments. The graphs are presented as the mean and the standard error of the mean (SEM) was calculated from triplicate samples in most cases.

 

  1. RESULTS

3.1 Characterising the expression of CD36

3.1.1 The U937 cell line produces detectable basal levels of CD36

Prior research exploring the down regulation of CD36 has focused mostly on the down regulation of this scavenger receptor after an up regulation event (Draude & Lorenz, 2000; J. Han et al., 2000). In contrast this research instead focuses on the down regulation of CD36 from the basal levels that naturally occur in the U937 cell line. Before research could commence it first had to be determined at what levels basal CD36 could be detected using western blotting protocols. To determine this U937 cells were incubated in RPMI-1640 with 5% FBS for 24 hours. Due to basal CD36 levels being the only ones of interest, cells were incubated solely RPMI-1640 media with foetal calf serum. Cell lysate was then immunoblotted for CD36 with β-actin used as a loading control. It was found that a strong CD36 band could be detected at the ~100 kDa mark, with a weaker band at ~88 kDa (Figure 3.1.1). Different bands represent the different glycosylation isoforms and maturity levels of the CD36 protein. The 88 kDa band is reported to be incompletely glycosylated CD36 located in the Golgi apparatus, whereas the fully glycosylated 100 kDA band is mature CD36 residing on the plasma membrane (Alessio et al., 1996). The strength of the bands observed was considered to be suitable for detecting changes in basal CD36 levels upon treatment. It was decided that the main focus of this research would be on the 100 kDa isoform due to the strength of the band, and that cell surface CD36 is likely responsible for oxLDL binding leading to foam cell formation.

100 kDa

88 kDa

Figure 3.1.1 The U937 cell line produces detectable basal levels of CD36

U937 cells (0.5×10cells/mL) were incubated at 37oC for 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. Quantitative densitometry of western blots shows a strong band at ~100 kDa and a weaker band at ~88 kDa.

3.1.2 Triton X-100 as a lysing agent

Before further research could take place, appropriate experiments had to be under taken in order to determine an appropriate lysing agent for efficient use with the U937 cell line. Previous research using human monocyte derived macrophages (HMDM) in the Free Radical Biochemistry (FRB) lab group used a HEPES and NaCl based lysing agent. Initial experiments showed that this failed to efficiently lyse U937 cells. To overcome this Triton X-100 was considered for use. Triton X-100 is a nonionic surfactant detergent successfully used previously in working with the U937 cell line. In order to re-examine its use here U937 cells were incubated with increasing concentrations of Triton X-100 of between 0 and 1.0% concentration for 10 minutes at 37oC. Cell viability and lysation was determined via a light microscope in order to determine the appropriate amount of Triton X-100 required for complete lysation. Cell images of the Triton X-100 treatments showed that 0.05% Triton X-100 was the minimum concentration required to lyse 100% of viable cells (Figure 3.1.2).

Triton Control 0 0

Control

0.01% Triton X-100

0.1% Triton X-100

0 1

0.5% Triton X-100

1.0% Triton X-100

Figure 3.1.2 0.5% TritonX-100 effectively lysis cells

Images of U937 cells (0.5×10cells/mL) after incubation with 0–1% Triton X-100 for 10 minutes at 37oC. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera. The population density is a 500,000 cells per mL.

3.1.3 Effect of PMA of U937 expression

After confirmation that the U937 cell line produced detectable basal levels of CD36 it was of interest to further characterise CD36 expression. One aim was to see whether expression could be increased and if so to what extent. CD36 expression is known to increase as monocytes differentiate into macrophages (Chanput, Peters, & Wichers, 2015). Phorbol-12-myristate-13-acetate (PMA) is commonly used as a stimuli in the differentiation process and was considered an appropriate choice to use here. In order to examine the effect of this differentiation agent on CD36 levels U937 cells were incubated in RPMI-1640 with 5% FBS and 25nM PMA over a 24 hour time course. After incubation with PMA cell lysate was immunoblotted for CD36 with β-actin used as a loading control.

Results showed that at the 6 hour time point there appeared to be an increase in CD36 expression to 129.66% of control (Figure 3.1.3). This increase however was non-significant and had dropped down to 87.93% of control by the 12 hour time point. CD36 expression then held relatively steady till time course completion at 24 hours at which it was found to be present at 92.62% of control. Overall the results showed that at 25nM PMA caused no significant change in CD36 expression over a 24 hour time course.

A

B

100 kDa

β-actin

  Time (Hours)         0               6                 12             24

Figure 3.1.3 PMA has a limited effect on U937 expression

U937 cells (0.5×10cells/mL) were treated with 25nM of PMA and incubated at 37oC over a 24 hour time course in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of duplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (**) p ≤ 0.001. B) Quantitative densitometry of western blots shows PMA having a limited effect on CD36 relative to controls.

 

 

3.2 Characterization of CD36 down regulation via 7,8-NP

3.2.1 7,8-NP reduces the expression of the CD36 scavenger receptor in the U937 cell Line

Previous research has shown that the macrophage derived antioxidant 7,8-dihydroneopterin (7,8-NP) caused a decreased in CD36 expression at concentrations up to 250µM in HMDM cells (S. P. Gieseg, Z. Amit, Y.-T. Yang, A. Shchepetkina, & H. Katouah, 2010). While the U937 cell line is considered to be a good model for researching monocytes, 7,8-NP induced down regulation needed to be confirmed here. To achieve this U937 cells were incubated for 24 hours with 7,8-NP concentrations ranging from 0-150µM. Cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control. Down regulation was observed beginning at 50µM 78-NP with CD36 expression found to be 54.77% of control (Figure 3.2.1). Expression then decreased further at both 100µM and 150µM concentrations (44.43% and 42.73% of control respectively). Significance was observed at all three data points, with the highest level of significance occurring at 150µM. No significant changes were observed between the different concentrations. The strength of this down regulation showed that the effect of 7,8-NP on CD36 in the U937 cell line was stronger to that seen with HMDM’s and that 150µM 7,8-NP was the most appropriate concentration to use in further experimentation due to it having the most significant effect.

A

B

100 kDa

β-actin

7,8-NP (µM)       0                    50                  100            150

 

Figure 3.2.1 7,8-NP reduces the expression of the CD36 scavenger receptor in the U937 cell line

U937 cells (0.5×10cells/mL) were treated with 0-150µM 7,8-NP and incubated at 37oC over a 24 hour time course in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots shows 7,8-NP reduces CD36 expression between 50-150µM relative to controls.

3.2.2 7,8-NP induced down regulation occurs in both adherent and suspension well plates

Two common cell culture plates used in experimental work are adherent and suspension plates. In adherent plates cells will interact closely with the plate, forming an adherent layer on the well surface, where as in suspension plates cells are suspended within the media minimising well interaction. It is possible that the type of cell culture plate used may have an effect on the morphology and differentiation of U937 cells, and subsequently production of the heavily glycosylated 100 kDa CD36 band. This is more likely to occur when adherent plates are used due to closer interaction between the cells and the plate, which may be recognised as a foreign body and trigger the differentiation process. In order to rule out any effect due to the type of plate used on CD36 levels, and the subsequent effects of 7,8-NP, both adherent and suspension plates were examined for use in CD36 down regulation experiments. U937 cells were incubated with increasing concentrations of 7,8-NP ranging from 0-150µM for 24 hours in both adherent and suspension plates. Subsequently the cells were washed and lysed. Cell lysate was immunoblotted for CD36 with β-actin used as a loading control.

It was found that CD36 could be detected at strong basal levels when using both types of plates and that a concentration related decrease could be observed with increasing concentrations of 7,8-NP in both cases (Figure 3.2.2). These results show that any U937 interaction with the type of plate use has no or a very limited effect on basal CD36 levels and 7,8-NP induced down regulation. As such all further experimentation looking at this effect will take place using suspension well plates owing to lesser cell-plate interaction and easier use in extracting cells from the wells.

100 kDa

β-actin

Adherent

100 kDa

β-actin

Suspension

 

 

 

 

 

 

Figure 3.2.2 7,8-NP down regulation can be observed using both adherent and suspension plates

U937 cells (0.5×10cells/mL) were treated with increasing concentrations of 7,8-NP (0-150µM) and incubated at 37oC for 24 hours in RPMI-1640 with phenol red using both adherent and suspension well plates. Cell lysate was assessed for CD36 protein expression via western blot. Figure shows the quantitative densitometry of western blots and that increasing 7,8-NP concentration reduces CD36 expression relative to controls in both plate types.

 

 

 

 

 

 

 

3.2.3 7,8-NP down regulation can be observed using both wet and semi-dry protein transfer protocols

Western blotting involves the transfer of proteins after polyacrylamide gel electrophoresis over to a membrane suitable for antibody probing such as nitrocellulose. Two commonly used transfer techniques (wet and semi-dry) were explored for their suitability in experimental work looking at the 7,8-NP induced down regulation of CD36. The wet transfer technique system uses a larger volume of transfer buffer and is run over a longer time period, whereas the semi-dry transfer system uses less buffer and is run over a shorter time. The wet transfer system is usually considered to be better suited for larger molecular weight proteins, whilst the semi-dry method is thought to be more ideal for smaller proteins. To explore any differences in their suitability for this work U937 cells were incubated with increasing concentrations of 7,8-NP ranging from 0-150µM with protein transfer occurring using either a wet or semi-dry protocol. It was found that both systems were able to detect 7,8-NP induced down regulation of CD36 to a similar extent as confirmed via antibody probing (Figure 3.2.3). Both systems also transferred similar amounts of protein from the SDS page gel onto the nitrocellulose membrane as detected via both ponceau staining of the nitrocellulose membranes after transfer, along with coumassise blue staining of the SDS gel itself (results not shown). However the semi-dry method was found to produce a more reliable transfer with almost no air bubbles forming during the transfer process compared to the wet transfer method. Overheating of the nitrocellulose membranes was also cut back when the semi-dry method was used. Due to these factors the semi-dry system was the method of choice for all future transfers.

100 kDa

β-actin

Wet

100 kDa

β-actin

Semi-Dry

Figure 3.3.3 7,8-NP down regulation can be observed using both wet and semi-dry transfer protocols

U937 cells (0.5×10cells/mL) were treated with increasing concentrations of 7,8-NP (0-150µM) and incubated at 37oC for 24 hours in RPMI-1640 with phenol red using both wet and semi-dry transfer methods. Cell lysate was assessed for CD36 protein expression via western blot. Figure shows quantitative densitometry of western blots and that increasing 7,8-NP concentration reduces CD36 expression relative to controls in protein transfer systems.

 

 

 

 

 

 

3.2.4 Time course of 7,8-NP induced CD36 down regulation

Once CD36 down regulation was confirmed in the U937 cell line a time-course was performed in order to investigate the speed at which 7,8-NP has its effect. To achieve this U937 cells were incubated with 150µM 7,8-NP over a 24 hour time course. During incubation the cells were extracted from the wells at 0, 6, 12 and 24 hours and lysed in the presence of protease inhibitor. Cell lysate was kept at -80oC until analysis. Once the time course was complete the cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control.

Down regulation was observed beginning at 6 hours with CD36 expression found to be at 55.12% of control (Figure 3.2.2). CD36 expression then continued to decrease at both 12 and 24 hour time points (43.56% and 29.16% of control respectively).  Change compared to control was significant at all time points, being the most significant at the 24 hour time point. No significant changes were observed between the different time points. The results suggest that CD36 down regulation occurs rapidly within the first six hours of incubation with 7,8-NP but continues on to some extent to the 24 hour time point.

C:UsersyeandleaDesktopUniversity Masters ThesisTHESIS-RESULTSCompleteU937 + SB202U937 +SB202.tif

A

B

100 kDa

β-actin

 Time (Hours)        0                  6                 12                 24

                        

 

 

 

 

 

 

 

Figure 3.2.4 7,8-NP down regulates CD36 over 24 hours

U937 cells (0.5×10cells/mL) were treated with increasing concentrations of 7,8-NP (0-150µM) and incubated at 37oC for 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A)  Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots shows 150µM 7,8-NP decreasing CD36 expression over 24 hours relative to controls.

3.2.5 Recovery of CD36 expression after 7,8-NP removal

Once the 7,8-NP induced down regulation of CD36 had been established the next step was to examine the recovery of CD36 after 7,8-NP removal. Investigation into the recovery of the cells after treatment is an important factor into the characterization of 7,8-NP’s effect on CD36. It also provides evidence as to whether or not cells are suitable for further experimentation after initial down regulation. Previous work examining the recovery of HMDMs after the removal of 7,8-NP was limited and only examined recovery up to a six hour time point (Schepetkina, 2013). Here we look at recovery up to 48 hours after 7,8-NP removal. U937 cells were incubated for 24 hours with 150µM 7,8-NP in a 75 cm3 cell culture flask in order to allow CD36 down regulation to take place. The cells were then centrifuged at 500x g to pellet the cells and the supernatant discarded to remove 7,8-NP. The cells were then re-suspended in fresh media before transfer into a 24 well suspension plate and incubated at 37oC. Cells were extracted from the wells at 0, 12, 24 and 48 hour time points and lysed in the presence of protease inhibitor and kept at       -80oC until analysis. Once the time course was complete the cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control.

The results show that after a 24 hour incubation period with 150µM 7,8-NP CD36 levels were down-regulated to a significant 50.22% of control (Figure 3.2.5). Recovery was then observed 12 hours after 7,8-NP removal where CD36 expression had recovered to a non-significant 63.17% of control, and 24 hours after removal CD36 expression had recovered to a significant 96.02% of control. Between the 24 and 48 hour time points CD36 expression began to decrease again to 30.91% of control at 48 hours. Recovery of CD36 between the 12 and 24 hour time points, while showing that recovery does occur, limits time for any subsequent experimental work to between 0 and 12 hours after removal to ensure levels of CD36 are still down regulated.

A

B

100 kDa

β-actin

       Time (Hours)      Control       0             12              24              48

 

Figure 3.2.5 CD36 expression recovers 24 hours after 7,8-NP removal

U937 cells (0.5×10cells/mL) were treated 150µM 7,8-NP and incubated at 37oC over 24 hours in RPMI-1640 with phenol red to allow for down regulation to occur. 7,8-NP was then removed and the cells plated for an additional 48 hour time course. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots shows 7,8-NP CD36 expression recovering 24 hours after 7,8-NP removal relative to controls.

3.2.6 Cellular viability of the recovery experiment

During the recovery experiment the U937 cells underwent down regulation via 7,8-NP for 24 hours before being plated for up to an additional 48 hours. The viability of this experiment was investigated at each time point to ensure that the results observed were not influenced by the loss of cellular viability over the experimental time course. At each time point (0, 12, 24 and 48 hours) prior to full cellular extraction and lysation, 50 µL of cells were taken from each well and incubated with 0.4% trypan blue at a ratio of 1:1 for 2 minutes before viability analysis using a heamocytometer. Results showed that the recovery experiment had very little effect on cell viability (Figure 3.2.6). Viability immediately after 7,8-NP removal was found to be at 96.37%. This level then held relatively steady over the entire time course with viability after 48 hours measured at 95.11%. No significant changes were observed between the different time points.

Cells were also viewed under a light microscope at 20x magnification and images taken to observe morphological changes. Changes in cell morphology during 7,8-NP induced down-regulation and after removal appear to be limited. Control cells show a typical spherical morphology which continued on through to 48 hours after 7,8-NP removal.

A

B

Control012

Control

0 Hours

12 Hours

 

2448

24 Hours

48 Hours

Figure 3.2.6 CD36 expression recovers 24 hours after 7,8-NP removal

U937 cells (0.5×10cells/mL) were treated with 150µM of 7,8-NP at 37oC over 24 hours. 7,8-NP was then removed and cells incubated for a further 48 hour time course.  A) Cell viability was measured by tyrpan blue and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B. Images of U937 cells during each time point of recovery. B) Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera. The population density is a 500,000 cells per mL.

3.2.7 Effect of neopterin on CD36 expression in the U937 cell line

Neopterin is the oxidised product of 7,8-NP, and like 7,8-NP is also a member of the pteridine family. Biologically it can be used to provide information on cellular immune activation and as an inflammatory disease marker (Murr, Widner, Wirleitner, & Fuchs, 2002). Once the down regulation effect of 7,8-NP was characterised in the U937 cell line it was of interest to measure any effect that neopterin may have on CD36 levels. To achieve this U937 cells were incubated with increasing levels of neopterin ranging from 0-100µM for 24 hours. Afterwards the cells were extracted from the wells and lysed. The cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control.

The results show that in the 0-100µM concentration range neopterin had no significant effect of CD36 expression compared to control levels (Figure 3.2.7). At 25µM & 75µM neopterin CD36 expression was 8.14% and 19.43% lower than control respectively, while at 50µM and 100µM CD36 expression was 6.50% and 12.29% higher than control. The fact that none of these changes were found to be significant or followed a discernible pattern indicates that unlike 7,8-NP, neopterin has no effect on CD36 expression.

Cells were also viewed under a light microscope at 20x magnification and images taken to observe any morphological changes upon treatment. Changes in cell morphology during treatment with neopterin are limited with control cells showing a typical spherical morphology comparable to that seen up to and including 150µM neopterin (Figure 3.2.7).

A

B

100 kDa

β-actin

Neopterin (µM)  0            25             50       75            100

Figure 3.2.7 Neopterin has a limited effect on CD36 expression

U937 cells (0.5×10cells/mL) were treated with increasing concentrations of Neopterin (0-100µM) at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots shows limited effect on CD36 expression decreasing with increasing neopterin concentration relative to controls.

 

control2550

Control

25µM Neopterin

50µM Neopterin

 

75100

75µM Neopterin

100µM Neopterin

 

Figure Error! No text of specified style in document..17 Images of U937 cells after treatment with increasing concentrations of neopterin

Images of U937 cells after treatment with increasing concentrations of neopterin for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera. The population density is a 500,000 cells per mL.

 

 

3.3 Transcriptional control of CD36

3.3.1 Effect of PPARγ inhibitor GW9662 on CD36 expression

Previous publications have shown that CD36 is under control of a heterodimer formed between the nuclear receptors PPARγ and RXR (S. Han & Sidell, 2002). A kinase binding site located on the PPARγ component makes it a likely target for exploration into the 7,8-NP induced down regulation pathway. Yet before this could commence it first had to be confirmed that CD36 was indeed under control of this heterodimer in the U937 cell line. To achieve this GW9662, an irreversible antagonist of PPARγ was chosen for use. GW9662 selectively and covalently modifies the CYS285 residue of PPARγ, and is 10- and 600-fold less potent in binding experiments using PPARα and PPARδ isoforms (Leesnitzer et al., 2002).

U937 cells were incubated for 24 hours with increasing concentrations of GW9662 ranging from 0-15µM, with cells only used as a negative control and 150µM 7,8-NP as a positive control. At the 24 hour time point the cells were extracted and lysed. The cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control. GW9662 significantly decreased CD36 expression beginning at 5µM (66.47% of control) with a similar level of effect seen at 10µM (72.69% of control) (Figure 3.3.1). 15µM GW9662 decreased CD36 expression further still (41.57% of control). Change compared to control was significant at all concentrations, being the most significant at 15µM. No significant changes were observed between the different concentrations. 150µM 7,8-NP alone decreased CD36 expression to 60.89% of control, comparable most to that seen with 5µM. The results confirm that CD36 is under control of PPARγ, and also our choice of PPARγ as a target in the exploration of the 7,8-NP induced down regulation pathway.


A

B

100 kDa

β-actin

Figure 3.3.1 PPARγ inhibitor GW9662 decreases CD36 expression

U937 cells (0.5×10cells/mL) were treated with increasing concentrations of GW9662 (0-15µM) at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots shows CD36 expression decreasing with increasing GW9662 concentration relative to controls.

3.3.2 Effect of GW9662 on cellular viability

The PPARγ transcription factor alongside its role in CD36 regulation, is involved in controlling a wide variety of genes over several pathways related to lipid metabolism. These include fatty acid transport, catabolism and storage (Hardwick, Osei-Hyiaman, Wiland, Abdelmegeed, & Song, 2010). The use of the GW9662 inhibitor would interfere with these processes with a possible negative effect on cellular viability. To confirm the reliability of the results of the previous experiments cell viability was measured using the propidium iodide assay. This assay can determine the ratio of alive cells which have excluded the dye (PI negative) to non-viable cells (PI positive). Cells were incubated for 24 hours with increasing concentrations of GW9662 of between 5-15µM in RPMI-1640 without phenol red as the use of this colored pH indicator has the potential to interfere with assay results. Results show that all treatments had a limited effect on cellular viability with all GW9662 concentrations used non-significantly reducing viability in a range of between 1.56-5.75% (Figure 3.3.2). The low impact that GW9662 had on viability, even at the higher concentrations, indicate cell viability loss was most likely not a factor in the GW9662 triggered decrease in CD36 expression.

Changes in cell morphology after GW992 treatment was examined using a light microscope (Leitz Wetzlab, Germany). Changes in cell morphology during treatment with GW9662 are limited with control cells showing a typical spherical morphology comparable with that seen up to and including 15µM GW9662 (Figure 3.2.2).

 

 

 

 

 

 

A

B

Control (G)5 (G)

Control

5µM GW9662

 

10 (G)15 (G)

10µM GW9662

15µM GW9662

 

Figure 3.3.2 Effect of GW9662 on cellular viability using propidium iodide

U937 cells (0.5×106) were treated with between 0-15µM GW9662 at 37oC over 24 hours in non-phenol red RPMI-1640. A) Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Images of U937 cells after treatment with increasing concentrations of GW9662 for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera. The population density is a 500,000 cells per mL.

3.3.3 Effect of GW9662 in conjunction with 7,8-NP on CD36 expression

Once it was confirmed that both 7,8-NP and GW9662 were able to down regulate observed levels of CD36, both were given in conjunction to U937 cells to determine if any additive effect on CD36 could be seen. To achieve this U937 cells were incubated for 24 hours with 5µM GW9662 in combination with 150µM 7,8-NP. A 5µM concentration of GW9662 was chosen as this level was shown to down-regulate CD36 levels to a similar extent as 150µM 7,8-NP. GW9662 and 7,8-NP were also incubated with the cells in isolation to act as positive controls. At the 24 hour time point the cells were extracted from the wells and lysed. The cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control. The results show that while both treatments down regulated observed CD36 levels to comparable levels, no additive effect could be seen. 7,8-NP in conjunction with GW9662 down regulated CD36 expression to 62.00% of control. This result is comparable to 7,8-NP (58.55%) and GW9662 alone (61.49%)  (Figure 3.3.3). This indicates that if 7,8-NP is having its effect via PPARγ that a maximum level of down regulation can be reached where any additional treatment will have an un-proportional effect.

 

 

 

 

 

 

 

A

B



100 kDa

β-actin

Figure 3.3.3 GW9662 and 7,8-NP do not have an additive effect on 7,8-NP expression

U937 cells (0.5×10cells/mL) were treated with 5µM GW9662 in conjunction with 150µM 7,8-NP at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of quadruplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots shows no additive effect on CD36 expression with both GW9662 and 7,8-NP relative to controls.

3.3.4 Effect of increasing concentrations of cycloheximide on cellular viability

Cycloheximide produced by the bacterium Streptomyces griseus acts as an inhibitor of protein synthesis in Eukaryotic organisms by interfering with the translocation process (Jones & Cushman, 1989). It is used in determining protein half-life via the use of time-course experiments in conjunction with western-blotting. This enables the detection of changes in protein bound signal over time. Due to its effect on protein synthesis the use of cycloheximide is likely to have an effect on cellular viability at higher levels that would confound any experimental results. In order to examine what levels are suitable for experimental work involving the U937 cell line a cell viability time course was examined using various concentrations of cycloheximide. U937 cells were incubated for 24 hours with increasing concentrations of cycloheximide ranging between 0-10µM in RPMI-1640 without phenol red. At the 24 hour time point cells were extracted from cells and incubated with PI in the dark for 10 minutes before viability was measured. Results showed that increasing concentrations of cycloheximide had a negative effect on cellular viability (Figure 3.3.4). 2-5µM cycloheximide decreased viability in a range of 6.15-7.87% lower than control after 24 hours, however this decrease was non-significant. 10µM cycloheximide had a bigger impact, decreasing cellular viability to a significant 22.99% lower than that of control. No significant changes were observed between the different concentrations

Changes in cell morphology after Cycloheximide treatment were examined using a light microscope (Leitz Wetzlab, Germany). U937 cells in the absence of cycloheximide showed a typical spherical morphology. Cells treated with 2µM cycloheximide appear lower in density but still display a healthy morphology albeit with a small amount of debris present. At 5µM continuing on to 10µM density decreased and cellular debris increased with pronounced blebbing becoming apparent. (Figure 3.3.4)

 

 

A

B

Control2

2µM Cycloheximide

Control

 

510

5µM Cycloheximide

10µM Cycloheximide

 

Figure 3.3.4 Effect of increasing concentrations of cycloheximide on cellular viability

U937 cells (0.5×10cells/mL) were treated with 0-10µM of cycloheximide at 37oC over 24 hours in non-phenol red RPMI-1640. A) Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Images of U937 cells after treatment with increasing concentrations of cycloheximide for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera. The population density is a 500,000 cells per mL.

3.3.5 Effect of cycloheximide on CD36 expression

After completion of the cell viability experiment a concentration of 1µM cycloheximide was decided as best suitable for use. Next a cycloheximide time course was undertaken in order to measure the natural half-life of CD36 and to explore whether 7,8-NP additionally has any effect on promoting CD36 degradation.

U937 cells were plated in 24 well suspension plates and incubated with either 1) 150µM of 7,8-NP, or 2) 150µM of 7,8-NP in conjunction with 1µM cycloheximide. After incubation cells were extracted from the wells at 0, 3, 6 and 9 hour time points, lysed in the presence of protease inhibitor and kept at -80oC until analysis. Once the time course was complete the cell lysate was then immunoblotted for the CD36 protein with β-actin used as a loading control. A 3 hour incubation with 1µM cycloheximide triggered a decrease in CD36 expression to 83.61% of control levels while cycloheximide + 7,8-NP triggered a decrease to 73.61% of control levels. At 6 hours CD36 expression had further decreased with both cycloheximide, and cycloheximide + 7,8-NP (49.93% and 53.53% respectively). While at 9 hours the detected CD36 levels were 32.91% and 36.19% respectively. Significance was detected for both cycloheximide and cycloheximide + 7,8-NP treatments after 6 and 9 hours of incubation but not at 3 hours. There was no significance between treatments at each respective time point. The results indicate that the half-life for CD36 is at around 6 hours, and that as there was lack of any significant difference between the treatments, that 7,8-NP is not enhancing the degradation of CD36 but rather is effecting CD36 transcription, complimenting previous mRNA data (Shchepetkina, 2013).

A

B

100 kDa

β-actin

Cycloheximide

Time:                  0                3               6               9

C

Cycloheximide + 7,8-NP

100 kDa

β-actin

 

 

 

Time:             0                3             6               9

 

 

 

 

Figure 3.3.5 Effect of cycloheximide on CD36 expression

U937 cells (0.5×106) were treated with 1µM of cycloheximide at 37oC over 12 hours in non-phenol red RPMI-1640. A) Results were normalised by β-actin and displayed as mean ± SEM of quadruplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control B) Quantitative densitometry of western blots shows effect of 1µM of cycloheximide on the degradation of CD36 over 9 hours. C) Quantitative densitometry of western blots shows effect of cycloheximide + 7,8-NP on the degradation of CD36 over 9 hours.  Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001.

3.3.6 Cellular viability of the cycloheximide experiment

During the 1µM cycloheximide time course the viability of this experiment was investigated at each time point to ensure that the results observed were not influenced by the loss of cellular viability. At each time point (0, 3, 6 and 9 hours) prior to full cellular extraction and lysation 50 µL of cells were taken from each well and incubated with 0.4% trypan blue at a ratio of 1:1 for 2 minutes before analysis using a heamocytometer. Results show both cycloheximide, and cycloheximide + 7,8-NP treatments had a limited effect on cellular viability over the 9 hour time course (Figure 3.3.6). 1µM cycloheximide decreased viability to a non-significant 6.86% lower than control by the 9 hour time point where as cycloheximide + 7,8-NP decreased viability to a similar 5.36 % of control. The results indicate that loss of viability was not an issue in the cycloheximide experiment.

 

 

             

A

Cycloheximide

B

           

Cycloheximide + 7,8-NP

Figure 3.3.6 Effect of cycloheximide and cycloheximide+7,8-NP on U937 cell viability using trypan blue

U937 cells (0.5×106) were treated with 1µM of cycloheximide (A) or cycloheximide + 7,8-NP (B) at 37oC over 12 hours in non-phenol red RPMI-1640. Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001.

3.4 Effect of MAP kinase inhibitors on CD36 down regulation

3.4.1 Effect of JNK inhibition via SP600125 on 7,8-NP induced down regulation in the U937 cell line

Phosphorylation of PPARγ leads to a decrease in its CD36 transcriptional activity which in turn lowers CD36 protein expression (Adams et al., 1997; Hu et al., 1996). This phosphorylation effect makes protein kinase pathways a possible target for 7,8-NP induced down regulation. PPARγ’s kinase site has been shown to recognise and bind two different MAP kinases, c-Jun N-terminal kinase (JNK) and extracellular signal–regulated kinases (ERK) 1/2. To explore the effect on inhibiting the JNK MAP kinase pathway on CD36 expression the cell permeable and selective inhibitor SP600125 was used. SP600125 works by competing with JNK’s ATP binding site which halts the flow of the kinase cascade, thus preventing the signal from progressing (Bennett et al., 2001). U937 cells where pre-incubated with 50µM SP600125 in a 24 well suspension plate. 150µM 7,8-NP in conjunction with SP600125 was additionally added to measure whether any additive effect could be observed. 150µM 7,8-NP alone was used as a positive control and cells only as a negative control. At the 24 hour time point cells were extracted and lysed in the presence of a protease inhibitor. Cell lysate was immunoblotted for CD36 with β-actin used as a loading control.

Results showed that SP600125 did not block 7,8-NP’s effect on CD36 expression but rather was able to decrease CD36 expression to 32.65% of control alone (Figure 3.4.1). This decrease was more pronounced than the 7,8-NP control which down regulated CD36 expression to 64.86% of control. When given in conjunction SP600125 and 7,8-NP lowered CD36 expression to 42.61% of control. All results were significant.

A

B

100 kDa

β-actin

 

 

 

150µM 7,8-NP

50µM SP600125

 

 

Figure 3.4.1 JNK inhibition via SP600125 does not alleviate CD36 down regulation

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 50µM of SP600125, and both in conjunction at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots showed that SP600125 did not rescue the effect of 7,8-NP on CD36 expression relative to controls.

3.4.2 Effect of JNK inhibition on cellular viability

JNK kinase cascades are involved in mediating the cellular apoptotic response to pro-inflammatory cytokines, genotoxins and environmental stress, along with having a role in the regulation of cell proliferation, survival and differentiation (C. Davies & Tournier, 2012). Use of the SP600125 inhibitor would interfere with these processes with possible negative effects on cellular viability. To confirm the reliability of the results of the previous experiment cell viability was measured using the propidium iodide assay. Cells were incubated for 24 hours with 50µM SP600125, along with 50µM SP600125 in conjunction with 150µM 7,8-NP, 150µM 7,8-NP alone and a cell only control. At the 24 hour time point cells were extracted and cell viability measured by PI using flow cytometry.

Results show that at 50µM SP600125 had a limited effect on U937 cellular viability. After a 24 hour incubation viability decreased to 7.16% lower than that of control (Figure 3.4.2). Even though this decrease was found to be significant, the low level of change means that it was unlikely to have had a major impact on the observed results. In contrast after incubation with 150µM 7,8-NP viability was 1.37% lower than control and 7,8-NP + SP600125 in combination 3.41% lower than control.

Changes in cell morphology after SP600125 treatment was examined using a light microscope (Leitz Wetzlab, Germany). Changes in cell morphology during treatment with SP600125 are non-existent with all cellular treatments showing a typical healthy morphology (Figure 3.4.2B). In treatments with SP600125 sharp yellow strands can be seen which appear to be the inhibitor precipitating out of solution.

 

 

 

 

B

A

B

Control78

150µM 7,8-NP

Control

 

 SPSP78

50µM SP600125

SP600125 + 7,8-NP

 

Figure 3.4.2 Effect of SP600125 on U937 cell viability using propidium iodide

U937 cells (0.5×106) were treated with 150µM 7,8-NP, 50µM of SP600125 and both in conjunction at 37oC over 24 hours in non-phenol red RPMI-1640. A.) Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Images of U937 cells after treatment with 150µM 7,8-NP, 50µM of SP600125 and both in conjunction for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera. The population density is a 500,000 cells per mL.

3.4.3 Effect of MEK inhibition via PD98059 on 7,8-NP induced down regulation in the U937 cell line

PPARγ’s kinase site, alongside recognising JNK, has also been shown to recognise and bind the ERK1/2 MAP kinase (Burns, 2007). To explore the effect on inhibiting the ERK1/2 MAP kinase pathway on CD36 expression the inhibitor PD98059 was used. PD98059 has its effect by selectively inhibiting the MAP kinase kinases MEK1 and MEK2 by binding to their inactive form (S. P. Davies, Reddy, Caivano, & Cohen, 2000). This prevents any activators from binding and effectively blocks downstream activation of ERK1/2 and the signalling cascade. U937 cells where pre-incubated with 25µM PD98059 before 150µM 7,8-NP was added to selected wells to measure whether any additive effect between the two could be observed. 150µM 7,8-NP in isolation was used as a positive control and cells only as a negative control. At the 24 hour time point cells were extracted, washed in PBS and lysed in the presence of a protease inhibitor. Cell lysate was immunoblotted for CD36 with β-actin used as a loading control. Results show that PD98059 did not block 7,8-NP’s effect on CD36 expression (Figure 3.4.3).  150µM 7,8-NP was found to significantly reduce CD36 expression to 41.20% of control, while 7,8-NP in conjunction with PD98059 reduced expression to a similar 48.66% of control. The differences between these two treatments were non-significant indicating that 7,8-NP did not down-regulate CD36 expression via the ERK 1/2 pathway. PD98059 treatment alone did not have a major effect on CED36 expression with a non-significant reduction to 75.17% of control, much less than was seen with other MAP kinase inhibitors.

 

 

 

 

 

A

B

 

 

 

 

 

 

 

100 kDa

β-actin


 

 

Figure 3.4.3 MEK inhibition via PD98059 does not alleviate CD36 down regulation

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 20µM of PD98059 and both in conjunction at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots showed that SP600125 did not block the effect of 7,8-NP on CD36 expression to controls.

3.4.4 Effect of MEK inhibition on cellular viability

ERK1/2 cascades are involved in a wide variety of cellular functions including in adhesion, progression, migration, survival, differentiation, metabolism, proliferation, and transcription (Roskoski, 2012). Use of the PD98059 inhibitor would interfere with these processes with possible negative effects on cellular viability.  To confirm the reliability of the results of the previous experiment cell viability was measured using the propidium iodide assay. Cells were incubated for 24 hours with either 25µM PD98059, 25µM PD98059 in conjunction with 150µM 7,8-NP, 150µM 7,8-NP alone or a cell only control. At the 24 hour time point cells were extracted and cell viability measured by PI using flow cytometry. Results show that at 25µM PD98059 had a limited effect on U937 cellular viability. After a 24 hour incubation viability had decreased to 0.67% lower than control (Figure 3.4.4). In contrast 150µM 7,8-NP viability was 1.33% higher than control and the two in combination 0.94% lower than control.

Changes in cell morphology after PD98059 treatment was examined using a light microscope (Leitz Wetzlab, Germany). Changes in cell morphology during treatment with PD98059 are non-existent with all cellular treatments showing a typical healthy morphology (Figure 3.4.4B).

 

 

 

 

 

A

B

control  78

Control

150µM 7,8-NP

M  m78

25µM PD98059

PD98059 + 7,8-NP

 

Figure 3.4.4 Effect of PD98059 on U937 cell viability using propidium iodide

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 50µM of PD98059 and both in conjunction at 37oC over 24 hours in non-phenol red RPMI-1640. A) Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Images of U937 cells after treatment with 150µM 7,8-NP, 50µM of PD98059  and both in conjunction for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera.

3.4.5 Effect of p38 inhibition via SB202190 on 7,8-NP induced down regulation in the U937 cell line

Alongside the JNK and ERK1/2 kinases several other MAP kinases are involved in cell signalling and are candidates for involvement in 7,8-NP induced down regulation. The p38 MAP kinase and its signalling cascades are activated by stress and play a role in immune response, cellular survival and differentiation (Cuadrado & Nebreda, 2010). To explore the role of p38 in relation to 7,8-NP and CD36 the p38 inhibitor SB202190 was used. SB202190 is a selective, potent and cell permeable inhibitor of the p38 MAP kinase including both p38α and β isoforms. It works by binding within the ATP pocket of the active kinase preventing further signal transduction (S. P. Davies et al., 2000).

U937 cells where pre-incubated with 20µM SB202190 before 150µM 7,8-NP was added to selected wells to measure whether any additive effect between the two could be observed. 150µM 7,8-NP in isolation was used as a positive control and cells only as a negative control. At the 24 hour time point cells were extracted and lysed. Cell lysate was immunoblotted for CD36 with β-actin used as a loading control.

Results show that SB202190 did not block 7,8-NP’s effect on CD36.  Rather like SP600125 it was able to decrease CD36 expression alone (Figure 3.4.5). This decrease was found to be 42.32% of control, slightly less than that of 7,8-NP which down regulated CD36 expression to 55.37% of control. When given in conjunction SB202190 and 7,8-NP lowered CD36 expression to 30.71% of control, which although is a larger decrease, is not significant compared to the two in isolation. All results were found to be significant in relation to control.

 

A

B

C:UsersyeandleaDesktopUniversity Masters ThesisTHESIS-RESULTSCompleteU936 + 7,8-NP TC78np dr tc.tif

100 kDa

β-actin


 

Figure 3.4.5 p38 inhibition via SB202190 does not alleviate CD36 down regulation

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 20µM of SB202190 and both in conjunction at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots showed that SP600125 did not rescue the effect of 7,8-NP on CD36 expression relative to controls.

3.4.6 Effect of p38 inhibition on cellular viability

Due to the various roles that p38 plays in cellular signalling it is possible that the use of SB202190 would have an effect of cellular viability. To confirm the reliability of the results of the previous experiment cell viability was measured using the propidium iodide assay. Cells were incubated for 24 hours with either 20µM SB202190, 20µM SB202190 in conjunction with 150µM 7,8-NP, 150µM 7,8-NP alone or a cell only control. At the 24 hour time point cells were extracted and cell viability measured by propidium iodie using flow cytometry. Results show that 50µM SB202190 has a limited effect on U937 cellular viability. After a 24 hour incubation viability had non-significantly decreased to 2.02% lower than control (Figure 3.4.6). In contrast with the 150µM 7,8-NP treatment viability was 1.37% lower than control and with the two in combination viability was 2.26% lower than control.

Changes in cell morphology after GW992 treatment was examined using a light microscope (Leitz Wetzlab, Germany). Changes in cell morphology during treatment with SB202190 were limited with control cells showing a typical spherical morphology comparable to that seen with all other treatments (Figure 3.4.6).

A

B

Control78

Control

150µM 7,8-NP

sb202sb78

SB202190 + 7,8-NP

20µM SB202190

 

Figure 3.4.6 Effect of SB202190 on U937 cell viability using propidium iodide

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 50µM of SB202190 and both in conjunction at 37oC over 24 hours in non-phenol red RPMI-1640. A) Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (**) p ≤ 0.001. B) Images of U937 cells after treatment with 150µM 7,8-NP, 50µM of SB202190 and both in conjunction for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera.

3.4.7 Effect of NF-kB inhibition via BAY 11-7082 on 7,8-NP induced down regulation in the U937 cell line

MAP kinases are not alone in the mediation of cellular signalling. The transcription factor NF-kB has been shown to be present in macrophages, and is involved in the transcription of numerous cytokines which play a role in cell growth and differentiation (Brasier, 2006). It was of interest here to explore the role of NF-kB inhibition on CD36 expression and whether this would have any effect on 7,8-NP induced down regulation. To achieve this U937 cells where pre-incubated with 2µM BAY 11-7082 before 150µM 7,8-NP was added to selected wells to measure whether any additive effect between the two could be observed. 150µM 7,8-NP in isolation was used as a positive control and cells only as a negative control. At the 24 hour time point cells were extracted and lysed in the presence of a protease inhibitor. Cell lysate was immunoblotted for CD36 with β-actin used as a loading control.

Results show that 2µM BAY 11-7082 did not alleviate 7,8-NP’s effect on CD36 expression (Figure 3.4.7).  150µM 7,8-NP was found to significantly reduce CD36 expression to 53.66% of control, while 7,8-NP in conjunction with 1µM BAY 11-7082  reduced expression to a similar 62.34% of control. The differences between these two treatments were non-significant indicating that 7,8-NP did not down-regulate CD36 expression via the NF-kB. 2µM BAY 11-7082 treatment alone was found to have an effect on CD36 expression with a significant reduction to 55.55% of control, similar to what was seen with the MAP kinase inhibitors.

 

 

 

A

B

100 kDa

β-actin

 

150µM 7,8-NP

2µM BAY 11-7082

Figure 3.4.7 NF-kB inhibition via BAY 11-7082 does not alleviate CD36 down regulation

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 2µM of BAY 11-7082 and both in conjunction at 37oC over 24 hours in RPMI-1640 with phenol red. Cell lysate was assessed for CD36 protein expression via western blot. A) Results were normalised by β-actin and displayed as mean ± SEM of triplicate experiments. Data is expressed as percentage of the respective control (cell only control) and significance is indicated from this control. Significance levels are: (*) p ≤ 0.05, (**) p ≤ 0.01 and (***) p ≤ 0.001. B) Quantitative densitometry of western blots showed that SP600125 did not rescue the effect of 7,8-NP on CD36 expression relative to controls.

3.4.8 Effect of NF-kB Inhibition on cellular viability

NF-kB plays an important role in the mediation of stress responses and inflammation. Use of the BAY 11-7082 inhibitor would interfere with these processes with possible negative effects on cellular viability. To confirm the reliability of the results of the previous experiment cell viability was measured using the propidium iodide assay. Cells were incubated for 24 hours with 2 µM BAY 11-7082, along with 2µM BAY 11-7082 in conjunction with 150µM 7,8-NP, 150µM 7,8-NP alone and a cell only control. At the 24 hour time point cells were extracted and cell viability measured by PI using flow cytometry under the FL-3 filter. Results show that 2µM BAY 11-7082 has a limited effect on U937 cellular viability after a 24 hour incubation in which viability had non-significantly decreased to 3.19% lower than control (Figure 3.4.8). In contrast with the 150µM 7,8-NP treatment viability was 1.37% lower than control and the two in combination 2.80% lower than control.

Changes in cell morphology after BAY 11-7082 treatment was examined using a light microscope (Leitz Wetzlab, Germany). Changes in cell morphology during treatment with BAY 11-7082 were limited with control cells showing a typical spherical morphology comparable to that seen with all other treatments (Figure 3.4.8).

 

 

 

 

 

 

 

A

B

Control (Bay) 78 (Bay)

150µM 7,8-NP

Control

Bay (Bay) Bay 78 (Bay)

2µM BAY 11-7082

BAY 11-7082 + 7,8-NP

Figure 3.4.8 Effect of BAY 11-7082 on U937 cell viability using propidium iodide

U937 cells (0.5×10cells/mL) were treated with 150µM 7,8-NP, 2 µM of BAY 11-7082 and both in conjunction at 37oC over 24 hours in non-phenol red RPMI-1640. A) Cell viability was measured by PI and the data was expressed as a percentage of the respective control (cell only control). Results are displayed as mean ± SEM of triplicates from a single experiment, representative of three separate experiments. Significance levels are indicated as: (*) p ≤ 0.05, (**) p ≤ 0.01 and (**) p ≤ 0.001. ) Images of U937 cells after treatment with 150µM 7,8-NP, 2µM of BAY 11-7082  and both in conjunction for 24 hours. Cells were viewed in tissue culture plates through an inverted microscope (20x magnification). The images were taken using a LEICA DMIL microscope with a LEICIA DFC290 camera.

3.5 CD36 flow cytometry

3.5.1 CD36 flow cytometry protocol design



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