View Focusing on the “Dark Side” Of

CONTROL OF ANTIOXIDATIVE RESPONSE BY THE TUMOR SUPPRESSOR PROTEIN PML THROUGH REGULATING NRF2 ACTIVITY

SHUANG GUO

Submitted in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

Department of Biochemistry

CASE WESTERN RESERVE UNIVERSITY

January, 2015 CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Shuang Guo

candidate for the degree of Doctor of Philosophy.

Committee Chair

David Samols

Committee Member

Hung-Ying Kao

Committee Member

Yu-Chung Yang

Committee Member

Monica Montano

Committee Member

Shigemi Matsuyama

Date of Defense

November 20, 2014

*We also certify that written approval has been obtained for any proprietary material contained therein. TABLE OF CONTENTS LIST OF TABLES ...... 5 LIST OF FIGURES...... 6 ACKNOWLEDGEMENTS ...... 8 LIST OF ABBREVIATIONS ...... 9 ABSTRACT ...... 14 CHAPTER 1. INTRODUCTION ...... 15 1.1 Oxidative Stress and ROS ...... 15 1.1.1 ROS generation and sources ...... 15 1.1.2 Mitochondrial complex II as a source of ROS ...... 17 1.1.3 Physiological roles of ROS ...... 18 1.1.4 Pathological roles of ROS ...... 20 1.1.5 ROS in the vasculature ...... 22 1.2 Nrf2-mediated Antioxidative Defense System ...... 24 1.2.1 Nrf2 as a critical transcription factor ...... 25 1.2.2 Domain structures of Nrf2 and Keap1 ...... 26 1.2.3 The Keap1-Nrf2-ARE pathway ...... 29 1.2.4 Keap1-independent Nrf2 regulation ...... 32 1.2.5 Cytoprotective functions of Nrf2 downstream targets ...... 33 1.2.6 The role of Nrf2 in cancer ...... 34 1.3 PML: the Tumor Suppressor and Stress Sensor ...... 35 1.3.1 Domain structure and regulation of PML ...... 35 1.3.2 PML nuclear bodies ...... 38 1.3.3 PML as a tumor suppressor ...... 39 1.3.4 PML as a stress sensor ...... 42 1.3.5 Functions of cytoplasmic PML ...... 43 1.3.6 PML in the endothelium ...... 44 1.4 Sulforaphane: the Multi-functional Drug ...... 45 1.4.1 SFN activities mediated by Nrf2 ...... 46

1.4.2 SFN activities mediated by apoptosis-related proteins ...... 50 1.4.3 SFN activities mediated by NF-κB ...... 54 1.4.4 SFN-related clinical trials ...... 56 CHAPTER 2. CONTROL OF ANTIOXIDATIVE RESPONSE BY THE TUMOR SUPPRESSOR PROTEIN PML THROUGH REGULATING NRF2 ACTIVITY ...... 60 2.1 Abstract ...... 60 2.2 Introduction ...... 61 2.3 Results...... 66 2.3.1 Loss of PML increases Nrf2 protein abundance and stability ...... 66 2.3.2 PML inhibits nuclear accumulation and trans-activating capacity of Nrf2 ...... 73 2.3.3 ROS play a role in PML-mediated Nrf2 regulation ...... 84 2.3.4 Sulforaphane (SFN) alters subcellular distribution of PML ...... 90 2.3.5 PML mediates multiple cellular functions of SFN ...... 97 2.4 Discussion ...... 105 2.5 Materials and Methods ...... 111 Cell Culture and Medium ...... 111 Chemicals and Antibodies ...... 111 siRNA Transfection ...... 112 Microarray Analysis ...... 112 Mouse Embryonic Fibroblast and Liver Tissue Isolation ...... 113 Plasmid Construction and Transfection ...... 114 Immunoblotting ...... 114 Co-Immunoprecipitation (Co-IP) ...... 115 Subcellular Fractionation...... 115 Immunofluorescence Microscopy ...... 116 Total RNA extraction, RT-PCR and Quantitative Real-time PCR ...... 117 Luciferase Reporter Assay ...... 117 Chromatin Immunoprecipitation (ChIP) Assay ...... 118 ROS Assay ...... 118

Complex II Enzyme Activity Assay ...... 119 Cell Proliferation Assay ...... 120 Wound-Healing Assay ...... 120 In Vitro Capillary Tube Formation Assay ...... 121 CHAPTER 3. STUDIES OF INTERACTIONS BETWEEN NRF2 AND PML AND POTENTIAL REGULATIONS OF NRF2 ON PML ...... 122 3.1 Abstract ...... 122 3.2 Introduction ...... 123 3.3 Results...... 125 3.3.1 Identification of Nrf2 as a PML-interacting protein ...... 125 3.3.2 Mapping of interacting domains on Nrf2 and PML ...... 130 3.3.3 Regulations of Nrf2 on PML ...... 135 3.4 Discussion ...... 138 3.5 Materials and Methods ...... 140 Plasmid Construction and Transfection ...... 140 GST Pull-down Assay ...... 140 CHAPTER 4. MECHANISTIC INVESTIGATION OF SFN-MEDIATED SUBCELLULAR ALTERATIONS OF PML ...... 141 4.1 Abstract ...... 141 4.2 Introduction ...... 142 4.3 Results...... 145 4.3.1 The effects of SFN on PML isoforms ...... 145 4.3.2 Putative reactive cysteine residues in PML that are important for SFN-mediated PML regulations ...... 148 4.3.3 Microscopic examination of PML localization in response to SFN treatment ...... 153 4.3.4 Crosstalk between SFN-dependent PML regulation and TGF-β signaling ...... 162 4.4 Discussion ...... 166 4.5 Materials and methods ...... 169 Chemicals and Antibodies ...... 169

Plasmid Construction and Transfection ...... 169 Con-focal microscopy ...... 169 CHAPTER 5. DISCUSSION, IMPLICATION AND FUTURE DIRECTIONS . 173 5.1 How PML and Nrf2 coordinate in cancer development? ...... 174 5.2 How does PML function in liver? ...... 175 5.3 Is PML a direct target of SFN? ...... 176 5.4 What is the role of cytoplasmic PML in SFN-mediated multiple cellular functions? ...... 177 5.5 Is TGF-β signaling involved in SFN-mediated multiple cellular functions? ...... 177

BIBLIOGRAPHY ...... 179

LIST OF TABLES

Table 1. Sequences of primers used in PCR and qRT-PCR…………….……171

LIST OF FIGURES

Figure 1-1. Domain structures of Nrf2 and Keap1………………………………28

Figure 1-2. Model of Keap1-Nrf2-ARE signaling pathway……………………..31

Figure 1-3. Domain structure of PML…………………………………………….37

Figure 1-4. PML as a tumor suppressor…………………………………………41

Figure 1-5. SFN is an Nrf2 activator………………………………………...……48

Figure 1-6. Mechanisms for SFN actions…………………………….………….58

Figure 2-1. PML negatively regulates Nrf2 protein abundance and its downstream target genes…………………………………………………………68

Figure 2-2. Characterization of Nrf2 protein species………………………...…71

Figure 2-3. PML inhibits nuclear accumulation of Nrf2…………………………75

Figure 2-4. Subcellular localizations of PML mutants……………………..……78

Figure 2-5. The effects of a nuclear mutant of PML4 overexpression on co-transfected Nrf2 protein abundance in HeLa cells…………………..………79

Figure 2-6. PML antagonizes transactivating activity of Nrf2……………….…81

Figure 2-7. ROS accumulation due to mitochondrial defects accounts for

PML-mediated Nrf2 regulations…………………………………………..………87

Figure 2-8. The effects of SFN on abundance and subcellular distribution of

PML…………………………………………………………………………….……92

Figure 2-9. mRNA level of PML is not affected by SFN treatment……………95

Figure 2-10. Subcellular distribution of PML is regulated by SFN in lung carcinoma cancer cell line A549 cells……………………………………………96

Figure 2-11. SFN-mediated Nrf2 activation is PML-dependent………………98

Figure 2-12. PML is required for SFN-mediated anti-proliferation, anti-migration and anti-angiogenesis activity……………………………..……102

Figure 3-1. Identification of Nrf2 as a PML interacting protein………….……126

Figure 3-2. Mapping of PML and Nrf2 interacting domains……………..……131

Figure 3-3. Regulations of Nrf2 on PML………………………………..………136

Figure 4-1. The effects of SFN on PML isoforms 1, 4, and 6…………………146

Figure 4-2. The effect of SFN on certain cysteine mutants of PML…….……150

Figure 4-3. Partial co-localization of PML with mitochondria, ER, and early

endosome………………………………………………………………………....155

Figure 4-4. Crosstalk between SFN-dependent PML regulation and TGF-β

signaling………………………………………………………...…………………164

ACKNOWLEDGEMENTS

I would like to thank

Dr. Hung-Ying Kao for his mentorship, advice, and encouragement;

Dr. David Samols for his comments on manuscript preparation and thesis writing;

My thesis committee for their advice and suggestions during annual committee meetings;

My current and past colleagues in Kao lab for their contribution and assistance on a daily basis;

My friends for their help, encouragement, and companionship;

My parents Yuwen Guo and Xianglin Zeng for their unconditional support and love.

LIST OF ABBREVIATIONS

β-TrCP, β-transducin repeat-containing protein

AIF, apoptosis inducing factor

A L P, alkaline phosphatase

ALT, alanine transaminase

APL, acute promyelocytic leukemia

AR, androgen receptor

ARE, antioxidant response element

ASK1, apoptosis signal-regulated kinase 1

AST, aspartate transaminase

Bcl-XL, B-cell lymphoma-extra large

BLM, bloom syndrome protein

BTB, bric-a-brac

bZip, basic region-leucine zipper

CBP, CREB-binding protein

Cdc25C, cell division cycle 25C

Cdk, cyclin-dependent kinase

ChIP, chromatin immunoprecipitation

CHK2, checkpoint kinase-2

CHX, cycloheximide

CK2, casein kinase 2

CNC, cap’n’collar

Co-IP, co-immunoprecipitation

CTR, C-terminal regions

DAPI, 4',6-diamidino-2-phenylindole

Daxx, death-associated protein 6

DCFH-DA, 2’,7’-dichlorodihydrofluorescin diacetate

DSBs, double strand breaks

ECs, endothelial cells

EEA1, early endosome antigen 1

EPC, endothelial progenitor cells

ER, endoplasmic reticulum

ERK, signal-regulated kinase

ETC, electron transport chain

FAD, flavin adenine dinucleotide

GPx, glutathione peroxidase

GSH, glutathione

GSK-3β, glycogen synthase kinase 3β

GST, glutathione-S-transferase

HBMECs, human brain microvascular endothelial cells

HDAC, histone deacetylase

HIF1/2-α, hypoxia-inducible factor 1/2 alpha

HO-1, heme oxygenase 1

HSP90, heat shock protein 90

HUVECs, human umbilical vein endothelial cells

IFNs, interferons

IKK, IκB kinase

IL-1β, interleukin-1 beta

iNOS, inducible nitric oxide synthase

I/R, ischemia-reperfusion

IVR, intervening region

JNK, c-Jun N-terminal kinase

Keap1, Kelch-like ECH-associated protein 1

LC3, microtubule-associated protein 1A/1B-light chain 3

LC-MS/MS: liquid chromatography–mass spectrometry/mass spectrometry

LPS, lipopolysaccharide

MAMs, mitochondria-associated membranes

MAPK, mitogen-activated protein kinase

MAPKKK, mitogen-activated protein kinase kinase kinase

MCP-1, monocyte chemoattractant protein-1

Mdm2, mouse double minute 2 homolog

MEFs, mouse embryonic fibroblasts

MKK, mitogen-activated protein kinase kinase

MMP, matrix metalloproteinases

NAC, N-acetyl cysteine

NBs, nuclear bodies

Neh, Nrf2-ECH homology

NES, nuclear export sequence

NF-κB, nuclear factor kappa-light-chain-enhancer of activated B cells

NLS, nuclear localization sequence

Noxs, NADH/NADPH oxidases

NQO1, NAD(P)H dehydrogenase [quinone] 1

Nrf2, nuclear factor erythroid 2–related factor 2

PDI, protein disulfide isomerase

PI3K, phosphoinositide 3-kinase

PKC, protein kinase C

PML, promyelocytic leukemia protein pRB, retinoblastoma protein

Prdx, peroxiredoxins

RARα, retinoic acid receptor α

RING, really interacting new gene

RLR, RIG-I-like receptor

ROS, reactive oxygen species

SARA, Smad anchor for receptor activation

SDH, succinate dehydrogenase

SFN, sulforaphane

SOD, superoxide dismutase

SUMO, small ubiquitin-like modifier

TβRI/II, transforming growth factor β type I/II receptor

TDKI, transforming growth factor β type I receptor kinase domain inhibitor

TGF-β, transforming growth factor β

TGIF, transforming growth interacting factor

TLR, Toll-like receptor

TNFs, tumor necrosis factors

Trx, thioredoxin

UV, ultra violet

VCAM-1, vascular cell adhesion molecule-1

VEGF, vascular endothelial growth factor

VSMCs, vascular smooth muscle cells

Control of antioxidative response by the tumor suppressor protein PML

through regulating Nrf2 activity

ABSTRACT

SHUANG GUO

Oxidative stress is a consequence of an imbalance between reactive oxygen species (ROS) production and the ability of cytoprotective system to detoxify the reactive intermediates. We report that the tumor suppressor promyelocytic leukemia protein (PML) functions as a stress sensor to regulate antioxidative response by controlling nuclear factor erythroid 2-related factor 2 (Nrf2) activity.

Our findings reveal that loss of PML results in ROS accumulation and subsequent activation of Nrf2 antioxidative pathway. We have also identified

PML as an interacting partner of Nrf2 and Nrf2 as a modulator of PML. Of note, we also demonstrate that sulforaphane (SFN), an antioxidant and an Nrf2 activator, regulates the subcellular distribution of PML. We are able to partially dissect the underlying mechanism and show that multiple cellular activities of

SFN are PML-dependent. Taken together, we have established a novel antioxidative mechanism by which PML regulates cellular oxidant homeostasis through controlling Nrf2 activity, and identified PML as an indispensable mediator of SFN activity.

CHAPTER 1. INTRODUCTION

1.1 Oxidative Stress and ROS

Oxidative stress is a consequence of an imbalance between reactive oxygen

species (ROS) production and the ability of cytoprotective systems to detoxify

the reactive intermediates. Oxidative stress is associated with many

pathological conditions, such as atherosclerosis, diabetes, hypertension, and heart failure (Heistad et al., 2009).

1.1.1 ROS generation and sources

Under physiological conditions, oxygen molecules O2 barely react with

non-radical molecules. However, it is easily attacked by a single electron to

.- .- form the superoxide anion, O2 . O2 can be supplied with another electron and

protonated to produce hydrogen peroxide, H2O2. Reactions between H2O2

and reduced metal ions such as Fe2+ give rise to the production of hydroxyl

. .- radicals, OH (Lenaz, 2012). Collectively, molecules including O2 , H2O2, and

OH. are thus defined as reactive oxygen species (ROS) because of their

extremely high reactivity toward biological molecules.

ROS are produced by both exogenous and endogenous sources. Exogenous

sources of ROS include tobacco smoke, fatty acids from diets, ultra violet (UV)

and visible lights, ionizing radiation, heavy and transition metals, xenobiotics,

15 and chlorinated compounds. Endogenous sources of ROS include mitochondrial respiration, xanthine oxidase, nitric oxide (NO) synthases,

NADH/NADPH oxidases (Noxs), cytochrome p450s, lipoxygenase, cyclooxygenase, peroxidases, and inflammatory cells (Cai and Harrison, 2000;

Martin and Barrett, 2002; Seddon et al., 2007; Franco et al., 2008). Among all those sources, mitochondrial respiration is accepted as the major and most important source of endogenous ROS.

Historically, the major sites for mitochondrial ROS generation were believed to be complex I and complex III of the electron transport chain (ETC). However, complex II is now thought to be an equally important site, and we will extensively discuss this in section 1.1.2. Superoxide anions are produced by complex I in vitro when NADH is supplied as a substrate, and this process is accelerated by the presence of the complex I inhibitor rotenone (Takeshige and Minakami, 1979; Turrens and Boveris, 1980). Multiple subunits of complex

I have been suggested as superoxide generators (Kushnareva et al., 2002;

Treberg et al., 2011), however, the exact sites responsible for ROS production are still unclear due to the massive complexity of complex I. Complex III produces superoxide via ubisemiquinone or cytochrome b, which is strongly stimulated by inhibitors antimycin A and myxothiazol (Nohl and Jordan, 1986;

Starkov and Fiskum, 2001; Chen et al., 2003).

In summary, ROS are generated by a series of chemical reactions. Sources of

ROS are either exogenous or endogenous. Among endogenous sources of

ROS, mitochondrial complexes are of great importance.

1.1.2 Mitochondrial complex II as a source of ROS

In addition to complex I and III, complex II is a significant contributor to

mitochondrial ROS generation. Complex II (succinate dehydrogenase, SDH) is

the only mitochondrial complex that participate in both ETC and the citric acid

cycle. It functions to oxidize succinate to fumarate in citric acid cycle and

reduce ubiquinone in ETC. Complex II consists of four subunits that are

encoded by the nuclear genes SDHA, SDHB, SDHC, and SDHD (Rutter et al.,

2010). SDHA is a hydrophilic flavoprotein subunit where the flavin adenine dinucleotide (FAD) cofactor is covalently attached and succinate binds. SDHB is a hydrophilic iron-sulfur protein subunit harboring three iron-sulfur clusters.

SDHC and SDHD are hydrophobic membrane anchor subunits that contain the heme moiety (Sun et al., 2005).

Complex II-dependent electron transport from succinate to ubiquinone is defective in C.elegans carrying a mutation in succinate dehydrogenase cytochrome b. This leads to an accumulation of superoxide, hypersensitivity to oxygen, and premature aging (Ishii et al., 1998). Different mutants of Sdhb in yeast have also been studied and they are associated with ROS production 17

(Smith et al., 2007; Szeto et al., 2007; Goffrini et al., 2009). A transgenic

mouse cell line with a mutated Sdhc gene generates more superoxide anions

and shows high levels of DNA damage and genomic instability. When injected

in nude mice, this cell line also contributes to tumorigenesis (Ishii et al., 2005).

Similar phenotypes have been observed in hamster fibroblasts with an Sdhc

mutation (Slane et al., 2006). SDHB knockdown in cells leads to an elevation

of normoxic ROS generation, ROS-dependent stabilization of

hypoxia-inducible factor 1/2 alpha (HIF1/2-α), and an abnormal cell

proliferation rate (Guzy et al., 2008). Additionally, complex II contributes to

hypoxic ROS accumulation in pulmonary vasculature (Paddenberg et al.,

2003). Direct evidence showing the production of superoxide by complex II

under in vitro conditions was provided in 1998 using isolated bovine heart

mitochondria (Zhang et al., 1998). In recent years, people have found that

complex II generated ROS at the flavin site occurs at a rate that is comparable

to complex I and III (Orr et al., 2012; Quinlan et al., 2012).

1.1.3 Physiological roles of ROS

Since ROS are continuously produced through different sources endogenously,

the physiological roles of ROS can be significant. Basically, ROS affect a wide

range of cellular processes originating from simple chemical reactions. The most common reaction occurs between H2O2 and thiol groups (-SH) on

reactive cysteine residues, leading to the formation of sulphenic acid (-SOH). 18

Sulphenic acid (-SOH) can further react with thiol groups (-SH) to form a

disulfide bond (-SS-) (Finkel, 2012). These post-translational modifications alter the activity of target proteins, and thus indirectly affect essential signaling pathways in diverse physiological systems.

ROS are able to regulate transcription. For instance, mitochondria are redistributed and cluster in perinuclear regions in response to hypoxia. ROS

are then produced to modify promoter regions of HIF-1 target genes including

vascular endothelial growth factor (VEGF), which further enhances the

association of HIF-1 with gene expression (Al-Mehdi et al., 2012). Moreover,

ROS play an important role in the immune system. ROS could eliminate

pathogens directly in a highly oxidative environment, or indirectly by

stimulating signaling downstream of Toll-like receptors (TLRs), RIG-I-like

receptors (RLRs), and inflammatory cytokines (Sena and Chandel, 2012).

Multiple cellular events also require the presence of ROS. One example is

autophagy, which is a lysosomal process for the degradation and turnover of

various cellular components including long-lived proteins and organelles

(Mizushima et al., 2008). During starvation, the phosphoinositide 3-kinase

(PI3K) signaling pathway is activated, which triggers ROS accumulation in

cells. ROS further bind to cysteine protease, HsAtg4, to inactivate its function,

which concomitantly leads to autophagy (Scherz-Shouval et al., 2007).

Furthermore, ROS regulate multiple signaling pathways. For example, once

19 cysteine residues on thioredoxin are oxidized by ROS, thioredoxin associating protein apoptosis signal-regulated kinase 1 (ASK1) becomes activated (Fujino et al., 2007). Acting as an upstream mitogen-activated protein kinase (MAPK) kinase kinase (MAPKKK), ASK1 then regulates c-Jun N-terminal kinase (JNK) and p38 MAPK pathways, causing apoptosis (Ichijo et al., 1997). ROS also inhibit phosphatases that inactivate JNK, by enhancing JNK signaling (Kamata et al., 2005).

Taken together, ROS have been well studied to regulate protein post-translational modifications, transcription, and a series of signaling pathways via basic reactions toward target proteins. By doing so, ROS gain their indispensable part in multiple physiological processes such as pathogen clearance, autophagy, and apoptosis.

1.1.4 Pathological roles of ROS

In living organisms, ROS act as a double-edged sword. Besides the physiological benefits they provide, ROS can cause lots of pathological damages. Many biological molecules are targets of ROS and undergo oxidative modifications, leading to protein oxidation, lipid peroxidation, and

DNA damage (Diplock, 1994).

In the presence of ionizing radiation and metal ions, ROS are able to attack the 20

polypeptide backbone of proteins leading to cleavage of peptide bonds

(Stadtman, 1992). Among all amino acids, methionine and cysteine are the most vulnerable for ROS attack, with methionine forming methionine sulphoxide Met-SO and cysteines forming S-S cross-links (Lenaz, 2012). Lipid peroxidation occurs mainly on polyunsaturated fatty acids that are enriched in subcellular membranes. The major end products are malondialdehyde and

4-hydroxy-2-nonenal (Balazy and Nigam, 2003; Negre-Salvayre et al., 2008).

DNA molecules easily react with hydroxyl radicals to form single- or

double-strand breaks, nucleobase modifications, and cross-links. The most

widely used biomarker to detect oxidative DNA damage is one of the end

products 8-hydroxydeoxy-guanine (8-OH-dG) (Valko et al., 2006).

By oxidizing biomolecules to interfere with their proper functions, ROS are

ultimately linked to various disease conditions such as diabetes, hypertension,

neurodegeneration, atherosclerosis, heart failure, and cancer. Pathogenesis of

diabetes requires communications among different tissues that are affected by

ROS. In pancreatic β cells, ROS inhibit the initial insulin secretion by

suppressing GAPDH activity. ASK1 and JNK signaling are activated by ROS in

liver causing insulin resistance. And glomerular hyperfiltration in kidney is also

dependent on mitochondrial ROS accumulation (Nishikawa and Araki, 2007).

Polyunsaturated fatty acids that are enriched in neuronal membrane lipids are

readily attacked by ROS. Additionally, tight junction protein synthesis and

matrix metalloproteinases (MMP) activation are increased by ROS to “loosen”

the blood-brain barrier. Both of these activators allow neurodegenerating

events to take place (Halliwell, 2006). In the heart, ROS activate a wide variety

of hypertrophy related transcription factors and signaling molecules, stimulate

MMPs, up-regulate cardiac fibroblasts proliferation, and modify proteins

essential for excitation-contraction coupling. All of these events lead to

myocardial remodeling and heart failure (Tsutsui et al., 2011). In terms of

cancer, 8-OH-dG generated by the reaction between DNA and ROS has been

shown to be carcinogenic. And multiple ROS-related signaling pathways are

implicated in tumor progression (Sosa et al., 2013).

In summary, basal level of ROS is required to maintain normal functionality of

cell, however, excess ROS cause damages on biomolecules including proteins

and DNA. Pathological conditions arise as a consequence.

1.1.5 ROS in the vasculature

In the vasculature, ROS can be generated in endothelial cells (ECs), vascular smooth muscle cells (VSMCs), and fibroblasts. Nox family proteins are the major sources of ROS in vessel, whose activity is stimulated by environmental stresses, cytokines, hormones, and vasoactive agents, and are tightly regulated to maintain redox homeostasis. They control basal levels of reactive intermediates which are critical to relay signal transduction and maintain 22

vascular function. However, excessive production of ROS leads to oxidative stress and is correlated with the onset and progression of cardiovascular diseases. Pathological conditions characterized by the imbalance between

ROS production and clearance have become the focus of cardiovascular research during the past decades. One best-characterized process disrupted by oxidative stress is the vasoprotective NO signaling. ROS-mediated

inactivation of NO leads to an increase of endothelial transcytosis, a defect of vascular relaxation, and an alteration of EC fibrinolytic activity (Ryoo et al.,

2006; Manea et al., 2010). Additionally, ROS accumulation leads to activation of key signaling proteins in ECs, causing pathological alterations of endothelium. One such signaling protein, nuclear factor

kappa-light-chain-enhancer of activated B cells (NF-κB) is subject to activation

by mitochondrial derived H2O2 in vessels of aged rats, contributing to a

pro-inflammatory endothelial phenotype (Ungvari et al., 2007). Furthermore,

ROS also oxidizes and activates MMPs, which are responsible for vascular

remodeling (Brown and Griendling, 2009).

In summary, cardiovascular diseases arise when ROS are excessively

produced. Mitochondrial DNA damage and membrane lipid peroxidation

resulting from ROS accumulation leads to dysfunction or apoptosis of ECs and

VSMCs, contributing to the initiation of atherosclerosis. Thereafter,

accumulation of apoptotic VSMCs and macrophages cause atherosclerotic

lesion progression and potential plaque rupture (Madamanchi and Runge,

2007). In patients with hypertension, a correlation between high blood

pressure and vascular ROS elevation has been established. Additionally,

application of antioxidant drugs has been shown to down-regulate blood

pressure in multiple animal models of hypertension (Schulz et al., 2011).

1.2 Nrf2-mediated Antioxidative Defense System

In order to combat ROS accumulation, mammalian cells have developed

multiple antioxidative defense systems to produce various antioxidants to

neutralize or scavenge ROS. Common antioxidants include superoxide

dismutase (SOD), catalase, glutathione (GSH), and peroxiredoxins (Prdx).

SOD functions to convert superoxide into H2O2 and O2, and H2O2 is further

reduced to H2O and O2 by catalase (Mates et al., 1999; Landis and Tower,

2005). Glutathione is able to counteract H2O2 by forming glutathione disulfide

(GSSG) in a reaction catalyzed by glutathione peroxidase (GPx). Glutathione

is also used to detoxify xenobiotics and lipid hydroperoxidases, acting as a

cofactor for glutathione-S-transferase (GST) (Mari et al., 2009). Additionally,

peroxides are majorly removed by Prdx (Rhee et al., 2005). Due to the many

pathways antioxidants utilize in reducing ROS levels, redox regulation of transcription factors that control antioxidant gene expression becomes very

important. One of the most widely studied transcription factors that target

antioxidant genes is nuclear factor erythroid 2–related factor 2 (Nrf2), which 24

will be comprehensively discussed in this section.

1.2.1 Nrf2 as a critical transcription factor

In 1990s, several proteins were identified that activate transcription of β-globin by recognizing the NF-E2/AP-1 motif on its promoter (Moi et al., 1994). Nrf2 is one of such proteins and belongs to the family of cap’n’collar (CNC)-related basic region-leucine zipper (bZip) transcription factors. This family also includes p45-NFE2, Nrf1, Nrf3, Bach1, and Bach2 in mammals (Chan et al.,

1993a, b; Moi et al., 1994; Oyake et al., 1996; Kobayashi et al., 1999). These proteins act as heterodimeric transcription factors by associating with other bZip proteins such as small-Mafs (Igarashi et al., 1994). Interestingly, compared with other CNC family proteins, Nrf2 is highly expressed in tissue contacting the environment for detoxification (Chan et al., 1996). Moreover, the binding sequence of Nrf2 was found to be very similar to the antioxidant response element (ARE) identified within promoter of the NAD(P)H dehydrogenase [quinone] 1 (NQO1) gene, which was the first hint suggesting a role of Nrf2 in the antioxidative response (Venugopal and Jaiswal, 1996).

To analyze how Nrf2 is involved in antioxidative regulation, the gene encoding

Nrf2 was knocked out in mice and numerous animal studies followed. First of all, Nrf2 has been shown to be dispensable for murine embryonic development.

Nrf2-/- mice are able to reach adulthood normally and are fertile (Chan et al., 25

1996). Secondly, induction of phase 2 detoxifying enzymes by a phenolic

antioxidant was largely eliminated in Nrf2-/- mice. In the same study, Nrf2 was first characterized to directly bind to the ARE of mouse Nqo1 gene for

activation after heterodimerization with MafK (Itoh et al., 1997). Thirdly, several

antioxidative or detoxifying genes were identified to be Nrf2 targets, including

heme oxygenase 1 (HO-1) and GSTs (Hayes et al., 2000; Ishii et al., 2000).

Indeed, based on current microarray studies, hundreds of genes are regulated

in an Nrf2-dependent manner (Lee et al., 2003; Kimura et al., 2007).

1.2.2 Domain structures of Nrf2 and Keap1

Comparison of Nrf2 sequences from different species reveals six evolutionarily

conserved domains, named Nrf2-ECH homology (Neh) domains (Figure 1-1A).

The C-terminal Neh3 domain is required for transcriptional activation of Nrf2

through an interaction with a chromo-ATPase/helicase DNA binding protein

(Nioi et al., 2005). The Neh1 domain is highly conserved and comprises the

CNC homology region and bZip region. The latter is able to bind DNA and

heterodimerize with small Maf proteins (Itoh et al., 1999). Neh6 is required for

redox-insensitive degradation of Nrf2 in the nucleus via binding to β-transducin

repeat-containing protein (β-TrCP) (McMahon et al., 2004; Chowdhry et al.,

2013). The major transactivation activity of Nrf2 lies in Neh4 and Neh5. They

cooperatively bind CREB-binding protein (CBP) and induce transcription of

target genes in a synergistic manner (Katoh et al., 2001). The N-terminus 26

Neh2 domain is another highly conserved domain that encodes a bZip region,

and once deleted, Nrf2 activity is markedly elevated, indicating that Neh2

domain is essential for inhibiting Nrf2 activity. In a yeast two-hybrid screen

using Neh2 as bait, an Nrf2 repressor was identified, which is Kelch-like

ECH-associated protein 1 (Keap1) (Itoh et al., 1999).

Functional domains of Keap1 are depicted in Figure 1-1B. The C-terminal

regions (CTR) and six Kelch domains of Keap1 are required for its interaction

with Nrf2, which occur at DLG and ETGE motifs on the Neh2 domain of Nrf2

(Tong et al., 2006). The bric-a-brac (BTB) domain is required for homodimerization and its association with Cullin 3 to form an ubiquitin E3 ligase (Zipper and Mulcahy, 2002). On the other hand, a distinguishing feature of Keap1 is its large number of cysteine residues. Some of the cysteines are highly reactive since their locations are adjacent to basic amino acids, suggesting that Keap1 is a direct sensor for electrophiles and ROS (Itoh et al.,

1999). The intervening region (IVR) of Keap1 is where several important reactive cysteine residues are located.

Figure 1-1. Domain structures of Nrf2 and Keap1. (A) Nrf2 protein consists

of six Neh domains. They are Neh2, Neh4, Neh5, Neh6, Neh1, and Neh3 from

N-terminus to C-terminus. Neh2 is responsible for Keap1 binding, Neh4 and

Neh5 are critical for CBP binding during transactivation, and Neh1 is required

for dimerization with small Maf and DNA binding. (B) Keap1 protein consists of

BTB, IVR, six Kelch-repeats, and CTR from the N-terminus to C-terminus. BTB domain is responsible for Cullin 3 binding, IVR domain is enriched in reactive cysteine residues, and Nrf2 binding requires the Kelch-repeats and CTR domains.

1.2.3 The Keap1-Nrf2-ARE pathway

Keap1 is normally localized in the cytoplasm by associating with the

actin-cytoskeleton (Kang et al., 2004). It has been shown that homodimerization is obligatory for Keap1 to act as an adaptor protein for

protein ubiquitination and degradation. In the case of Nrf2, it recognize two

sites in the Neh2 domain, the high affinity ETGE motif and the low affinity DLG

motif (Tong et al., 2006). Under unstressed conditions, Nrf2 is constantly

sequestered by Keap1 in cytoplasm, and degraded by a poly-ubiquitination

and proteasomal degradation pathway. However, during oxidative stress,

electrophiles or ROS modify reactive cysteines of Keap1, which alters its

conformation. Therefore, poly-ubiquitination and degradation of Nrf2 ceases,

and the stabilized Nrf2 accumulates in the nucleus, where it heterodimerizes

with small Maf proteins and transactivates target gene expression via binding

to AREs in their promoter regions (Taguchi et al., 2011) (Figure 1-2).

A number of reactive cysteine residues of Keap1 have been identified by mass

spectrometry in vitro, with different electrophilic reagents resulting in different

patterns of Keap1 modification (Dinkova-Kostova et al., 2002; Hong et al.,

2005b). Among those reactive cysteines, C151, C273 and C288 have been

well-characterized in vivo. C273 and C288 are required for Keap1-dependent

ubiquitination and proteasomal degradation of Nrf2, and C151 is essential for

oxidative stress-mediated Nrf2 activation (Zhang and Hannink, 2003; Zhang et

29 al., 2004). This model of Nrf2 regulation is supported by cell- and animal-based studies. In one study, nuclear Nrf2 extensively accumulated in

Keap1-/- mouse embryonic fibroblasts (MEFs), resulting in high level of cytoprotective enzymes (Wakabayashi et al., 2003). In another study, an Nrf2 mutant lacking the ETGE motif which no longer interacts with Keap1, showed no response to electrophiles (McMahon et al., 2004).

Figure 1-2. Model of Keap1-Nrf2-ARE signaling pathway.

Under unstressed conditions, Nrf2 is expressed at low steady-state levels and undergoes rapid turnover due to ubiquitination and degradation by proteasome in a Keap1-dependent manner. However, when oxidative or electrophilic stress inactivates Keap1 through conjugating to certain cysteine residues, Nrf2 is then stabilized and nuclear translocation of Nrf2 is promoted. Once residing in the nucleus, Nrf2 is able to recruit small Mafs and the general transcriptional machinery, and activate target genes for cytoprotection through binding to

AREs.

1.2.4 Keap1-independent Nrf2 regulation

Indeed, Nrf2 is subjected to phosphorylation that is mediated by multiple kinases including protein kinase C (PKC), glycogen synthase kinase 3β

(GSK-3β), casein kinase 2 (CK2), JNK, and the tyrosine kinase Fyn (Huang et al., 2002; Salazar et al., 2006; Xu et al., 2006b; Apopa et al., 2008).

PKC-dependent phosphorylation of Nrf2 at Ser 40 facilitates release of Nrf2 from Keap1 and enhances transcriptional activity of Nrf2 (Huang et al., 2002).

Similarly, CK2- and JNK-mediated phosphorylation also potentiates nuclear accumulation and activity of Nrf2 (Xu et al., 2006b; Apopa et al., 2008).

However, not all phosphorylation events are related with Nrf2 activation.

Phosphorylation of Nrf2 mediated by GSK-3β and Fyn lead to nuclear export and degradation of Nrf2 (Jain and Jaiswal, 2006; Salazar et al., 2006).

It has also been shown that p21 is able to directly interact with the DLG motif of

Nrf2 and competes with Keap1 for Nrf2 binding, resulting in Nrf2 stabilization.

This stabilization is required for the cytoprotective activity of p21 (Chen et al.,

2009a). In addition, the polyubiquitin-interacting protein p62 interacts with

Keap1 at the Nrf2 binding site, which competes with Nrf2 and inhibits its degradation. Cytoprotective genes are then induced when p62 accumulates, especially when autophagy-mediated p62 clearance is inhibited (Komatsu et

al., 2010).

1.2.5 Cytoprotective functions of Nrf2 downstream targets

In response to oxidative stress, Nrf2 up-regulates a broad spectrum of genes

that act in a concerted manner to protect cells from pathological damages

caused by ROS. Genes induced by Nrf2 are involved in many survival

pathways, such as antioxidative response, xenobiotic detoxification, and

proteome maintenance. Enzymes of the GSH and thioredoxin (Trx) redox

systems are subjected to regulation by Nrf2. Reduced GSH and Trx are

produced by the coordinated reactions of enzymes in the GSH and Trx

systems, with NADPH providing as the reducing equivalent. Hydroperoxides

and peroxynitrites can further be reduced by passing electrons through the

GSH and Trx pathways ultimately to NADPH (Kensler et al., 2007).

Detoxifying enzymes NQO1 and HO-1 are usually considered as markers for

the activation of Nrf2 signliang. NQO1 is a member of the NAD(P)H dehydrogenase (quinone) family that possesses cytoplasmic 2-electron

reductase activity. This enzyme catalyzes reduction and detoxification of highly

reactive quinones that can cause oxidative stress (Jaiswal et al., 1988). HO-1

is an essential enzyme that protects cells against oxidant-mediated cellular injury and inflammation, and cleaves heme to form biliverdin, which is subsequently converted to radical scavenger bilirubin by biliverdin reductase

(Siow et al., 1999). Bilirubin has been reported to directly inhibit NAD(P)H oxidase by interrupting the assembly and activation of enzyme (Jiang et al.,

2006).

Additionally, a series of proteins in the proteasomal pathway are activated by

Nrf2. When proteins that have been modified by oxidants and electrophiles accumulate in cells, normal cellular function is antagonized and apoptosis is initiated. This altered proteasomal system is related to several neurodegenerative diseases, such as Alzheimer’s disease and Parkinson’s disease (Sherman and Goldberg, 2001).

1.2.6 The role of Nrf2 in cancer

Initially, studies utilizing the Nrf2-/- mouse model for carcinogen challenges reveals a chemopreventive function for Nrf2. For example, a higher incidence of intestinal cancer was observed in Nrf2-/- mice exposed to azoxymethane followed by dextran sulfate (Osburn et al., 2007). Additionally,

7,12-dimethylbenz(a)anthracene-induced skin tumor number was elevated in

Nrf2-/- mice compared with wild type (Xu et al., 2006a). Similarly, benzo[a]pyrene-induced gastric neoplasia was exacerbated in Nrf2-/- mice

(Fahey et al., 2002).

However, there has been an emerging view focusing on the “dark side” of

Nrf2-mediated cytoprotection in recent years. Mutations in Nrf2 that stabilize it have been identified in subsets of human head-and-neck, lung, and 34

gall-bladder cancers (Hayes and McMahon, 2009). Moreover, multiple Keap1

mutants are observed in lung cancer tissue and cell lines, which lose their

ability to inhibit Nrf2 and result in constitutive Nrf2 activation (Singh et al.,

2006). Furthermore, a recent study also suggests that Nrf2-mediated

adaptation to ROS stress is crucial for early stages of tumor development, and

if blocked, may slow down cancer progression (DeNicola et al., 2011).

1.3 PML: the Tumor Suppressor and Stress Sensor

The promyelocytic leukemia protein (PML) was originally identified in acute

promyelocytic leukemia (APL)-related studies. In a large number of APL patients, the PML gene on chromosome 15 fuses with the retinoic acid receptor α (RARα) gene on chromosome 17, which is referred as the t(15;17)

chromosomal translocation of APL (Melnick and Licht, 1999). The fusion protein PML-RARα then acts in an oncogenic manner that correlates with leukemia incidence (Piazza et al., 2001). However, by itself, the PML gene is a tumor suppressor and will be extensively discussed in this section.

1.3.1 Domain structure and regulation of PML

The N-terminus of PML is characterized by the distinctive zinc-finger domain

called really interacting new gene (RING) domain, followed by two additional

zinc-finger domains called B-boxes and an α-helical coiled-coil domain. They

35 are collectively referred to as RBCC domain, which promotes homo-multimerization and formation of macromolecular complexes (Jensen et al., 2001). The C-terminus is less structured and subjected to alternative splicing, resulting in formation of seven different isoforms of PML (Figure 1-3).

Different protein partners are recruited to the different isoforms due to their diverse binding interfaces and functional specificities (Jensen et al., 2001).

Figure 1-3. Domain structure of PML.

There are seven isoforms of PML, which are generated by alternative splicing at the C-terminus. PML1 is the longest isoform. It and PML4 are the most abundant isoforms of PML in humans. Most well-studied functions of PML are localized in the N-terminal RING domain, two B-boxes and the coiled-coil domain. They are referred to as R, B, and CC in the figure.

Several kinases have been observed to phosphorylate PML on Tyr and Ser

residues, including extracellular signal-regulated kinase (ERK), checkpoint kinase-2 (CHK2), and CK2 (Yang et al., 2002; Hayakawa and Privalsky, 2004;

Scaglioni et al., 2006). However, the best-studied post-translational modification on PML is sumoylation. Conjugation of small ubiquitin-like modifier (SUMO) on Lys residues of PML is mediated by SUMO-conjugating enzyme UBC9. SUMO1, SUMO2, and SUMO3 of the SUMO family are all involved in sumoylation of PML (Bernardi and Pandolfi, 2007).

1.3.2 PML nuclear bodies

Under physiological conditions, PML is localized to distinct and punctate

nuclear structures called PML nuclear bodies (NBs). PML NBs are ring-like

protein structures that are tightly bound to the nuclear matrix. They have a

diameter of 0.2-1.0 µm and the number of NBs in a single nucleus varies

depending on cell type and conditions (Dellaire and Bazett-Jones, 2004).

Contact with surrounding chromatin is critical for their integrity and positional

stability (Eskiw et al., 2004). Besides PML, more than 30 proteins either

constitutively or transiently localize to PML NBs, including SUMO-1, Sp100,

Sp140, CBP, bloom syndrome protein (BLM), death-associated protein

6 (Daxx), retinoblastoma protein (pRB), and p53 (Zhong et al., 2000).

PML NBs have been proposed to act as storage depots for protein 38 sequestration, degradation, and post-translational modifications. By regulating

PML NB partners, several physiological functions including DNA-damage response, angiogenesis, cellular senescence, and apoptosis affected

(Bernardi and Pandolfi, 2007). All of these functions are consistent with PML being a tumor suppressor.

1.3.3 PML as a tumor suppressor

Among all the functions of PML identified so far, the pro-apoptotic or tumor suppressive activity of PML is the best characterized. Pml-/- mice are viable, but loss of PML renders the mice sensitive to spontaneous and chemically-induced tumorigenesis, including dimethylbenzanthracene

-induced skin papillomagenesis, lymphomas, fibrohistiocytomas, and

γ-irradiation-induced leukemogenesis (Wang et al., 1998a; Wang et al., 1998b).

Cells derived from Pml-/- mice are protected from diverse apoptotic stimuli such as CD95/Fas, ceramide, UV irradiation, γ-irradiation, interferons (IFNs), tumor necrosis factors (TNFs), and chemotherapeutic drugs (Wang et al., 1998b;

Louria-Hayon et al., 2003; Wu et al., 2003; Bernardi et al., 2004). In contrast, overexpression of PML often results in senescence, cell cycle arrest, and apoptosis (Mu et al., 1997). Utilizing tumor samples from cancer patients, low expression levels of PML have been demonstrated in germ cell tumors, lymphomas, breast carcinomas, lung carcinomas, gastric carcinomas, colon adenocarcinomas, and prostate adenocarcinomas (Koken et al., 1995; 39

Gambacorta et al., 1996; Gurrieri et al., 2004a).

The mechanism by which PML regulates apoptosis primarily relies on the

regulation of its associating partners. This occurs by recruiting or releasing

them for transcriptional or post-translational activity (Figure 1-4). The

best-known partner of PML is the tumor suppressor p53. Full activity of p53

upon oncogene stimulation is dependent on its recruitment to PML NBs and its

acetylation which occurs in the PML NBs (Ferbeyre et al., 2000; Pearson et al.,

2000). Additionally, PML inhibits the ubiquitination and degradation of p53 by mouse double minute 2 homolog (Mdm2), which is achieved via sequestering

Mdm2, or physically disrupting the association of p53 and Mdm2, or by promoting phosphorylation of p53 (Kurki et al., 2003; Louria-Hayon et al., 2003;

Bernardi et al., 2004). Another well-studied partner of PML is Daxx, which is a transcriptional repressor that interacts with sumoylated transcription factors.

Once Daxx is sequestered by PML NBs, the transcriptional repression mediated by Daxx is reduced (Li et al., 2000). Furthermore, sequestration of

Daxx by PML also regulates apoptosis. However, the exact role of Daxx in apoptosis remains unclear since both pro-apoptotic and anti-apoptotic activities of Daxx have been observed by different groups (Torii et al., 1999;

Meinecke et al., 2007).

Figure 1-4. PML as a tumor suppressor.

Endogenously, PML forms discrete structure in the cell nucleus called PML

NBs. They are complexes containing PML that are tightly bound to the nuclear matrix. PML NBs regulate a number of transcription factors by sequestration, degradation, or post-translational modification, which can lead to apoptosis and ultimately cause tumor suppression.

1.3.4 PML as a stress sensor

Instead of being stable, PML NBs are highly dynamic structures in response to various cellular events and stress conditions. During the cell cycle, PML NBs increase in number by a fission mechanism in S phase and undergo reformation during the M/G1 transition (Dellaire et al., 2006a; Dellaire et al.,

2006b). The biochemical composition of PML NBs in mitosis is fundamentally different from PML NBs in interphase. The former become de-sumoylated and lose their association with Daxx and Sp100 (Everett et al., 1999; Dellaire et al.,

2006b). Moreover, PML NBs disperse into numerous small NBs in response to different DNA-damaging agents such as UV irradiation and cisplatin (Seker et al., 2003; Salomoni et al., 2005). Exposure to actinomycin D or doxorubicin causes PML to accumulate around the nucleolus (Bernardi et al., 2004;

Janderova-Rossmeislova et al., 2007).

Bach2 is a member of the CNC family and a partner of small Mafs. It is recruited to PML NBs upon oxidative stress (Tashiro et al., 2004). ROS levels are linked to protein aggregation and misfolding when cells are maintained in culture for prolonged periods (Farout and Friguet, 2006). ROS-dependent aggregation of proteins including PML has been identified in certain cell lines

(Moran et al., 2009). Moreover, PML is regulated by oxidative stress inducers in some cell types. For example, PML is subjected to ubiquitin- and

SUMO-dependent degradation by As2O3 treatment. PML is the target for the

therapeutic effects of As2O3 in APL patients (de The et al., 2012;

Lallemand-Breitenbach et al., 2012). Direct binding of As2O3 to specific cysteines of PML has also been reported and is suggested to play a significant role in As2O3-mediated PML degradation (Jeanne et al., 2010; Zhang et al.,

2010). High doses of H2O2 stabilize PML protein, which in turn mediates

H2O2-induced apoptosis in breast cancer cells (Reineke et al., 2008). In

contrast, PML protein accumulation is down-regulated in cells treated with low

concentrations of H2O2 in human umbilical vein endothelial cells (HUVECs)

(Han et al., 2010).

1.3.5 Functions of cytoplasmic PML

Based on previous and current studies, PML is localized in both the nucleus

and the cytoplasm (Giorgi et al., 2010; Carracedo et al., 2011). Although we

usually associate PML with nuclear functions, in recent years, the activity of

cytoplasmic PML has been discovered. For example, cytoplasmic forms of

PML have been shown to be critical for transforming growth factor β (TGF-β)

signaling. The tumor suppressive activity of TGF-β including growth arrest,

apoptosis, and cellular senescence are largely abolished in Pml-/- cells. This

observation is explained by the association of PML, with the TGF-β receptor,

the TGF-β downstream effector protein Smad, and the Smad anchor for

receptor activation (SARA) in the cytoplasm (Lin et al., 2004). Consistent with

this observation, the antagonizing effects of tumor growth interacting factor 43

(TGIF) on TGF-β signaling is attenuated by translocation of PML from the

nucleus to the cytoplasm, confirming the capacity of cytoplasmic PML in

boosting TGF-β signaling (Faresse et al., 2008). Moreover, cytoplasmic PML is

implicated in playing a part in Ca2+ release from endoplasmic reticulum (ER).

PML is enriched at ER and the mitochondria-associated membranes (MAMs)

that connect ER and mitochondria. Under ER stress condition, this enrichment

of PML is required for Ca2+ release from ER and downstream apoptotic effects

(Giorgi et al., 2010). Therefore, PML in the cytoplasm is suggested to behave

as a tumor suppressor based on the studies mentioned above.

However, earlier studies indicate that cytoplasmic forms of PML could be

oncogenic. High levels of cytoplasmic PML have been reported in

hepatocellular carcinomas by different groups (Terris et al., 1995; Chan et al.,

1998). Two mutants of PML that are associated with cytoplasm accumulated

have been identified in patients suffering from aggressive APL. These mutants

facilitate the translocation of PML from the nucleus to the cytoplasm. This

shuttling inhibits p53 signaling (Gurrieri et al., 2004b; Bellodi et al., 2006). Thus, the elusive property of cytoplasmic PML in cancer will require further investigation.

1.3.6 PML in the endothelium

The level of PML expression varies depending on tissue and cell type. PML is 44

highly expressed in endothelium, epithelia, and tissue macrophages, but

weakly expressed in liver (Flenghi et al., 1995). However, while the role of PML

and its regulation have been intensively studied in cancer cells, its role in ECs

remains largely unknown. Our lab has recently shown that PML is

indispensable for TNF-α- and IFN-α-mediated inhibition of EC migration, in

vitro network formation, and in vivo angiogenesis (Cheng et al., 2012). A

microarray gene expression analysis has also been carried out in our lab by

knocking down PML using two independent siRNAs in HUVECs. A broad

spectrum of genes involved in multiple biological pathways and diseases was identified as being regulated by PML under both basal state and TNF-α-treated conditions (Cheng and Kao, 2012).

1.4 Sulforaphane: the Multi-functional Drug

Sulforaphane (SFN) is a naturally occurring isothiocyanate that was originally

discovered in 1992 by a screen intended to identify chemical inducers of detoxification enzymes. In this system, mouse hepatoma cells were grown in a microtiter plate with wells exposed to a variety of candidate inducers. The potency of the candidates was determined by measuring the activity of quinone reductase in cells. Extracts of vegetables from the crucifer family showed great capacity to induce this detoxification enzyme (Prochaska et al.,

1992). SFN was then isolated and identified in broccoli by the same system as a major inducer of detoxification enzymes. Synthesis of SFN based on its 45

naturally occurring structure was also accomplished (Zhang et al., 1992).

1.4.1 SFN activities mediated by Nrf2

Among cytoprotective genes induced by SFN, the best-studied branch is

Nrf2-regulated downstream target genes. A microarray analysis carried out in

2002 using small intestine derived from Nrf2+/+ and Nrf2-/- mice was the first

study to show that a number of Nrf2 downstream target genes are induced by

SFN treatment (Thimmulappa et al., 2002). Subsequently, high-throughput studies continued to support the notion that the Nrf2-related signaling pathway is regulated by SFN as more clusters of SFN targets participating in cellular defense and cell cycle regulation were identified (Hu et al., 2004; Hu et al.,

2006).

Several cysteines on Keap1 have been identified by mass spectrometry to

associate with SFN with the highest reactivity occurring on C38, C151, C368,

and C489 (Hu et al., 2011). Cell-based studies also suggest that C151 of

Keap1 is essential for SFN-mediated Nrf2 activation (Zhang and Hannink,

2003; Zhang et al., 2004). By conjugating SFN to cysteine residues of Keap1,

the drug was shown to cause dissociation of Nrf2 from Keap1 and subsequent

activation of Nrf2 as a transcription factor (Figure 1-5). On the other hand,

Keap1-indepdent mechanisms have also been established. For example, SFN

has been shown to inhibit the phosphorylation of mitogen-activated protein 46

kinase kinase (MKK) 3/6 and its downstream p38 MAPK isoforms p38γ and p38δ. These isoforms are normally able to facilitate the binding between Nrf2 and Keap1 and reduce Nrf2 nuclear translocation (Keum et al., 2006).

Figure 1-5. SFN is an Nrf2 activator.

SFN is able to conjugate to certain cysteine residues of Keap1. High reactivity has been detected at C38, C151, C368, and C489. Due to the conformational change of Keap1 caused by the modifications, Nrf2 dissociates from Keap1 and translocates to the nucleus. Once residing in the nucleus, Nrf2 is able to recruit small Maf, the general transcriptional machinery, and activate target genes for cytoprotection through binding to AREs in their promoter regions.

Due to the carcinogenic property of cellular toxins, inducers of detoxification

enzymes usually exhibit anti-cancer effects. Thus, earlier studies of SFN were

mostly focused on its anti-cancer property. It has been shown that in several

chemical-induced carcinogenesis models, SFN treatment results in attenuation

of tumorigenesis. For example, in rats exposed to the carcinogen 9,

10-dimethyl-1, 2-benzanthracene, the incidence, weight, and progression of

mammary tumors was blocked when SFN was administered (Zhang et al.,

1994). This observation was supported by an in vitro study using cultured

mammary glands (Gerhauser et al., 1997). Additionally, the incidence of

forestomach tumors in mice induced by benzo[a]pyrene was decreased by

SFN (Fahey et al., 2002). Similar to its role in rodents, SFN was also able to

antagonize the genotoxicity of N-nitrosodimethylamine in Salmonella

typhimurium (Barcelo et al., 1996). Indeed, the Nrf2-mediated classical

detoxifying and antioxidative pathway is partially responsible for

SFN-dependent chemoprevention, due to the established roles of toxins and

oxidants in tumorigenesis. For example, the incidence of

7,12-dimethylbenz(a)anthracene-induced skin cancer was decreased by

topical application of SFN in Nrf2+/+ mice, but not in Nrf2-/- mice (Xu et al.,

2006a).

ROS are critical in mediating ischemia-reperfusion (I/R)-related damages on various organs. Thus, elimination and detoxification of ROS by Nrf2-modulated

antioxidative defense becomes extremely important for organ protection. In a

renal I/R model, SFN treatment was able to protect rats from kidney injury

through inducing detoxifying enzymes in an Nrf2-dependent manner (Yoon et

al., 2008). Similarly, I/R injury of heart was attenuated in Sprague-Dawley rats

received SFN, which was also partially mediated by Nrf2 pathway (Piao et al.,

2010). Preconditioning of rats with SFN treatment prior to heart transplantation

protected the heart from I/R injury (Li et al., 2013). In an intestinal I/R model,

intestinal and liver injury was reduced by SFN due to activation of Nrf2

signaling (Zhao et al., 2010).

1.4.2 SFN activities mediated by apoptosis-related proteins

In addition of the antioxidant pathway, apoptosis-associated proteins are also regulated by SFN. The level of apoptosis regulator Bax is up-regulated by SFN in human leukemia cells, prostate cancer cells, colon cancer cells, HUVECs, and endothelial progenitor cells (EPCs) (Gamet-Payrastre et al., 2000;

Fimognari et al., 2002; Singh et al., 2004a; Asakage et al., 2006; Nishikawa et al., 2009). In addition, caspase-3, -8, and -9 are activated by SFN, and

SFN-mediated apoptosis is blocked if cells are pretreated with inhibitors for caspase-8 and -9 (Pham et al., 2004; Singh et al., 2004a; Nishikawa et al.,

2009). Direct evidence showing the involvement of SFN in classical apoptosis pathways comes from studies in Bax-/- and Bak-/- mice. MEFs isolated from

those strains are protected from SFN-dependent apoptotic events such as 50

cytochrome c release and caspase activation, which ultimately leads to

reduced apoptosis (Choi and Singh, 2005). Moreover, key proteins during cell

cycle are subject to SFN regulation. CHK2 is activated by SFN, which leads to

rapid and sustained phosphorylation of cell division cycle 25C (Cdc25C) and

cell cycle arrest (Singh et al., 2004b). The cyclin-dependent kinase (Cdk)

inhibitor p21 is induced by SFN at the transcriptional level to act against cell

proliferation (Kim et al., 2010). By contrast, p53 expression is not affected by

SFN treatment in prostate cancer cells (Gamet-Payrastre et al., 2000).

Additionally, in immortalized fibroblasts from p53+/+ mice, p53-/- mice, or mice

expressing an inactive mutant form of p53, their sensitivity to SFN-induced apoptosis was the same, suggesting SFN exerts its pro-apoptotic action in a

p53-independent manner (Fimognari et al., 2005).

The role of SFN as an inhibitor for histone deacetylase (HDAC) was first

suggested in 2004 (Myzak et al., 2004). In that study, reduction of HDAC activity and a global increase of histone acetylation was observed when SFN was applied to the human embryonic kidney 293 cells and the human colorectal cancer cell line HCT116. The HDAC inhibitory effect presumably resulted from SFN metabolites SFN-cysteine and SFN-N-acetylcysteine

(Myzak et al., 2004). An in vivo study in mice given SFN in the drinking water showed down-regulation of HDAC activity in colonic mucosa and up-regulation of acetylated histones in this tissue (Myzak et al., 2006a). Furthermore, the

HDAC inhibition was demonstrated to be partially responsible for

SFN-mediated cell cycle arrest and apoptosis in human prostate cancer cells

BPH-1, LnCap, and PC3 (Myzak et al., 2006b). In the context of prostate cancer, SFN was able to alternatively destabilize the androgen receptor (AR) by inhibiting HDAC6-mediated inactivation of the AR chaperone heat shock protein 90 (HSP90) (Gibbs et al., 2009). Interestingly, SFN showed preferences for individual HDACs in terms of inhibition. Several class II HDACs including HDAC5, HDAC7, and HDAC9 are not affected by SFN treatment, whereas HDAC3 and HDAC6 are selectively inhibited by SFN in a time- and dose-dependent manner. This may account for the global effects of SFN on histone acetylation (Rajendran et al., 2011).

Furthermore, apoptosis-related cellular processes are also directly regulated by SFN. First, microtubule dynamics during cell cycle was arrested by SFN in both cancer cells and ECs, but the microtubule mass was unaltered (Jackson et al., 2007; Azarenko et al., 2008). This property of SFN is common to many chemotherapeutic drugs, which result in cell cycle arrest and anti-proliferation.

Second, autophagy is elevated by SFN treatment. Markers of autophagy including a number of autophagosomes in the cytoplasm and incorporation of microtubule-associated protein 1A/1B-light chain 3 (LC3) to autophagosomes are up-regulated by SFN (Herman-Antosiewicz et al., 2006). Interestingly, co-treatment of SFN with the autophagy inhibitors bafilomycin A1 or

3-methyladenine potentiates the pro-apoptotic activity of SFN, indicating a

cytoprotective feature of autophagy in this context (Kanematsu et al., 2010;

Nishikawa et al., 2010a). Third, DNA double strand breaks (DSBs), an initial cause of cell death upon chemical- or ionizing radiation-induced challenges, was accelerated by SFN in HeLa cells (Sekine-Suzuki et al., 2008).

As a consequence, mice carrying tumor transplants were protected by SFN

application. Mice injected with mouse mammary carcinoma cell line F3II,

human prostate cancer cell line PC-3, or human pancreatic carcinoma cell line

PANC-1 developed smaller tumors when treated with SFN (Jackson and

Singletary, 2004b; Pham et al., 2004; Singh et al., 2004a). Additionally, UV

irradiation-initiated skin tumor development was blocked by SFN. When

SFN-containing broccoli extracts was applied topically on the backs of mice

that were intermittently exposed to UV light, the incidence and burden of skin

tumor were significantly reduced (Dinkova-Kostova et al., 2006). Cell-based

studies have also been carried out for more than a decade. For example, SFN

has been shown to dose- or time-dependently promote cell cycle arrest and

apoptosis of immortalized human T lymphocyte line Jurkat, mouse mammary

carcinoma cell line F3II, human prostate cancer cell line PC-3 and LNCap, human colon cancer cell line HT-29, human pancreatic carcinoma cell line MIA

PaCa-2 and PANC-1, human epithelial colorectal adenocarcinoma cell line

Caco-2, and human ovarian cancer cell line SKOV3 (Gamet-Payrastre et al.,

2000; Fimognari et al., 2002; Jackson and Singletary, 2004b; Pham et al.,

2004; Singh et al., 2004a; Jakubikova et al., 2005; Herman-Antosiewicz et al.,

2006; Chaudhuri et al., 2007).

Indeed, the pro-apoptotic and anti-proliferative activity of SFN is reserved in

primary cells as well, such as HUVECs. This notion was confirmed by carrying out proliferation assays, migration assays, and TUNEL assays in HUVECs treated with SFN. Interestingly, the tube forming potential of HUVECs, which

represents in vivo angiogenesis capacity, was also abolished by SFN

(Asakage et al., 2006). Similar observations were made in EPCs and human

brain microvascular endothelial cells (HBMECs) (Annabi et al., 2008;

Nishikawa et al., 2009). Due to the essential role of angiogenesis during cancer progression and metastasis, SFN-related studies in ECs have implications for chemoprevention and chemotherapy.

1.4.3 SFN activities mediated by NF-κB

SFN has been shown to antagonize NF-κB activity in a dose- and

time-dependent manner thereby exerting anti-inflammatory effects. The

phosphorylation and degradation of NF-κB inhibitor IκB-α by IκB kinase (IKK)

is blocked by SFN in both cancer cells and ECs, leading to a reduction in

downstream target gene expression including VEGF, cyclin D1, and B-cell

lymphoma-extra large (Bcl-XL) (Xu et al., 2005; Liu et al., 2008). This activity of 54

SFN is dependent on cellular glutathione levels in ECs (Liu et al., 2008).

Additionally, it has also been reported that SFN is able to decrease transcriptional activity of NF-κB without altering its nuclear translocation and

IκB-α degradation (Heiss et al., 2001; Woo and Kwon, 2007). Consequently,

TNF-α-induced expression of inflammatory genes such as vascular cell adhesion molecule-1 (VCAM-1) and monocyte chemoattractant protein-1

(MCP-1) in ECs is inhibited by SFN (Chen et al., 2009b). It has also been suggested that the inhibitory role of SFN on lipopolysaccharide (LPS)-induced expression of inducible nitric oxide synthase (iNOS) and Cox-2 is mediated through p38, ERK, and JNK (Shan et al., 2009; Brandenburg et al., 2010; Shan et al., 2010; Shibata et al., 2010). UVB-induced inflammation of skin is also alleviated by feeding mice SFN (Shibata et al., 2010). Indeed, anti-inflammatory effects of SFN correlate with its potential in treating several diseases. In a skin tumor mouse model, SFN down-regulated pro-inflammatory cytokines including TNF-α, interleukin-1 beta (IL-1β), IL-6, and suppressed their immune response (Thejass and Kuttan, 2007). Moreover,

TNF-α-mediated adhesion of monocytes to HUVECs, a key step during the initiation of atherosclerosis, is inhibited by SFN. Also, SFN is able to protect the endothelial lining of the aorta and block neointima formation in mice (Kwon et al., 2012; Nallasamy et al., 2014). Furthermore, the insulin secretion activity of rat pancreatic islets, challenged by pro-inflammatory cytokine exposure, is restored by SFN treatment. Finally, diabetic development is antagonized by

SFN in mice treated with the β cell damaging agent streptozotocin (Song et al.,

2009).

1.4.4 SFN-related clinical trials

Clinical trials have been carried out by several groups to evaluate the potential

of SFN as a therapeutic agent in treating various types of diseases. In healthy

people taking broccoli sprout extracts as a source of SFN, no clinic-related adverse effects were observed while evaluating the safety and

pharmacokinetics of SFN (Shapiro et al., 2006). In one study carried out in

Qidong, China, residents who were exposed to carcinogenic environmental

pollutants, were given drinking water infused with broccoli sprouts. Excretion of

carcinogens in urine was reduced in these residents, which may lower the

incidence of cancer development in this population (Kensler et al., 2005). In

another study, when a single oral dose of broccoli sprout extract containing

active SFN were given to healthy women undergoing reduction mammoplasty,

metabolites of SFN were quickly detected in their breast tissue isolated after

surgery (Cornblatt et al., 2007). This study developed a strategy for oral

administration of SFN in human and established the fact that SFN is able to

reach the mammary gland with high efficiency. In healthy human volunteers

with certain spots on skin exposed to UV irradiation, topical application of

SFN-containing broccoli extracts alleviated erythema conditions, indicating a

role of SFN in preventing UV-caused skin damages (Talalay et al., 2007). 56

In summary, through interfering with classical apoptotic signaling, ROS accumulation, the Nrf2-mediated antioxidative pathway, NF-κB-dependent inflammatory signaling, and HDAC activity, SFN is involved in numerous critical cellular processes (Figure 1-6). Due to its ability to confer chemoprevention and other therapeutic effects, SFN is currently a promising agent for treating disease either alone or in combination with other drugs to provide synergistic effects. However, most of the related mechanistic studies are not fully dissected and in need for further investigation. Also, more human trials are encouraged to initiate in order to pave the way for additional clinical applications of SFN.

Figure 1-6. Mechanisms for SFN actions.

SFN is involved in multiple signaling pathways including the

ROS-Nrf2-Keap1-ARE antioxidative axis, apoptotic pathways mediated by caspase activation, HDAC inhibition, and anti-inflammatory signaling mediated by NFκB inhibition. All of these account for therapeutic effects of SFN in treating cancer and non-cancer diseases.

Following this introduction, our observations regarding the role of PML in

regulating Nrf2-mediated signaling pathways and how PML is regulated by

SFN will be presented and discussed in Chapter 2. Studies of physical interactions between PML and Nrf2, and feedback regulations of Nrf2 on PML will be described and discussed in Chapter 3. Mechanistic studies of

SFN-dependent PML regulations will be summerized and discussed in

Chapter 4.

CHAPTER 2. CONTROL OF ANTIOXIDATIVE RESPONSE BY

THE TUMOR SUPPRESSOR PROTEIN PML THROUGH

REGULATING NRF2 ACTIVITY

This chapter has been published in Molecular Biology of the Cell (Guo et al.,

2014).

2.1 Abstract

Oxidative stress is a consequence of an imbalance between reactive oxygen

species (ROS) production and the ability of cytoprotective system to detoxify

the reactive intermediates. The tumor suppressor promyelocytic leukemia

protein (PML) functions as a stress sensor. Loss of PML results in impaired

mitochondrial complex II activity, increased ROS, and subsequent activation of

nuclear factor erythroid 2-related factor 2 (Nrf2) antioxidative pathway. We also

demonstrate that sulforaphane (SFN), an antioxidant, regulates Nrf2 activity

through controlling abundance and subcellular distribution of PML and that

PML is essential for SFN-mediated ROS increase, Nrf2 activation,

anti-proliferation, anti-migration and anti-angiogenesis. Taken together, we have uncovered a novel antioxidative mechanism by which PML regulates cellular oxidant homeostasis through controlling complex II integrity and Nrf2 activity, and identified PML as an indispensable mediator of SFN activity.

2.2 Introduction

Oxidative stress is a consequence of an imbalance between reactive oxygen

species (ROS) production and the ability of the cytoprotective system to

detoxify the reactive intermediates. Basal levels of reactive intermediates are

critical for relaying signal transduction and the maintenance of cellular function

(Sena and Chandel, 2012). However, excessive production of ROS leads to

oxidative stress, and is correlated with the onset and progression of many

diseases such as atherosclerosis, diabetes, neurodegeneration and cancer

(Andersen, 2004; Nishikawa and Araki, 2007; Heistad et al., 2009;

Trachootham et al., 2009). Nuclear factor erythroid 2-related factor 2 (Nrf2) is a

member of the cap’n’collar-related basic region-leucine zipper transcription

factors that plays a pivotal role in the cellular defense system against oxidative

stress (Moi et al., 1994; Itoh et al., 1997). In resting cells, Nrf2 resides in the cytosol through association with Kelch-like ECH-associated protein 1 (Keap1).

Keap1, complexes with Cullin 3 to form a ubiquitin E3 ligase which acts as a substrate adaptor that binds Nrf2 and promotes its ubiquitination-dependent proteasomal degradation (Itoh et al., 1999). Upon ROS exposure, several reactive cysteine residues in Keap1 are covalently modified, resulting in Keap1 inactivation and subsequent stabilization and nuclear translocation of Nrf2

(Zhang and Hannink, 2003). Nuclear Nrf2 heterodimerizes with a small Maf protein and binds to antioxidant response elements (AREs) in the promoter regions of a subset of antioxidative genes that include NAD(P)H

61 dehydrogenase [quinone] 1 (NQO1) (Venugopal and Jaiswal, 1996; Tong et al.,

2006; Yamamoto et al., 2008). Induction of Nrf2 by ROS leads to a marked increase in the expression of target genes that protect the cells from oxidative stress-mediated cytotoxicity (Itoh et al., 1997). In addition to antioxidative enzymes, Nrf2 target genes also include cytoprotective genes involved in several protective systems, such as conjugating/detoxification enzymes, molecular chaperones, transporters, and anti-inflammatory signaling molecules (Kensler et al., 2007).

Sulforaphane (SFN) belongs to the isothiocyanate family that is widely used as an antioxidant supplement and applied in cancer chemoprevention. The precursor of SFN, glucoraphanin, is abundant in cruciferous vegetables with the highest concentration found in broccoli (Zhang et al., 1992). Among known

SFN targets, Keap1 is most extensively studied, which serves as an adaptor protein for a Cullin 3-dependent E3 ubiquitin ligase complex-mediated degradation of substrates such as Nrf2 (Zhang et al., 2004). SFN reacts with the thiol groups on several cysteine residues of Keap1, thereby disrupting its association with Cullin 3 and relieving its inhibitory activity on its target proteins such as Nrf2. Inactivation of Keap1 leads to nuclear translocation of Nrf2 and subsequent induction of its target genes (Thimmulappa et al., 2002; Hong et al., 2005a). Notably, many reports have suggested that SFN can impinge on apoptotic signaling pathways in cancer cells with various origins

(Gamet-Payrastre et al., 2000; Fimognari et al., 2002; Pham et al., 2004; Singh et al., 2004a; Mi et al., 2007; Pledgie-Tracy et al., 2007) and possesses anti-proliferative and anti-angiogenic activity in endothelial cells (ECs)

(Asakage et al., 2006; Nishikawa et al., 2010b).

The promyelocytic leukemia protein (PML) was originally identified as a fusion partner with human retinoic acid receptor alpha due to a chromosomal translocation found in patients with acute promyelocytic leukemia (APL)

(Melnick and Licht, 1999). PML knockout mice are viable, but loss of PML renders the mice sensitive to spontaneous and chemically-induced tumorigenesis (Wang et al., 1998a). In contrast, overexpression of PML often results in senescence, cell cycle arrest and apoptosis (Mu et al., 1997). PML is primarily found in discrete nuclear structures referred to as PML nuclear bodies (NBs) (Stuurman et al., 1992). PML NBs are dynamic structures that are the targets of various extracellular stimuli (Reineke and Kao, 2009;

Lallemand-Breitenbach and de The, 2010). In most cell types, the number and size of PML NBs increase in response to cellular stresses, which has led people to believe that PML NBs are stress-responsive structures (Maul et al.,

1995; Eskiw et al., 2003; Seker et al., 2003; Salomoni et al., 2005).

However, while the role of PML and its regulation have been intensively studied in cancer cells, its role in ECs remains largely unknown. Our lab has

recently shown that PML is required for tumor necrosis factor alpha- and

interferon alpha-mediated inhibition of EC migration and in vitro network

formation (Cheng et al., 2012). In the vasculature, ROS can be generated in

ECs. ROS levels are linked to protein aggregation and misfolding when cells

are maintained in culture for prolonged periods (Farout and Friguet, 2006).

ROS-dependent aggregation of proteins including PML has been identified in

certain cell lines (Moran et al., 2009). Recent studies have indicated that PML

is an oxidative stress sensor and its abundance is regulated by oxidative

stress inducers. For example, PML is subjected to ubiquitin- and small

ubiquitin-like modifier (SUMO)-dependent degradation by As2O3 treatment,

which serves as a mechanism for the therapeutic effects of As2O3 in APL

patients (de The et al., 2012; Lallemand-Breitenbach et al., 2012). Direct

binding of As2O3 to specific cysteines of PML has also been reported and is suggested to play a significant role in As2O3-mediated PML degradation

(Jeanne et al., 2010; Zhang et al., 2010). High doses of hydrogen peroxide

(H2O2) stabilize PML protein, which in turn mediates H2O2-induced apoptosis

in breast cancer cells (Reineke et al., 2008). High doses of H2O2 also induce

nuclear accumulation and deacetylation of PML in HeLa cells (Guan et al., in

press). In contrast, PML protein accumulation is down-regulated in cells

treated with low concentrations of H2O2 in HUVECs (Han et al., 2010). In this study, we report that PML plays a crucial role in regulating abundance, nuclear accumulation and trans-activating capacity of Nrf2 by regulating ROS

64 accumulation. We also identify PML as an integral component of SFN-induced

Nrf2 activation, anti-proliferation, anti-migration and anti-angiogenic activities.

2.3 Results

2.3.1 Loss of PML increases Nrf2 protein abundance and stability

Using an siRNA knockdown approach and microarray gene expression

analysis, we found that the expression of a cluster of genes related to

antioxidative pathways was altered in PML knockdown human umbilical vein

endothelial cells (HUVECs). Many of the antioxidant genes up-regulated in

PML knockdown cells were known downstream targets of Nrf2 (Figure 2-1A).

This observation was further confirmed by immunoblotting of HUVEC extracts

which showed that the protein abundance of Nrf2 and NQO1 were elevated in

PML knockdown cells (Figure 2-1B). We detected multiple bands of Nrf2 which

appear to be distinct isoforms, because the intensity of all the bands detected

between 65 and 120 kDa was significantly decreased by two independent Nrf2

siRNAs (Figure 2-2A). Also, we suggest that the slowest migrating species of

Nrf2 is a phosphorylated form since it disappears in phosphatase-treated

samples (Figure 2-2B). Furthermore, cell-based sumoylation assays indicated

that the slower migrating Nrf2 species was not sumoylated Nrf2 (Figure 2-2, C

and D). Consistent with our observations in HUVECs, increased protein levels

of Nrf2 and its target NQO1 were also observed in Pml-/- mouse embryonic

fibroblasts (MEFs) compared to those in Pml+/+ MEFs (Figure 2-1C). Because of the essential role of liver in antioxidative defense and detoxification, we examined liver isolated from Pml+/+ and Pml-/- mice and found that Pml-/- liver

expressed slightly higher levels of Nrf2 and NQO1 than those in the Pml+/+ liver

(Figure 2-1D). In contrast, Nrf2 and NQO1 were down-regulated in HeLa cells

when PML isoforms 1 and 4 were ectopically overexpressed (Figure 2-1E),

and additive effects on Nrf2 were observed when PML1 and PML4 were

co-expressed (Figure 2-1F). Similarly, exogenously expressed Nrf2 was

down-regulated following PML overexpression (Figure 2-1G), suggesting that

the stability of Nrf2 might be affected by PML abundance. To test this, an Nrf2

expression plasmid was co-transfected with or without a PML4 expression

plasmid and the cells were treated with cycloheximide (CHX), an inhibitor of

protein synthesis. We found that the half-life of Nrf2 was decreased in PML overexpressing cells compared to the control (no PML overexpression) (Figure

2-1H). Collectively, our evidence points to a role for PML as a negative regulator of Nrf2 activity by reducing its protein accumulation and stability.

Figure 2-1. PML negatively regulates Nrf2 protein abundance and its downstream target genes. (A) A heat map of significantly altered genes

(>2-fold, p<0.01) involved in antioxidant pathways identified by microarray

gene expression analysis in PML knockdown HUVECs. siControl represents

control siRNA, whereas siPML-1 and siPML-2 represent two independent PML siRNAs targeting different regions of PML mRNA. Asterisks mark known Nrf2 target genes. (B) The effects of PML knockdown on Nrf2 and NQO1 protein abundance in HUVECs. HUVECs were transfected with a non-targeting siRNA

or two independent PML siRNAs for 72 hrs. Cell extracts were analyzed by

immunoblotting with the indicated antibodies. β-actin was used as a loading

control. Relative intensities of the bands are normalized to both loading control

and siControl. L.E., longer exposure; S.E., shorter exposure. (C) Nrf2 and

NQO1 protein expression in Pml+/+ and Pml-/- mouse embryonic fibroblasts

(MEFs). Relative intensities of the bands are normalized to both loading

control and Pml+/+. (D) Nrf2 and NQO1 protein expression in Pml+/+ and Pml-/-

liver. Liver homogenates were prepared from 3 Pml+/+ and 3 Pml-/- mice.

Relative intensities of the bands are normalized to both loading control and

Pml+/+ #1. (E) The effects of PML1 or PML4 overexpression on endogenous

Nrf2 protein abundance in HeLa cells. HeLa cells were transfected with

plasmids expressing HA-tagged PML1 or PML4. Relative intensities of the

bands are normalized to both loading control and vector control. (F) The

effects of PML1 and PML4 co-expression on endogenous Nrf2 protein

abundance in HeLa cells. HeLa cells were transfected with plasmids

expressing HA-tagged PML1, PML4 or PML1 combined with PML4. (G) The

effects of PML1 or PML4 overexpression on co-transfected Nrf2 protein

abundance in HeLa cells. HeLa cells were transfected with plasmids

expressing FLAG-tagged Nrf2, GFP and different amounts of HA-tagged

PML4. GFP was co-transfected and used as a transfection and loading control.

(H) The effects of PML4 overexpression on the half-life of co-transfected Nrf2 in HeLa cells. HeLa cells were transfected with plasmids expressing

FLAG-tagged Nrf2 and GFP with or without HA-tagged PML4. After 24 hrs, cells were re-plated for CHX treatments. CHX was added to the medium at 20

µg/ml for 0, 15, 30, 60, 90 and 120 minutes. Relative intensities of the bands are normalized to both loading control and 0 min.

Figure 2-2. Characterization of Nrf2 protein species. (A) Immunoblotting

analysis of Nrf2 species in HUVECs. HUVECs were transfected with a

non-targeting siRNA or two Nrf2 targeting siRNAs for 72 hrs. Cell lysates were

analyzed by immunoblotting with the indicated antibodies. β-actin was used as

a loading control. Note that the intensity of all bands was significantly decreased in Nrf2 knockdown cells. (B) Immunoblotting analysis of Nrf2 in

HeLa cells treated with SFN. HeLa cells were treated with SFN for 1 hr at 0 or

40 µM final concentrations. Cell lysates were prepared, treated with λ

phosphatase and analyzed by immunoblotting with the indicated antibodies. (C)

HeLa cells were transfected with HA-tagged Nrf2 or PML (positive control)

together with FLAG-tagged empty vector, SUMO1 or SUMO2. Cell lysates

were prepared and followed by Co-Immunoprecipitation (Co-IP) with

anti-FLAG antibody and immunoblotting with anti-HA and anti-FLAG

antibodies as indicated. Arrow heads, FLAG-SUMO1; arrows, FLAG-SUMO2;

*, IgG; L.E., longer exposure; S.E., shorter exposure. (D) Immunoblotting

analysis of Nrf2 in HeLa cells transfected with SENP1. HeLa cells were

transfected with HA-tagged SENP1. Cell lysates were analyzed by

immunoblotting with the indicated antibodies.

2.3.2 PML inhibits nuclear accumulation and trans-activating capacity of Nrf2

Since the stability of Nrf2 is directly linked to its nuclear accumulation, we

performed subcellular fractionation in Pml+/+ and Pml-/- MEFs and found that

the nuclear fraction contained significantly more Nrf2 in Pml-/- MEFs than in

Pml+/+ MEFs, while the amount in the cytoplasmic fraction was only slightly

altered (Figure 2-3A). Interestingly, the slower migrating Nrf2 species were

predominantly found in the nucleus. Immunofluorescence microscopy also

indicated that Nrf2 accumulated to a greater extent in nuclei of Pml-/- MEFs

when compared to Pml+/+ MEFs (Figure 2-3B). In contrast, nuclear Nrf2 was

minimally detectable when PML was overexpressed in HeLa cells compared to

control cells, while cytoplasmic Nrf2 was also decreased by PML

overexpression as indicated by immunoblotting and immunofluorescence

microscopy (Figure 2-3, C and D). A similar observation was obtained in

HUVECs (Figure 2-3E). We conclude that the nuclear accumulation of Nrf2 is

significantly up-regulated in Pml-/- MEFs, while reduced in cells overexpressing

PML.

To further dissect whether nuclear or cytoplasmic PML affects Nrf2 protein abundance, we employed a PML mutant, K487R, which is constitutively localized in the cytoplasm (Figure 2-4). Overexpression of PML (K487R) had no effect on Nrf2 protein abundance, while restoring localization of this mutant to the nucleus by adding a nuclear localization sequence (NLS) resulted in

similar effects as wild type (Figure 2-3F and 2-4). Exogenous overexpressed

Nrf2 was also subjected to reduction by the nuclear mutant of PML (Figure 2-5).

These data suggest that decreases in nuclear Nrf2 accumulation were

primarily mediated by nuclear forms of PML.

Figure 2-3. PML inhibits nuclear accumulation of Nrf2. (A) Subcellular

fractionation and immunoblotting analysis of Pml+/+ and Pml-/- MEFs. Nuclear

and cytoplasmic fractions prepared from Pml+/+ and Pml-/- MEFs were

subjected to immunoblotting analysis with the indicated antibodies. Lamin B

and α-tubulin were used as loading controls for nuclear and cytoplasmic

fractions respectively. Relative intensities of the bands are normalized to both

loading control and Pml+/+. N, nucleus; C, cytoplasm. (B) Immunofluorescence

analysis of MEFs. Cells were immunostained with anti-PML and anti-Nrf2 antibodies, and images were taken by a fluorescence microscope.

DAPI-stained nuclei (a and d); PML (b and e); endogenous Nrf2 (c and f).

Scale bar, 20 µm. (C) Subcellular fractionation and immunoblotting analysis of

HeLa cells with PML overexpression. HeLa cells were transfected with plasmids expressing HA-tagged PML4. Nuclear and cytoplasmic fractions prepared from transfected HeLa cells were subjected to immunoblotting analysis with the indicated antibodies. Relative intensities of the bands are normalized to both loading control and vector control. N, nucleus; C, cytoplasm.

(D) Immunofluorescence analysis of HeLa cells with PML1 or PML4 overexpression. HeLa cells were transfected with plasmids expressing

HA-tagged PML1 or PML4. Cells were immunostained with anti-HA and anti-Nrf2 antibodies, and images were taken on a fluorescence microscope.

DAPI-stained nuclei (a, d and g); HA-tagged PML (b, e and h); endogenous

Nrf2 (c, f and i). The arrows mark cells expressing transfected PML. Scale bar,

20 µm. (E) Immunofluorescence analysis of HUVECs with PML1 or PML4 overexpression. The experiments were performed as described in (D). (F) The

effects of nuclear and cytoplasmic mutants of PML4 overexpression on

endogenous Nrf2 protein abundance in HeLa cells. HeLa cells were

transfected with plasmids expressing HA-tagged PML4 (wild type), PML4

(K487R) and NLS-PML4 (K487R). Dividing line marks edges of different parts

of the same gel.

Figure 2-4. Subcellular localizations of PML mutants. HeLa cells were transfected with plasmids expressing HA-tagged PML4 (K487R) or NLS-PML4

(K487R) followed by immunostaining with anti-HA antibodies. Cell nuclei were stained with DAPI, and images were taken on a fluorescence microscope.

Figure 2-5. The effects of a nuclear mutant of PML4 overexpression on co-transfected Nrf2 protein abundance in HeLa cells. HeLa cells were transfected with plasmids expressing FLAG-tagged Nrf2, GFP and HA-tagged empty vector, PML4 or NLS-PML4. Cell lysates were analyzed by immunoblotting with the indicated antibodies. GFP was co-transfected and used as a transfection and loading control.

Since nuclear Nrf2 activates its target genes through binding to

ARE-containing promoter regions, we next examined Nrf2 target gene expression. As expected, a battery of Nrf2 target genes which contain AREs in their promoter regions, were up-regulated in Pml-/- MEFs (Figure 2-6A) and liver tissue isolated from Pml-/- mice (Figure 2-6B). We further tested the promoter activity of an ARE-containing luciferase reporter under conditions of

PML overexpression and found the promoter activity was significantly down-regulated by PML in a dose-dependent manner (Figure 2-6C). Using chromatin immunoprecipitation (ChIP) assays, we observed that there was less binding of Nrf2 on the AREs of the NQO1 (Figure 2-6D) and HO-1 promoters (Figure 2-6E) in PML overexpressing HeLa cells. Furthermore, recruitment of Nrf2 to the NQO1-ARE was increased in PML knockdown

HUVECs (Figure 2-6F). These results lead to the conclusion that a loss of PML leads to an accumulation of nuclear Nrf2 and an elevation in the trans-activating capacity of Nrf2 on ARE-driven gene expression.

Figure 2-6. PML antagonizes transactivating activity of Nrf2. (A)

Quantitative real-time PCR (qRT-PCR) analysis of the mRNA levels of Nrf2 target genes and Pml in Pml+/+ and Pml-/- MEFs. Total RNA was prepared from

Pml+/+ and Pml-/- MEFs and reverse transcribed into cDNA, which was used as

a template for qRT-PCR analysis. Values were normalized to the amount of each mRNA in Pml+/+ MEFs. Data are presented as the mean ± s.d. from

triplicates. *p<0.05; **p<0.01; ***p<0.001. (B) Quantitative real-time PCR

(qRT-PCR) analysis of the mRNA levels of Nrf2 target genes and Pml in liver

tissue isolated from Pml+/+ and Pml-/- mice. Similar procedures were performed

as in (A). Data are presented as the mean ± s.d. from triplicates. *p<0.05;

**p<0.01; ***p<0.001. (C) Luciferase reporter assay and analysis of the effects

of PML4 on an ARE-containing reporter. Different amounts of PML4

expression plasmid were transfected into CV-1 cells along with pNQO1-ARE

reporter plasmid. At 24 hrs after transfection, the luciferase activity was

measured in accordance with the instructions provided by the manufacturer.

Data are presented as the mean ± s.d. from triplicates. *p<0.05; ***p<0.001.

Cell extracts were analyzed by immunoblotting with the indicated antibodies.

β-actin was used as a loading control. (D) The effects of PML4 on the

recruitment of Nrf2 to the NQO1 promoter. HeLa cells were transfected with

plasmids expressing HA-tagged PML4 and harvested for ChIP assays using

anti-Nrf2 antibodies or anti-HA antibodies as a control. Percentages of

amplified DNA amounts from precipitates as normalized to 10% of input are

shown. Data are presented as the mean ± s.d. from triplicates. ***p<0.001.

(E) The effects of PML4 on the recruitment of Nrf2 to the HO-1 promoter.

Similar procedures were performed as in (D). ***p<0.001. (F) The effects of

PML knockdown on the recruitment of Nrf2 to the NQO1 promoter. HUVECs

were transfected with a non-targeting siRNA or a PML targeting siRNA for 72

hrs and harvested for ChIP assays using anti-Nrf2 antibodies or anti-HA antibodies as a control. Percentages of amplified DNA amounts from precipitates as normalized to 10% of input are shown. ***p<0.001.

2.3.3 ROS play a role in PML-mediated Nrf2 regulation

Previous studies have demonstrated that nuclear Nrf2 is induced in response

to increases in ROS accumulation (Zhang and Hannink, 2003; Yamamoto et

al., 2008). We hypothesized that loss of PML may impair redox homeostasis, leading to an accumulation of excess ROS. To test this, we measured ROS levels in Pml+/+ and Pml-/- MEFs via a fluorescence-based assay and observed

a two-fold increment of ROS accumulation in Pml-/- MEFs compared to those in

Pml+/+ MEFs (Figure 2-7A). Similarly, ROS levels were up-regulated in PML

knockdown HUVECs (Figure 2-7B). Conversely, ROS production was reduced

by PML overexpression in a dose-dependent manner (Figure 2-7C).

Furthermore, only wild type or nuclear PML (HA-NLS-PML4 (K487R)) were

capable of reducing ROS accumulation (Figure 2-7D). Since the major source

of endogenous ROS is mitochondrial respiration (Raha and Robinson, 2000),

we asked whether mitochondrial respiration was defective in Pml-/- mice.

Mitochondrial complex I, II and III contribute to ROS accumulation

(Paddenberg et al., 2003; Calkins et al., 2005; Ishii et al., 2005; Guzy et al.,

2008). Using liver tissue derived from Pml+/+ and Pml-/- mice, we performed

complex I and complex II enzyme activity assays. We observed little or no

difference in complex I activity between Pml+/+ and Pml-/- liver tissue (Cheng et

al., 2013), but a dramatic reduction of complex II enzymatic activity was

observed in liver derived from Pml-/- mice (Figure 2-7E). In addition, we

detected a global reduction of genes encoding subunits of complex II (Sdha-d)

or factors (Sdhaf1 and Sdhaf2) required for complex II assembly in liver from 84

Pml-/- mice (Figure 2-7F), suggesting that complex II dysfunction observed in

Pml-/- mice is, in part, due to reduced complex II gene expression. We further

examined whether PML-mediated Nrf2 regulation was related to alterations in

ROS. To test this, PML knockdown HUVECs were treated with N-acetyl

cysteine (NAC) to eliminate ROS. We were unable to detect up-regulations of

Nrf2 by PML knockdown in NAC-treated samples (Figure 2-7G), suggesting a

requirement of ROS in PML-mediated regulation on Nrf2. Notably, Nrf2 levels

were down-regulated by NAC treatment alone, likely due to the fact that NAC

chelates ROS, which is required for Nrf2 activation.

The cysteine residues, C212/213, C77/80, and C88/91 of PML, have been

reported to bind As2O3 and play an important role in As2O3-induced

sumoylation and degradation (Jeanne et al., 2010; Zhang et al., 2010). PML

may thus function as a ROS scavenger via these cysteine residues. As such, it

is possible that the PML-mediated ROS decrease may also depend on these

cysteines. To test this, we generated mutants, C212/213A, C77/80A, and

C88/91A, and tested their effects on cellular ROS accumulation and their

ability to decrease Nrf2 protein abundance. Figure 2-7H shows that these

mutants behaved similarly to the wild type protein in reducing ROS

accumulation and Nrf2 protein level. Taken together, we conclude that a loss of

PML results in complex II deficiency and ROS accumulation, which in turn

induces nuclear Nrf2 accumulation and subsequently its target gene

85 expression.

Figure 2-7. ROS accumulation due to mitochondrial defects accounts for

PML-mediated Nrf2 regulations. (A) Reactive oxygen species (ROS)

accumulation in Pml+/+ and Pml-/- MEFs. Equal number of Pml+/+ and Pml-/-

MEFs were cultured in a 96-well plate. The ROS levels were measured

according to manufacturer’s instruction. Data are presented as the mean ±

s.d. from triplicates. **p<0.01. (B) ROS accumulation in PML knockdown

HUVECs. HUVECs were transfected with a non-targeting siRNA or a PML

siRNA for 72 hrs and ROS assays were carried out as described in (A).

*p<0.05. (C) ROS accumulation in HeLa cells overexpressing PML4. Different

amounts of an expression plasmid for PML4 were transfected into HeLa cells

and ROS assays were carried out as described in (A). *p<0.05; ***p<0.001. (D)

The effects of nuclear and cytoplasmic PML4 mutants on ROS accumulation in

HeLa cells. Equal amounts of HA-tagged empty vector, wild type PML,

cytoplasmic and nuclear PML mutants were transfected into HeLa cells and

ROS assays were carried out as described in (A). **p<0.01; ***p<0.001; n.s.,

not significant. (E) Complex II enzyme activity assay of Pml+/+ and Pml-/-

mouse liver. The rate of enzyme activity was determined according to the

manufacturer’s instruction. n=3 per group. ***p<0.001; n, number of mice. (F)

Quantitative real-time PCR (qRT-PCR) analysis of the mRNA levels of genes encoding components of complex II in liver tissue isolated from Pml+/+ and

Pml-/- mice. Fold change on mRNA abundance was normalized to the amount of each mRNA in Pml+/+ tissues. n=3 per group. *p<0.05; **p<0.01; n.s., not

significant; n, number of mice. (G) The effects of N-acetyl cysteine (NAC) on

Nrf2 protein accumulation in PML knockdown HUVECs. HUVECs were

transfected with a non-targeting siRNA or a PML siRNA for 48 hrs. Cells were

then re-plated and treated with NAC for 24 hrs at 0 or 10 mM final

concentration. Cell extracts were prepared and analyzed by immunoblotting

with the indicated antibodies. β-actin was used as a loading control. (H) The

effects of C212/213A, C77/80A, and C88/91A mutations of PML on ROS

accumulation and Nrf2 protein abundance. Equal amounts of HA-tagged

empty vector, wild type PML, and PML mutants were transfected into HeLa cells and split for ROS assays and immunoblotting. *p<0.05; **p<0.01;

***p<0.001; n.s., not significant.

2.3.4 Sulforaphane (SFN) alters subcellular distribution of PML

SFN is known to react with the thiol groups of Keap1 and form thioacyl adducts,

thereby promoting dissociation of Keap1 from Nrf2 and subsequent induction

of ARE-containing genes (Thimmulappa et al., 2002; Hong et al., 2005a). To our surprise, SFN also induced PML protein levels in both a dose- and time-dependent manner in HUVECs (Figure 2-8, A and B). However, this regulation of SFN on PML was not at the transcriptional level (Figure 2-9). We thus wondered whether the nuclear or cytoplasmic distributed PML was being affected by SFN. Unexpectedly, following SFN exposure we found that nuclear

PML was significantly reduced while cytoplasmic PML was concomitantly increased, all of which occurred at early time points accompanied by an induction of Nrf2 in both the nucleus and the cytoplasm (Figure 2-8C). We also observed that slower migrating bands of Nrf2 were more responsive to SFN treatment. Loss of nuclear PML upon SFN treatment was also confirmed by immunofluorescence microscopy. The treatment resulted in a reduction in the number of PML NBs per cell and nuclear fluorescence intensity (Figure 2-8D).

These observations suggest a reverse correlation between Nrf2 and nuclear forms of PML, which is consistent with our previous observations (Figure 2-3).

Our analysis thus far clearly shows that SFN induces an accumulation of cytoplasmic PML while reducing nuclear PML.

Keap1 binds to Nrf2 and promotes its proteasomal degradation (Itoh et al.,

1999). Our observation that PML deficiency induces accumulation of nuclear 90

Nrf2 prompted us to further investigate whether Keap1 regulates PML protein

accumulation. Using siRNA knockdown, we found that depletion of Keap1 had little or no effect on total PML protein abundance (Figure 2-8E). However, an

increase in nuclear PML and a decrease in cytoplasmic PML were observed in

Keap1 knockdown cells (Figure 2-8F). As expected, knockdown of Keap1

resulted in accumulation of nuclear Nrf2. We next addressed whether Keap1

play a role in SFN-mediated regulation of PML. We found that SFN-induced

nuclear reduction and cytoplasmic accumulation of PML were unchanged in

both control and Keap1 knockdown cells (Figure 2-8G). Consistent with this

observation, we found that SFN was capable of inducing total and cytoplasmic

accumulation of PML in lung carcinoma cancer cell line A549 (Figure 2-10),

where Keap1 is mutated, inactive and expressed at an extremely low level

(Singh et al., 2006; Ohta et al., 2008). Taken together, these data indicate that

SFN-mediated regulation of PML is independent of Keap1 activity.

Figure 2-8. The effects of SFN on abundance and subcellular distribution of PML. (A) Immunoblotting analysis of HUVECs treated with different concentrations of sulforaphane (SFN). HUVECs were treated with SFN for 1 hr at 0, 10, 20, 40, or 80 µM final concentrations. Cell extracts were analyzed by immunoblotting with the indicated antibodies. β-actin was used as a loading control. (B) Immunoblotting analysis of HUVECs treated with SFN. HUVECs were treated with vehicle control (DMSO) or SFN at 10 or 20 µM for 0.5, 1, 4, or 8 hrs. Relative intensities of the bands are normalized to both loading control and DMSO within each time point. (C) Subcellular fractionation and immunoblotting analysis of HUVECs treated with SFN. HUVECs were treated with vehicle control (DMSO) or SFN at 10 or 20 µM for 0.5, 1, 4, or 8 hrs.

Nuclear and cytoplasmic fractions prepared were subjected to immunoblotting analysis with the indicated antibodies. Lamin B and α-tubulin were used as loading controls for nuclear and cytoplasmic fractions respectively. Relative intensities of the bands are normalized to both loading control and DMSO within each time point. (D) Immunofluorescence analysis of HUVECs treated with SFN. HUVECs were treated with vehicle control (DMSO) or SFN at 10 µM

for 1 hr. Cells were immunostained with anti-PML antibodies and images were taken on a fluorescence microscope. Nuclei were stained with DAPI and PML

NBs were shown in green. Statistical analysis of the PML NB number and nuclear fluorescence intensity are shown. n>150 per group. *p<0.05; **p <

0.01; n, number of cells. Scale bar, 20 µm. (E) The effects of Keap1

knockdown on PML protein abundance in HUVECs. HUVECs were transfected

with a non-targeting siRNA or two independent Keap1 siRNAs for 72 hrs. Cell extracts were analyzed by immunoblotting with the indicated antibodies. (F)

The effects of Keap1 knockdown on subcellular distribution of PML in HUVECs.

HUVECs were subjected to Keap1 knockdown as described in (E), and followed by subcellular fractionation as described in (C). *, non-specific band.

(G) Immunoblotting analysis of Keap1 knockdown HUVECs treated with SFN.

HUVECs were transfected with a non-targeting siRNA or Keap1 siRNA for 72 hrs. and re-plated, treated with vehicle control (DMSO) or SFN at 10 µM for 1 hr and harvested for subcellular fractionation as described in (C).

Figure 2-9. mRNA level of PML is not affected by SFN treatment.

Quantitative real-time PCR (qRT-PCR) analysis of the mRNA level of the PML gene in DMSO- and SFN-treated HUVECs. Fold change of PML mRNA was normalized to that in vehicle treated cells. Data are presented as the mean ± s.d. from triplicates.

Figure 2-10. Subcellular distribution of PML is regulated by SFN in lung

carcinoma cancer cell line A549 cells. (A) Immunoblotting analysis of A549 cells treated with different concentrations of SFN. HUVECs were treated with

SFN for 1 hr at a final concentration of 0, 10, 20, 40 or 80 µM. Cell lysates were analyzed by immunoblotting with the indicated antibodies. β-actin was used as a loading control. (B) Subcellular fractionation and immunoblotting analysis of

A549 cells treated with SFN. A549 cells were treated with vehicle control

(DMSO) or SFN at 40 µM for 1 hr. Nuclear and cytoplasmic fractions were subjected to immunoblotting analysis with the indicated antibodies. Lamin B and α-tubulin were used as loading controls for nuclear and cytoplasmic fractions respectively. N, nucleus; C, cytoplasm.

2.3.5 PML mediates multiple cellular functions of SFN

We have demonstrated that loss of nuclear PML induces Nrf2 accumulation

(Figure 2-3) through up-regulating ROS levels (Figure 2-7), and that SFN

treatment results in a decrease in PML accumulation in the nucleus (Figure 2-8,

C and D). We thus tested the hypothesis that the SFN-mediated subcellular

alteration of PML accounts for its ability to potentiate Nrf2 activity. Since it has

previously been shown that SFN treatments lead to ROS accumulation (Singh

et al., 2005; Choi et al., 2008), we first tested whether PML is involved in this

regulation by SFN. Using HUVECs transiently transfected with control or PML

siRNA followed by SFN treatment, we were able to show that the

SFN-mediated ROS increase was undetectable following PML knockdown

(Figure 2-11A). This result indicates that SFN-dependent nuclear loss of PML

contributes to ROS production, and that Nrf2 is anticipated to be activated due

to this elevation in ROS. As expected, when PML was knocked down, we

found that the induction of Nrf2 by SFN was largely abolished at the protein

level (Figure 2-11B), and that NQO1 was no longer induced by SFN at the

transcript level (Figure 2-11C). Moreover, SFN-induced recruitment of Nrf2 on the NQO1-ARE was drastically reduced in PML knockdown cells (Figure

2-11D). These data indicate that SFN-induced Nrf2 accumulation is in part, mediated by PML redistribution in a ROS-dependent manner.

Figure 2-11. SFN-mediated Nrf2 activation is PML-dependent. (A) ROS accumulation in PML knockdown HUVECs. HUVECs were transfected with a non-targeting siRNA or a PML siRNA for 72 hrs. Equal number of HUVECs were re-plated in a 96-well plate and treated with vehicle control (DMSO) or

SFN at 40 µM for 1 hr. The ROS level was measured according to the manufacturer’s instruction. Data are presented as the mean ± s.d. from triplicates. *p<0.05; **p < 0.01; n.s., not significant. (B) Immunoblotting analysis of PML knockdown HUVECs treated with SFN. HUVECs were transfected with a non-targeting siRNA or a PML targeting siRNA for 48 hrs.

Cells were then re-plated and treated with vehicle control (DMSO) or SFN at

20 µM for 1 hr. Cell extracts were analyzed by immunoblotting with the indicated antibodies. β-actin was used as a loading control. Dividing line marks edges of different parts of the same gel. (C) Quantitative real-time PCR

(qRT-PCR) analysis of the mRNA levels of NQO1 in PML knockdown HUVECs with SFN treatment. HUVECs were subjected to PML knockdown and SFN treatment as described in (B), and harvested for qRT-PCR. Fold change of

NQO1 mRNA was normalized to that in vehicle treated cells within each group.

Data are presented as the mean ± s.d. from triplicates. **p < 0.01; n.s., not significant. (D) The effects of PML on the recruitment of Nrf2 to the NQO1 promoter following SFN treatment. HUVECs were subjected to PML knockdown and SFN treatment as described in (B), and harvested for ChIP assays using anti-Nrf2 antibodies or anti-HA antibodies as a control.

Percentages of amplified DNA amounts from precipitates as normalized to 10% of input are shown. Data are presented as the mean ± s.d. from triplicates.

***p<0.001.

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We have previously shown that PML is anti-proliferative and anti-angiogenic in

ECs (Cheng et al., 2012). In addition to its ability to activate Nrf2, SFN

possesses anti-proliferation and anti-angiogenesis activity in ECs (Asakage et

al., 2006; Nishikawa et al., 2010b). However, the mechanism underlying these

activities remains largely unknown. The observation that SFN affects the

abundance and subcellular distribution of PML prompted us to investigate

whether PML plays a role in SFN-mediated effects on proliferation and

angiogenesis. To do so, we carried out proliferation assays in control and PML

knockdown HUVECs and observed that the proliferation rate was no longer

decreased by SFN in PML knockdown cells compared to control cells. In fact it

increased (Figure 2-12A). Additionally, in scratch wound healing assays,

HUVECs were less affected by SFN when PML was knocked down (Figure

2-12B). Furthermore, we performed an in vitro capillary tube formation assay

by analyzing the branch points of sprouting vessels, which represent the

angiogenic capacity of ECs. We observed that the number of branch points per

field was unchanged upon SFN treatment when PML was deficient but

down-regulated in cells transfected with a control siRNA (Figure 2-12C). Taken

together, our results unequivocally demonstrate that multiple pharmacological functions of SFN including ROS elevation, Nrf2 activation, anti-proliferation, anti-migration and anti-angiogenesis are all dependent on PML.

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102

Figure 2-12. PML is required for SFN-mediated anti-proliferation, anti-migration and anti-angiogenesis activity. (A) The effects of PML knockdown on proliferation of SFN-treated HUVECs. HUVECs were transfected with a non-targeting siRNA or a PML siRNA for 48 hrs. Cells were then re-plated and treated with vehicle control (DMSO) or SFN at 10 µM for 3 hrs. A cell proliferation assay was performed in accordance with the instructions provided by the manufacturer. Cell numbers were determined at 0,

24, 48, and 72 hrs and normalized to the 0 hr group. Data are presented as the mean ± s.e.m. from triplicates. *p<0.05; **p < 0.01; n.s., not significant. (B)

The effects of PML knockdown on migration of SFN-treated HUVECs.

HUVECs were transfected with a non-targeting siRNA or a PML siRNA for 48 hrs. Cells were then re-plated and treated with vehicle control (DMSO) or SFN at 10 µM for 4 hrs. Wounds were generated using a pipet tip and images were taken at 0 hr and 12 hrs afterwards. Wound widths were measured and shown as percentages of migration. n=6 per group. Data are presented as the mean

± s.d. from triplicates.***p < 0.001; n.s., not significant; n, number of fields. (C)

The effects of PML knockdown on capillary tube formation of SFN-treated

HUVECs. HUVECs were subjected to PML knockdown and SFN treatment as described in (B). In vitro capillary tube formation assays were performed

according to manufacturer’s instructions. Images were taken 21 hrs after

HUVECs seeded on Matrigel. Statistical analysis was performed by counting

branch points per field. n=5 per group. Data are presented as the mean ±

103 s.e.m. from triplicates. **p < 0.01; n.s., not significant; n, number of fields.

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2.4 Discussion

As stress-responsive structures, PML NBs are dynamic and their size and

components are constantly changing depending on the cellular environment

(Eskiw et al., 2003; Dellaire and Bazett-Jones, 2004). These changes in PML levels and the composition of PML NBs are critical for the ability of cells to adapt to environmental cues and maintain cellular homeostasis. Indeed, several studies on the effects of As2O3 on PML and PML NBs have implied

that PML is sensitive to redox disturbance (Jeanne et al., 2010; Zhang et al.,

2010; Lallemand-Breitenbach et al., 2012). In this study, loss of PML resulted

in the inability of cells to control ROS levels and subsequent increases in ROS

(Figure 2-7, A and B). Loss of Nrf2 also results in increased ROS (Frohlich et

al., 2008). Collectively, these observations support a mechanism in which both

PML and Nrf2 are parts of the same pathway critical for ROS homeostasis. An

alternative mechanism was proposed that PML promoted Nrf2 degradation in

PML NBs by recruiting the E3 ubiquitin ligase RNF4 to sumoylated Nrf2

(Malloy et al., 2013). However, based on our observations, Nrf2 is not

subjected to sumoylation (Figure 2-2, C and D), we thus suggest Nrf2 may not

be a RNF4 substrate. Paradoxically, RNF4 is also a PML E3 ubiquitin ligase

(Lallemand-Breitenbach et al., 2008; Tatham et al., 2008; Weisshaar et al.,

2008; Percherancier et al., 2009). If this model is correct, one would anticipate that loss of PML would result in decreased ROS due to increased Nrf2 in Pml-/-

cells. However, our data unequivocally show that loss of PML increases

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cellular ROS levels.

Complex II is the only complex that participates in both citric acid cycle as well

as in mitochondrial oxidative phosphorylation. Although complex II has not generally been considered a major source of mitochondrial ROS, emerging evidence has demonstrated its indispensable role in ROS production

(Paddenberg et al., 2003; Calkins et al., 2005; Ishii et al., 2005; Guzy et al.,

2008). In this study, we have identified PML as a critical regulator of

mitochondrial function, in part, by regulating complex II gene expression and

activity. As PML has been shown to regulate transcription factors, it is likely

that nuclear PML controls the activity of transcription factors that directly or

indirectly modulate promoter activity of complex II genes. The mechanism

underlying PML’s control of complex II gene expression warrants further

investigation.

Mechanisms other than transcriptional dysregulation of genes encoding components of mitochondrial complex II likely contribute to the increased

accumulation of ROS observed in PML deficient cells. It is noteworthy that

As2O3 directly binds to PML on C77/80 and C88/91, which leads to its

enhanced sumoylation and ultimately degradation (Zhang et al., 2010).

Another study reported that C212/213 is also involved in As2O3 binding

(Jeanne et al., 2010). Thus, PML may function as a ROS scavenger via

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reactive cysteine residues. If this hypothesis is correct, one would anticipate

that mutations of these cysteine residues will abolish the ability of PML to

decrease ROS and subsequent Nrf2 accumulation. However, this is clearly not

the case (Figure 2-7H). Additionally, metabolic roles of PML that have recently

been appreciated may link PML to ROS homeostasis in indirect ways. PML

has been shown to regulate PPAR signaling pathways and downstream events

including fatty acid metabolism (Kim et al., 2011; Carracedo et al., 2012; Ito et al., 2012; Cheng et al., 2013). Furthermore, our lab has demonstrated that the energy sensor AMPK, which is also implicated in regulation of cellular ROS, is activated in Pml-/- muscle and liver (Cheng et al., 2013). Since some of the

transcription factors regulated by PML are also involved in ROS regulation, we

cannot rule out the possibility that PML controls cellular ROS levels indirectly

through its regulation of transcription factors or energy sensors.

It has been proposed that SFN induces antioxidative responses by modifying

reactive cysteine residues and subsequent inactivation of Keap1, therefore

facilitating nuclear translocation of Nrf2 and transactivation of its antioxidant

genes (Thimmulappa et al., 2002; Hong et al., 2005a). Indeed, the ability of

SFN and oxidative stress to activate Nrf2 depends on Keap1 Cysteine 151

(C151) (Zhang et al., 2004). As such, one would predict that because SFN and

ROS modify Keap1 C151, treatment of SFN in a ROS-enriched environment

would not induce Nrf2 accumulation. Consistent with this notion, our data

107 indicated that SFN has little or no effect on Nrf2 accumulation in PML knockdown cells, in which elevated ROS was observed.

Interestingly, while knockdown of Keap1 has little effect on total PML protein abundance, it significantly induces nuclear PML accumulation while reducing the cytoplasmic PML abundance. However, Keap1 is not required for effects of

SFN on the subcellular distribution of PML. These results indicate that SFN and knockdown of Keap1 regulate the subcellular distribution of PML through distinct mechanisms. Several studies have reported that NF-κB and histone deacetylases (HDACs) are also SFN targets (Heiss et al., 2001; Myzak et al.,

2004). These data further provide alternative pathways by which SFN promotes its antioxidative activity. Our data show that PML protein accumulation is regulated by SFN in a dose- and time-dependent manner in

HUVECs. At 1 hr, we observed a 2-3 fold increase in PML protein levels without changes in PML mRNA. Interestingly, this induction is accompanied by a significant loss of nuclear PML and an accumulation of cytoplasmic PML.

These effects were observed as early as 0.5 hour after SFN administration.

Further investigation is needed to dissect the mechanism by which SFN regulates PML accumulation and subcellular distribution.

Using an siRNA knockdown approach (Figure 2-2) and subcellular fractionation (Figures 2-3 and 2-8), we demonstrated that in un-stimulated

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HUVECs, a major Nrf2 isoform predominantly localizes in the cytoplasm and migrates faster than the 75 kDa marker. The predicted molecular weight of

Nrf2 is 68 kDa, so we assume that the fastest migrating species is the unmodified Nrf2. In contrast, SFN induced accumulation of nuclear Nrf2, which migrated close to the 100 kDa marker. We speculate that the slower migration

(100 kDa) of the nuclear Nrf2 species is due to a post-translational modification. Indeed, phosphatase treatment resulted in the disappearance of the slowest migrating Nrf2 species (Figure 2-2B), which is consistent with previous reports suggesting that Nrf2 is phosphorylated by PKC, GSK-3β, CK2, and JNK (Huang et al., 2002; Salazar et al., 2006; Xu et al., 2006b; Apopa et al., 2008).

The anti-angiogenesis activity of SFN has been attributed to its ability to inhibit

EC proliferation (Jackson et al., 2007) and VEGF or transcription factors such as HIF-1α and c-Myc (Bertl et al., 2006). We are the first group to establish

SFN-induced cytoplasmic accumulation of PML as an important action in its anti-angiogenic activity. Moreover, as PML NBs possess well-established pro-apoptotic and anti-proliferative properties, recent studies have started to focus on the tumor suppressor activity of cytoplasmic PML. One study has shown that cytoplasmic PML is able to activate Ca2+ release from ER, which contributes to the apoptotic property of PML (Giorgi et al., 2010). Another study suggests that cytoplasmic PML expression is induced by TGF-β, which

109 plays an essential role in TGF-β signaling-mediated growth arrest of cells (Lin et al., 2004). Thus, one should not equate all of the anti-proliferative, anti-angiogenic and pro-apoptotic activity of SFN to decreases in nuclear PML and reduced number of PML NBs. It is possible that SFN cellular activity is mediated by increased cytoplasmic PML. The specific functions of nuclear and cytoplasmic PML isoforms and differentially modified PMLs in SFN-treated cells require further elucidation. Nonetheless, our findings that PML is indispensable for SFN-mediated ROS generation, Nrf2 activation, anti-proliferation, anti-migration, and anti-angiogenesis activity provide a novel and plausible mechanism for the mysterious chemopreventive property of

SFN.

In summary, PML participates in oxidative responses, in part, via controlling the Nrf2-dependent antioxidative pathway. The idea that PML serves as a mediator for multiple SFN actions opens up the significance of its involvement in antioxidative responses. Our study has provided additional insights into the mechanisms regulating ROS homeostasis and implications in therapeutic designs for cancer treatments.

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2.5 Materials and Methods

Cell Culture and Medium

Human umbilical vein endothelial cells (HUVECs, Lonza, C2519A) were grown

in Endothelial Cell Growth Medium-2 (Lonza, CC-4176) supplemented with fetal bovine serum (FBS) and growth factors (Lonza). Cells of less than 5 passages were used in this study. HeLa, CV-1, and A549 cells were maintained in 1X Dulbecco’s modified Eagle’s medium with 4.5 g/liter glucose,

L-glutamine, and sodium pyruvate supplemented with 10% charcoal-stripped

FBS (Sigma, F0926) and 100X Penicillin-Streptomycin Solution (Cellgro,

30-002-CI). Mouse embryonic fibroblasts (MEFs) were grown in the same medium as HeLa and CV-1 cells except supplemented with 55°C heat-inactivated FBS. All cells were maintained in a humidified 37°C, 5% CO2

incubator.

Chemicals and Antibodies

Sulforaphane (SFN, Sigma, S4441) and cycloheximide (CHX, Sigma, 01810)

were dissolved in dimethyl sulfoxide (DMSO, Fisher, D128-1), which was also

used as a vehicle control. All SFN treatments in HUVECs and A549 cells were

performed in basal medium with no supplements. N-Acetyl-L-Cysteine (Sigma,

A8199) was dissolved in double distilled water. The commercial antibodies used in this study were: anti-human PML (Santa Cruz, sc-562), anti-mouse

PML (Chemicon, MAB3738), anti-human PML peptide antibody generated

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against sequence PSTSKAV-S(PO3)-PPHLDGPP (Affinity BioReagents), anti-Nrf2 (Santa Cruz, sc-13032), anti-NQO1 (Abcam, ab28947), anti-Keap1

(Santa Cruz, sc-15246), anti-HA (Santa Cruz, sc-7392; Roche, 12013819001), anti-FLAG (Sigma, F3165), anti-GFP (Santa Cruz, sc-9996), anti-β-actin

(Sigma, A5441), anti-Lamin B (Santa Cruz, sc-6216; sc-365962), anti-α-tubulin

(Sigma, T5168); anti-Mouse IgG conjugated with HRP (Santa Cruz, sc-2005), anti-Rabbit IgG conjugated with HRP (Millipore, 12-348), anti-Goat IgG conjugated with HRP (Santa Cruz, sc-2033); Alexa Fluor 488 μm goat anti-rabbit (Invitrogen, A-11008), Alexa Fluor 488 μm goat anti-mouse

(Invitrogen, A-11001), Alexa Fluor 594 μm goat anti-mouse (Invitrogen,

A-11005).

siRNA Transfection

HUVECs were grown to 60% confluency and then treated with either a non-targeting small interfering RNA (siRNA) oligonucleotide (Dharmacon,

D-001810-01-50) or siRNAs against PML (Dharmacon, J-006547-05 and

J-006547-07), NRF2 (Dharmacon, J-003755-10 and J-003755-11), or KEAP1

(Dharmacon, J-012453-05 and J-012453-06) according to the manufacturer’s protocol using DharmaFECT1 (Thermo Scientific, T-2001).

Microarray Analysis

Procedures for analyzing microarray data have been published elsewhere

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(Cheng et al., 2012). Briefly, HUVECs were transiently transfected with control

siRNA or two different PML siRNAs (siPML-1 and siPML-2) for 72 hrs followed

by total RNA extraction using PrepEase RNA spin kit (USB/Affymetrix, 78766).

The microarray hybridization was carried out by the Genomics Core at

Cleveland Clinic Foundation and HumanRef-8_V2_0_R0_11223162_A chip

(Illumina) was used. Each sample had technical duplicates. All statistical analysis was done in R/Bioconductor. The raw data were transformed by the

Variance-Stabilizing Transformation method and then normalized using the

Robust Spline Normalization package. The significantly changed gene list was

retrieved by a general linear model and empirical Bayes method through the

Linear Models for Microarray Data (limma) package with the false discovery

rate adjusted by Benjamini and Hochberg’s method. The data was represented

in a heat map with the row z-score mapped to a green-black-red color scheme.

Mouse Embryonic Fibroblast and Liver Tissue Isolation

Pml+/+ and Pml-/- mice were maintained in the 129S1/SvImJ background in the

Health Science Animal Facility of Case Western Reserve University. MEFs

were isolated from pregnant female mice 13.5 days post coitum. Genotyping

was performed by PCR using extracted DNA from head tissue of embryos.

MEFs with Pml+/+ and Pml-/- background were used in this study. Liver tissue

+/+ -/- was isolated from Pml and Pml mice sacrificed via euthanasia with CO2.

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Plasmid Construction and Transfection

pCMX-HA-PML1, pCMX-HA-PML4 and pCMX-GFP were previously described

(Reineke et al., 2008). pCMX-HA-PML4-K487R, pCMX-ATG-NLS-HA-PML4-K487R, pCMX-ATG-NLS-HA-PML4, pCMX-HA-PML4-C212/213A, pCMX-HA-PML4-C77/80A, pCMX-HA-PML4-C88/91A and pCMX-FLAG-Nrf2 were generated by PCR and site-directed mutagenesis. All constructs were verified by DNA sequencing.

The primer sequences are listed in Table 1. HeLa, HUVECs and CV-1 cells were transfected with plasmids using Lipofectamine 2000 (Invitrogen, P/N

52887) following the manufacturer’s protocol. Amount of plasmid transfected in each sample was kept constant by adding vector controls.

Immunoblotting

Whole cell extracts were prepared by incubating harvested cells for 30 min in radioimmune precipitation assay buffer (1X PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, and 0.1% SDS) supplemented with protease inhibitors.

Insoluble components were removed by centrifugation at 13,000 rpm for 15 min at 4°C and whole cell extracts were subjected to SDS-PAGE. Proteins were transferred to PVDF membranes (Millipore, IPVH00010) and products were visualized by immunoblotting. Membranes were blocked in 10% nonfat milk in 1X PBS with 0.1% Tween-20 (PBST) for 30 min at room temperature and primary antibodies were added in 5% milk/PBST solution for 2-3 hrs at room temperature or overnight at 4°C. Membranes were then washed with 114

PBST and secondary antibodies were added in 5% milk/PBST solution for 30

min at room temperature. After three washes with PBST, detection was

performed using ECL detection kits (Thermo, 34080; LPS, K-12045-D2).

Intensities of the bands were quantified by ImageJ (v1.46r, NIH).

Co-Immunoprecipitation (Co-IP)

Co-IP was performed according to our published protocol (Su et al., 2013).

Briefly, whole cell extracts were prepared by incubating harvested cells for 30 min in NETN buffer (20 mM Tris-HCl, pH 8.0, 100 mM NaCl, 1 mM EDTA, 10% glycerol, 1 mM dithiothreitol, 0.1% Nonidet P-40) supplemented with protease inhibitors and NEM, followed by sonication and centrifugation.

Immunoprecipitations were carried out using anti-FLAG antibody-conjugated agarose (Sigma, F2426) for overnight at 4°C. Beads were washed with NETN buffer 5 times, and immunoprecipitates were subjected to SDS-PAGE and immunoblotting.

Subcellular Fractionation

Subcellular fractionations of MEFs, HUVECs, HeLa and A549 cells were carried out according to a published protocol (Li et al., 2004). Briefly, harvested cell pellets were resuspended and lysed in cytoplasmic lysis buffer, and incubated for 15 min on ice. Cytoplasmic fractions were acquired by centrifugation at 3,000 rpm for 5 min at 4°C. Nuclear pellets were washed once

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with cytoplasmic lysis buffer followed by centrifugation at 3,000 rpm for 5 min

at 4°C. Nuclear lysis buffer was then added to nuclear pellets and the mixture

was incubated on ice for 30 min with occasional stirring. Nuclear fractions were

acquired by centrifugation at 3,000 rpm for 15 min at 4°C. Both nuclear and

cytoplasmic fractions were subjected to SDS-PAGE and immunoblotting

analysis.

Immunofluorescence Microscopy

Immunofluorescence microscopy was performed as previously described (Gao

et al., 2008). Briefly, MEFs, HUVECs or HeLa cells, plated on glass cover slips in a 12-well plate, were transfected with the indicated plasmids or treated with

SFN with the indicated doses. The cells were fixed in 3.7% paraformaldehyde by incubating for 30 min at room temperature and washed with 1XPBS for three times. Permeabilization was performed by incubating cells with 1XPBS

supplemented with 1% Triton X-100 and 10% goat serum for 10 min. The plate was covered to avoid light since this step. Cells were then washed with 1XPBS for three times and blocked in 1XPBS supplemented with 0.1% Tween-20 and

10% goat serum for 1 hr. Primary antibodies were added to the cells for 2 hrs

followed by three washes with 1XPBS. Secondary antibodies conjugated with

Alexa Fluor were added for 30 min followed by three washes with 1XPBS.

Cover slips were mounted on slides using Vecta shield mounting medium with

4',6-diamidino-2-phenylindole (DAPI) (Vector Laboratories, H-1200) to

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visualize nuclei. Images were captured by a Leica immunofluorescence

microscope (Leica, Wetzlar, Germany) using a 40X lens. Images were

captured by a computer connected with a digital camera at room temperature.

All images were taken under same microscope settings. Intensity of integrated nuclear fluorescence was determined by ImageJ (v1.46r, NIH).

Total RNA extraction, RT-PCR and Quantitative Real-time PCR

Cells or liver tissue was harvested and total RNAs were extracted using

PrepEase RNA spin kit (USB/Affymetrix, 78766) and quantified by A260/A280 spectrometry. Total RNA was used for RT-PCR. The cDNA pool was generated from each RNA sample using iScript Reverse Transcription Supermix kit

(Biorad, 170-8841) with a PCR program of 25°C for 5 min, 42°C for 30 min and

85°C for 5 min. Quantitative real-time PCR (qRT-PCR) was performed using an iCycler (Bio-Rad) platform with 2X iQ SYBR Green Supermix (Bio-Rad,

170-8880). The PCR program is 1 cycle of 95°C for 3 min, 50 cycles of 94°C for 15 sec, 57°C for 20 sec, and 72°C for 30 sec. The relative abundance of an mRNA was normalized to the level of 18S rRNA and presented as the mean ± s.d. from three experiments. The primer sequences are listed in Table 1.

Luciferase Reporter Assay

Luciferase reporter assay was performed according to a published protocol

(Cheng and Kao, 2009). Briefly, CV-1 cells were co-transfected with

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pNQO1-ARE-Luc (gift from Dr. Yamamoto) (Kang et al., 2004) and pCMX-β-gal, with or without pCMX-HA-PML4. Cells were harvested 24 hrs after transfection; luciferase and β-galactosidase activities were measured with kits according to the manufacturer’s protocol (Promega, E1501).

Luciferase activity was normalized to β-gal activity and fold change presented as the mean ± s.d. from three experiments. Unpaired two-tail t-tests were performed to determine significance.

Chromatin Immunoprecipitation (ChIP) Assay

ChIP assay was performed according to our published protocol (Cheng and

Kao, 2009). Briefly, HeLa cells were transfected with an empty vector or HA tagged PML4 for 48 hrs, HUVECs were transfected with control siRNA or siRNA targeting PML for 72 hrs. Anti-Nrf2 antibodies were used for immunoprecipitation with anti-HA antibodies as a control. qRT-PCR was performed to quantify retrieved DNAs and primer sequences are listed in Table

1. The results were calculated from three independent qRT-PCR experiments and presented as the mean ± s.d. of the relative fold enrichment as the percentage of 10% input signal. Unpaired two-tail t-tests were performed to determine significance.

ROS Assay

ROS assay was performed using OxiSelect Intracellular ROS Assay Kit (Cell

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Biolabs, STA-342). Briefly, Pml+/+ and Pml-/- MEFs, PML knockdown HUVECs, or transfected HeLa cells were cultured in a 96-well plate. Cell-permeable fluorogenic probe 2’,7’-Dichlorodihydrofluorescin diacetate (DCFH-DA) was added to wells followed by an 1 hr incubation in a cell incubator. The assay was terminated by adding cell lysis buffer and incubating for 5 minutes.

Fluorescence intensities were quantified by a SpectraMax M2 plate reader at

480 nm/530 nm with 530 nm cutoff. All samples were prepared in triplicate and data are presented as the mean ± s.d. from three experiments. Unpaired two-tail t-tests were performed to determine significance.

Complex II Enzyme Activity Assay

Complex II enzyme activity assay was performed using Complex II Enzyme

Activity Microplate Assay Kit (MitoSciences, MS241). Briefly, liver homogenates with equal amount of protein were prepared with buffers provided in the kit and loaded to the ELISA plate coated with anti-complex II antibody in triplicate. After 2 hours of incubation at room temperature, the plate was washed 2 times; enzyme substrates succinate and ubiquinone were added together with a blue-colored ubiquinol substrate DCPIP and incubated for 30 minutes. Kinetics of enzyme activity was measured for 2 hours at 1 minute intervals by monitoring the O.D. 600 absorbance using a SpectraMax

M2 plate reader.

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Cell Proliferation Assay

Cell proliferation assay was performed using CellTiter 96 AQueous One

Solution Cell Proliferation Assay Kit (Promega, G3580). Briefly, HUVECs were transfected with either control siRNA or PML siRNA for 48 hrs. Prior to the assay, cells were re-plated and pre-treated with SFN for 3 hrs at 0 or 10 µM final concentrations, and seeded to 5 96-well plates in triplicate with equal cell number (1X103 cells/well). After 0, 24, 48, 72, and 96 hrs, cells were treated with reagents provided in the kit and DNA content within each well was measured using a SpectraMax M2 plate reader at A490. Cell numbers were normalized to day 0 and presented as the mean ± s.d. Unpaired two-tail t-tests were performed to determine significance.

Wound-Healing Assay

HUVECs were transfected with either control siRNA or PML siRNA for 48 hrs, and re-plated on a 12-well plate in duplicate with equal cell number (6X104 cells/well). Prior to the wound-healing assay, cells were pre-treated with SFN for 4 hrs at 0 or 10 µM final concentrations. Wounds were then generated in each well using a 200 µl pipet tip. Images were taken at 0 and 12 hrs using a

Leica Wetzlar microscope. Wound widths were measured by ImageJ and presented as percentage of width difference between 0 and 12 hrs divided by width at 0 hr (n=6). Percentages of migration are presented as the mean ± s.d. and unpaired two-tail t-tests were performed to determine significance.

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In Vitro Capillary Tube Formation Assay

Capillary tube formation assays was performed using an In Vitro Angiogenesis

Assay Kit (Millipore, ECM625). Briefly, HUVECs were transfected with control

siRNA or PML siRNA for 48 hrs. Prior to the assay, cells were re-plated and

pre-treated with or without SFN for 4 hrs at 0 or 10 µM final concentration, and

then seeded to a 96-well plate coated with Matrigel with equal cell number

(1X103 cells/well). Images were taken after 3, 7, and 21 hrs in randomly picked fields (n=5) using a Leica Wetzlar microscope. Statistical analysis and images presented were based on branch point counts in images taken at 21 hrs. The branch points presented are the mean ± s.d. and unpaired two-tail t-tests were performed to determine significance.

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CHAPTER 3. STUDIES OF INTERACTIONS BETWEEN NRF2

AND PML AND POTENTIAL REGULATIONS OF NRF2 ON PML

3.1 Abstract

The cellular antioxidant gene induction mediated by Nrf2 is a major defense mechanism for oxidative stress in mammalian cells. PML functions as a tumor suppressor by regulating a number of pro-apoptotic activities in response to cellular stress and DNA damage. Here we report the identification of a physical interaction between PML and Nrf2 and that PML is mutually controlled by Nrf2 through mechanisms yet to be characterized.

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3.2 Introduction

Nrf2 is a member of the CNC-related bZip transcription factors that plays a pivotal role in cellular defense system against oxidative stress (Itoh et al.,

1997). In resting cells, Nrf2 is retained in the cytosol through an association

with Keap1. Keap1, complexes with Cullin 3 to form an ubiquitin E3 ligase and

acts as a substrate adaptor that binds to Nrf2 and promotes its

ubiquitination-dependent proteasomal degradation (Itoh et al., 1999). Upon

ROS stimulation, reactive cysteine residues in Keap1 are modified by

electrophiles, resulting in Keap1 inactivation and subsequent release and

stabilization of Nrf2 (Zhang and Hannink, 2003). Unbound Nrf2 is then able to

translocate into the nucleus, where it heterodimerizes with a small Maf protein

and binds to AREs upstream of the promoters of a set of antioxidant genes.

This Nrf2-dependent ROS detoxification system is present in several cell types

including ECs. Induction of Nrf2 in cultured ECs by ROS or electrophiles

causes a marked increase in antioxidant genes that function to protect the

cells from oxidative stress-mediated cytotoxicity (Itoh et al., 1999). In addition

to antioxidative enzymes, the Nrf2 target genes are involved in other protective

systems, such as conjugating/detoxification enzymes, transporters, and

molecular chaperones (Tong et al., 2006; Kensler et al., 2007; Hayes and

McMahon, 2009).

PML was originally identified as a fusion partner with RARα due to a

123 chromosomal translocation involved in APL (Melnick and Licht, 1999). PML knockout mice are viable, but exhibit increased susceptibility to spontaneous and chemical-induced carcinogenesis (Wang et al., 1998b). Overexpression of

PML results in cell arrest and apoptosis (Mu et al., 1997). PML is primarily localized in discrete nuclear structures referred to as PML nuclear bodies (NBs)

(Stuurman et al., 1992). PML NBs are dynamic structures that are found predominantly in the nucleus, tightly bound to the nuclear matrix and are the targets of various extracellular stimuli including RNA and DNA viruses, ultraviolet radiation, growth factors, and ionizing radiation. PML is highly expressed in endothelium (Flenghi et al., 1995). However, while the role of

PML and its regulation have been intensively studied in cancer cells, its role in

ECs remains largely unknown.

In this study, we demonstrate that Nrf2 and PML physically interact when they are either overexpressed in HeLa cells or under endogenous conditions in

HUVECs. Additionally, we have mapped the interacting domains on Nrf2 and

PML by GST pull-down assays. Furthermore, we have observed that Nrf2 differentially regulates nuclear and cytoplasmic forms of PML. Taken together, these observations imply a role for PML and Nrf2 in maintaining homeostasis of oxidative stress via physical interaction and mutual regulation.

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3.3 Results

3.3.1 Identification of Nrf2 as a PML-interacting protein

To explore the functions of PML in endothelial cells, we knocked down PML in

HUVECs followed by microarray gene expression analysis. A cluster of

antioxidant genes were up-regulated in HUVECs after PML knockdown, while

oxidative stress promoting genes were down-regulated (Chapter 2, Figure 2-1).

Interestingly, many of these antioxidant genes are Nrf2 targets that carry AREs.

This observation prompted us to investigate whether PML is able to regulate

Nrf2 via physical interaction. By performing immunoprecipitations in HeLa cells

co-transfected with HA-PML4 and FLAG-Nrf2, we found that FLAG-tagged

Nrf2 was immunoprecipitated by anti-HA antibody conjugated beads (Figure

3-1A). Additionally, endogenous Nrf2 was co-immunoprecipitated by HA beads in cells transfected with HA-tagged PML (Figure 3-1B). We also detected an

interaction between endogenous PML and endogenous Nrf2 in both HeLa cells

(Figure 3-1C) and HUVECs (Figure 3-1D). This interaction was detected in the nuclear fraction of HUVECs (Figure 3-1E). In MEFs isolated from Pml+/+ mice,

we observed partial co-localization between Nrf2 and PML NBs (Figure 3-1F).

Therefore, we conclude that the interaction between PML and Nrf2 occurs

under both exogenous and endogenous conditions.

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126

127

Figure 3-1. Identification of Nrf2 as a PML interacting protein. (A)

FLAG-tagged Nrf2 and/or HA-tagged PML4 expression plasmids were

expressed in HeLa cells by transient transfection. Cell lysates were prepared

and immunoprecipitated with anti-HA antibodies. The resulting

immunoprecipitates were subjected to SDS-PAGE and analyzed by

immunoblotting with anti-FLAG and anti-HA antibodies. The bands

corresponding to HA-PML4 and FLAG-Nrf2 are indicated. (B) HA-tagged

PML4 expression plasmid was expressed in HeLa cells and cell lysates

prepared followed by immunoprecipitation with anti-HA antibodies. The resulting immunoprecipitates were subjected to SDS–PAGE and analyzed by

immunoblotting with anti-Nrf2 antibodies. The bands corresponding to

HA-PML4 and endogenous Nrf2 are indicated. (C) Cell lysates prepared from

HeLa cells were immunoprecipitated with an anti-PML antibody or an anti-IgG

antibody (negative control) followed by immunoblotting with antibodies against

Nrf2 and PML. The bands corresponding to endogenous Nrf2 and PML are

indicated. (D) Cell lysates prepared from HUVECs were immunoprecipitated

with anti-PML antibody or anti-HA antibody (negative control) followed by

immunoblotting with antibodies against Nrf2 and PML. (E) Similar procedures

were performed as in (D) except using the nuclear fractions of HUVECs. (F)

Pml+/+ MEFs were immunostained with anti-PML and anti-Nrf2 antibodies, and

images were taken by fluorescence microscopy. Nuclei were stained with DAPI,

PML NBs were stained in red, Nrf2 were stained in green, yellow indicates

128 co-localization. The arrows mark PML NBs in which Nrf2 is also present.

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3.3.2 Mapping of interacting domains on Nrf2 and PML

In order to map the interaction domains, we performed GST pull-down assays

using immobilized, purified GST-tagged PML fusion protein and whole cell

extracts prepared from cells transfected with different Nrf2 fragments. All

C-terminal deletions inclusive of the Neh2 domain bind to PML (Figure 3-2A).

By serial deletion of Nrf2 from the N-terminus, we surprisingly found that

mutants with Neh6 domain deletion lost PML interaction (Figure 3-2B). A schematic presentation of our deletion series mutants of Nrf2 is shown in

Figure 3-2C. Mutants showing PML interaction are also indicated. Moreover, we used truncated forms of GST-PML4 for GST pull-down assays under the same experimental condition. The results indicated that the interacting region on PML is found between amino acids 181-350, which contains the second

B-box domain and the coiled-coil domain (Figure 3-2D). Interestingly, a faster migrating band of Nrf2 interacted with GST-PML4 under this experimental condition. A schematic presentation of truncated GST-PML4 constructs is shown in Figure 3-2E. Mutants showing Nrf2 interaction are also indicated.

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131

132

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Figure 3-2. Mapping of PML and Nrf2 interacting domains. (A) GST-PML4

pulls down FLAG-Nrf2 expressed in HeLa cells. Full-length or C-terminal deletions of FLAG-tagged Nrf2 expression plasmids were expressed in HeLa

cells. Cell lysates were prepared for pull-down assays with immobilized GST or

GST-PML4 fusion proteins. The pull-down fractions were analyzed by immunoblotting with anti-FLAG antibodies. Bands of interests are indicated.

CBS, coomassie blue staining. (B) Similar procedures were performed as in

(A), except using N-terminal deletions of FLAG-tagged Nrf2 expression plasmids. (C) A schematic presentation of deletion series mutants of Nrf2 used in (A) and (B) with functional domains marked. (D) Similar procedures were performed as in (A), except using full-length FLAG-Nrf2 and truncation

mutants of GST-PML4. (E) A schematic presentation of the truncation mutants

of GST-PML4 used in (D) with functional domains marked.

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3.3.3 Regulations of Nrf2 on PML

Based on the observations that PML and Nrf2 physically interact (Figure 3-1) and PML is able to regulate Nrf2 (Chapter 2), we hypothesize that PML and

Nrf2 may mutually regulate each other. Interestingly, total PML protein accumulation was reduced when NRF2 was knocked down in HUVECs.

Down-regulation of Nrf2 target proteins NQO1 and SQSTM1 (also referred to as p62) served as controls for efficient knockdown of NRF2 (Figure 3-3A).

Unexpectedly, the intensity and number of PML NBs were up-regulated in

NRF2 knockdown HUVECs (Figure 3-3B). Additionally, the intensity and number of PML NBs were both significantly reduced when FLAG-Nrf2 was overexpressed in HUVECs (Figure 3-3C). Taken together, we conclude that

Nrf2 differentially modulates PML by negatively regulating nuclear PML, while up-regulating total PML accumulation.

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Figure 3-3. Regulations of Nrf2 on PML. (A) HUVECs were transiently

transfected with a non-targeting siRNA or siRNAs targeting two different regions of the NRF2 transcript. Cells were harvested and whole cell extracts prepared for immunoblotting using the indicated antibodies. (B) HUVECs were transiently transfected with a non-targeting siRNA or a siRNA targeting the

NRF2 transcript. Cells were immunostained with anti-PML and anti-Nrf2 antibodies. Nuclei were stained with DAPI, and images were taken on a fluorescence microscope. (C) HUVECs were transfected with plasmids expressing FLAG-Nrf2, followed by immunostaining with anti-FLAG and anti-PML antibodies. Nuclei were stained with DAPI, and images were taken on a fluorescence microscope.

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3.4 Discussion

In Chapter 2, we have established a negative regulation of PML on Nrf2. Thus,

combined with our observations in this chapter, we conclude that PML and

Nrf2 mutually regulate each other. First, when expressed exogenously, PML

and Nrf2 interacted and co-localized in PML NBs. Second, endogenous immunoprecipitation was employed to confirm that endogenous Nrf2 and PML can also interact. Third, GST pull-down assay was used to map their interaction domains. Fourth, total PML protein level was down-regulated in

HUVECs when NRF2 was knocked down. Fifth, number of PML NB was negatively regulated by Nrf2.

We have identified PML as a novel Nrf2 interacting protein and mapped the binding region to the Neh2 and Neh6 domains of Nrf2. Neh2 has been shown to be a redox-sensitive domain for Nrf2 degradation in the cytoplasm in a

Keap1-dependent manner, and Neh6 is defined as a redox-insensitive domain

that is involved in degradation of Nrf2 in the nucleus (Katoh et al., 2001). It

therefore seems plausible that PML plays a role in Nrf2 degradation. In order

to dissect the mechanism by which PML negatively regulates Nrf2, we will

generate Nrf2 mutants with either Neh2 or Neh6 deletions and use these

constructs to test the effects of PML on the stability and activity of Nrf2.

At this point, the mechanism for mutual regulation between PML and Nrf2 is

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unknown. The observed Nrf2-mediated PML regulation may either be directed

through physical interactions or through the downstream target genes of Nrf2.

However, it is known that NQO1 contributes to p53 stability and inhibition of

NQO1 activates an Mdm-2 and ubiquitin-independent proteasomal

degradation pathway of p53 (Asher et al., 2001; Asher et al., 2002).

Additionally, it has long been known that PML can be induced by p53 at the transcriptional level (de Stanchina et al., 2004). Thus, it is premature to suggest that downstream target genes are directly involved in Nrf2-mediated

PML regulation until we dissect the mechanism of these intriguing observations.

In summary, physical interaction and mutual regulation exist between PML and

Nrf2. The idea that PML and Nrf2 may constitute parts of the same pathway which regulates oxidant homeostasis uncovers the potential significance of their involvement in antioxidative responses.

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3.5 Materials and Methods

Plasmid Construction and Transfection

Truncation mutants GST-PML (1-633, 2-180, 181-350, 351-565, 500-633) and

series deletion mutants FLAG-Nrf2 (1-605, 1-386, 1-134, 1-86, 112-605,

336-605, 435-605) were generated by PCR. The primer sequences are listed in Table 1. HeLa cells were transfected with plasmids using Lipofectamine

2000 (Invitrogen, P/N 52887) following the manufacturer’s protocol. The total amount of plasmid transfected in each sample was kept constant by adding empty vector.

GST Pull-down Assay

GST pull-down assays were performed according to our published protocol

(Reineke et al., 2008). For whole-cell lysate preparation, HeLa cells were transfected with an expression plasmid encoding FLAG-Nrf2 (1-605, 1-386,

1-134, 1-86, 112-605, 336-605, 435-605) or (1-605), harvested, and incubated with immobilized GST-PML (1-633) or (1-633, 2-180, 181-350, 351-565,

500-633) for 1 hr at 4°C. After several washes with NETN buffer, retained fractions were subjected to immunoblotting with anti-FLAG antibodies.

Additional materials and methods used in this chapter can be referred to

Chapter 2, section 2.5.

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CHAPTER 4. MECHANISTIC INVESTIGATION OF

SFN-MEDIATED SUBCELLULAR ALTERATIONS OF PML

4.1 Abstract

The tumor suppressor PML is a stress sensor and an essential constituent of

the highly dynamic PML nuclear bodies (NBs). SFN is a family member of

isothiocyanates that functions as both antioxidant and chemopreventive agent.

We have previously shown that nuclear species of PML is reduced by SFN,

whereas cytoplasmic PML is up-regulated by SFN. In this study, we report that

several cysteine residues on PML are essential for this observation and TGF-β

signaling is likely involved in this regulation. We have also observed that

cytoplasmic PML is partially localized to early endosomes.

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4.2 Introduction

The tumor suppressor PML is primarily found in discrete nuclear structures referred to as PML nuclear bodies (NBs) (Stuurman et al., 1992). PML NBs are dynamic structures that are the targets of various extracellular stimuli (Reineke and Kao, 2009). In most cell types, the number and size of PML NBs increases in response to cellular stress, which has led people to believe that PML NBs are stress-responsive structures (Maul et al., 1995; Eskiw et al., 2003; Seker et al., 2003; Salomoni et al., 2005). Of note, PML is highly expressed in endothelium (Flenghi et al., 1995). Our lab has recently shown that PML is indispensable for TNFα- and IFNα-mediated inhibition of EC migration, in vitro network formation, and in vivo angiogenesis (Cheng et al., 2012). Bach2, a member of the CNC family and a partner with small Maf proteins, is recruited around PML NBs after oxidative stress (Tashiro et al., 2004). By sequestering

Bach2, PML NBs are thought to inhibit antioxidative responses. Elevated ROS levels cause protein aggregation and misfolding when cells are maintained under these conditions for prolonged periods (Farout and Friguet, 2006).

ROS-dependent aggregation of proteins, including PML, has been identified in some cell lines (Moran et al., 2009). Moreover, PML abundance is regulated by oxidative stress inducers. For example, As2O3 induces initial rapid PML

accumulation followed by its degradation in Chinese hamster ovary cells

(Lallemand-Breitenbach et al., 2001). Our lab has previously reported that high

doses of H2O2 stabilize PML protein and that PML mediates H2O2-induced

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apoptosis in breast cancer cells (Reineke et al., 2008). In contrast, PML protein

accumulation is down-regulated in cells treated with low concentrations of

H2O2 in HUVECs (Han et al., 2010). Thus, we speculate that PML may be

susceptive to agents that modulate cellular oxidant state, such as SFN.

SFN is an isothiocyanate that is widely used in cancer chemoprevention and

chemotherapy. The precursor of SFN, glucoraphanin, is abundant in broccoli,

cauliflower, cabbage, and kale with the highest concentration found in broccoli

(Zhang et al., 1992). As a chemopreventive agent, SFN functions via inhibiting

Phase 1 enzymes that convert procarcinogens to carcinogens. It also acts by

inducing Phase 2 enzymes that detoxify carcinogens and facilitate their

excretion from the body. Importantly, the induction of Phase 2 detoxifying

genes such as NQO1 after SFN treatment is dependent on Nrf2 (Clarke et al.,

2008). SFN is known to react with the thiol groups of Keap1 and form thionacyl

adducts, thereby promoting dissociation of Nrf2 from Keap1, Nrf2 nuclear

translocation, and subsequent induction of ARE-containing genes (Clarke et al., 2008). Since cellular toxins are tumorigenic, earlier studies of SFN were mostly focused on its anti-cancer property. Multiple chemical-induced carcinogenesis has been shown to be alleviated by SFN treatment in animal models (Zhang et al., 1994; Gerhauser et al., 1997; Fahey et al., 2002). For

example, mice injected with cancer cells from different origins are protected by

SFN treatment from tumorigenesis (Jackson and Singletary, 2004a; Pham et

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al., 2004; Singh et al., 2004a). Furthermore, SFN is known to promote cell cycle arrest and apoptosis in many types of cancer cells (Gamet-Payrastre et

al., 2000; Jackson and Singletary, 2004a; Singh et al., 2004a; Jakubikova et

al., 2005; Herman-Antosiewicz et al., 2006; Chaudhuri et al., 2007).

Of note, the pro-apoptotic and anti-proliferative properties of SFN are

maintained in HUVECs. Angiogenic potential of HUVECs is also consistently

inhibited by SFN (Asakage et al., 2006; Davis et al., 2009; Nishikawa et al.,

2010b). In Chapter 2, we have demonstrated that PML is an integral

component of SFN-induced Nrf2 activation as well as its anti-proliferation,

anti-migration, and anti-angiogenic activities in HUVECs. SFN regulates the

subcellular distribution of PML by increasing its cytoplasmic abundance and

reducing its nuclear accumulation. However, the detailed mechanism of

SFN-dependent subcellular re-distribution of PML and SFN modified cysteines

are still unclear. Here, we report that C212/213, C77/80, and C88/91 at

N-terminus of PML are critical for SFN-induced cytoplasmic accumulation of

PML. Once localized in the cytoplasm, PML is recruited to early endosomes.

Also, a potential crosstalk between the TGF-β signaling pathway and

SFN-mediated subcellular re-distribution of PML will be described.

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4.3 Results

4.3.1 The effects of SFN on PML isoforms

In Chapter 2, we have shown that SFN is able to alter the subcellular

distribution of PML by reducing its nuclear accumulation and up-regulating its

cytoplasmic accumulation. In order to study the underlying mechanism for this

observation, we would like to first determine which part of PML is responsible for this differential regulation of SFN. To test this, we transfected HeLa cells with plasmids carrying PML1, PML4, or PML6, which are the most widely-studied isoforms of PML. Each of them has a common N-terminus and a distinct C-terminus. Interestingly, the protein levels of these isoforms were

generally unchanged in response to SFN treatments (Figure 4-1A). This is in contrast to our observations in HUVECs, in which endogenous total PML was slightly increased by SFN (Chapter 2, Figure 2-8). However, the nuclear and

cytoplasmic distribution of the three PML isoforms tested was indeed regulated

by SFN in HeLa cells (Figure 4-1B). Nuclear PML1, PML4, and PML6 were significantly down-regulated by SFN. On the other hand, all three isoforms were up-regulated by SFN in the cytoplasm, suggesting an involvement of

PML N-terminus in modulating SFN-dependent re-distribution of PML (Figure

4-1B).

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Figure 4-1. The effects of SFN on PML isoforms 1, 4, and 6. (A) HeLa cells

were transiently transfected with plasmids expressing HA-PML1, HA-PML4, or

HA-PML6 for 24 hrs. Cells were trypsinized, equal number of cells re-plated

and treated with DMSO or SFN (10 µM) for 1 hr, harvested, and whole cell

extracts prepared. Immunoblotting analysis was carried out with the indicated antibodies. β-actin was used as loading control. (B) Nuclear and cytoplasmic fractions were prepared after the cells had been treated as in (A).

Immunoblotting analysis was carried out with the indicated antibodies. Lamin B and α-tubulin were used as loading controls for nuclear and cytoplasmic fractions respectively. S.E., short exposure; L.E., long exposure.

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4.3.2 Putative reactive cysteine residues in PML that are important for

SFN-mediated PML regulations

SFN is capable of modifying reactive cysteine residues in proteins thereby altering protein-protein interactions and/or activity. For example, SFN activates

Nrf2 by altering the conformation of Keap1 through modification of cysteine residue C151 to form a thioacyl adduct (Zhang et al., 2004). In our case, the

N-terminus of PML is rich in cysteine residues. It is thus plausible that SFN is able to react with certain cysteines at the N-terminus of PML, which may account for its cytoplasmic accumulation. C212/213, C77/80, and C88/91 are of particular interest since they have been reported to bind As2O3 and play an

important role in As2O3-induced sumoylation and degradation of PML (Jeanne

et al., 2010; Zhang et al., 2010). We postulate that these potential reactive

cysteine residues are able to react with SFN and lead to cytoplasmic PML

accumulation. To test this, we generated double mutants, C212/213A,

C77/80A, and C88/91A, and a series of combination mutants. For combination

mutants C212/213A plus C77/80A, C212/213A plus C88/91A, and C77/80A

plus C88/91A, nuclear accumulation was reduced by SFN to a similar extent

as wild type. However, SFN-mediated cytoplasmic up-regulation of PML was

completely lost in these mutants (Figure 4-2A). Indeed, the triple combination

mutant was not regulated by SFN in the cytoplasm (Figure 4-2B). Additionally,

any single pair of cysteine mutants lost the SFN effects in the cytoplasm

(Figure 4-2C). Taken together, these results indicate that C212/213A, C77/80A,

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and C88/91A in PML are putative SFN targets that are critical for

SFN-dependent accumulation in the cytoplasm.

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151

Figure 4-2. The effect of SFN on PML mutants, C212/213A, C77/80A, and

C88/91A. (A) HeLa cells were transiently transfected with plasmids expressing

HA-PML4, HA-PML4-C212/213A-C77/80A, HA-PML4-C212/213A-C88/91A, or

HA-PML4-C77/80A-C88/91A for 24 hrs. Cells were trypsinized, equal number of cells re-plated and treated with DMSO or SFN (10 µM) for 1 hr, harvested, and nuclear and cytoplasmic fractions prepared. Immunoblotting analysis was carried out with the indicated antibodies. Lamin B and α-tubulin were used as loading controls for nuclear and cytoplasmic fractions respectively. (B) Similar

procedures were performed as in (A) except here cells were transfected with

HA-PML4 and HA-PML4-C212/213A-C77/80A-C88/91A. (C) Similar procedures were performed as in (A) except here cells were transfected with

HA-PML4, HA-PML4-C212/213A, HA-PML4-C77/80A, and

HA-PML4-C88/91A.

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4.3.3 Microscopic examination of PML localization in response to SFN

treatment

To validate our immunoblotting observations, we employed fluorescence microscopy using exogenously expressed GFP-PML1 (Figure 4-3A) and

GFP-PML4 (Figure 4-3B) to show their accumulation in the cytoplasm after

SFN treatment. Previously, we showed that the K487R mutant of PML is

constitutively localized in the cytoplasm (Chapter 2, Figure 2-4). However, we wondered whether cells with these forms of PML were localized in the same cytoplasmic compartment after SFN treatment. To investigate this, we transfected HeLa cells with the HA-PML1-K487R mutant followed by immunofluorescence designed to examine co-localization between PML and different organelles. Interestingly, we observed partial co-localization between

PML and the mitochondrial marker, apoptosis inducing factor (AIF) (Figure

4-3C), the ER marker, protein disulfide isomerase (PDI) (Figure 4-3D), and the early endosome marker, early endosome antigen 1 (EEA1) (Figure 4-3E).

Since co-localization between PML and EEA1 has previously been reported

(Lin et al., 2004; Jul-Larsen et al., 2010), we focused on this co-localization by utilizing confocal microscopy. We observed limited co-localization between

HA-PML-K487R and EEA1 in HeLa cells under basal condition, however, this co-localization was dramatically induced by SFN treatment (Figure 4-3F). Of note, both PML and EEA1 were recruited to the boundaries of cells when they co-localized after SFN exposure. This result was recapitulated in HUVECs with

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endogenous PML examined (Figure 4-3G).

Due to the complexity of the information that we have gathered so far, we decided to examine the response of cells to SFN treatment in real-time by live

imaging. For this purpose, HeLa cells were transfected with GFP-PML1 and

treated with SFN. Images were obtained every 4 minutes during the hour-long experiment. The results show that nuclear PML, as represented by intensity

and number of NBs, was significantly reduced in response to SFN treatment

during the 1-hour session (Figure 4-3H). Cytoplasmic PML was too diffuse to

be detected during this experiment.

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Figure 4-3. Partial co-localization of cytoplasmic PML with mitochondria,

ER, and early endosomes. (A) HeLa cells were transfected with plasmids expressing GFP-PML1 for 24 h, re-plated, and treated with SFN for 1 hr at 0,

10, 20, 40, or 80 µM final concentrations. Nuclei were stained with DAPI and images were captured on a fluorescence microscope. (B) Similar procedures were performed as in (A) except cells were transfected with GFP-PML4. (C)

HeLa cells were transfected with plasmids expressing HA-PML1-K487R for 24 hrs, re-plated, and treated with DMSO or SFN (5 µM) for 1 hr. Immunostaining was carried out using anti-HA and anti-AIF antibodies. Nuclei were stained with

DAPI, and images were captured on a fluorescence microscope. (D) Similar procedures were performed as in (C) except anti-HA and anti-PDI antibodies were used. (E) Similar procedures were performed as in (C) except anti-HA and anti-EEA1 antibodies were used. (F) HeLa cells were transfected with plasmids expressing HA-PML1-K487R for 24 hrs, re-plated, and treated with

DMSO or SFN (20 µM) for 1 h. Immunostaining was carried out using anti-HA and anti-EEA1 antibodies. Nuclei were detected by far-red emitting fluorescent

DNA dye DRAQ5, and images were taken on a con-focal microscope. The images from a single plane and from all planes merged are shown. (G)

HUVECs were treated with DMSO or SFN (20 µM) for 1 h. Immunostaining was carried out using anti-PML and anti-EEA1 antibodies. Nuclei were detected by far-red emitting fluorescent DNA dye DRAQ5, and images were captured on a confocal microscope. The images from a single plane and from

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all planes merged are shown. (H) GFP-PML1 was transiently transfected into

HeLa cells. Live cell imaging microscopy was performed prior to (0’) and

continued for 1 hr of SFN (10 µM) treatment. Images were obtained every 4 minutes. DAPI stained nuclei. Note that these images did not show cytoplasmic GFP-PML1.

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4.3.4 Crosstalk between SFN-dependent PML regulation and TGF-β

signaling

Canonical TGF-β signaling involves a cascade of events that include binding of

TGF-β to type II receptors (TβRII), stabilization of TβRI and TβRII

heterodimers, phosphorylation of TβRI by TβRII, phosphorylation of Smad2/3

by TβRI, nuclear translocation of phosphorylated Smad2/3 (p-Smad2/3), and

induction of Smad2/3 target genes (Xu et al., 2012). Interestingly, it has been

reported that cytoplasmic forms of PML are critical for TGF-β signaling. The

tumor suppressive activity of TGF-β including growth arrest, apoptosis, and

cellular senescence are significantly reduced in Pml-/- MEFs. PML is able to

form a complex with TβR, SARA, and TGF-β downstream effector protein

Smad2/3 in the early endosomes. Phosphorylation and nuclear translocation of Smad2/3 are activated by TGF-β signaling through dissociating PML from

SARA and Smad2/3 (Lin et al., 2004). Our observations that SFN is capable of

inducing cytoplasmic accumulation of PML (Chapter 2, Figure 2-8) and partial

recruitment of PML to early endosomes (Figure 4-3), suggested that there

might be crosstalk between the SFN-PML axis and TGF-β signaling pathway.

In order to test this possibility, we first treated HUVECs with TGF-β followed by

subcellular fractionation. As expected, TGF-β induced accumulation of nuclear

p-Smad2/3, while cytoplasmic p-Smad2/3 was concomitantly reduced.

However, little or no change in the abundance of cytoplasmic PML was

observed in TGF-β-treated cells (Figure 4-4A). These results indicate that

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TGF-β promotes phosphorylation and nuclear translocation of Smad2 without increasing cytoplasmic PML abundance. We further studied whether TGF-β signaling is involved in SFN-mediated PML regulation by inhibiting the kinase activity of TβRI with the TβRI kinase domain inhibitor (TKDI). We observed that in the absence of TKDI, upon SFN exposure, p-Smad2 was significantly reduced in the nucleus, and concomitantly increased in the cytoplasm. As expected, TKDI completely blocked phosphorylation and nuclear translocation of Smad2. Remarkably, TKDI treatment abolished SFN-dependent accumulation of cytoplasmic PML and reduction in nuclear PML (Figure 4-4B).

This result suggests that there is crosstalk between TGF-β signaling pathway and SFN-mediated subcellular re-distribution of PML.

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164

Figure 4-4. Crosstalk between SFN-dependent PML regulation and TGF-β

signaling. (A) HUVECs were treated with TGF-β (5 ng/ml) for 1 hr and

harvested. Nuclear and cytoplasmic fractions were prepared followed by

immunoblotting analysis with the indicated antibodies. Lamin B and α-tubulin

were used as loading controls for nuclear and cytoplasmic fractions,

respectively. (B) HUVECs were treated with vehicle control (DMSO) or TKDI

(200 nM) for 1 hr. DMSO or SFN were then added to the medium at a final

concentration of 20 µM and incubated for another 1 hr. Nuclear and

cytoplasmic fractions were prepared followed by immunoblotting with the

indicated antibodies. Lamin B and α-tubulin were used as loading controls for nuclear and cytoplasmic fractions respectively.

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4.4 Discussion

SFN has been used as a dietary supplement to induce antioxidative responses

by modifying reactive cysteines on Keap1. Nrf2 is then released and

antioxidant targets are transactivated (Clarke et al., 2008). In Chapter 2, we

showed that PML was induced by SFN in a dose- and time-dependent manner

and this induction was also accompanied by marked accumulation of

cytoplasmic PML. This observation was also examined for its contribution to

SFN-mediated ROS elevation, Nrf2 activation, anti-proliferation, anti-migration,

and anti-angiogenesis activities. In this chapter, we have studied the

underlying mechanism based on a series of new observations. First, the

N-terminus of PML is critical for SFN-induced subcellular re-distribution of PML.

Second, potential reactive cysteine residues C212/213, C77/80, and C88/91 of

PML are essential for SFN-dependent PML accumulation in the cytoplasm.

Third, PML is partially recruited to early endosomes following SFN treatment.

Fourth, TGF-β signaling pathway is involved in the SFN-mediated subcellular re-distribution of PML. In sum, we have both partially dissected the underlying mechanism of SFN-dependent cytoplasmic accumulation of PML and extended our observations to interfere with TGF-β signaling.

At this point, it is still unclear whether the nuclear import and export of PML is directly affected by SFN. It has previously been shown that PML shuttles from the nucleus to the cytoplasm via a Crm1 (exportin-1)-dependent mechanism in

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cells infected with human immunodeficiency virus (HIV) (Turelli et al., 2001). In

order to test if this is applicable to our case, the Crm1 inhibitor leptomycin B or

a PML1 plasmid with nuclear export sequence (NES) region mutated that is

defective in nuclear export, will be utilized to examine whether SFN alters

nuclear export of PML. On the other hand, SFN is unlikely to be a general

inhibitor of nuclear import based on the fact that certain proteins including Nrf2

accumulation in nucleus when SFN is applied (Chapter 2, Figure 2-8). In

addition, there are no cysteine residues flanking the nuclear localization

sequence (NLS) region of PML that can be directly targeted by SFN. However,

it is still possible that binding of SFN on cysteine residues in the N-terminus of

PML alters the protein structure, which blocks recognition of the NLS by the nuclear import machinery. Future studies are needed to test this possibility.

Since loss of PML significantly inhibits the recruitment of TGF-β downstream components to endosomes and tumor suppressor activity of TGF-β (Lin et al.,

2004), we postulate that localization of PML to endosomes might be required to mediate TGF-β signaling. Interestingly, recruitment to endosomes is lost by mutating C57/60 of PML (Jul-Larsen et al., 2010). Thus, in the future, we should be able to test the hypothesis that endosomal localization is a key event in mediating the SFN effects by using K487R and C57/60A mutants of PML1.

In summary, our observations combined with current views provide insights on

167 the understanding of the dynamics of nuclear and cytoplasmic PML species upon SFN-mediated re-distribution. In-depth mechanistic studies will be carried out in the future to clarify how tumor suppressor PML is involved in therapeutic applications of SFN.

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4.5 Materials and methods

Chemicals and Antibodies

TGF-β, TKDI, anti-Smad2 antibody, and anti-p-Smad2 antibody were generous

gifts from Dr. David Danielpour. DRAQ5 was a generous gift from Dr. Minh

Lam. Alexa Fluor 555 μm goat anti-mouse (A21422) was purchased from Life

Technologies.

Plasmid Construction and Transfection

HA-PML4-C212/213A-C77/80A, HA-PML4-C212/213A-C88/91A,

HA-PML4-C77/80A-C88/91A, HA-PML4-C212/213A-C77/80A-C88/91A, and

HA-PML1-K487R were generated by site-directed mutagenesis. All constructs were verified by DNA sequencing. The primer sequences are listed in Table 1.

HeLa cells were transfected with plasmids using Lipofectamine 2000

(Invitrogen, P/N 52887) following the manufacturer’s protocol. The amount of plasmid transfected in each sample was kept constant by adding empty vector.

Con-focal microscopy

HeLa cells transfected with HA-PML1-K487R or HUVECs were plated on glass bottom microwell dishes (MatTek, P35G-1.5-14-C). SFN was introduced prior to fixation at 20 µM for 1 hr. The cells were fixed in 3.7% paraformaldehyde for

30 min at room temperature and washed with 1XPBS three times.

Permeabilization was performed by incubating cells with 1XPBS

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supplemented with 1% Triton X-100 and 10% goat serum for 10 min. The plate was covered to avoid light exposure since this step. Cells were then washed with 1XPBS for three times and blocked in 1XPBS supplemented with 0.1%

Tween-20 and 10% goat serum for 1 hr. Primary antibodies were added to the cells for 2 hrs followed by three washes with 1XPBS. Secondary antibodies conjugated with Alexa Fluor and far-red emitting fluorescent DNA dye DRAQ5

(ab108410) were added for 30 min followed by three washes with 1XPBS. All

confocal images were acquired using a Zeiss LSM 510 inverted laser-scanning

confocal microscope. All images were taken at the same time using same

settings.

Additional materials and methods used in this chapter can be referred to

Chapter 2, Section 2.5.

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Table 1. Sequences of primers used in PCR and qRT-PCR.

Primer Names Sequences (5'-3') hNRF2.Forward CATCTCGAGGGTACCATGATGGACTTGGA GCTGCCG hNRF2.Reverse CATTGGCCAGCTAGCCTAGTTTTTCTTAAC ATCTGG qRT.mPml.Forward CTGCGCTGCCCGAGCTGCCAGG qRT.mPml.Reverse CAGCGCAGGGTTGCGGTGGTTGG qRT.mNqo1.Forward GGCATCCTGCGTTTCTGTGGCT qRT.mNqo1.Reverse TTGCCCAGGTGATGGCCCAC qRT.mHmox1.Forward CCCCTGCAGAGACACCCCGA qRT.mHmox1.Reverse ACAGGGAGTGGGCTAGGGACC qRT.mFtl1.Forward TCTGCATGCCCTGGGTTCTGC qRT.mFtl1.Reverse GGGGAGCCCCTTGGAAGGTACA qRT.mTxn1.Forward AAGCTCCGTTGGGCGCCTTG qRT.mTxn1.Reverse GGCCTCCTGAAAAGCTTCCTTGCT qRT.mTxn2.Forward CAGCCCAGCCCGGACAGTAC qRT.mTxn2.Reverse GGGCCACACCACTGTGCATGA qRT.mTxnrd1.Forward AGACGATGAACGTGTCGTGGGC qRT.mTxnrd1.Reverse GCACAGACCGGGTGGATGCC qRT.mGclm.Forward ACAGCTACTGACTCACAATGACCCG qRT.mGclm.Reverse AGCCACAGCGGCACCCAATC qRT.mPrdx1.Forward TCCAGGCCTTCCAGTTCACTGAC qRT.mPrdx1.Reverse CTGGCTGCTCAATGCTGCCTGG qRT.mSdha.Forward GCTCTTTCCTACCCGATCACA qRT.mSdha.Reverse TCGGAGCCTTTCACAGTGTC qRT.mSdhb.Forward AGTGCGGACCTATGGTGTTG qRT.mSdhb.Reverse AGACTTTGCTGAGGTCCGTG qRT.mSdhc.Forward TCCCGCTCATGTACCACTCA qRT.mSdhc.Reverse ACAACACAGCAAGAACCACG qRT.mSdhd.Forward GGTGTGGTACCCAGCACATT qRT.mSdhd.Reverse GCAGCCAGAGAGTAGTCCAC qRT.mSdhaf1.Forward ATGGTGGTGCCCCAAAGAAT qRT.mSdhaf1.Reverse CGCCTGTGCTCTCTCTTTGA qRT.mSdhaf2.Forward TGCACAACATGACAGAGAAGCA qRT.mSdhaf2.Reverse CCAAGGTCAGTGCCAGAAGC qRT.mRn18s.Forward TAGTGAGGCCCTCGGATCGGC qRT.mRn18s.Reverse CCTTCCGCAGGTTCACCTACG qRT.hNQO1.Forward CCGTGGATCCCTTGCAGAGA qRT.hNQO1.Reverse AGGACCCTTCCGGAGTAAGA qRT.hPML.Forward GAATCAACGAATGAATGGCT qRT.hPML.Reverse CCAGGGACTCAGAATACAGG qRT.h18S rRNA.Forward CGTCTGCCCTATCAACTTTCG 171 qRT.h18S rRNA.Reverse CCTTGGATGTGGTAGCCGTT ChIP.hNQO1.Forward AAGTGTGTTGTATGGGCCCC ChIP.hNQO1.Reverse TCGTCCCAAGAGAGTCCAGG ChIP.hHmox1.Forward CCATCTGGCGCCGCTCTGC ChIP.hHmox1.Reverse GAGCAGCTGGAACTCTGAGGA Neh2(1-86).Forward CATGGTACCCTCGAGATGGACTTGGAGCT GCCGCCG Neh2(1-86).Reverse CATCTCGAGAATTGGGAGAAATTCACCTGT Neh4(112-134).Forward CATGGTACCCTCGAGGATGCTTTGTACTTT GATGAC Neh4(112-134).Reverse CATCTCGAGAACCTCATTGTCATCTACAAA Neh5(183-201).Forward CATGGTACCCTCGAGGACATTGAGCAAGT TTGGGAG Neh5(183-201).Reverse CATCTCGAGATTAAGACACTGTAACTCAGG Neh6(336-386).Forward CATCTCGAGACAGCAGAATTCAATGATTCT Neh6(336-386).Reverse CATGCTAGCTCCAGGGGCACTATCTAGCTC Neh1(435-562).Forward CATGAATTCGGTCATCGGAAAACCCCATTC Neh1(435-562).Reverse CATGCTAGCGAGATATAAGGTGCTGAGTTG Neh3(562-605).Forward CATGAATTCCTCGAAGTTTTCAGCATGCTA Neh3(562-605).Reverse CATGCTAGCCTAGTTTTTCTTAACATCTGG hPML.C77C80A.Forward CCTTGTCTGCACACGCTGGCCTCAGGAGC CCTGGAGGCGTCGGGC hPML.C77C80A.Reverse GCCCGACGCCTCCAGGGCTCCTGAGGCC AGCGTGTGCAGACAAGG hPML.C88C91A.Forward GCGTCGGGCATGCAGGCCCCCATCGCCC AGGCGCCCTGGCCC hPML.C88C91A.Reverse GGGCCAGGGCGCCTGGGCGATGGGGGCC

TGCATGCCCGACGC

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CHAPTER 5. DISCUSSION, IMPLICATION AND FUTURE

DIRECTIONS

In summary, we provide several lines of evidence demonstrating that PML plays a critical role in the maintenance of ROS homeostasis. Using PML knockdown HUVECs, Pml-/- MEFs, and overexpression approaches in HeLa cells, we have established that the abundance and subcellular distribution of

PML regulates ROS accumulation as well as Nrf2 activity. We also demonstrated that mitochondrial complex II activity was markedly reduced and genes encoding complex II proteins were down-regulated in PML deficient cells. Because impaired mitochondrial complex II results in increased ROS, which in turn activates Nrf2, we propose a model in which the Nrf2 antioxidant pathway is activated in response to increased ROS due to mitochondrial complex II dysfunction in PML deficient cells. Moreover, we are able to show that Nrf2 and PML physically interact with each other when they are either overexpressed in HeLa cells or under physiological conditions in HUVECs. We have mapped the interacting domains on Nrf2 and PML by GST pull-down assays. Additionally, we have observed that Nrf2 differentially regulates nuclear and cytoplasmic forms of PML. These observations suggest that PML and Nrf2 maintain homeostasis following oxidative stress via physical interaction and mutual regulation. Moreover, we show that SFN, a dietary supplement with antioxidative activity, exerts its cellular activity including ROS elevation, Nrf2 activation, anti-proliferation, anti-migration, and 173

anti-angiogenesis in a PML-dependent manner. Specifically, SFN alters the

subcellular distribution of PML by increasing its cytoplasmic accumulation

while reducing its nuclear abundance. The N-terminal region of PML and

potential reactive cysteine residues within the N-terminus are of particular

importance in mediating SFN-dependent cytoplasmic increase of PML. Once

localized in the cytoplasm, PML become partially recruited to early endosomes

as demonstrated by confocal microscopy.

5.1 How PML and Nrf2 coordinate in cancer development?

Based on our previous observations, PML and Nrf2 are part of a pathway that

maintains the homeostasis of cellular oxidant status. Cancer cells are exposed

to high levels of oxidative stress and are more resistant to oxidant-related

apoptotic cell death. Thus, how PML and Nrf2 affect tumorigenesis becomes

very important. PML has long been recognized as a tumor suppressor.

However, unlike PML, the role of Nrf2 in cancer remains illusive. In recent

years, there has been an emerging view that focuses on the “dark side” of

Nrf2-mediated cytoprotection. Mutations in Nrf2 that stabilize it have been

identified in subsets of human head-and-neck, lung, and gall-bladder cancers

(Hayes and McMahon, 2009). A recent study also suggest that Nrf2-mediated

adaptation to ROS stress is crucial for early stages of tumor development, and

if blocked, may slow down cancer progression (DeNicola et al., 2011). Our observations that increased protein abundance and activation of Nrf2 in 174

Pml-null or PML knockdown backgrounds, which are prone to tumorigenesis, shed new light on future studies regarding oncogenic potentials of Nrf2.

Specifically, carcinogens can be introduced to Pml+/+ and Pml-/- mice and tumorigenesis monitored. We anticipate that Pml-/- mice will develop tumors more rapidly. To test whether Nrf2 plays a role in oncogenesis in Pml-/- mice, liver-directed adenoviruses carrying Nrf2 shRNAs will be injected into the Pml-/- mice to determine whether tumorigenesis can be alleviated. Alternatively,

Pml-/- mice and Nrf2-/- mice can be crossed to examine whether their responses to carcinogens is comparable to their wild-type counterparts.

5.2 How does PML function in liver?

Our observations that PML is capable of reducing ROS generation and its regulation by the antioxidant drug SFN support the possibility that PML itself might be an antioxidant gene. Thus, in order to dissect the role of PML in the oxidative response, animal studies should be pursued with a primary focus on liver. Being the most important organ for metabolism and detoxification, liver has long been employed as an essential site for antioxidative investigations. To test our hypothesis, first, primary mouse hepatocytes will be isolated from

Pml+/+ and Pml-/- mice via liver perfusion to provide a more physiological setting for cell-based study. They will be used for functional studies including

ROS assays and cell proliferation assays. Second, liver homogenates prepared from Pml+/+ and Pml-/- mice will be used to analyze mRNA and 175

protein levels of Nrf2 and downstream cytoprotective genes. Third,

immunohistochemical staining and liver function tests will be performed using

either liver sections or blood isolated form Pml+/+ and Pml-/- mice. It has been shown that hyperactivated Nrf2 leads to severe liver injury in mice with autophagy deficiency (Komatsu et al., 2007; Komatsu et al., 2010). Since we expect more activated Nrf2 in Pml-/- mice, we would examine both liver lobule structures and levels of alanine transaminase (ALT), aspartate transaminase

(AST), and alkaline phosphatase (ALP) in circulating blood to evaluate liver function.

5.3 Is PML a direct target of SFN?

We have shown that C212/213, C77/80, and C88/91 of PML are critical for

SFN-mediated cytoplasmic accumulation of PML. However, it is not clear

whether SFN directly modifies these residues, although they can bind to

As2O3 (Jeanne et al., 2010; Zhang et al., 2010). Given the fact that there are

20 cysteine residues in the RING domain and B-boxes of PML, it is plausible

that some of these cysteine residues directly react with SFN. Because there is

no sequence or structure specificity established for SFN targets, it is difficult to

predict which cysteines on PML could be direct targets of SFN. Therefore, we

would like to thoroughly map SFN reactive cysteine residues on PML via liquid

chromatography–mass spectrometry/mass spectrometry (LC-MS/MS).

Plasmids carrying GST-PML and GST-Keap1 (positive control) will be 176

expressed in bacteria and subject to purification. Purified GST-PML will be incubated with SFN for varying times. Samples will then be prepared and analyzed by LC-MS/MS to identify SFN-modified cysteines in PML. Once

reactive cysteines are mapped, targeted LC-MS/MS will be performed using

extracts prepared from HUVECs treated with SFN. This will help us to confirm

whether in vitro reactivity between SFN and cysteines occurs in vivo.

5.4 What is the role of cytoplasmic PML in SFN-mediated multiple cellular functions?

We have demonstrated that PML is indispensable for several SFN activities in

cells. However, it is unknown whether cytoplasmic PML is sufficient to mediate

SFN activity even though it is dramatically induced by SFN. In order to test this, aortic ECs derived from Pml+/+ and Pml-/- mice will be utilized. Pml-/- ECs will be

transfected with either vector control or HA-PML1-K487R (cytoplasmic mutant).

The cells will be treated with or without SFN and used for cell proliferation

assays, would-healing assays, and in vitro network formation assays to

examine whether the cytoplasmic mutant of PML is capable of rescuing the

loss of PML and mediate SFN activity.

5.5 Is TGF-β signaling involved in SFN-mediated multiple cellular functions?

177

Our observation that inhibition of TGF-β signaling abrogates SFN-mediated

PML cytoplasmic accumulation and that PML is required for multiple cellular

functions of SFN, indicates that TGF-β signaling probably interferes with the

cellular activity of SFN. In order to examine this possibility, HUVECs will be

treated with TKDI, TGF-β, and SFN and followed by cell proliferation assay,

would-healing assay, and in vitro network formation assays to test whether

TGF-β signaling is involved. Alternatively, knockdown of TβR1 or

overexpression of a dominant negative mutant TβR1 will be employed under

the same experimental conditions as above to confirm our findings.

Taken together, we anticipate that our proposed study will shed light on our understanding of the mechanisms regulating ROS homeostasis and the mechanisms by which SFN exerts cellular functions. SFN is currently a promising agent for treating disease either alone or in combination with other drugs. However, most of the related mechanistic studies are not fully understood and in need for further clarification. Results from our study will also have implications in therapeutic designs for the treatment of diseases and may

pave the way for clinical applications of SFN.

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