Mechanisms of Aryl Hydrocarbon Receptor-Mediated Regulation of Gene Expression and the Cell Cycle

UNIVERSITY OF CINCINNATI

Date:______

I, ______, hereby submit this work as part of the requirements for the degree of: in:

It is entitled:

This work and its defense approved by:

Chair: ______

Mechanisms of Aryl Hydrocarbon Receptor-Mediated Regulation of Gene Expression and the Cell Cycle

A dissertation submitted to the

Division of Research and Advanced Studies of the University of Cincinnati

In partial fulfillment of the requirements for the degree of

DOCTORATE OF PHILOSOPHY (Ph.D.)

In the Department of Environmental Health of the College of Medicine

November 28, 2006

Jennifer L. Marlowe

B.S., Miami University, 1999

Committee Chair: Alvaro Puga, Ph.D. Professor Department of Environmental Health University of Cincinnati

Committee: Dr. Timothy Dalton Dr. Mary Beth Genter Dr. Erik Knudsen Dr. Ying Xia

The focus of this dissertation is the discovery of novel mechanisms and pathways of gene regulation by the aryl hydrocarbon receptor (AHR), primarily regarding the role of this protein in modulating cell cycle progression. The AHR is a member of the PAS (Per-Arnt-Sim) superfamily of receptors, which mediate responses to environmental stresses such as hypoxia and circadian rhythm, and control basic physiologic processes like vascular development, learning, and neurogenesis. The AHR protein was discovered by virtue of its high affinity interaction with the persistent environmental contaminant 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD), and is now known as the primary mediator of the toxic effects of this and hundreds of other HAH and

PAH ligands. The mechanisms by which the AHR acts to mediate toxicity of these compounds include the activity of the AHR as a potent transcriptional activator. The ligand-bound AHR, with its dimerization-binding partner ARNT, upregulates the expression of a battery of genes that function in the metabolism of PAH and HAH compounds. However, the diversity of toxic responses mediated by compounds such as TCDD are not adequately explained by the expression of this battery of genes. One of the primary roles of the AHR, both from a physiological and toxicological standpoint, is the control of cell cycle progression. The AHR may affect cell proliferation, differentiation, or apoptosis depending on the cell type examined, and the mechanisms of these effects remain unclear. Literally hundreds of genes have been implicated as being regulated either directly or indirectly by the AHR, and many of these genes are related to aspects of cell cycle control. The goal of this dissertation is to explore mechanisms by which the AHR modulates the cell cycle through the investigation of novel gene targets of the receptor. Chapter 2 summarizes the current body of knowledge regarding the AHR, its ligands, and perturbations of the cell cycle. Chapter 3 investigates a mechanism whereby the AHR is able to repress the expression of specific cell cycle-regulated genes through its interaction with

iii

the retinoblastoma protein, a tumor suppressor and major component of the G1/S checkpoint control mechanism. Chapter 4 explores an interaction between the AHR and E2F proteins, also major regulatory components of S-phase progression and DNA replication, and the constitutive activity of the AHR in maintaining basal expression levels of a large number of E2F-regulated

genes. Finally, Chapter 5 outlines the identification of novel promoter targets of the AHR using

chromatin immunoprecipitation and promoter tiling arrays. The results presented throughout this

work show the diversity of AHR functions related both to toxicological endpoints and normal

cell physiology, and illustrate the ability of this important transcription factor to regulate the

expression of a large number of genes by a variety of distinctive mechanisms.

Acknowledgements

My sincerest gratitude goes to Dr. Alvaro Puga for inviting me to join his lab as a graduate student, after hiring me as a technician with almost no experience in molecular or cell biology. He believed in me all those years ago, and the decision I made to complete my thesis work in his lab has led me to follow paths I only dreamed of going down before. I have learned almost everything I know about research, and a lot about life as well, from Alvaro, and am forever grateful for both the personal and professional experience of working in his lab.

I am indebted to all of the members of my dissertation committee for their dedication, insight, and criticisms that have helped to produce this thesis. Many thanks to Drs. Timothy Dalton, Mary Beth Genter, Erik Knudsen, and Ying Xia for their help and encouragement.

I would like to extend a special thanks to the classmates and lab partners who have helped me in countless ways over the years. Thank you to Xiaoqing Chang for both her friendship and willingness to help in any way I ever asked of her. From cell maintenance to western blots, from coimmunoprecipitations to PCR reactions, Xiaoqing was always there when I needed her. Thanks to Michael Schnekenberger for his tireless assistance and valuable scientific insight. Thank you as well to Mingya Huang, I have missed your companionship dearly since you left the lab, but have not forgotten your kind and generous spirit. I would also like to acknowledge the many terrific friends I have made through the years, those who have been there to commiserate with me on everything from the hardships of graduate school, relationships and family, to the drudgery of lab retreat planning. I am especially grateful for the support of Chris Curran, Scott Schneider, and Li Peng, always there when I needed to complain.

I would like to acknowledge the contribution of all of the faculty members and students in the Department of Environmental Health’s Division of Toxicology. I feel I have received excellent training in this stimulating and critical environment, and I will undoubtedly use the skills I have learned here in many new and exciting ways.

Thank you to my friends and family, past and present, for your invaluable perspective and support through this most trying of times. You have truly kept me from going off the deep end.

Thanks to the many collaborators for providing various reagents. Their contribution is acknowledged within each chapter.

Thanks to grant numbers 2R01 ES06273 and P30 ES06096 of the National Institutes of Environmental Health Sciences (NIEHS) for providing the funding for this research. Also thanks to the University of Cincinnati for support during my first two years of graduate school as a predoctoral Functional Genomics Fellow, and to The Ryan Foundation as well for fellowship support during my last years as a student.

Table of Contents

List of Tables…………………………………………………………………………………... 10

List of Figures………………………………………………………………………………….. 11

Abbreviations…………………………………………………………………………………... 13

Chapter I

Introduction…………………………………………………………………………………….. 16

AHR Ligands, Exposure, and Human Health………………………………………………….. 17

TCDD and the AHR……………………………………………………………………………. 21

Physiological Roles of the AHR……………………………………………………………….. 23

The AHR Signaling Pathway…………………………………………………………………... 25

Expanding Roles for the AHR in Gene Expression and the Cell Cycle……………………….. 30

Chapter II

Aryl Hydrocarbon Receptor: Cell Cycle Regulation, Toxicity and Tumorigenesis

Introduction…………………………………………………………………………………….. 34

AHR ligand-dependent activation of signal transduction pathways…………………………… 35

AHR agonists activate immediate-early response genes………………………………………. 38

Ligand-independent cell cycle control through the AHR……………………………………… 39

AHR-mediated inhibition of apoptosis………………………………………………………… 40

Cell cycle arrest induced by AHR ligands……………………………………………………... 42

Conclusions…………………………………………………………………………………….. 45

References……………………………………………………………………………………… 48

Chapter III

The Aryl Hydrocarbon Receptor Displaces p300 from E2F-Dependent Promoters and Represses S-Phase Specific Gene Expression

Introduction…………………………………………………………………………………….. 61

Materials and Methods…………………………………………………………………………. 64

Results………………………………………………………………………………………….. 71

Discussion……………………………………………………………………………………… 80

References……………………………………………………………………………………… 85

Chapter IV

Modulation of E2F-dependent gene expression by an interaction between the AHR and E2F proteins

Introduction……………………………………………………………………………………. 105

Materials and Methods………………………………………………………………………… 108

Results…………………………………………………………………………………………. 118

Discussion……………………………………………………………………………………... 129

References……………………………………………………………………………………... 135

Chapter V

ChIP-on-chip microarray analysis of AHR promoter binding sites in Hepa-1c1c7 cells

Introduction……………………………………………………………………………………. 157

Materials and Methods………………………………………………………………………... 160

Results………………………………………………………………………………………… 164

Discussion……………………………………………………………………………………... 168

References……………………………………………………………………………………... 172

Chapter VI

Conclusion…………………………………………………………………………………...... 192

References for Introduction and Discussion Sections………………………………………… 200

Appendix……………………………………………………………………………………… 212

List of Tables

Chapter III

Table 1 Gene-specific primer sets for real-time PCR analysis of relative mRNA expression levels……………………………………………………………….. 92

Table 2 Promoter-specific primer sets for chromatin immunoprecipitation analysis…... 92

Chapter IV

Table 1 Gene-specific primer sets for real-time PCR analysis of relative mRNA expression levels………………………………………………………………. 141

Table 2 Cell cycle-regulated genes and their canonical E2F and AHR binding sites…. 142

Chapter V

Table 1 Fifty AHR target gene hits with the highest signal ratios from DMSO-treated cells……………………………………………………………………………. 178

Table 2 Fifty AHR target gene hits with the highest signal ratios from TCDD-treated cells……………………………………………………………………………. 180

Table 3 Number of AHR and E2F binding sites of the annotated genes in the top 50 AHR gene target hits from DMSO-treated cells…………………………………….. 182

Table 4 Number of AHR and E2F binding sites of the annotated genes in the top 50 AHR gene target hits from TCDD-treated cells……………………………………... 183

Table 5 Comparison of AHR target genes identified using Chip-on-chip with available expression array data…………………………………………………………... 184

List of Figures

Chapter I

Figure 1 Examples of the diversity of aryl hydrocarbon receptor ligands………………. 18

Figure 2 Model for the aryl hydrocarbon receptor signaling pathway…………………... 26

Chapter II

Figure 1 Postulated ying-yang role of the Ah receptor in cell cycle regulation…………. 58

Chapter III

Figure 1 TCDD inhibits S-phase progression in MCF-7 and Hepa-1 cell lines…………. 97

Figure 2 TCDD inhibits expression of E2F-dependent, S-phase specific genes………… 98

Figure 3 AHR-mediated gene repression does not require transactivation…………….... 99

Figure 4 ARNT dimerization is not required for AHR-mediated gene repression……... 100

Figure 5 AHR represses E2F-dependent gene expression by blocking p300 coactivator activity…………………………………………………………………………. 101

Figure 6 Recruitment of AHR to E2F-regulated promoters results in loss of p300…….. 102

Chapter IV

Figure 1 AHR represses E2F-dependent transcription in multiple cell lines…………… 148

Figure 2 AHR binds RB and E2F in vitro with equal affinities………………………… 149

Figure 3 In vivo evidence for a direct interaction between the AHR and E2F1 transcription factors…………………………………………………………………………. 150

Figure 4 AHR interacts with E2F in gel shift assays and displaces RB from RB-E2F-DP1 complexes……………………………………………………………………... 151

Figure 5A AHR-mediated induction of Cyp1a1 gene expression is lost in AHR-/- MEF cells……………………………………………………………………………. 152

Figure 5B AHR expression is required for basal expression of many E2F-regulated genes…………………………………………………………………………... 153

Figure 6 The constitutive and TCDD-bound AHR binds differentially to E2F-regulated promoters……………………………………………………………………… 154

Chapter VI

Figure 1 Basic components of the ChIP-on-chip methodology………………………… 188

Figure 2 Verification of AHR target enrichment in ChIP samples……………………... 189

Figure 3 Verification of AHR target enrichment in LM-PCR-amplified ChIP samples... 190

Figure 4 Map of the top 20 genes hits found on Chromosome 2 by Chip-on-chip……... 191

Chapter V

Figure 1 Multiple models for the interplay of AHR in cell cycle pathways……………. 194

Appendix

Figure 1 AHR inhibits E2F1-induced apoptosis in transfected Saos-2 cells…………… 225

Figure 2 Co-expression of the AHR inhibits E2F-1-overexpression-induced apoptosis in Soas-2 cells……………………………………………………………………. 226

Figure 3 RB inhibits the expression of genes that are necessary for E2F-dependent apoptosis………………………………………………………………………. 227

Figure 4 AHR inhibits the expression of genes that are necessary for E2F-dependent apoptosis………………………………………………………………………. 228

Abbreviations

7-AAD, 7-amino-actinomycin D

AH, aromatic or aryl hydrocarbon

AHR, Ah receptor

AHRE, AHR response element

AHRR, AHR repressor

AIP, AHR interacting protein

ALDH3A1, aldehyde dehydrogenase 3

APAF1, apoptotic peptidase activating factor 1

ARA9, AHR associated protein 9

ARNT, aromatic hydrocarbon nuclear translocator

B[a]P, Benzo[a]pyrene bHLH, basic helix-loop-helix bp, base pairs

BrdU, bromodeoxyuridine

BRG, brahma-related gene

CDK, cyclin-dependent kinase

Cdt1, chromatin licensing and DNA replication factor 1

CERCLA, Comprehensive Environmental Response, Compensation, and Liability Act

ChIP, chromatin immunoprecipitation

CKI, Cyclin-dependent kinase inhibitor

Ct, cycle threshold

CYP, cytochrome P450

DEN, Diethylnitrosamine

DHFR, dihydrofolate reductase

DME, drug metabolizing enzyme

DMSO, dimethyl sulfoxide (Me2SO)

Dnmt1, DNA methyltransferase (cytosine-5) 1

DRE, dioxin response element

E2F, E2 factor

EGF, Epidermal growth factor

EGFR, Epidermal growth factor receptor

ERK, Extracellular signal-regulated kinase

GST, glutathione-s-transferase

GSTA1-Ya, glutathione transferase Ya

HAH, halogenated aromatic hydrocarbons

HDAC, histone deacetylase

HIF-1β: hypoxia-inducible factor 1β

HSP, heat shock protein

IARC, International Agency for Research on Cancer

IgG, Immunoglobulin G

JNK, Jun N-terminal kinase kb, kilobase pair

LM-PCR, ligation mediated PCR

MAPK, Mitogen activated protein kinase

MAPKK, MAPK kinase

MAPKKK, MAPKK kinase

Mcm3, minichromosome maintenance deficient 3

Me2SO, dimethyl sulfoxide

MEF, Mouse embryo fibroblasts

NQO1, NAD(P)H:quinone oxidoreductase

NTP, National Toxicology Program

PAH, polycyclic aromatic hydrocarbon

PAS, Per-Arnt-Sim

PCB, polychlorinated biphenyl

PCDD, polychlorinated dibenzo-p-dioxin

PFU, plaque-forming unit

Pol α, DNA polymerase alpha

RB, retinoblastoma protein

SDS, sodium dodecyl sulfate

SRE, Serum response element

TCDD, 2,3,7,8-tetrachlorodibenzo-p-dioxin

UGT1A6, UDP glucuronosyltransferase 1A6

XME, xenobiotic metabolizing enzyme

XRE, xenobiotic response element

Chapter I

Introduction

Organisms have evolved a variety of molecular mechanisms to adapt to both endogenous stimuli and exogenous insults. Xenobiotic exposures may specifically or non-specifically induce signal transduction events that lead to various physiological responses, including homeostatic

perturbations and cellular responses such as proliferation, differentiation, apoptosis, or necrosis,

among others. Organisms attempt to minimize the damage caused by such exposures through

the expression of a variety of diverse drug or xenobiotic metabolizing enzymes (DMEs or

XMEs). DMEs play a central role in the detoxification and/or biotransformation of xenobiotic

compounds, and are present in abundance either at the basal level or induced in most tissues and

organs following an exposure (1). The response of the aryl hydrocarbon receptor (AHR) signal

transduction pathway is implicated as one mechanism whereby organisms respond to

environmental xenobiotics, primarily through the ability of the AH receptor to induce or

upregulate drug metabolizing enzymes.

The existence of a single genetic locus was initially hypothesized to explain the

differential induction of aryl hydrocarbon hydroxylase (AHH, now known as Cyp1a1) in two

strains of mice, DBA/2 and C57BL/6 (2), exposed to various PAH inducers. This difference was

thought to lie in the enzyme responsible for AHH activity. However, AHH was equally

inducible in both strains of mice in the presence of the persistent environmental contaminant

2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD or dioxin) (3). Thus, the AHR was eventually

discovered as a component of the hepatic cytosol that bound with specific and high affinity to

TCDD, and to ultimately control induction of AHH activity (4-6). Currently, the AHR is viewed

as playing a role in a number of clearly defined biological processes, including the adaptive

metabolic response to exogenous chemicals, specifically polycyclic aromatic hydrocarbons

(PAHs), the pleiotropic toxic responses associated with exposure to more potent agonists such as

TCDD, and vascular remodeling of the developing embryo (7;8). However, the AHR appears to

play a relatively broad role in development outside of the vasculature, although this role is still

being defined (9;10). In addition, evidence continues to emerge concerning the multiple roles

played by the AHR in fundamental aspects of cell biology and physiology (11-15).

Significantly, discovery of the AHR was the first step in the ultimate identification of the PAS

(Per-Arnt-Sim) superfamily of receptors. In general, PAS receptors mediate responses to various

environmental stresses, such as hypoxia and circadian rhythm, and control basic physiologic

processes and functions such as vascular development, learning, and neurogenesis (16).

AHR Ligands, Exposures, and Human Health

Ligands of the AHR generally fall into two main categories: those that are formed as the

result of anthropogenic or nonbiological activity, and those that occur naturally or are formed in

biological systems as a result of natural processes. The anthropogenic forms tend to be the most

potent of the AHR-binding compounds, having both the highest affinities for the receptor, as

well as the most obvious and profound biological or toxicological effects. These high affinity ligands of the aryl hydrocarbon receptor are most likely to be hydrophobic, planar, and of a

defined size, all properties which facilitate interaction with the hydrophobic ligand-binding

pocket of the AHR (17). However, the range of compounds able to interact with the AHR

suggests that its ligands may diverge widely from the structural requirements for maximal

binding affinity (see Fig. 1). Although many studies have shown that the AHR can interact with

and be activated by a structurally diverse range of chemicals (18), the best-characterized high affinity ligands for the AHR include a wide variety of ubiquitous and hydrophobic

2,3,7,8-tetrachlorodibenzo-p-dioxin Prostaglandin G2 Tryptamine

Benzo[a]pyrene Indirubin

Cl Cl

Cl Bilirubin Cl PCB 126 Omeprazole

NH2

NH2 3-methylcholanthrene 7-ketocholesterol 1,5-diaminonapthalene

Fig. 1. Examples of the diversity of aryl hydrocarbon receptor ligands. Classical inducers of the AHR are represented by TCDD, B[a]P, PCB 126, and 3-methylcholanthrene. Naturally occuring or dietary sources of AHR ligands are represented by prostaglandin G2, bilirubin, 7-ketocholesterol, tryptamine, and indirubin. Additional nonclassical AHR ligands include omeprazole and 1,5-diaminonapthalene. environmental contaminants such as HAHs (polychlorinated dibenzo-p-dioxins (PCDDs), dibenzofurans and co-planar members of the polyhalogenated biphenyl family) and PAHs (such

as benzo[a]pyrene, 3-methylcholanthrene, benzanthracenes, benzoflavones, aromatic amines and

related chemicals) (19). The HAHs represent the most potent class of AHR ligands and are

relatively metabolically stable, with binding affinities in the pM to nM range, whereas the PAHs,

being metabolically unstable in comparison, and bind with lower affinity (nM to µM range) (20).

Of the top 10 substances listed in the 2005 CERCLA (Comprehensive Environmental

Response, Compensation, and Liability Act, commonly known as Superfund) Priority List of

Hazardous Substances, five are agonists of the AHR, including polychlorinated biphenyls

(PCBs), benzene, PAHs, benzo[a]pyrene (B[a]P), and benzo[b]fluoranthene (21). A variety of exposure outcomes has been reported for both humans and animals for each of these environmentally important chemicals. The PCBs are synthetic organic chemicals comprising

209 individual chlorinated biphenyl congeners. Although banned in the U.S. since 1979, PCBs were once widely used in many industrial applications. PCBs degrade slowly in the environment and hence can bioaccumulate in the food chain. Consumption of fish and other meat products

with high PCB levels is thought to put individuals at risk for adverse reproductive effects,

neurobehavioral and developmental deficits (especially among children exposed in utero), systemic effects including in the thyroid and immune systems, and some forms of cancer (22;23).

However, there is continuing controversy over the human health effects of environmental exposure to PCBs (24;25). Additionally, the polycyclic aromatic hydrocarbons, such as

benzo[a]pyrene, constitute a portion of the carcinogenic components of cigarette smoke, smog,

and some over-cooked foods. Very high levels of PAH exposure, such as in occupational

settings, have long been associated with increased cancer risk, although questions still remain as

to the relative risk associated with PAH exposures in the ambient environment (26). Most

importantly, the health effects observed upon exposure to a majority of the aforementioned

chemicals requires activation of the AHR signal transduction pathway, described in detail in the

following sections.

It has long been hypothesized that in addition to the AHR’s role in the induction of drug

metabolism by HAHs and PAHs, the receptor is also required for endogenous functions critical to life processes (27). Studies using AHR knock-out mice have shown that this is indeed the case (28-30), and although no physiologically relevant endogenous ligand for AHR has been

unequivocally established, it is presumed that some if not all of the biological functions performed by the AHR occur as the result of ligand-receptor interactions. The existence of endogenous physiological AHR ligands has been suggested by numerous studies in which the

AHR signaling pathway is active in the absence of exogenous ligands (31-34), and that this activity is likely due to the presence of endogenous ligands (35). Numerous endogenous chemicals have in fact been identified as either binding to or activating the AHR, including UV- induced photoproducts of tryptophan (36;37) and histidine (38), indirubin and indigo (39), arachidonic acid metabolites, especially prostaglandin G2 (40), and the tetrapyrole heme degradation products bilirubin and biliverdin (41). Despite the large number of potential endogenous AHR ligands identified to date, one of the major unanswered questions in AHR biology is the specific compounds that activate the AHR at normal physiological concentrations.

However, the promiscuous ligand binding activity of the AHR is suggestive of multiple, and probably tissue-specific, endogenous ligands (42).

In addition to endogenous ligands, a variety of naturally-occurring dietary compounds, including a number of flavonoids, carotinoids, and phenolics, activate the AHR despite being relatively weak ligands (43-46). Conversion of dietary indoles and trytophan in the mammalian digestive tract can produce potent AHR ligands (47;48). Extracts of a large diversity of vegetables, teas, fruits, and natural herbal products also have AHR agonist and/or antagonist activity, likely due to their high concentration of flavonoid chemicals (49;50). As plant-derived extracts in general commonly contain AHR ligands or compounds that are readily converted into

AHR ligands, dietary sources represent the largest class of natural AHR ligands to which humans are exposed (20). It may be that naturally occurring ligands of the AHR are an essential key to understanding the evolution and function of this receptor in biological systems. Importantly,

many of the compounds represented in this class of AHR ligands are AHR antagonists, and may be able to inhibit the deleterious effects of several of the exogenous AHR ligands of

environmental concern (51). Although it appears that some AHR ligands have potential as

anticancer therapeutics, the totality of AHR signaling is complicated by a number of regulatory

and crosstalk mechanisms which are still poorly understood (52).

TCDD and the AHR

The polychlorinated dibenzo-p-dioxins (PCDDs or dioxins) have been recognized as

highly toxic and persistent environmental contaminants since the early 1900s (53). PCDDs are

produced in nature from the incomplete combustion of organic materials, such as during forest

fires or volcanic activity. However, the most abundant sources of PCDDs are the unwanted

byproducts of industrial, municipal, and domestic incineration of materials containing chlorine, which include plastic, treated wood products, and pesticides (54). TCDD is the most potent member of the PCDD compounds (42), and is also the most widely studied of the AHR ligands.

It is broadly accepted that the diverse biochemical, biological, and toxicological responses observed following TCDD exposure, both in vitro and in vivo, are mediated by the AHR (55).

TCDD is an extremely persistent, non-metabolizable AHR agonist, quite distinct from the myriad of metabolizable dietary, endogenous, and exogenous AHR ligands mentioned previously. The most puzzling aspect of TCDD toxicity may be that its effects vary greatly depending on cell type, tissue, age, sex, species, and the timing and duration of exposure. In fact, toxicity can vary greatly even within the same cell, depending upon the culture conditions and stage of growth (53). Although there remains a great deal of controversy over the human health effects of TCDD (56;57), it is currently listed as a definitive human carcinogen by the

National Toxicology Program (NTP) and the International Agency for Research on Cancer

(IARC) (58). The basis for this designation includes limited evidence in humans, sufficient evidence in animals, and extensive mechanistic data that TCDD acts through the AHR pathway, present in both humans and animals (57).

Current exposures to TCDD generally occur through soil, dust, smoke and ambient air, although the primary route of exposure is through the diet (59;60), accounting for over 90% of the body burden of PCDDs in the general population (background is estimated at 5 ppt or 2 pg/g body fat (54;57)). Once exposure occurs, dioxin remains in human tissues, particularly fatty tissues, for extended periods of time (the half-life of dioxin in humans is calculated to be 7 to 10 years) (61), one reason for the persistent concern over relatively low levels of TCDD in the environment. Levels of dioxins in the ambient environment continue to decrease as the major sources of PCDD contaminants have been eliminated, including production of the herbicide

2,4,5-trichlorophenoxyacetic acid and the antiseptic hexachlorophene, and the bleaching of paper

and pulp products using free chlorine (53).

Much of the data on the toxicological effects of TCDD in humans comes from 3 main

sources: industrial cohorts of chlorophenol/phenoxy herbicide chemical workers, in which

TCDD is produced as a byproduct, Vietnam-era military personnel exposed to the TCDD-

contaminated defoliant Agent Orange, and community-based studies of accidentally exposed

populations. The range of estimated exposures among these groups varies widely, but is without

question significantly higher than current background levels, often by as much as 1000-fold (54).

Acute symptoms of TCDD exposure include the onset of chloracne, symptoms of acquired

porphyria cutanea tarda, possible transient liver toxicity, fatigue, general weakness, and weight

loss (54;62). Long-term effects of dioxin exposure include an increased risk for atherosclerosis

(63;64), hypertension and ischemic heart disease (62;65), retinopathies, neurological

abnormalities (54), diabetes (66;67), hormonal perturbations (68;69), and cancer (57;62). This last observation, namely the ability of dioxin exposure to promote carcinogenesis in humans, has been a primary focus of research into the impact of TCDD on human health. TCDD exposure results in small but significant increases in lung, lymphopoietic, rectal, liver, breast and skin cancers in humans (57). Animal studies have confirmed most of the adverse effects observed in humans (53;70). Significantly, TCDD acts as a potent non-genotoxic tumor promoter in multiple organs of laboratory mice and rats, including the liver, lung, thyroid, adrenal cortex, and skin

(71). While animal studies in multiple species have also shown that TCDD affects the developing immune, nervous, reproductive, and cardiovascular systems, the same effects have not been documented in humans (70). However, other potent ligands of the AHR are known to exhibit developmental toxicities, particular the polyhalogenated biphenyl compounds (72). It should be noted that humans appear relatively resistant to TCDD toxicity as compared to other mammalian species, likely due the relatively low dioxin-binding affinity of the human AHR

(73).

Physiological Roles of the AHR

A general hypothesis is emerging that the diversity of tissue-specific, TCDD-mediated toxicities observed in humans and animals is due to sustained and inappropriate AHR activation, resulting in the deregulation of physiologic functions normally performed by the receptor (74).

Therefore, the toxicological profile of TCDD as outlined above may provide clues as to the nature of these physiological functions. Other clues are born out of physiological defects observed in AHR knockout mice, as well as the nature of receptor homologs in other vertebrate and non-vertebrate species.

Single homologs of the AHR have been discovered in many invertebrate species, including the nematode Caenorhabditis elegans (ahr-1) (75), the fly Drosophila melanogaster

(spineless) (76), the chordate Ciona intestinalis (77), the clam Mya arenaria (78), and the mussels Dreissena polymorpha and Mytilus edulis (79). The AHR homolog found in Drosophila is involved in the development of antennae and legs (76), while the C. elegans homolog ahr-1 is involved in nervous system development, specifically the fate of GABAergic neurons (80).

However, none of these receptors exhibit high-affinity binding of TCDD or other classical AHR ligands, suggesting that the physiological functions of the receptor during development are ancestral to the adaptive functions apparent in higher order organisms. Among vertebrate species, fishes, amphibians, and birds are known to possess multiple AHR isoforms, whereas mammalian species harbor only one AHR. Mammals vary widely in their sensitivity to dioxin toxicity, at least in part due to differences in the physicochemical properties of the AHR (81).

The AHR is ubiquitous throughout a diversity of mammalian tissues. While levels of the receptor may vary widely across different tissues and among different cell types, it has been found in abundance in liver (82), lung, small intestine, kidney, thymus, pancreas, heart (83;84), placenta (85), vasculature (86), ovaries and uterus (87), and brain (88). Significant clues as to the physiological function of the receptor in mammalian systems have come through the study of

AHR knockout mice. Notably, these mice display a number of defects in many of the organ systems which are directly impacted by TCDD exposure. In initial studies of these AHR-/- mice, pups were born and survived at much lower rates than their heterozygous counterparts, likely due to increased susceptibility to infection, and lower lymphocyte counts were observed in immature and older knockout mice (28). However, studies on AHR-/- mice developed independently and in a separate laboratory indicated no impact of the AHR on survival, but that mice lacking the

receptor grew slower and weighed less then wild-type controls (30). Both strains of knockout mice showed signs of liver fibrosis, generally smaller livers, and minor effects in the spleen, while only one strain appeared to exhibit fibrosis of the lung. Basal expression of several drug metabolism genes was also lost, suggesting a role for the AHR in both inducing these genes as well as maintaining basal expression levels (29). Additional studies have refined the AHR knockout phenotype even further. Defects in the livers of AHR-null mice have been attributed to the failure of a patent developmental structure known as the ductus venosus to properly close

(89), resulting in compromised blood perfusion, hepatic necrosis, and ultimately liver deformation persisting through adulthood (90). AHR-/- mice also exhibit cardiac hypertrophy and increased blood pressure under conditions of mild hypoxia, correlated with elevated angiotensin II and endothelin-1 levels (91). The AHR appears also to play a role in both male sexual maturation, namely for development of the prostate and seminal vesicles (92), and in female reproduction, by regulating the expression of ovarian P450 aromatase (Cyp19), a key enzyme in estrogen synthesis (13). In addition, loss of the AHR results in defects in ovarian and uterine functions concomitant with poor reproductive success at a relatively young age (93).

Importantly, AHR knockout mice are resistant to toxicity associated with xenobiotic exposures

(94;95), confirming its central role in mediating the adverse effects of toxic AHR ligands.

The AHR Signaling Pathway

The previous sections have focused on the nature of the AHR protein, its ligands, and the adverse effects of the xenobiotic ligands. At the heart of these toxicological effects is the aryl hydrocarbon signal transduction pathway, of which the AHR itself is absolutely required for these adverse outcomes, particularly those associated with TCDD exposure. Much is known about the molecular biology of the AHR signal transduction pathway, as several components of

this pathway are well defined and relatively invariable with respect to the particular chemical under investigation (Fig. 2). In its unliganded, inactive form, the AHR exists as a cytosolic protein in complex with a dimer of the 90 kD heat shock protein (hsp90) (96-98), the hsp90- associated co-chaperone p23 protein (99;100), and the immunophilin homolog XAP2 (also called

ligand (TCDD etc.)

p23 p23 Hsp90 Hsp90 Hsp90 Hsp90 XAP2 R XAP2

XAP2 AHR pleiotropic effects AH cytoplasm nucleocytoplasmic importin shuttling p23 Hsp90 R Ahrr Hsp90 H p27 p23 A Hsp90 Cyp1a1 Hsp90 p23 importin p23 Hsp90 R

? XAP2 H Hsp90 p/CIP/ Hsp90 CRM1 Hsp90 RIP140 A export XAP2 CRM1 p300

export T nucleus AHR ARN XRE

NT Cyp1a1, Nqo1, RNT Ahrr, p27, etc. AR A RR NT AHRR AR AH

Fig. 2. Model for the aryl hydrocarbon receptor signaling pathway. Adapted from Mimura and Fujii-Kuriyama (2003) and Petrulis and Perdew (2002).

AIP (AHR interacting protein) or ARA9 (AHR associated protein 9)) (101-103). Hsp90 binds both the ligand-binding (PAS) domain of the receptor, as well as the basic helix-loop-helix

(bHLH) DNA-binding domain, and functions to aid in proper folding and to enhance stability of the AHR protein (104;105). The XAP2 protein also functions to maintain AHR stability, as well as subcellular localization, and the association of XAP2 with the AHR prevents dynamic nucleocytoplasmic shuttling of the receptor in its unliganded state (106). The significance of

AHR shuttling in the absence of ligand is not yet understood, but it is hypothesized that the AHR may serve some ligand-independent function in the nucleus, possibly through interactions with other proteins (107). Upon ligand binding, the entire AHR complex translocates to the nucleus with the aid of a conformational change resulting in exposure of a nuclear localization sequence on the AHR (108). In the nucleus, the AHR forms a heterodimer with the aryl hydrocarbon receptor nuclear translocator (ARNT) protein (also called HIF-1β) (109), concurrent with the loss of the hsp90/p23/XAP2 complex (110). The AHR-ARNT heterodimer binds to DNA at dioxin response elements (DREs) or xenobiotic response elements (XREs), containing the core recognition motif 5’-GCGTG-3’ (111), to regulate the transcription of a number of target genes

(112). The best-characterized of these target genes are members of the AHR “gene battery,” and include Phase I drug metabolizing enzymes of the CYP1 family, namely CYP1A1, CYP1A2, and

CYP1B1, and the Phase II enzymes NAD(P)H:quinone oxidoreductase (NQO1), aldehyde dehydrogenase 3 (ALDH3A1); UDP glucuronosyltransferase (UGT1A6); and a glutathione transferase (GSTA1,Ya) (113). Notably, these enzymes exhibit an obvious preference for substrates that are AHR agonists, suggesting that the activation of this signaling pathway acts as a defense system aimed at the elimination of the inducer and its metabolites (52). The AHR also

regulates the transcription of a number of genes that are not metabolizing enzymes, including

p27Kip1 (114), N-myristoyltransferase 2 (115), and Bax (116), among others. The AHR repressor

(AHRR) protein is also inducible in an AHR-dependent manner (Fig. 2), and it acts to antagonize

the activity of the AHR/ARNT heterodimer (117;118). Essentially, the AHRR functions as a

negative regulator of AHRR by competing with AHR for ARNT binding, suggesting that AHR

and AHRR form a regulatory feedback loop (112). Following gene transactivation, the AHR is

exported to the cytoplasm from the nucleus through binding of a nuclear export sequence to

CRM-1, and is subsequently ubiquinated and degraded by the 26S proteasome pathway

(106;119).

The mechanism by which AHR/ARNT activates gene transcription has been studied primarily for genes of the AHR gene battery, chiefly CYP1A1, but the mechanisms are thought to apply to most genes that are activated by the AHR through DRE binding. The ability of the

AHR/ARNT heterodimer to activate transcription of these genes is enhanced by its interaction with a number of basal transcription factor proteins, including TFIIE (120), TFIIB (121), and the

TATA binding protein, as well as the transcriptional coactivators p300, p/CIP, RIP140, BRG-1,

GRIP1, and SRC1 (122). The AHR also physically associates with the endogenous

TRAP/DRIP/ARC/Mediator complex, specifically the Med220 subunit, which is known to interact with several nuclear hormone receptors, at the CYP1A1 enhancer (123). In contrast, inhibition of transcriptional activation by AHR/ARNT may occur through interactions with corepressor proteins such as SMRT (124) and SHP (125). The extent to which the AHR/ARNT complex activates its targets genes varies widely depending on the particular gene or cell type examined. This variability depends upon a number of elements, including the specific

nucleotides surrounding the core DRE sequence, expression of the AHR repressor (AHRR)

protein, relative expression levels of co-activator and co-repressor proteins, as well as the

relative levels of other dimerization partners for ARNT (126). For example, in cells such as

fibroblasts, in which the AHR/ARNT heterodimer shows little ability to activate gene

expression, there exist very high expression levels of the AHRR protein, which acts as a

transcriptional repressor at DRE sequences through its interaction with ARNT (127). In

addition, considering that a majority of detailed studies on AHR-interacting proteins, particularly

within the chaperone complex, were carried out in one specific cell type (Hepa-1), it is likely that

other proteins interact with the AHR in other cell types or in other species, and modify its activity by thus far unknown mechanisms (17). For example, multiple observations signify that the AHR and AHR/ARNT dimers interact with a diversity of transcription factors, suggesting an additional mechanism by which the AHR signaling pathway alters gene expression (126).

Interactions between the AHR or ARNT and such transcription factors represent a potentially important mechanism by which AHR ligands such as TCDD block the ability of proteins to upregulate their respective target genes and block cellular events involved in progression of the cell cycle (via RB) (128;129), the actions of estrogens (via ERα, COUP-TF1, and ERRα1)

(130), and in the inflammatory response (via NF-κΒ) (131).

In addition to its interaction with coactivator and corepressor proteins, the ability of the

AHR to transactivate genes is also controlled through mechanisms that regulate AHR protein

levels and its DNA- or ligand-binding affinity. The regulation of AHR levels and activity is

complex and depends upon the ligand and system under study. For example, treatment of cells

with TCDD over hours or days results in decreased AHR protein levels, as well as decreased

ligand and DNA binding activities. In contrast, in vivo dioxin treatment over days or weeks

causes increases in AHR protein and mRNA levels, and increased ligand binding activity.

However, decreases in AHR expression are also seen in vivo at very early time points or at later

time points if the dioxin dose is very high. Other ligands of the AHR generally cause an increase

in AHR levels and molecular activities, whether in cells or in vivo, but this also varies with the

cell or tissue type examined. Additionally, chemicals that are not AHR ligands have been

reported to increase its expression, including phenobarbitol and non-coplanar PCBs. Other

ligand-independent conditions where increased AHR mRNA or protein levels are evident include

cell differentiation or senescence, increasing degrees of cell transformation (including tumors in

vivo), and pre-pubertal development. Decreased receptor levels have been noted in growth factor-deprived cell cultures and in aging animals (132). The intrinsic regulation of AHR levels during normal physiological processes again implies an important role for the receptor in these processes. Controlling the levels of activated or nuclear AHR is clearly important, as blockage of the AHR degradation pathway leads to potentiation of gene induction (133;134), and expression of a constitutively active AHR in mice results in reduced life span, stomach tumors

(135), thymic involution (136), inflammatory skin lesions (137), and the promotion of hepatocarcinogenesis (138).

Expanding Roles for the AHR in Gene Expression and the Cell Cycle

It is clear that the AHR plays a role in many biological processes, and that its aberrant activation by exogenous ligands results in the disruption of these processes. One aspect of AHR biology receiving increasing attention is the apparent role played by the receptor in the cell cycle, both through ligand-dependent and ligand-independent mechanisms, and the relevance of this role to the myriad toxicities observed following TCDD exposure in particular. It is known that activation of the AH receptor by high-affinity ligands such as TCDD may result in arrest of the cell cycle, cell proliferation, apoptosis, or differentiation, depending on the cell type examined

(139). The fact that AHR activation results in such a variety of outcomes indicates that the AHR modulates molecular pathways unrelated to the induction of the phase I and phase II gene battery for detoxification of foreign compounds. At least some of these pathways are likely to involve components of cell cycle regulation, and evidence from multiple investigators indicates that this is the case (reviewed in Chapter 2). Previous results from the Puga laboratory and others suggested that the AHR may function to perturb cell cycle progression through its interaction with a major regulator of the cell division process, namely the retinoblastoma (RB) protein. The

overarching themes of the following work are thus two-fold. First is the idea that the AHR may control the expression of a host of genes that are unrelated to the phase I and phase II enzymes for drug metabolism, and are involved in cell cycle regulation. Second is the idea that the consequence for the expression of a particular gene as controlled by the AHR is dependent upon the specific manner in which the receptor is recruited to that gene, namely through DRE sequences or through alternative mechanisms.

The goal of this dissertation is to explore novel mechanisms by which the AHR modulates cell cycle progression, in order to address one aspect of the toxicity observed in cells exposed to exogenous AHR ligands, as well as to shed light on possible physiological roles of the AHR in alternative signal transduction pathways. Specifically, the following chapters aim to characterize a role for the AHR in modifying cell cycle progression through its interaction with cell cycle regulatory proteins, and by regulating the expression of genes through novel mechanisms involving these interactions. Chapter 2 provides a summary of current research into the role of the AHR in regulating cell cycle progression through a number of mechanisms, and speculates as to the ways in which the AHR affects one of these pathways, namely those controlling the G1 to S-phase transition. Chapter 3 explores an interaction between the AHR and

the RB proteins, and a mechanism by which the AHR can act as a repressor of gene expression.

Chapter 4 discerns a newly-identified interaction between the AHR and E2F family of proteins,

and outlines an additional mechanism whereby the AHR controls cell cycle progression. Finally,

Chapter 5 aims to identify novel AHR-binding genes through the use of promoter microarrays, in

order to show that AHR targets and regulates the transcription of many unrecognized genes, not

only through DRE sequences, but also through its interactions with other transcription factors or

through as-yet uncharacterized mechanisms.

Chapter II

Aryl Hydrocarbon Receptor: Cell Cycle Regulation, Toxicity and Tumorigenesis

Jennifer L. Marlowe and Alvaro Puga*

*Corresponding author

Department of Environmental Health and Center for Environmental Genetics University of Cincinnati Medical Center P.O. Box 670056 Cincinnati, OH 45267-0056 Phone:(513) 558-0916 Fax:(513) 558-0925 email: [email protected]

KEYWORDS: AH receptor, Xenobiotic ligands, Signal Transduction, Retinoblastoma protein

Abstract

Most effects of exposure to halogenated and polycyclic aromatic hydrocarbons are mediated by the aryl hydrocarbon receptor (AHR). It has long been recognized that the AHR is a ligand-activated transcription factor that plays a central role in the induction of drug- metabolizing enzymes and hence in xenobiotic detoxification. Of late, it has become evident that outside this well-characterized role, the AHR also functions as a modulator of cellular signaling pathways. In this review, we discuss the involvement of the AHR in pathways critical to cell cycle regulation, mitogen-activated protein kinase cascades, immediate-early gene induction, and the functions of the RB protein. Ultimately, the toxicity of AHR xenobiotic ligands may be

intrinsically connected with the perturbation of these pathways and depend on the many critical

signaling pathways and effectors with which the AHR itself interacts.

Introduction

Exposure to halogenated aromatic hydrocarbons (HAHs) and polycyclic aromatic hydrocarbons (PAHs) results in a wide range of toxic and carcinogenic responses in animals and

in humans. It is widely accepted that most of these exposure effects are mediated by the aryl

hydrocarbon receptor (AHR), a cytosolic ligand-activated transcription factor that upon ligand

binding, translocates to the nucleus where it complexes with ARNT (a.k.a. HIF-1β).

AHR/ARNT heterodimers bind to specific consensus DNA sites in the regulatory domains of genes coding for many Phase I and Phase II drug-metabolizing enzymes and activate the transcription of these genes (1). During the last 8-10 years, it has also become evident that the

AHR has a second function, involving promotion of cell cycle progression, and that this function is accomplished in the absence of an exogenous ligand. In contrast, activation of the AH receptor by high-affinity HAH or PAH ligands such as TCDD and B[a]P has been known for many years to result in a wide range of cell cycle perturbations, including G0/G1 and G2/M arrest, diminished

capacity for DNA replication, and inhibition of cell proliferation [reviewed in (2)]. These two

outcomes are diametrically opposed and raise questions for which we do not have satisfactory

answers at present. For example, how does the unliganded cytosolic AH receptor influence a

nuclear function such as cell cycle progression? Does this effect involve nuclear translocation? If

so, do liganded and unliganded nuclear translocation events have different molecular outcomes?

What makes dioxin carcinogenic? In these and similar questions, we need to bear in mind that it

is only our current state of ignorance that allows us to apply the term unliganded to an AH

receptor that may be liganded by an as yet uncharacterized endogenous ligand.

The fact that the AHR is involved in such a variety of outcomes indicates that it is able to

modulate diverse molecular pathways in concert with the induction of Phase I and Phase II genes

for the detoxification of foreign compounds. Many studies have shown that the AHR functions in the direct and indirect modulation of transcriptional programs, at least in part by associating with additional transcription factors (3;4), coactivators or corepressors (5;6), and by altering signal transduction cascades (7;8). In doing so, the unliganded receptor may modulate components critical for the regulation of cell cycle progression. In this case, the presence of a high affinity ligand might cause a toxic response because a particular regulatory function would not be carried out by virtue of the receptor's engagement with ligand. Alternatively, the unliganded AHR might be fully quiescent and its activation by ligand might simply elicit detoxification and adaptive responses. In this case, the toxicity of the ligand may be determined by the functions that the

AHR would carry out by virtue of its engagement with ligand. In recent years, experimental

evidence has accumulated in favor of the first alternative, whereas classical toxicological

research has provided much evidence in favor of the second. In all likelihood, both alternatives

are correct.

AHR ligand-dependent activation of signal transduction pathways

Exposure to exogenous ligands of the AHR, such as TCDD and B[a]P, causes the

activation of multiple signaling pathways, although the mechanisms that connect these toxic

agents to their effects on a particular signaling pathway have not been characterized. Thus, even

though the available information indicates that the interaction of the ligand-activated AHR with

signal transduction events enhances the toxicity of the ligand, the understanding of the

connections between agent, signaling mechanism and toxic outcome remains poor.

Exposure of multiple rodent cell lines to TCDD results in an essentially

immediate increase in protein kinase C levels and activity in cellular membranes (9-12). TCDD

specifically activates tyrosine kinases associated with the EGFR (11;13) and induces the

association of adaptor proteins such as SHC, GRB2, and SOS with EGFR (14). While TCDD is not an EGFR ligand (15), it seems that cross-talk between AHR and EGFR signaling is critical for TCDD-induced developmental toxicity (16) and hepatocarcinogenicity (17). Farther downstream in the MAP kinase pathway, TCDD activates the expression of the HRAS gene coding for the small GTPase p21RAS, probably as the result of the transmission of receptor and

non-receptor phosphotyrosine kinase signals from the EGFR adaptor complex (18). As a

consequence, RAS GTP binding activity is increased in adipose tissues treated with TCDD. In a

different system altogether, consisting of rat vascular smooth muscle cells in culture, Hras

expression is induced by exposure to B[a]P, yet a different AHR ligand (19), perhaps in a tissue-

specific manner (20). Gene expression analyses in human hepatoma cells using global

microarray experiments confirm the activation of the RAS MAP kinase pathway by TCDD (21).

Additionally, AHR-independent induction of K-RAS activity by TCDD in mouse lung tissues

suggests that there are multiple mechanisms by which AHR ligands influence signal transduction

pathways (22).

The ERK family of proteins, in addition to the JNKs and p38, are serine/threonine

kinases of the MAP kinase family. ERK activity is stimulated in human epithelial cells treated

with B[a]P (23) and in endocervical cells from TCDD-exposed macaque monkeys (24). TCDD

also induces ERK and JNK phosphorylation in cell lines lacking a functional AHR, with kinetics

indicative of a so-called non-genomic effect (7). Upstream signaling cascades of MAPKKKs

and MAPKKs regulate the downstream MAP kinases (25). MEK-1 and -2 are the upstream

MAPKKs for ERK proteins, while the MEKs themselves are regulated by the upstream

MAPKKK RAF-1, among others. AHR agonists may stimulate ERK activity by inducing EGFR

activation and the subsequent activation of RAS; however, an AHR-dependent interaction of

RAS with RAF-1, which would be responsible for RAF-1 activation and progression of the signal to MEK and ERK (26), has yet to be found. Other EGFR downstream signaling molecules involved in ERK activation, such as PK-C, PLC-γ, and PI-3K, may also be responsive to TCDD. In fact, several studies have shown that activation of the PK-C pathway is required for AHR activation and CYP1A1 expression (12;27-29). Exposure of cell cultures to either

TCDD or B[a]P leads to increases in intracellular calcium levels as well as extracellular calcium fluxes (29-31), possibly through activation of PLC-γ, a major regulator of intracellular calcium stores. In addition to the effects of AHR ligands on MAPK signaling cascades, recent studies have shown that AHR transcriptional activity is dependent upon ERK and JNK activation in a cell-lineage and gene-specific manner (8).

Still other signaling pathways are affected following exposure to AHR ligands. The tyrosine kinase activity of c-SRC is triggered in multiple in vitro and in vivo systems in response to AHR activation. SRC may be activated through several alternative pathways, including by signals initiated from cell surface receptors such as EGFR, or from G protein-coupled receptors and intracellular receptors (32;33), as well as via ligand-induced disruption of the AHR-HSP90 complex, to which SRC may be functionally associated (34-36). TCDD fails to suppress the differentiation of c-SRC-deficient MEF cells, but not of wild type cells (37;38). Furthermore, the overall toxicity of TCDD is dependent in part on the activation of c-SRC, as SRC-deficient,

TCDD-treated mice exhibit reduced toxicity (36). Lastly, a signal transduction pathway that involves the c-SRC protein tyrosine kinase has been shown to be responsible for the activation of the AHR by pharmacologic agents in a ligand-independent manner by mechanisms unlike those exerted by high-affinity ligands (39).

AHR agonists activate immediate-early response genes

As outlined above, many AHR ligands activate signaling cascades initiated and propagated by trans-membrane and intracellular ion fluxes, and by protein kinase and phosphatase activation. Transduction of such signals to the nucleus of quiescent cells induces the expression of multiple immediate-early response genes, including MYC, MYB, and members

of the FOS and JUN families, which coordinate the expression of additional genes required for

subsequent cell cycle progression (40). The untimely expression of such genes and the

successive cycling of normally quiescent cells may in part explain the ability of TCDD and other

ligands of the AHR to act as powerful tumor promoters and carcinogens (41).

AHR ligand-dependent activation of the c-MYC gene in human breast cancer cells results

from the binding of an AHR-RelA protein complex to an NF-κB DNA binding element in the c-

MYC promoter (42). The AHR may therefore contribute indirectly in this context to entry of

these cells into the cell cycle. NF-κB controls many physiological functions adversely affected

by PAHs, and the formation of AHR-RelA complexes may also help to explain some of the

adverse toxicological outcomes of AHR ligands such as immune suppression, thymic involution,

hyperkeratosis, and carcinogenesis. In addition to the effect of AHR activation on NF-κB-

mediated transcriptional activity, the formation of AHR-RelA complexes also results in the

functional repression of AHR/ARNT activities (43;44). Increased p50 homodimer binding to

NF-κB sites may also be explained by sequestration of RelA by the AHR (45).

The expression of additional immediate-early genes, specifically members of the FOS

and JUN families of protooncogenes, has been shown to be induced by AHR agonists, with a

resulting increase in AP-1 DNA-binding activity. This effect has been observed in multiple liver

cell types (24;46;47), but not in all (48). In other cells, such as LPS-activated B cells, TCDD

downregulates AP-1 expression (49), suggesting a cell type-dependent effect of AHR ligands on immediate-early protooncogene induction. The induction of c-Jun and Jun-D expression by the

AHR appears to result from AHR-complex binding sites in the promoters of these genes. In contrast, c-Fos induction by TCDD is dependent on a SRE motif in its promoter, and is not dependent on the presence of the AHR (50). Activation of the ERK MAP kinases leads to ELK-

1 phosphorylation and to binding of the ternary ELK-1/TCF complex to the SRE motif (51), potentially connecting ERK activation by TCDD to AHR-independent downstream effects on immediate-early gene expression (8); however, neither ELK-1 phosphorylation nor formation of the ternary complex have been observed after AHR activation.

Ligand-independent cell cycle control through the AHR

Cell cycle progression, through the controlled process of DNA replication and cell division, is initiated in quiescent cells by mitogen stimulation. Typically, eukaryotic cells progress through cell cycle stages by the activities of cyclins, CDKs, and CKIs, which are

responsible for the ordered transition from one phase of the cycle to the next. Their expression

and activities are in turn controlled and modulated by members of the RB and E2F families of

proteins (52-54).

It has long been recognized that the AHR plays a role in cell cycle regulation. AH

receptor-null mice exhibit epidermal hyperplasia and hyperproliferation of hair follicles,

hyperproliferation of liver blood vessels, and an age-dependent incidence of adenocarcinomas of

liver and lung; paradoxically, these mice also show accelerated rates of apoptosis in the liver

(55). Fetal fibroblasts from AHR-null mice show slower proliferation rates and increased

apoptosis, concomitant with the accumulation of cells in G2/M, possibly due to altered expression of the G2/M kinases CDC2 and PLK. The increase in apoptosis of AHR-null cells

was attributed to increased levels of TGF-β, an inhibitor of cell proliferation (56). Retinoic acid

levels are increased in the livers of AHR-null mice, possibly due to the absence of some AHR-

regulated P450 enzyme, and this elevation in retinoic acid content is thought to cause the higher levels of TGF-β (55). However, other studies in AHR-null embryo fibroblasts have shown that p300-dependent stimulation of DNA synthesis by the adenovirus E1A protein does not take place in the absence of AHR, suggesting the possibility that the AHR exerts its influence on cell cycle regulation by other mechanisms (57).

Evidence that the absence of the AHR results in prolongation of the cell cycle has grown in recent years. Both AHR-negative mouse hepatoma Hepa1c1c7 cell variants (58), and human

HepG2 hepatoma cells transfected with AHR siRNA (59) show a slower progression through the cell cycle, attributed to a delay in the transition from G1 to S. These results suggest that the AHR plays an endogenous role in the promotion of cell cycle progression and that this role is independent of activation by exogenous ligands. This conclusion is significantly strengthened by

the findings that, in the absence of ligand, expression of a constitutively active AHR variant in

transgenic mice causes pro-proliferative effects, such as induction of stomach tumors (60), and

promotion of hepatocarcinogenesis (61). Paradoxically, expression of the same variant AHR in

human Jurkat cells causes growth inhibition and apoptosis (62).

AHR-mediated inhibition of apoptosis

Paradoxically, TCDD is one of the most potent tumor promoters known in animal model

systems, including the liver and the skin, in two-stage carcinogenesis assays (63;64). Tumor

promoters are believed to act by affecting the rate of proliferation, terminal differentiation, or

death of tumor precursor cells. One widely accepted mechanism of tumor promotion/progression

is the inhibition of apoptosis (65). The capacity of AHR ligands such as TCDD to act as tumor

promoters, particularly in rodent liver, has been attributed to their ability to inhibit the apoptotic elimination of initiated cells bearing genotoxic lesions (41). However, the precise mechanisms responsible for this effect remain elusive, and likely differ with the organism, tissue, or cell type

examined. In DEN-initiated rats, both acute and chronic TCDD treatment results in an

approximate 10-fold decrease in the rate of apoptosis in preneoplastically transformed liver foci,

with no effect on the background rate of apoptosis in normal hepatocytes (66). The overall effect

of TCDD in this system is thus to accelerate the rate at which DNA-damaged cells convert to a

neoplastic phenotype (67). Stimulation of cell division in these assays is negligible (68); hence,

the primary effect of TCDD is the inhibition of apoptosis, which has also been shown to occur in

Myc transgenic mice (41) and in the promotion of ovarian tumors in rats (69). Absence of tumor

promotion by TCDD treatment in rat strains lacking a functional AHR suggests that the AH

receptor is required for this effect, which has been shown to include activation of MDM2 and

attenuation of p53 probably by increased ubiquitination (70;71).

Effects of TCDD on apoptosis have also been documented in cultured cells. TCDD

inhibits apoptosis in hepatocytes treated with UV light or 2-acetylaminofluorene, an effect that

was also attributed in part to attenuation of p53 activity (72;73). Apoptosis induced by growth

factor withdrawal in human epithelial cells is inhibited by TCDD treatment, in correlation with

activation of the EGF signaling pathway (74). Studies with AHR-null mice confirm the

importance of the AHR in tissue homeostasis, as hepatocytes from these mice exhibit accelerated

rates of apoptosis associated with increased production of TGF-β (55). In vitro cell populations

lacking the AH receptor also have higher rates of apoptotic death (56).

Cell cycle arrest induced by AHR ligands

There is a large body of evidence showing that exogenous AHR ligands, especially

TCDD, actually inhibit cell proliferation and induce cell cycle arrest in normally cycling cell populations [reviewed in (2)]; however, the mechanisms controlling this effect remain indistinct and ill-defined. TCDD was shown to inhibit DNA synthesis in confluent mouse epithelial cells

(75), in partially hepatectomized rat liver (76), and in rat primary hepatocytes (77) by mechanisms that were independent of TGF-β or the mitogenic activity of EGF. Similar observations were found in mice, where TCDD treatment inhibits liver regeneration by 50% to

75% following partial hepatectomy, concurrent with decreased CDK2 activity and cyclin E-

CDK2 binding (78). TCDD also inhibited 17β-estradiol-induced growth of MCF-7 human breast cancer cells, concomitantly with decreases in RB phosphorylation, cyclin D1 protein levels, and CDK-dependent kinase activities (79). In mouse intrathymic progenitor cells, TCDD blocked S-phase progression and caused persistent thymic atrophy (80). TCDD also inhibited proliferation of the fish hepatocellular carcinoma PLHC-1 cell line (81) and the androgen- induced proliferation of G0/G1-synchronized human prostate cancer LNCaP cells (82).

Treatment with TCDD also induced the AHR-dependent G1 arrest of SK-N-SH human neuronal

cells concomitant with the increased expression of p27 and the hypophosphorylation of RB (83).

Similar effects on the cell cycle and cell proliferation have been observed with AHR

ligands other than TCDD. B[a]P suppressed cell proliferation in Swiss mouse 3T3 cells (84).

Treatment of rat 5L hepatoma cells with several different flavonoids known to be AHR ligands

resulted in G1 arrest (85). The novel AHR agonist 2-(4-amino-3-methylphenyl)-5-

fluorobenzothiazole also induced an AHR-dependent cell cycle arrest in MCF-7 cells (86).

These results are consistent with findings in mouse skin carcinogenesis assays that showed that

TCDD pretreatment is anticarcinogenic under conditions of low doses of the tumor inducer

DMBA (87). Two components of essential oils, eugenol (4-allyl-2-methoxyphenol) and isoeugenol (2-methoxy-4-propenylphenol), induce a cell cycle arrest in HaCaT cells mediated by the AHR (88). Interestingly, eugenol has been found to induce growth suppression in a malignant melanoma cell line through the inhibition of E2F-1 transcriptional activity (89). AHR is also expressed at high levels in pancreatic cancer tissues, and TCDD and other AHR agonists

inhibit pancreatic cancer cell growth (90).

Like treatment with flavonoids, TCDD treatment of asynchronously growing 5L

hepatoma cell cultures leads to a delay in G1 to S-phase progression (91). This effect depends on the presence of the AH receptor, since variant 5L clones lacking AHR expression do not show delayed G1 to S progression (91-93), and expression of ectopic AHR in these variant cell lines

reconstitutes the ability of TCDD to delay cell cycle progression (94). Several observations may

explain these findings. TCDD was found to induce expression of the p27Kip1 CDK inhibitor in an

AHR-dependent manner, as the effect was lost in cells lacking AHR expression (95). The observed induction of p27Kip1 occurred concurrently with reduced cell proliferation, which was reversed by transient expression of a Kip1 antisense RNA. Independent studies have shown that

genes of the AHR gene battery are regulated in a cell cycle-dependent manner, and that the

greatest induction of CYP1A1 by TCDD occurs during late G1 to early S-phases (96). Serum-

mediated release of G0/G1-synchronized 5L cells into the cell cycle results in transient activation

of the AHR and subsequent CYP1A1 expression, followed by progression of the cells into S-

phase (97). This is in contrast to treatment of the same cells with TCDD, which results in

Kip1 sustained AHR activation, increased p27 expression, and G1 arrest. Simultaneous treatment

of G0/G1-synchronized 5L cells with serum and the CYP1A1 suicide substrate 1-PP triggers

sustained AHR activation and p27Kip1 induction, similar to the action of TCDD alone. Thus,

CYP1A1 activity appears to negatively regulate the length of AHR activation through the metabolism of an as yet unknown AHR agonist and CYP1A1 substrate, allowing cells to progress through the cell cycle in response to serum stimulation. It is possible that the lack of metabolism of TCDD and other persistent AHR agonists is responsible not only for the sustained induction of AHR activation but also for the induction of cell cycle arrest.

While these observations provide a plausible mechanistic rationale for the role of AHR in cell cycle regulation, additional data suggest that other mechanisms are equally important.

Several reports have shown that the AHR forms complexes with the RB protein, detected by yeast two-hybrid assays as well as by co-immunoprecipitation (3;98). RB acts as a negative regulator of cell cycle progression by preventing the expression of genes required for cell cycle entry through the inhibition of E2F-dependent transcriptional activity. At least two AH receptor domains interact with RB, including an LXCXE motif common to many RB-interacting proteins

(99), and a glutamine-rich region within the transactivation domain of the receptor. Further analysis of the biological consequences of this interaction revealed that the AHR acts in synergy with RB to repress E2F-dependent gene expression and to slow down cell cycle progression, particularly in the G1-to-S phase transition (98). In addition, work in human C33A cells, which are insensitive to RB-mediated active gene repression, has shown that the combination of RB with AHR or BRG-1 restores repression of CDK2 and cyclin A and causes cell cycle arrest

(100). These results suggest that AHR activation may inhibit cell cycle progression not only by inducing p27Kip1 expression to directly inhibit CDK2 activity and therefore RB inactivation, but

also by directly interacting with RB to repress expression of genes required for entry into S-

phase and cell cycle progression. Recent data from our laboratory ((101) and data presented in

Chapter 3) and from Elferink and colleagues (102) have confirmed this conclusion. AHR activation in cycling Hepa-1c1c7 cells results in the accumulation of cells in G1 (see Chapter 3 and ref. (101)). Preceding this effect there is a significant increase in the expression of p27Kip1 and reduction in the expression of specific E2F-regulated genes, including Cyclin E, CDK2,

DNA polymerase α, and DHFR. Chromatin immunoprecipitation assays showed that activation of the AHR by ligand causes it to be recruited to the promoters of these genes, while excluding p300 from the same promoters. These data suggest a novel mechanism by which the AHR, a potent transcriptional activator, may act as a repressor of transcription through the formation of specific protein-protein interactions, and in doing so, induce G1 arrest in normally cycling cells by preventing the expression of genes required for S-phase progression.

Conclusions

As briefly described above, the AH receptor pathway cross-talks with many cellular signal transduction cascades and ultimately leads cells in alternative directions of proliferation, cell cycle arrest, or apoptosis. It appears that whether one or the other outcome is reached might depend on the presence or absence of ligand, but the specific mechanisms by which ligands inhibit apoptosis or promote proliferation in preneoplastic liver cells, or in any system for that matter, are unknown. Results presented in Chapters 3 and 4 of this dissertation indicate that interactions of the AHR with the RB/E2F axis are a critical element of these mechanisms. RB suppresses apoptosis as a result of the repression of a distinct set of proapoptotic E2F target genes, which includes Apaf-1 and several caspases (103). Excess E2F-1 expression unchecked by RB forces the cells to enter S-phase and promotes p53-dependent and independent apoptosis

(103). This is a specific function of E2F-1 that depends on its binding to RB at a second binding site located entirely in the C-terminal domain of RB. E2F-1/RB complexes formed through this

site have low affinity for DNA, but their interaction is sufficient for RB to repress E2F-1-induced apoptosis. It follows that in cells lacking RB, E2F-1 proapoptotic activity is unchecked

(104;105). In addition, E2F-1 becomes proapoptotic in response to DNA damage as a result of phosphorylation by ATM/Chk2, thereby blocking its interaction with the RB C-terminal site and causing the stabilization of E2F-1 (106).

Interaction of AHR with RB mediates active repression of E2F-responsive genes, thereby cooperating in the inhibition of cell cycle progression. In addition, the activated AHR affects

E2F transcriptional activity in the absence of RB binding (see Chapter 4 and ref. (101). The proapoptotic activity of E2F-1 suggests that its deregulation may constitute an oncogenic stress that targets pre-malignant cells to undergo apoptosis, thus preventing tumor development

(105;107;108). It is therefore proposed that, by signaling through the RB/E2F-1 axis, which can induce cell cycle progression and proliferation, cell cycle arrest, apoptosis, and DNA damage repair, the activated AH receptor plays a critical role in all of these processes. This working hypothesis, summarized in Figure 1, suggests that the AH receptor has a yin-yang role in cell cycle regulation, whereby under some circumstances its activation is pro-proliferative and under others, anti-proliferative. For example, if environmental mitogens induce unscheduled cell cycle progression, activation of the AHR would cause its translocation to the nucleus, where it would function as an environmental checkpoint in cooperation with RB, inhibit S-phase gene expression by interacting with RB/E2F-1/DP1 complexes, and promote cell cycle arrest. This aspect of AHR biology is explored in Chapter 3 as it relates to RB, and in Chapter 4 as it relates to E2F proteins. On the other hand, under abnormal conditions of loss of RB or of DNA damage, stabilization of E2F-1 would upregulate its pro-apoptotic functions, but activation of

AHR under these conditions would cause it to interact with E2F-1 and block its apoptotic

properties, causing a pro-proliferative, anti-apoptotic response. This yin-yang activity of the

AHR and its ligand could be at the heart of the abnormal proliferative and apoptotic responses that characterize the carcinogenicity of TCDD and other AH receptor ligands. While the anti- apoptotic activities of the AHR are not the primary focus of the bulk of the following thesis, data relevant to this aspect of the overall working hypothesis are presented in the Appendix.

Acknowledgments

We thank Dr. Ying Xia for a critical reading of the manuscript.

References

1. Hankinson O. 1995. The aryl hydrocarbon receptor complex. Annu.Rev.Pharmacol.Toxicol. 35: 307-340

2. Puga A, Xia Y, and Elferink C. 2002. Role of the aryl hydrocarbon receptor in cell cycle regulation. Chem.Biol.Interact. 141: 117-130

3. Ge N-L and Elferink CJ. 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblatoma protein. J.Biol.Chem. 273: 22708-22713

4. Wang F, Wang W, and Safe S. 1999. Regulation of constitutive gene expression through interactions of Sp1 protein with the nuclear aryl hydrocarbon receptor complex. Biochemistry 38: 11490-11500

5. Nguyen TA, Hoivik D, Lee JE, and Safe S. 1999. Interactions of nuclear receptor coactivator/corepressor proteins with the aryl hydrocarbon receptor complex. Arch.Biochem.Biophys. 367: 250-257

6. Wang S and Hankinson O. 2002. Functional involvement of the Brahma/SWI2-related gene 1 protein in cytochrome P4501A1 transcription mediated by the aryl hydrocarbon receptor complex. J.Biol.Chem. 277: 11821-11827

7. Tan Z, Chang X, Puga A, and Xia Y. 2002. Activation of mitogen-activated protein kinases (MAPKs) by aromatic hydrocarbons: role in the regulation of aryl hydrocarbon receptor (AHR) function. Biochem.Pharmacol. 64: 771-780

8. Tan Z, Huang M, Puga A, and Xia Y. 2004. A Critical Role for MAP Kinases in the Control of Ah Receptor Complex Activity. Toxicol.Sci.

9. Bombick DW, Jankun J, Tullis K, and Matsumura F. 1988. 2,3,7,8-tetrachlorodibenzo-p- dioxin causes increases in expression of c-erb-A and levels of protein-tyrosine kinases in selected tissues of responsive mouse strains. Proc.Natl.Acad.Sci.USA 85: 4128-4132

10. Bombick DW, Madhukar BV, Brewster DW, and Matsumura F. 1985. TCDD (2,3,7,8- tetrachlorodibenzo-p-dioxin) causes increases in protein kinases particularly protein kinase C in the hepatic plasma membrane of the rat and the guinea pig. Biochem.Biophys.Res.Comm. 127: 296-302

11. Madhukar BV, Ebner K, Matsumura F, Bombick DW, Brewster DW, and Kawamoto T. 1988. 2,3,7,8-Tetrachlorodibenzo-p-dioxin causes an increase in protein kinases associated with epidermal growth factor receptor in the hepatic plasma membrane. J.Biochem.Toxicol. 3: 261-277

12. Carrier F, Owens RA, Nebert DW, and Puga A. 1992. Dioxin-dependent activation of murine Cyp1a-1 gene transcription requires protein kinase C-dependent phosphorylation. Mol Cell Biol 12: 1856-1863

13. Madhukar BV, Brewster DW, and Matsumura F. 1984. Effects of in vivo-administered 2,3,7,8-tetrachlorodibenzo-p- dioxin on receptor binding of epidermal growth factor in the hepatic plasma membrane of rat, guinea pig, mouse, and hamster. Proc.Natl.Acad.Sci.USA 81: 7407-7411

14. Park R, Kim DH, Kim MS, So HS, Chung HT et al. 1998. Association of Shc, Cbl, Grb2, and Sos following treatment with 2,3,7,8- tetrachlorodibenzo-p-dioxin in primary rat hepatocytes. Biochem.Biophys.Res.Commun. 253: 577-581

15. Sewall CH, Clark GC, and Lucier GW. 1995. TCDD reduces rat hepatic epidermal growth factor receptor: comparison of binding, immunodetection, and autophosphorylation. Toxicol.Appl.Pharmacol. 132: 263-272

16. Partanen AM, Alaluusua S, Miettinen PJ, Thesleff I, Tuomisto J et al. 1998. Epidermal growth factor receptor as a mediator of developmental toxicity of dioxin in mouse embryonic teeth. Lab Invest 78: 1473-1481

17. Sewall CH, Lucier GW, Tritscher AM, and Clark GC. 1993. TCDD-mediated changes in hepatic epidermal growth factor receptor may be a critical event in the hepatocarcinogenic action of TCDD. Carcinogenesis 14: 1885-1893

18. Enan E and Matsumura F. 1994. Significance of TCDD-induced changes in protein phosphorylation in the adipocyte of male guinea pigs. J.Biochem.Toxicol. 9: 159-170

19. Bral CM and Ramos KS. 1997. Identification of benzo[a]pyrene-inducible cis-acting elements within c- Ha-ras transcriptional regulatory sequences. Mol.Pharmacol. 52: 974- 982

20. Parrish AR, Fisher R, Bral CM, Burghardt RC, Gandolfi AJ et al. 1998. Benzo(a)pyrene- induced alterations in growth-related gene expression and signaling in precision-cut adult rat liver and kidney slices. Toxicol.Appl.Pharmacol. 152: 302-308

21. Puga A, Maier A, and Medvedovic M. 2000. The transcriptional signature of dioxin in human hepatoma HepG2 cells. Biochem.Pharmacol. 60: 1129-1142

22. Ramakrishna G and Anderson LM. 1998. Levels and membrane localization of the c-K- ras p21 protein in lungs of mice of different genetic strains and effects of 2,3,7,8- tetrachlorodibenzo-p-dioxin (TCDD) and Aroclor 1254. Carcinogenesis 19: 463-470

23. Jyonouchi H, Sun S, Iijima K, Wang M, and Hecht SS. 1999. Effects of anti-7,8- dihydroxy-9,10-epoxy-7,8,9,10- tetrahydrobenzo[a]pyrene on human small airway epithelial cells and the protective effects of myo-inositol. Carcinogenesis 20: 139-145

24. Enan E, El Sabeawy F, Scott M, Overstreet J, and Lasley B. 1998. Alterations in the growth factor signal transduction pathways and modulators of the cell cycle in endocervical cells from macaques exposed to TCDD. Toxicol.Appl.Pharmacol. 151: 283- 293

25. Cobb MH and Goldsmith EJ. 2000. Dimerization in MAP-kinase signaling. Trends Biochem.Sci. 25: 7-9

26. Barbacid M. 1987. Ras genes. Annu.Rev.Biochem. 56: 779-827

27. Chen YH and Tukey RH. 1996. Protein kinase C modulates regulation of the CYP1A1 gene by the aryl hydrocarbon receptor. J.Biol.Chem. 271: 26261-26266

28. Long WP, Pray-Grant M, Tsai JC, and Perdew GH. 1998. Protein kinase C activity is required for aryl hydrocarbon receptor pathway-mediated signal transduction. Mol.Pharmacol. 53: 691-700

29. Tannheimer SL, Barton SL, Ethier SP, and Burchiel SW. 1997. Carcinogenic polycyclic aromatic hydrocarbons increase intracellular Ca2+ and cell proliferation in primary human mammary epithelial cells. Carcinogenesis 18: 1177-1182

30. Puga A, Bohm J, Hoffer A, Leikauf GD, Shertzer HG, and Zhou S. 1995. Dioxin alters calcium homeostasis and the regulation of arachidonate metabolism in mouse hepatoma cells. Proc.15th Int.Symp.Chlor.Dioxins 25: 381-386

31. Puga A, Hoffer A, Zhou S, Bohm JM, Leikauf GD, and Shertzer HG. 1997. Sustained increase in intracellular free calcium and activation of cyclooxygenase-2 expression in mouse hepatoma cells treated with dioxin. Biochem.Pharmacol. 54: 1287-1296

32. Cao W, Luttrell LM, Medvedev AV, Pierce KL, Daniel KW et al. 2000. Direct binding of activated c-Src to the beta 3-adrenergic receptor is required for MAP kinase activation. J.Biol.Chem. 275: 38131-38134

33. Watters JJ, Chun TY, Kim YN, Bertics PJ, and Gorski J. 2000. Estrogen modulation of prolactin gene expression requires an intact mitogen-activated protein kinase signal transduction pathway in cultured rat pituitary cells. Mol.Endocrinol. 14: 1872-1881

34. Enan E and Matsumura F. 1995. Evidence for a second pathway in the action mechanism of 2,3,7,8- tetrachlorodibenzo-p-dioxin (TCDD). Significance of Ah-receptor mediated activation of protein kinase under cell-free conditions. Biochem.Pharmacol. 49: 249-261

35. Enan E, Dunlap DY, and Matsumura F. 1998. Use of c-Src and c-Fos knockout mice for the studies on the role of c- Src kinase signaling in the expression of toxicity of TCDD. J.Biochem.Mol.Toxicol. 12: 263-274

36. Dunlap DY, Ikeda I, Nagashima H, Vogel CF, and Matsumura F. 2002. Effects of src- deficiency on the expression of in vivo toxicity of TCDD in a strain of c-src knockout mice procured through six generations of backcrossings to C57BL/6 mice. Toxicology 172: 125-141

37. Vogel CF, Zhao Y, Wong P, Young NF, and Matsumura F. 2003. The use of c-src knockout mice for the identification of the main toxic signaling pathway of TCDD to induce wasting syndrome. J.Biochem.Mol.Toxicol. 17: 305-315

38. Vogel CF and Matsumura F. 2003. Interaction of 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) with induced adipocyte differentiation in mouse embryonic fibroblasts (MEFs) involves tyrosine kinase c-Src. Biochem.Pharmacol. 66: 1231-1244

39. Backlund M and Ingelman-Sundberg M. 2005. Regulation of aryl hydrocarbon receptor signal transduction by protein tyrosine kinases. Cell Signal. 17: 39-48

40. Kohn KW. 1999. Molecular interaction map of the mammalian cell cycle control and DNA repair systems. Mol.Biol.Cell 10: 2703-2734

41. Schwarz M, Buchmann A, Stinchcombe S, Kalkuhl A, and Bock K. 2000. Ah receptor ligands and tumor promotion: survival of neoplastic cells. Toxicol.Lett. 112-113: 69-77

42. Kim DW, Gazourian L, Quadri SA, Romieu-Mourez R, Sherr DH, and Sonenshein GE. 2000. The RelA NF-kappaB subunit and the aryl hydrocarbon receptor (AhR) cooperate to transactivate the c-myc promoter in mammary cells. Oncogene 19: 5498-5506

43. Ke S, Rabson AB, Germino JF, Gallo MA, and Tian Y. 2001. Mechanism of Suppression of Cytochrome P-450 1A1 Expression by Tumor Necrosis Factor-alpha and Lipopolysaccharide. J.Biol.Chem. 276: 39638-39644

44. Tian Y, Ke S, Denison MS, Rabson AB, and Gallo MA. 1999. Ah receptor and NF- kappaB interactions, a potential mechanism for dioxin toxicity. J.Biol.Chem. 274: 510- 515

45. Puga A, Barnes SJ, Chang C, Zhu H, Nephew KP et al. 2000. Activation of transcription factors activator protein-1 and nuclear factor-kappaB by 2,3,7,8-Tetrachlorodibenzo-p- dioxin. Biochem.Pharmacol. 59: 997-1005

46. Puga A, Nebert DW, and Carrier F. 1992. Dioxin induces expression of c-fos and c-jun proto-oncogenes and a large increase in transcription factor AP-1. DNA Cell Biol. 11: 269-281

47. Ashida H, Nagy S, and Matsumura F. 2000. 2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD)-induced changes in activities of nuclear protein kinases and phosphatases affecting DNA binding activity of c-Myc and AP-1 in the livers of guinea pigs. Biochem.Pharmacol. 59: 741-751

48. Gohl G, Lehmkoster T, Munzel PA, Schrenk D, Viebahn R, and Bock KW. 1996. TCDD-inducible plasminogen activator inhibitor type 2 (PAI-2) in human hepatocytes, HepG2 and monocytic U937 cells. Carcinogenesis 17: 443-449

49. Suh J, Jeon YJ, Kim HM, Kang JS, Kaminski NE, and Yang KH. 2002. Aryl hydrocarbon receptor-dependent inhibition of AP-1 activity by 2,3,7,8- tetrachlorodibenzo-p-dioxin in activated B cells. Toxicol.Appl.Pharmacol. 181: 116-123

50. Hoffer A, Chang C-Y, and Puga A. 1996. Dioxin induces fos and jun gene expression by Ah receptor dependent- and independent- pathways. Toxicol.Appl.Pharmacol. 141: 238- 247

51. Gille H, Kortenjann M, Strahl T, and Shaw PE. 1996. Phosphorylation-dependent formation of a quaternary complex at the c-fos SRE. Mol.Cell.Biol. 16: 1094-1102

52. Coqueret O. 2002. Linking cyclins to transcriptional control. Gene 299: 35-55

53. Murray AW. 2004. Recycling the cell cycle: cyclins revisited. Cell 116: 221-234

54. Trimarchi JM and Lees JA. 2002. Sibling rivalry in the E2F family. Nat.Rev.Mol.Cell Biol. 3: 11-20

55. Gonzalez FJ and Fernandez-Salguero P. 1998. The aryl hydrocarbon receptor: studies using the AHR-null mice. Drug Metab Dispos. 26: 1194-1198

56. Elizondo G, Fernandez-Salguero P, Sheikh MS, Kim GY, Fornace AJ et al. 2000. Altered cell cycle control at the G(2)/M phases in aryl hydrocarbon receptor-null embryo fibroblast. Mol.Pharmacol. 57: 1056-1063

57. Tohkin M, Fukuhara M, Elizondo G, Tomita S, and Gonzalez FJ. 2000. Aryl hydrocarbon receptor is required for p300-mediated induction of DNA synthesis by adenovirus E1A. Mol.Pharmacol. 58: 845-851

58. Ma Q and Whitlock JPJ. 1996. The aromatic hydrocarbon receptor modulates the Hepa 1c1c7 cell cycle and differentiated state independently of dioxin. Mol.Cell Biol. 16: 2144- 2150

59. Abdelrahim M, Smith R, III, and Safe S. 2003. Aryl Hydrocarbon Receptor Gene Silencing with Small Inhibitory RNA Differentially Modulates Ah-Responsiveness in MCF-7 and HepG2 Cancer Cells. Mol.Pharmacol. 63: 1373-1381

60. Andersson P, McGuire J, Rubio C, Gradin K, Whitelaw ML et al. 2002. A constitutively active dioxin/aryl hydrocarbon receptor induces stomach tumors. Proc.Natl.Acad.Sci.U.S.A 99: 9990-9995

61. Moennikes O, Loeppen S, Buchmann A, Andersson P, Ittrich C et al. 2004. A constitutively active dioxin/aryl hydrocarbon receptor promotes hepatocarcinogenesis in mice. Cancer Res. 64: 4707-4710

62. Ito T, Tsukumo S, Suzuki N, Motohashi H, Yamamoto M et al. 2004. A constitutively active arylhydrocarbon receptor induces growth inhibition of jurkat T cells through changes in the expression of genes related to apoptosis and cell cycle arrest. J.Biol.Chem. 279: 25204-25210

63. Pitot HC, Goldsworthy T, Campbell HA, and Poland A. 1980. Quantitative evaluation of the promotion by 2,3,7,8- tetrachlorodibenzo-p-dioxin of hepatocarcinogenesis from diethylnitrosamine. Cancer Res. 40: 3616-3620

64. Dragan YP and Schrenk D. 2000. Animal studies addressing the carcinogenicity of TCDD (or related compounds) with an emphasis on tumour promotion. Food Addit.Contam 17: 289-302

65. Roberts RA, Nebert DW, Hickman JA, Richburg JH, and Goldsworthy TL. 1997. Perturbation of the mitosis/apoptosis balance: a fundamental mechanism in toxicology. Fundam.Appl.Toxicol. 38: 107-115

66. Stinchcombe S, Buchmann A, Bock KW, and Schwarz M. 1995. Inhibition of apoptosis during 2,3,7,8-tetrachlorodibenzo-p- dioxin-mediated tumour promotion in rat liver. Carcinogenesis 16: 1271-1275

67. Luebeck EG, Buchmann A, Stinchcombe S, Moolgavkar SH, and Schwarz M. 2000. Effects of 2,3,7,8-tetrachlorodibenzo-p-dioxin on initiation and promotion of GST-P- positive foci in rat liver: A quantitative analysis of experimental data using a stochastic model. Toxicol.Appl.Pharmacol. 167: 63-73

68. Buchmann A, Stinchcombe S, Korner W, Hagenmaier H, and Bock KW. 1994. Effects of 2,3,7,8-tetrachloro- and 1,2,3,4,6,7,8- heptachlorodibenzo-p-dioxin on the proliferation of preneoplastic liver cells in the rat. Carcinogenesis 15: 1143-1150

69. Davis BJ, Mccurdy EA, Miller BD, Lucier GW, and Tritscher AM. 2000. Ovarian tumors in rats induced by chronic 2,3,7,8-tetrachlorodibenzo-p-dioxin treatment. Cancer Res. 60: 5414-5419

70. Viluksela M, Bager Y, Tuomisto JT, Scheu G, Unkila M et al. 2000. Liver tumor- promoting activity of 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in TCDD-sensitive and TCDD-resistant rat strains. Cancer Res. 60: 6911-6920

71. Paajarvi G, Viluksela M, Pohjanvirta R, Stenius U, and Hogberg J. 2005. TCDD activates Mdm2 and attenuates the p53 response to DNA damaging agents. Carcinogenesis 26: 201-208

72. Worner W and Schrenk D. 1996. Influence of liver tumor promoters on apoptosis in rat hepatocytes induced by 2-acetylaminofluorene, ultraviolet light, or transforming growth factor beta 1. Cancer Res. 56: 1272-1278

73. Schrenk D, Schmitz HJ, Bohnenberger S, Wagner B, and Worner W. 2004. Tumor promoters as inhibitors of apoptosis in rat hepatocytes. Toxicol.Lett. 149: 43-50

74. Davis JW, Melendez K, Salas VM, Lauer FT, and Burchiel SW. 2000. 2,3,7,8- Tetrachlorodibenzo-p-dioxin (TCDD) inhibits growth factor withdrawal-induced apoptosis in the human mammary epithelial cell line, MCF-10A. Carcinogenesis 21: 881- 886

75. Gierthy JF and Crane D. 1984. Reversible inhibition of in vitro epithelial cell proliferation by 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicol.Appl.Pharmacol. 74: 91-98

76. Bauman JW, Goldsworthy TL, Dunn CS, and Fox TR. 1995. Inhibitory effects of 2,3,7,8- tetrachlorodibenzo-p-dioxin on rat hepatocyte proliferation induced by 2/3 partial hepatectomy. Cell Prolif. 28: 437-451

77. Hushka DR and Greenlee WF. 1995. 2,3,7,8-Tetrachlorodibenzo-p-dioxin inhibits DNA synthesis in rat primary hepatocytes. Mutat.Res. 333: 89-99

78. Mitchell KA, Lockhart CA, Huang G, and Elferink CJ. 2006. Sustained aryl hydrocarbon receptor activity attenuates liver regeneration. Mol.Pharmacol. 70: 163-170

79. Wang W, Smith IR, and Safe S. 1998. Aryl hydrocarbon receptor-mediated antiestrogenicity in MCF-7 cells: modulation of hormone-induced cell cycle enzymes. Arch.Biochem.Biophys. 356: 239-248

80. Laiosa MD, Wyman A, Murante FG, Fiore NC, Staples JE et al. 2003. Cell proliferation arrest within intrathymic lymphocyte progenitor cells causes thymic atrophy mediated by the aryl hydrocarbon receptor. J.Immunol. 171: 4582-4591

81. Hestermann EV, Stegeman JJ, and Hahn ME. 2002. Relationships among the cell cycle, cell proliferation, and aryl hydrocarbon receptor expression in PLHC-1 cells. Aquat.Toxicol. 58: 201-213

82. Barnes-Ellerbe S, Knudsen KE, and Puga A. 2004. 2,3,7,8-Tetrachlorodibenzo-p-dioxin blocks androgen-dependent cell proliferation of LNCaP cells through modulation of pRB phosphorylation. Mol.Pharmacol. 66: 502-511

83. Jin DQ, Jung JW, Lee YS, and Kim JA. 2004. 2,3,7,8-Tetrachlorodibenzo-p-dioxin inhibits cell proliferation through arylhydrocarbon receptor-mediated G1 arrest in SK-N- SH human neuronal cells. Neurosci.Lett. 363: 69-72

84. Vaziri C and Faller DV. 1997. A benzo[a]pyrene-induced cell cycle checkpoint resulting in p53-independent G1 arrest in 3T3 fibroblasts. J.Biol.Chem. 272: 2762-2769

85. Reiners JJ, Jr., Clift R, and Mathieu P. 1999. Suppression of cell cycle progression by flavonoids: dependence on the aryl hydrocarbon receptor. Carcinogenesis 20: 1561-1566

86. Trapani V, Patel V, Leong CO, Ciolino HP, Yeh GC et al. 2003. DNA damage and cell cycle arrest induced by 2-(4-amino-3-methylphenyl)-5-fluorobenzothiazole (5F 203, NSC 703786) is attenuated in aryl hydrocarbon receptor deficient MCF-7 cells. Br.J.Cancer 88: 599-605

87. Lesca P, Perrot N, and Peryt B. 1994. Modulating effects of 2,3,7,8-tetrachlorodibenzo-p- dioxin on skin carcinogenesis initiated by the weak inducer 7,12- dimethylbenz(a)anthracene. Drug Metabol.Drug Interact. 11: 37-57

88. Kalmes M, Neumeyer A, Rio P, Hanenberg H, Fritsche E, and Blomeke B. 2006. Impact of the arylhydrocarbon receptor on eugenol- and isoeugenol-induced cell cycle arrest in human immortalized keratinocytes (HaCaT). Biol.Chem. 387: 1201-1207

89. Ghosh R, Nadiminty N, Fitzpatrick JE, Alworth WL, Slaga TJ, and Kumar AP. 2005. Eugenol causes melanoma growth suppression through inhibition of E2F1 transcriptional activity. J.Biol.Chem. 280: 5812-5819

90. Koliopanos A, Kleeff J, Xiao Y, Safe S, Zimmermann A et al. 2002. Increased arylhydrocarbon receptor expression offers a potential therapeutic target for pancreatic cancer. Oncogene 21: 6059-6070

91. Wiebel FJ, Klose U, and Kiefer F. 1991. Toxicity of 2,3,7,8-tetrachlorodibenzo-p-dioxin in vitro: H4IIEC3-derived 5L hepatoma cells as a model system. Toxicol.Lett. 55: 161- 169

92. Gottlicher M, Cikryt P, and Wiebel FJ. 1990. Inhibition of growth by 2,3,7,8- tetrachlorodibenzo-p-dioxin in 5L rat hepatoma cells is associated with the presence of Ah receptor. Carcinogenesis 11: 2205-2210

93. Göttlicher M and Wiebel FJ. 1991. 2,3,7,8-Tetrachlorodibenzo-p-dioxin causes unbalanced growth in 5L rat hepatoma cells. Toxicol.Appl.Pharmacol. 111: 496-503

94. Weiss C, Kolluri SK, Kiefer F, and Gottlicher M. 1996. Complementation of Ah receptor deficiency in hepatoma cells: negative feedback regulation and cell cycle control by the Ah receptor. Exp.Cell Res. 226: 154-163

95. Kolluri SK, Weiss C, Koff A, and Göttlicher M. 1999. p27kip1 induction and inhibition of proliferation by the intracellular Ah receptor in developing thymus and hepatoma cells. Genes Dev. 13: 1742-1753

96. Santini RP, Myrand S, Elferink C, and Reiners JJ, Jr. 2001. Regulation of Cyp1a1 induction by dioxin as a function of cell cycle phase. J.Pharmacol.Exp.Ther. 299: 718- 728

97. Levine-Fridman A, Chen L, and Elferink CJ. 2004. Cytochrome P4501A1 promotes G1 phase cell cycle progression by controlling aryl hydrocarbon receptor activity. Mol.Pharmacol. 65: 461-469

98. Puga A, Barnes SJ, Dalton TP, Chang C, Knudsen ES, and Maier MA. 2000. Aromatic hydrocarbon receptor interaction with the retinoblastoma protein potentiates repression of E2F-dependent transcription and cell cycle arrest. J.Biol.Chem. 275: 2943-2950

99. Chan HM, Smith L, and La Thangue NB. 2001. Role of LXCXE motif-dependent interactions in the activity of the retinoblastoma protein. Oncogene 20: 6152-6163

100. Strobeck MW, Fribourg AF, Puga A, and Knudsen ES. 2000. Restoration of retinoblastoma mediated signaling to Cdk2 results in cell cycle arrest. Oncogene 19: 1857-1867

101. Marlowe JL, Knudsen ES, Schwemberger S, and Puga A. 2004. The aryl hydrocarbon receptor displaces p300 from E2F-dependent promoters and represses S phase-specific gene expression. J.Biol.Chem. 279: 29013-29022

102. Huang G and Elferink CJ. 2005. Multiple mechanisms are involved in Ah receptor- mediated cell cycle arrest. Mol.Pharmacol. 67: 88-96

103. Nahle Z, Polakoff J, Davuluri RV, McCurrach ME, Jacobson MD et al. 2002. Direct coupling of the cell cycle and cell death machinery by E2F. Nat.Cell Biol. 4: 859-864

104. Hallstrom TC and Nevins JR. 2003. Specificity in the activation and control of transcription factor E2F-dependent apoptosis. Proc.Natl.Acad.Sci.U.S.A 100: 10848- 10853

105. Stevens C and La Thangue NB. 2004. The emerging role of E2F-1 in the DNA damage response and checkpoint control. DNA Repair (Amst) 3: 1071-1079

106. Dick FA and Dyson N. 2003. pRB contains an E2F1-specific binding domain that allows E2F1-induced apoptosis to be regulated separately from other E2F activities. Mol.Cell 12: 639-649

107. Ginsberg D. 2002. E2F1 pathways to apoptosis. FEBS Lett. 529: 122-125

108. Bell LA and Ryan KM. 2004. Life and death decisions by E2F-1. Cell Death.Differ. 11: 137-142

Fig. 1. Postulated yin-yang role of the AH receptor in cell cycle regulation. A. Under normal conditions, that is, in quiescent or in normal cycling cells, RB/E2F-1 interactions downregulate S-phase genes and mitogens activate cell cycle progression; under these conditions, activation of aryl hydrocarbon receptor (AHR) causes its translocation to the nucleus where it functions as an environmental checkpoint in cooperation with RB/E2F, inhibiting cyclin

D,E/cdk-dependent RB phosphorylation, promoting repression of S-phase specific genes and causing cell cycle arrest. B. Under abnormal conditions of loss of RB or DNA damage, E2F-1 is stabilized by ATM/ATR- and chk2-dependent phosphorylation and up-regulates expression of pro-apoptotic genes, promoting apoptosis; under these conditions, activation of AHR causes it to interact with E2F-1 and block its apoptotic properties, causing an antiapoptotic, pro-proliferative response.

Figure 1

(A) Mitogens Cyclin E

Cyclin D/cdk Cyclin E Pol α Cell cycle Cdk2 progression AHR DHFR Cyclin A AHR Activation G1/S arrest RR MCM7

(B) Loss of RB DNA damage ATM

Chk1(?) ATM/ATR

Chk2 ATM G2/M arrest/ Chk1 apoptosis ? AHR ? Apaf-1 AHR Activation p73 Antiapoptosis ARF ?

Chapter III

The Aryl Hydrocarbon Receptor Displaces p300 From E2F-Dependent Promoters and Represses S-Phase Specific Gene Expression

Jennifer L. Marlowe, Erik S. Knudsen, Sandy Schwemberger, and Alvaro Puga*

*Corresponding author:

Department of Environmental Health and Center for Environmental Genetics University of Cincinnati Medical Center P.O. Box 670056 Cincinnati, OH 45267-0056 Telephone: 513-558-0916 E-mail: [email protected]

Keywords: TCDD, E2F, p300/CBP, gene expression, chromatin immunoprecipitation

Abstract

The environmental contaminant TCDD causes a wide range of toxic, teratogenic, and

carcinogenic effects. TCDD is a ligand for the AHR, a ligand-activated transcription factor,

believed to be the primary mediator of these effects. Activation of the AHR by TCDD also

elicits a variety of effects on cell cycle progression, ranging from proliferation to arrest. In this report, we have further characterized the role of the activated AHR in cell cycle regulation. In human mammary carcinoma MCF-7 and mouse hepatoma Hepa-1 cells, TCDD treatment decreased the number of cells in S-phase and caused the accumulation of cells in G1. In Hepa-1 cells, this effect correlated with the transcriptional repression of several E2F-regulated genes required for S-phase progression. AHR-mediated gene repression was dependent on its interaction with RB but was independent of its transactivation function, since AHR mutants lacking DNA binding or transactivation domains repressed E2F-dependent expression as effectively as wild type AHR. Overexpression of p300 suppressed RB-dependent gene repression, and this effect was reversed by TCDD. Chromatin immunoprecipitation assays showed that TCDD treatment caused recruitment of the AHR to E2F-dependent promoters with the concomitant displacement of p300. These results delineate a novel mechanism whereby the

AHR, a known transcriptional activator, also mediates gene repression by pathways involving

combinatorial interactions at E2F-responsive promoters, leading to the repression of E2F-

dependent, S-phase-specific genes. The Ah receptor seems to act as an environmental

checkpoint that senses exposure to environmental toxicants and responds by signaling cell cycle

inhibition.

Introduction

2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD) is the prototypical compound of a class of

environmental contaminants that include many HAHs and PAHs, and is a model for the outcome

of exposure to these compounds as well as co-planar polychlorinated biphenyls. Exposure to

TCDD results in a plethora of toxic and carcinogenic responses in animals, including liver toxicity (1), immunosuppression (2;3), reproductive and developmental dysfunction (4-6),

endometriosis (7;8), and tumor promotion (9). The most immediate exposure outcome in

humans is chloracne (10). Long-term effects in humans range from immunological and

reproductive perturbations (11) to cardiovascular disease (12;13) and cancer (14-18). TCDD

itself is poorly metabolized, which leads to biological accumulation and production of sustained

effects (19).

While the in vivo effects of TCDD are wide-ranging, the in vitro effects are just as varied

and often contradictory, affecting cell proliferation, apoptosis, and differentiation, depending on

the cell type examined. TCDD acts as an endocrine disruptor in cell cultures, inhibiting several

estrogen-induced responses such as growth of human mammary and endometrial cancer cells

(20). TCDD induces the proliferation (21) and terminal differentiation (22;23) of human

keratinocytes, and both enhances (24) and inhibits (25;26) rat hepatocyte proliferation rates.

Induction of apoptosis has been reported for immature thymocytes from rats and mice treated

with TCDD in vivo (27-29). In cultures of rat hepatocytes, TCDD also induces apoptosis (24),

but inhibits UV-induced apoptosis (30).

The molecular basis for the biological effects of TCDD is largely speculative. The

majority of the effects of TCDD exposure are mediated by the AHR (31;32), a ligand-activated

transcription factor that, upon ligand binding, translocates to the nucleus where it complexes

with ARNT (33). The AHR/ARNT heterodimer stimulates the transcription of genes in the

cytochrome P450 CYP1 family, as well as several Phase II detoxification genes (34;35), via

transactivation through enhancer domains known as AHR-, dioxin-, or xenobiotic-response

elements (AhREs, DREs or XREs) (36). Gene transactivation mediated by the AHR represents

an adaptive response required for the detoxification of foreign compounds. However, this effect does not adequately explain the range of toxic outcomes resulting from TCDD exposure in different systems, particularly regarding gene repression effects as seen for TGF-β2 (37;38),

fibrinogen γ chain and plastin mRNAs (39), cyclin A (40), and others (41). Global gene expression studies indicate that the AHR participates in the direct and indirect modulation of the transcriptional program (41), at least in part by associating with additional transcription factors

(42-44) and coactivators or corepressors (45-47). Such associations may be partially responsible for the myriad effects of the ligand-activated AHR in the regulation of proliferation, apoptosis, differentiation, and signal transduction pathways (41).

Several lines of evidence indicate that AHR activation directly modulates normal cell

cycle regulation. Ma and Whitlock (48) noted that AHR-defective Hepa-1 cells exhibit slower

growth due to prolongation of G1. Rat hepatoma 5L cells that express wild-type AHR undergo

TCDD-mediated G1 arrest, while AHR-negative mutant 5L cells are refractory to growth

inhibition, suggesting a direct involvement of the AHR in this G1 arrest (49). Recent data has connected the role of the AHR in G1 regulation to a direct interaction with the retinoblastoma

(RB) protein (43;44). RB is a cell cycle regulated phosphoprotein that complexes with E2F

family proteins to control cell division. E2F family members control the transcription of a

variety of essential cell cycle control genes, including several cyclin-dependent kinases and their

inhibitors, RB and related pocket proteins, enzymes for nucleotide biosynthesis, and proteins

required for DNA replication (50). Hypophosphorylated RB binds E2F to inhibit transcriptional

activation and mediate active repression of E2F-responsive genes, thereby inhibiting cell cycle

progression from G1 into S-phase (51). Inactivation of RB by phosphorylation relieves this

repression, allowing cells to enter into S-phase. Previous observations from our laboratory have

shown that the activated AHR synergizes with RB to repress gene expression (44), which might

explain the involvement of the AHR in the G1 phase of the cell cycle. However, little is known

regarding the possible role that AHR-mediated transcriptional activity and ARNT interactions

might play in the repression and what role, if any, is played by chromatin remodeling factors,

known to mediate E2F-dependent transactivation, in the overall outcome of AHR-RB

interactions. The purpose of the present research was to address these central questions and to

test the hypothesis that through its interaction with RB, the AHR functions as a repressor of transcription. Our results help elucidate the complex role played by AHR signaling during the cell cycle and delineate a novel role for the Ah receptor, a known transactivator, in gene repression.

Materials and Methods

Cell Lines, Transfections, and Growth Conditions

The mouse Hepa-1c1c7 cell line, a subclone of the Hepa-1 hepatoma cell line (52) and

two of its mutant derivatives were used in these studies. The c35 derivative line expresses a

DNA-binding defective Ah receptor (53), and c4 cells lack ARNT (34). Exponentially growing

cells were cultured in minimal essential medium-α (Life Technologies, Inc.) supplemented with

5% fetal bovine serum (Life Technologies, Inc.), 1.0% 100X antibiotic/antimycotic solution

o (Sigma), and 26 mM NaHCO3 at 37 C in a humidified 5% CO2 atmosphere. The human breast

carcinoma cell line MCF-7 (54) was similarly cultured. The human cervical carcinoma cell line

C33A was cultured in the same medium containing 10% fetal bovine serum. When indicated,

cultures were treated with TCDD at a final concentration of 10 nM in DMSO vehicle, and

control cultures were treated with an equivalent volume of DMSO. C33A cells do not express

AHR, as determined from the lack of activity of a transfected AHR/ARNT-responsive luciferase

reporter plasmid and by immunoblot analysis. C33A cells harbor a mutation in the RB gene, and

express an unstable mutant RB protein (55).

Transient transfection experiments were performed using Lipofectamine Plus

(Invitrogen) on cells grown to 50-75% confluence in 24-well plates or 60-mm dishes. All

transfection mixtures were brought to the same amount of total DNA by the addition of the

appropriate amount of empty vector. To control for variations in transfection efficiency, cells

were also co-transfected with a pCMVβgal plasmid (Clontech) and transfections were normalized to β-galactosidase expression. Cells were transfected in serum-free medium, and 3 h

later one volume of medium containing twice the final concentration of culture serum was added.

After overnight incubation, cells were washed once with phosphate-buffered saline and cultured

in complete culture medium with TCDD or with vehicle. Between 24 - 48 h after transfection, cells were lysed in Reporter Lysis Buffer (Promega), and luciferase and β-galactosidase activities were measured. Each transfection was repeated a minimum of two times, and the results shown are the mean ± S.D. of one representative set of results. Where noted, the results shown are the mean ± SE of log-transformed results from multiple independent experiments.

Plasmid constructs

The plasmid pcDNAI/B6AHR, used to express the high affinity variant of murine AHR, has also been described (56). Mutant derivatives of the pcDNAI/B6AHR plasmid are schematically shown in figure 3A. Plasmid p∆495-805 was derived from pcDNAI/B6AHR by deletion of 933 nucleotides encoding the C-terminal 311 amino acids of the AHR peptide; this plasmid encodes a dominant-negative variant of the murine AHR, expressing a protein capable of ligand binding and dimerization with ARNT but lacking transactivation activity (56). The plasmid p∆323-494 contains a deletion of the LXCXE RB-binding motif and the PAS B domain, and plasmid p∆323-607 lacks the LXCXE motif, the PAS B domain, as well as part of the transactivation domain (57). Expression of the peptides encoded by each AHR truncation mutant was confirmed by Western blot.

The reporter p3×E2FLuc, containing three E2F-responsive elements, has been previously described (58). The reporter construct pAhRDtkLuc3 (56) is a luciferase reporter plasmid containing the AHR-responsive domain of the mouse Cyp1a1 gene promoter (from -1100 to -

869), harboring five AHR-responsive elements, fused to the HSV-1 thymidine kinase minimal promoter from -79 to +53 from which the Sp1 binding site had been removed. Other plasmids used were pCMVNeoRB, directing the expression of human RB (59) and pcDNAIneo/mARNT, directing the expression of mouse ARNT (60). pCMVhuHDACI, a plasmid directing the

expression of human histone deacetylase-1, was derived by BamHI digestion of the plasmid

pBJ5-HD1-F (ATCC) and insertion into the multiple cloning site of pCMVscript (Stratagene)

vector. Plasmids directing the expression of p300 and a dominant negative p300 were obtained

from commercial sources (Upstate Biotechnology, Inc.).

In Vitro Transcription/Translation

One microgram of plasmids encoding the wild type AHR or truncation mutants were

used in 50-µL in vitro transcription/translation reactions with a TNT coupled reticulocyte lysate

system (Promega), following the manufacturer's instructions and included 30 µCi of

[35S]methionine (Amersham Pharmacia Biotech). Reactions were incubated at 30oC for 1.5 h.

Translated proteins were used immediately for pull-down assays.

Pull-down Assays

Bacteria expressing a human RB protein tagged with His6 at the amino terminus were

lysed in 500 µL of in 6 M Guanidine-HCl, 0.1 M NaH2PO4, 0.01 M Tris (pH 8.0) and the lysate was incubated with a nickel-agarose slurry (Qiagen) in 2 mL of buffer containing 8 M urea, 0.1

M NaH2PO4, 0.01 M Tris (pH 8.0). The slurry was passed through a column, washed three times with 1 mL 20 mM Tris (pH 7.6), 150 mM NaCl, 0.1% Triton X-100, and stored in one volume of buffer at 4oC. Control and RB-bound columns were established using 100 µL aliquots of the pre-adsorbed slurry. Full-length and truncation peptides of the mouse AHR were in vitro translated with [35S]methionine and precleared through nickel-agarose columns. Equal volumes of AHR or truncated peptides were recycled 10 times through control and RB-bound columns.

Each column was washed twice with 2 column volumes of 20 mM Tris (pH 7.6), 150 mM NaCl,

0.1% Triton X-100. Bound proteins were eluted with 1.5 column volumes of 8 M urea, 0.1 M

NaH2PO4, and 0.01 M Tris (pH 6.3) to disrupt nickel coordination of the His6 tag. Eluted

fractions were denatured by boiling and analyzed by SDS-PAGE in 7.5% acrylamide gels.

Detection of [35S]AHR was by autoradiography. Radioactive quantitation was carried out with a

Storm 860 PhosphorImager (Molecular Dynamics, Inc.).

Flow Cytometry

For cell cycle studies, MCF-7 cells were maintained in serum-free minimal essential

medium-α supplemented with 1% antibiotic/antimycotic solution for 72 hours. Cells were

subsequently fed with medium containing 5% fetal bovine serum and 10 nM TCDD or an

equivalent volume of DMSO vehicle, and cultured for intervals ranging from 10 to 30 h. Cells

were pulse-labeled with BrdU (BD Pharmingen) for 1 h at the indicated time point and harvested

for cell cycle analysis using a Beckman Coulter Epics XL flow cytometer (Fullerton, CA). The

percentage of cells entering S phase was determined by gating BrdU positive cells and

subtracting background fluorescence using an unlabeled control. DNA content of the total cell

population was determined by 7-AAD staining and the fraction of cells in G1 was determined from the fraction of cells with 2N DNA content. Hepa-1 cells were similarly processed for analysis of BrdU incorporation. Cells were continuously cultured to approximately 80-90% confluence under standard conditions and treated with 10 nM TCDD or vehicle control for the time period indicated in Figure 1.

Total RNA isolation, reverse transcription, and real-time PCR

Hepa-1 cells were cultured in 10-cm plates to 80-90% confluence and treated under standard culture conditions with DMSO vehicle or with 1 µg/mL Actinomycin D for 1 hour.

Actinomycin-treated cells were further treated with DMSO vehicle or with 10 nM TCDD for 0,

4, or 8 h prior to harvesting of RNA. Total cellular RNA was isolated with Tri Reagent

(Molecular Research Center, Inc). Immediately following heating at 65°C for 10 min, first

strand cDNAs were synthesized from 15 µg of total RNA using SuperScript II reverse transcriptase (Life Technologies, Inc.) and random primers. Diluted cDNAs were subjected to

PCR amplifications with gene-specific primer sets for the various E2F-responsive and control

genes (Table 1). Reaction volumes of 25 µL contained 12.5 µL Brilliant SYBR Green QRT-

PCR Master Mix (Stratagene), 0.4 µM primers, and 2-4 µL cDNA template. Real-time quantitative PCR was performed using a Smart Cycler rapid thermal cycler (Cepheid) and fluorescence was measured after each of the repetitive cycles. A typical protocol included a 2 min denaturation step at 95oC followed by 40 cycles of 95oC denaturation for 30 sec, annealing for 30 sec at a primer-optimized temperature, and 72oC extension for 30 sec. Detection of the

fluorescent product was carried out during the 72oC extension period, and emission data were

quantified using threshold cycle (Ct) values. Ct values for all genes analyzed were determined 2 to 4 times, averaged and normalized to values for β-actin. The fold change from control Ct values (vector transfected, DMSO treated) was determined for each sample, as determined by the following calculation:

Fold change = 2-∆∆Ct

where ∆∆Ct = (CtTarget – CtActin)Time=t – (CtTarget – CtActin)Time=0. PCR product specificity from each

primer pair was confirmed using melting curve analysis and subsequent agarose gel

electrophoresis.

Chromatin immunoprecipitation (ChIP) assays

The ChIP protocol was performed with minor modifications as described previously by

others (61;62). Hepa-1 cells, bearing a neo-resistance cassette and cultured in 600 µg/mL G418, were grown in 15-cm plates to 90-100% confluence (approximately 2 x 107 cells). One plate of cells was used per treatment, per antibody. Cells were treated with DMSO vehicle or 10 nM

TCDD for 1.5 hours. Cross-linking was performed by adding formaldehyde directly to the

culture media to a final concentration of 1%, followed by a 10-minute incubation. The reaction

was quenched using glycine at a final concentration of 0.125 M. Cells were rinsed twice with

cold PBS, scraped from the dishes, pelleted and washed again with PBS containing 0.5 mM

PMSF. Cell pellets were resuspended in cell lysis buffer (5 mM PIPES (pH 8.0), 85 mM KCl,

0.5% NP-40, 0.5 mM PMSF, 5 µg/ml leupeptin, 5 µg/ml aprotinin) and incubated on ice for 10 min. Cells were then dounced with a B pestle 10 to 20 times on ice to aid in nuclei release.

Nuclei were pelleted and resuspended in nuclei lysis buffer (50 mM Tris-HCl (pH 8.1), 10 mM

EDTA, 1% SDS, 0.5 mM PMSF, 5 µg/ml leupeptin, 5 µg/ml aprotinin) and incubated on ice for

10 min. Chromatin was sonicated to an average length of 600 bp with 4 10-second pulses of 30

W, maintaining samples on ice for 30 seconds between pulses. Sonicated chromatin was then precleared for 15 min at 4°C with BSA- and salmon sperm DNA-saturated protein A agarose

(Upstate Biotechnology) in preparation for immunoprecipitation. The supernatant was divided equally among all samples and incubated overnight on a rotating platform at 4°C with 1 µg of the following antibodies: AHR (BioMol), E2F1 (Santa Cruz), HDAC-1 (Upstate), p300 (Upstate), and RB (Biomol). Protein A agarose slurry (20 µl) was added and incubated for 15 min at room temperature to allow for antibody binding. Agarose was pelleted and washed twice with 1X dialysis buffer (50 mM Tris-HCl (pH 8.0), 2 mM EDTA, 0.2% sarkosyl) and 4 times with IP wash buffer (100 mM Tris-HCl (pH 9.0), 500mM LiCl, 1% NP-40, 1% deoxycholic acid).

Immune complexes were eluted from the beads with elution buffer (50 mM NaHCO3, 1% SDS).

Reversal of crosslinks was carried out by heating the eluates at 67°C for 4-5 h. The eluates were

then digested with proteinase K at 56°C for 1.5 h. DNA was extracted with

phenol/chloroform/isoamyl alcohol, precipitated with ethanol and dissolved in TE buffer.

Samples were diluted between 5 and 10 times and subjected to PCR amplification with promoter-specific primer sets for the genes under study (Table 2). PCR products were separated for analysis in 10% polyacrylamide gels.

Results

TCDD inhibits S-phase progression in multiple cell lines

Results of previous work from our laboratory have shown that in RB- and AHR-negative

Saos-2 cells, ectopically overexpressed AHR and RB cooperate to suppress entry of transfected cells into S-phase. This effect was attributed to a reduction in E2F-dependent gene expression, as assessed using transfection of reporter constructs, required for DNA synthesis and cell cycle progression (44). Additional observations indicated that AHR activation by TCDD treatment in serum-starved, RB-positive cells resulted in a significant reduction in both RB phosphorylation and cyclin D1 expression (63). To determine whether physiologic levels of activated AHR and

RB could cooperate equally well in the inhibition of cell cycle progression, we used the RB- and

AHR-positive MCF-7 breast carcinoma cell line for analysis of BrdU incorporation. Cells were serum-starved for 3 days, followed by release into medium containing 5% fetal bovine serum and either DMSO vehicle or 10 nM TCDD for various time intervals ranging from 5 to 30 hours.

Cells were pulse-labeled with BrdU for 1 h at the indicated time points and harvested for flow cytometry analysis. The results indicate that TCDD treatment retards progression of cells into S- phase (Fig. 1A). The difference in the number of cells entering S-phase between control and

TCDD-treated cells increased significantly over the time course examined. As expected, the control group proceeded into a state of exponential growth, while S-phase progression in the

TCDD group lagged behind, with a significantly lower percentage of the total cell population.

Similar experiments were carried out using the Hepa-1c1c7 mouse hepatoma cell line.

Hepa-1 cells were cultured to 80-90% confluence and treated with DMSO vehicle or 10 nM

TCDD for 16 or 24 hours. Cells were pulse-labeled with BrdU for 1 h at the indicated time points and harvested for flow cytometry analysis. As in MCF-7 cells, there was a significant

10% decrease in the number of BrdU positive cells in cultures treated with TCDD, suggesting

that TCDD retarded DNA synthesis in Hepa-1 cells (Fig. 1B). The decrease in the number of

cells incorporating BrdU in response to TCDD treatment was accompanied by a corresponding

increase in the number of G1 cells and decrease in G2-M cells (Fig. 1C), indicating that TCDD acts primarily by promoting the accumulation of cells in G1 and blocking progression from G1 to

S phase. Similar results have been obtained in the rat 5L hepatoma cell line, where the effect of

TCDD on the cell cycle has been shown to be dependent on the presence of a functional AHR

(64;65). These results confirm those in both this and other cell lines, and indicate that the effect of AHR overexpression observed in transfection studies (44) is relevant to physiological conditions.

TCDD inhibits expression of E2F-dependent, S-phase specific genes

In order to investigate the mechanism by which TCDD slows down progression into S- phase, we measured the expression of several E2F-responsive, S-phase specific genes using real- time RT-PCR analysis. Cells were cultured in the presence of serum to 80-90% confluence, treated with DMSO vehicle, 10 nM TCDD, 1 µg/mL actinomycin D, or 1 µg/mL actinomycin D for 1 h followed by 10 nM TCDD, and harvested at 0, 4 and 8 hours after treatment. Treatment with actinomycin D provides a baseline for the level of mRNA remaining at any given time after inhibition of transcription. RNA was isolated from these cells and the mRNA levels of Cyp1a1, p27Kip1, and the E2F-regulated genes Cdk2, Cyclin E, Dhfr and DNA polymerase α were

determined by real-time RT-PCR and normalized to β-actin mRNA levels. We measured p27Kip1 mRNA levels because the effect of TCDD on cell cycle progression in 5L rat hepatoma cells has

been attributed to p27 accumulation (66). As expected, following 4 hours of TCDD treatment,

the level of Cyp1a1 mRNA increased 100-fold over control and this increase was maintained

throughout the 8-hour treatment period (Fig. 2). Between 0 and 4 hours of TCDD treatment, all

four E2F-regulated genes showed a similar pattern of time-dependent decrease in mRNA

expression relative to DMSO-treated controls. At this time, the pattern of inhibition by TCDD

mirrors that found for cells treated with actinomycin D or with actinomycin D plus TCDD, suggesting that TCDD treatment shuts-off completely the transcription of all four E2F-dependent genes tested. This pattern continued to be maintained for Dhfr and Pol α, but Cdk2 and Cyclin E

mRNAs began to recover toward control levels by 8 hours after TCDD treatment.

Notwithstanding, both Cdk2 and Cyclin E mRNA levels were still significantly decreased at the

8 h time point compared to vehicle-treated controls. In contrast to the results with E2F-regulated genes, p27 expression following TCDD treatment increased by approximately 2-fold (Fig. 2).

Transactivation is not required for AHR-mediated repression of gene expression

The observation that TCDD inhibits cell cycle progression in a variety of cell lines has been attributed to multiple different mechanisms, including the AHR-mediated up-regulation of genes that inhibit cell cycle progression (66) as well as a direct AHR-RB interaction resulting in enhanced down-regulation of genes required for cell cycle progression (44). The relative contribution of AHR-mediated transactivation to the down-regulation of E2F-regulated genes in cooperation with RB has never been investigated. Previous studies have shown that AHR and

RB interact in vitro, and that several domains of the AHR, including the glutamine-rich region of the carboxy-terminal transactivation domain and an RB-binding LXCXE motif, contribute to this interaction (43;44). To analyze specific regions of the AHR for their contribution to the interaction with RB, we tested AHR deletion mutants for their ability to bind RB in an in vitro pull-down assay. Several mutant AHR peptides were constructed containing deletions of the amino-terminal LXCXE motif at amino acid 325 of the PAS domain, and the glutamine-rich

sequence within the carboxy-terminal transactivation domain (Fig. 3A). In vitro synthesized

35 full-length or mutant [ S]AHR was passed through nickel-agarose columns saturated with His6- tagged RB prepared in E. coli. Control columns contained a lysate of host E. coli cells. Bound

[35S]AHR proteins were analyzed by electrophoresis in 7.5% SDS-polyacrylamide gels. The

fraction of labeled AHR bound to control columns was negligible for the full-length protein and

all mutant peptides. Quantitation of input and bound [35S]AHR from the RB-containing columns by PhosphorImager analysis showed that 39.5% of labeled full-length AHR protein input was bound by RB. The AHR deletion mutants bound RB to varying degrees, all significantly less than full-length AHR (Fig. 3B). These data suggest that not just two but possibly several domains, and perhaps proper folding and three-dimensional confirmation of those domains, are responsible for maximal interaction between AHR and RB as observed in these and other in vitro

experiments.

The binding data presented above suggests that the relative loss of RB binding by

truncated mutants of AHR may impair their ability to cooperate with RB to repress E2F-

regulated gene expression. We therefore used AHR truncation mutants (Fig. 3A) in transient

transfection assays and compared their ability to transactivate an AHR-dependent reporter gene

with their ability to repress an E2F-dependent reporter gene. Each truncated AHR peptide was

first tested for its ability to activate AHR-dependent transcription in the human cervical

carcinoma C33A cell line, which lacks expression of AHR, ARNT and a stable RB protein. As

expected, full-length AHR induced high levels of luciferase activity from the pAHRDtkluc3

reporter, driven by the AHR-responsive enhancer from the mouse Cyp1a1 gene (Fig. 3C).

Transactivation activity was retained by the ∆323-494 mutant, which is constitutively active but

unresponsive to TCDD stimulation, and was largely decreased for the ∆323-607 mutant.

E2F-dependent transcriptional activity was assayed using the luciferase reporter plasmid

p3×E2FLuc that responds to transactivation by E2F. Transfection of this plasmid into C33A

cells resulted in high luciferase expression levels (Fig. 3D). Ectopic expression of RB reduced

luciferase activity to 60% of control. As expected, simultaneous expression of RB and the full-

length AHR further reduced luciferase expression to approximately 30% of control, indicative of

the cooperative relationship between AHR and RB in the transcriptional repression of E2F-

dependent promoters. Additional AHR truncation mutants, including the transcriptionally

inactive mutants ∆495-805 and ∆323-607, were equally able to repress reporter activity in

cooperation with RB. In contrast, the transcriptionally active AHR mutant ∆323-494 was unable to repress E2F-dependent gene expression. These effects were independent of TCDD treatment, as ectopically expressed AHR is activated and localized to the nucleus in the absence of ligand

(56). These results suggest that the repressive effect on E2F-dependent gene expression does not require the maximal binding levels observed with full-length AHR in the in vitro binding experiments (see Fig. 3B). All of the truncated AHR peptides which cooperated with RB to repress E2F-dependent transcription showed little or no effect on luciferase expression in the

AHR-dependent gene expression assay, suggesting that the effect of AHR on E2F-dependent functions is an independent event unrelated to the ability of the AHR to transactivate gene expression. To insure that these results were indeed due to expression of the transfected proteins, whole cell extracts of transfected C33A cells were separated in 7.5% SDS-PAGE, transferred to a nitrocellulose membrane, and protein expression analyzed for all transfected plasmids by Western immunoblot. AHR, ARNT and RB proteins corresponding to all transfected plasmids were expressed at comparable levels (data not shown).

A transcriptionally competent AHR complex is not required for gene repression

In the nucleus, AHR forms a transcriptionally active heterodimer with ARNT/HIF-1β

(67), and the AHR/ARNT interaction is required for maximal transactivation of TCDD-inducible

genes (68). As the transactivation function of AHR is not required for the cooperative repression

of E2F-dependent gene expression by AHR-RB complexes, we hypothesized that ARNT might

also be dispensable for this effect. It is known that ARNT is not required for the interaction between AHR and RB (43), but its effect in mediating transcriptional repression by the AHR is

unknown. To determine the role played by ARNT in the AHR-dependent effects on gene

repression, we tested the effect of ARNT on E2F-dependent gene expression using c4 cells in

transient transfection assays with the p3×E2FLuc plasmid. The c4 cell line is derived from the

mouse Hepa-1 hepatoma cell line (34) and lacks expression of ARNT as well as RB. As shown

in Figure 4A, transfection of AHR into c4 cells enhanced RB-mediated repression of E2F-

dependent gene expression by about 50%, as previously reported in Hepa-1 cells (44). Co-

expression of AHR and wild type ARNT in these cells showed no further reduction in luciferase

expression as compared to AHR alone, indicating that interaction between AHR and ARNT is

not necessary for repression of E2F-dependent gene expression by AHR.

To verify that the endogenous AHR would repress gene expression independently of

DNA binding, we used the p3×E2FLuc plasmid in transient transfection assays in Hepa-1 cells,

containing a wild type Ah receptor, and in the c35 cell line, a Hepa-1 derivative that contains

normal levels of a mutant AHR that is incapable of binding to DNA (53). In Hepa-1 cells, in

agreement with previous results (44), AHR activation by TCDD treatment repressed E2F-

dependent gene expression, and more so in cooperation with overexpressed levels of RB (Fig.

4B, left panel). The same results were observed in the c35 cell line (Fig. 4B, right panel),

emphasizing the fact that AHR-mediated DNA-binding and transactivation is not a requirement for repression of E2F-regulated genes by TCDD.

TCDD reverses p300-mediated induction of E2F-dependent gene expression

E2F-dependent gene expression is regulated by the recruitment of a variety of corepressors and coactivators to both RB-bound and free forms of E2F. Active repression of certain E2F-responsive genes is mediated by the RB-dependent recruitment of corepressors, including histone deacetylase and chromatin remodeling factors, to RB-E2F-DP complexes

(reviewed in (50;69)). In addition, the ability of E2F to act as an activator of transcription depends in part on an interaction with the histone acetyl transferases p300/CBP and p/CAF

(70;71). A role for corepressor/coactivator activities is also recognized as important for the regulation of the AHR complex (46;72). We therefore tested the dependence of AHR transcriptional repression on specific corepressor and coactivator activities required for normal

E2F-dependent regulation of transcription. We used the p3×E2FLuc plasmid for transient transfection assays in Hepa-1 cells, containing a wild type AHR. As expected, AHR activation by TCDD treatment repressed E2F-dependent gene expression (Fig. 5). Maximal repression of luciferase activity was obtained by co-transfection of either HDAC-1 or BRG-1 with RB. There was a trend towards further repression in TCDD-treated cells co-transfected with HDAC-1, although the effect was not statistically significant. When p300 was included in the transfection, repression of luciferase activity by RB was abolished, an effect that was blocked by TCDD treatment. In contrast, expression of a dominant negative p300 did not reverse the repression of luciferase activity by RB. These results suggest that the activated AHR might prevent recruitment of p300 to the E2F complex, effectively enabling continued repression of gene

transcription by RB-bound corepressors. Similar results were obtained in c35 cells (data not shown).

Activation of the AHR by TCDD displaces p300 from E2F-dependent promoters

The transient transfection experiments reported above suggest the possibility that AHR activation resulting from TCDD treatment represses E2F-dependent gene expression by blocking the recruitment of p300 to E2F-dependent promoters. To test this hypothesis, we used ChIP assays to analyze protein constituents at native promoters. Hepa-1 cells were treated with

DMSO vehicle or 10 nM TCDD for 1.5 h and native chromatin was prepared from nuclear lysates. Immunoprecipitations were carried out using antibodies to AHR, E2F1, HDAC-1, p300 and RB. Controls containing no antibody were included to confirm the specificity of the precipitations. Figure 6 shows the results of PCR reactions from immunoprecipitated chromatin, analyzing AHR-binding sequences from the Cyp1a1 promoter, as well as sequences containing

E2F-binding sites from the promoters of Cdk2, Cyclin E, Dhfr, and Pol-α. Antibodies to AHR and p300 immunoprecipitated AHR-responsive enhancer sequences from the Cyp1a1 promoter in TCDD-treated cells, but not in vehicle-treated cells. There was no change observed between

DMSO- and TCDD-treated cells in the association of E2F1, RB, or HDAC-1 with the promoters of any of the S-phase-specific genes analyzed. However, while the p300 antibody immunoprecipitated the promoters of all four E2F-dependent genes in DMSO-treated cells,

TCDD treatment resulted in the loss of p300 from these promoters. Furthermore, the AHR antibody immunoprecipitated the promoters of these genes in TCDD-treated cells, but not in vehicle-treated cells. These results indicate that, following TCDD treatment, the recruitment of the activated nuclear AHR to E2F-regulated promoters, presumably through a direct interaction with RB and possibly E2F itself, is a critical step that leads to gene repression. In agreement

with the transfection studies (Fig. 5), these data point toward a mechanism of AHR-mediated gene repression that involves the inhibition of p300 recruitment to the E2F complex.

Discussion

In this paper, we show that AHR activation by TCDD inhibits cell cycle progression, accumulating cells in G1, and represses the transcription of E2F-dependent genes through

interaction with RB. Recruitment of the activated AHR to E2F-regulated promoters blocks the

recruitment of p300 and the concomitant up-regulation of E2F-dependent genes. We conclude

that the two processes are interrelated and that inhibition of p300 binding is one of the molecular

events responsible for the persistence of active gene repression by AHR-RB complexes. We

present evidence to show that the repressor activity of the AHR is fully independent of its

transcriptional activity, since it does not require its transactivation or DNA binding domains nor

interaction with ARNT. Hence, we propose that repression is the overall effect of ternary or

quaternary interactions taking place at E2F-regulated promoters.

Previous work has shown that the molecular interaction between the Ah receptor and the

retinoblastoma protein leads to enhanced repression of E2F-dependent genes and cell cycle

inhibition. RB-mediated transcriptional repression is dependent on two distinct protein-binding

sites in RB: the large A/B pocket, which binds E2F, and the A/B pocket, which binds LXCXE

peptide motifs (58). Deletion analysis of the Ah receptor reveals that neither the LXCXE motif

nor the glutamine-rich region of the AHR can mediate the maximal interaction levels between

the two proteins observed in vitro with full-length AHR. In experiments using RB deletion

mutants (see Chapter 4), we find that the large A/B pocket of RB is required for binding with the

AHR, confirming that RB sequences other than those that bind the LXCXE motif of AHR are

necessary for their interaction. These findings may explain previous observations that T-antigen

only partially relieves AHR effects on RB-mediated repression (44). However, maximal binding

to RB is not critical for enhanced repression of E2F-dependent genes by AHR, as evidenced by

transfection assays in which several AHR deletion mutants, none of which are capable of

maximal interaction with RB in vitro, can repress gene expression as efficiently as the wild-type

protein. This result is consistent with data showing that expression of a variant Ah receptor with a mutation in the LXCXE motif can still mediate G1 arrest (73), and that LXCXE-dependent interactions are not essential for RB to exert growth arrest (74). It is also notable that additional protein contacts may be involved in the interaction. This is underscored by the fact that the AHR can pull-down E2F1, and that ectopic expression or activation of the AHR by TCDD can mediate repression of E2F-dependent genes in the absence of RB (see Chapter 4).

Activation of the Ah receptor by TCDD blocks cell cycle progression in various cell lines and under a number of different conditions, including in pancreatic cell cultures (75), in specific sub-clones of PLHC-1 cells (76), and in the estrogen-induced proliferation of MCF-7 cells (63), among others. Inhibition of cyclin-dependent kinase activities, inhibition of RB

phosphorylation, and induction of p27Kip1 and p21WAF1 have been observed and proposed as potential mechanisms responsible for the block. Overexpression of AHR was also found to enhance RB-mediated inhibition of S-phase progression in Saos-2 cells when both proteins were ectopically expressed (44). We have determined that this effect is not merely the consequence of protein overexpression, as TCDD significantly retards S-phase progression in MCF-7 and in

Hepa-1 cells, both of which express endogenous physiologic levels of AHR and RB. This is consistent with recent observations in MCF-7 cells showing that ablation of endogenous AHR expression by use of siRNA results in enhanced progression of cells from G0/G1 into S phase

(77). In our experiments with TCDD-treated Hepa-1 cells, the number of cells in G1 increases by the same extent that the number of cells incorporating BrdU decreases, suggesting that the primary effect of AHR activation on cell cycle progression occurs during G1, as might result

from its interaction with RB. Importantly, our results in RB-expressing Hepa-1c1c7 cells show that the inhibition of S-phase progression following TCDD treatment coincides with a significant decrease in the expression of at least four genes that are positively regulated by E2F family members and necessary for S-phase progression (50). This decrease is accompanied by a doubling of p27 mRNA levels. Accumulation of p27 mRNA and protein in 5L rat hepatoma cells has been proposed as the major cause of the AHR-mediated cell cycle progression inhibition (66). Our results indicate that decreases in expression of E2F-dependent genes occurs concomitant with a moderate increase in p27 expression, suggesting that the two processes are linked and that the ultimate outcome on cell cycle progression might be the result of their cooperation. On the other hand, since transactivation by AHR and ARNT interaction are not necessary for AHR-dependent gene expression, it is unlikely that the increase in p27 mRNA is the direct result of AHR activation.

The data presented here eliminate several potential mechanisms by which the AHR acts to enhance repression of E2F-dependent genes. Gene repression mediated by the AHR is independent of its ability to transactivate gene expression, the as yet best characterized of AHR functions. Repression of E2F-dependent genes is also independent of DNA binding by the AHR, emphasizing the requirement for protein interactions in mediating this effect. While nuclear localization is required in vivo for the AHR/RB interaction, in vitro binding assays show that

ARNT is not needed for formation of the AHR/RB complex (44) nor is ARNT required for maximal repression of E2F-dependent gene expression by the AHR. Our studies, however, were not directed at analyzing the role of CYP1A1 expression on the extent of cell cycle inhibition and thus does not address recent work from Elferink and colleagues (78) that suggests that the

activity of the AHR-dependent CYP1A1 protein prevents AHR-mediated G1 arrest by metabolizing AHR ligands and negatively regulating the duration of AHR activation.

Binding of RB to E2F at E2F-responsive promoters recruits co-repressor proteins that mediate active transcriptional repression (51). Repression is relieved by RB phosphorylation and recruitment of coactivator proteins by E2F (71). p300/CBP activity is also required for cell cycle

progression and E2F activity (79), underscoring its necessity for up-regulation of E2F-dependent

transcription. We have previously shown that AHR attenuated both cdk2 protein levels as well

as cdk2-associated kinase activity (40) and thus helps maintain RB in a hypophosphorylated

state. It may be that in addition, the AHR blocks critical interaction sites required for the

recruitment of p300/CBP by E2F, and hence helps block transcriptional induction by also

preventing the reorganization of chromatin at E2F-responsive promoters. These two effects may

in fact be causally related. This concept is supported by our results showing an additive effect of

TCDD on HDAC-mediated repression of E2F-dependent gene expression. The ChIP data with

asynchronously growing cells shown here cannot properly address this issue, because it

represents the combination of results from cells at all stages of the cell cycle, and the fraction of

cells in S and G2-M mask the results specific for cells in G1.

There is no current paradigm that satisfactorily explains more than half of the gene regulatory effects of dioxin exposure, namely, the mechanisms responsible for its large effect as a repressor of gene expression (41). We believe that one of these mechanisms involves the

activated AHR-mediated repression of E2F-dependent gene transcription. As shown herein,

repression depends to a large extent on the interactions of AHR with RB and possibly with E2F itself. It appears that one of the critical functions of the activated Ah receptor is to serve as an

environmental checkpoint that senses exposure to damaging environmental toxicants and responds by signaling cell cycle inhibition.

Acknowledgements

We are grateful to Dr. Ying Xia for helpful suggestions and a critical reading of the manuscript and Dr. Oliver Hankinson for gifts of the c35 cell line and ARNT expression vectors. We thank

Brenda L. Schumann for technical assistance. This work was supported by NIEHS grants

ES06273 and P30 ES06096. JLM was supported by the University of Cincinnati predoctoral

Functional Genomics and Ryan Fellowships.

References

1. Mann PC. 1997. Selected lesions of dioxin in laboratory rodents. Toxicol.Pathol. 25: 72- 79

2. Oughton JA, Pereira CB, DeKrey GK, Collier JM, Frank AA, and Kerkvliet NI. 1995. Phenotypic analysis of spleen, thymus, and peripheral blood cells in aged C57Bl/6 mice following long-term exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin. Fund.Appl.Toxicol. 25: 60-69

3. Kerkvliet NI. 1995. Immunological effects of chlorinated dibenzo-p-dioxins. [Review]. Environmental Health Perspectives 103 Suppl 9: 47-53

4. Bjerke DL and Peterson RE. 1994. Reproductive toxicity of 2,3,7,8-tetrachlorodibenzo-p- dioxin in male rats: different effects of in utero versus lactational exposure. Toxicology & Applied Pharmacology 127: 241-249

5. Birnbaum LS. 1995. Developmental effects of dioxins. Environ.Health Perspect. 103(Suppl 7): 89-94

6. Mably TA, Bjerke DL, Moore RW, Gendron-Fitzpatrick A, and Peterson RE. 1992. In utero and lactational exposure of male rats to 2,3,7,8- tetrachlorodibenzo-p-dioxin. 3. Effects on spermatogenesis and reproductive capability. Toxicology & Applied Pharmacology 114: 118-126

7. Cummings AM, Metcalf JL, and Birnbaum L. 1996. Promotion of endometriosis by 2,3,7,8-tetrachlorodibenzo-p- dioxin in rats and mice: time-dose dependence and species comparison. Toxicology & Applied Pharmacology 138: 131-139

8. Rier SE, Martin DC, Bowman RE, Dmowski WP, and Becker JL. 1993. Endometriosis in rhesus monkeys (Macaca mulatta) following chronic exposure to 2,3,7,8- tetrachlorodibenzo-p-dioxin [see comments]. Fundam.Appl.Toxicol. 21: 433-441

9. Dragan YP and Schrenk D. 2000. Animal studies addressing the carcinogenicity of TCDD (or related compounds) with an emphasis on tumour promotion. Food Addit.Contam 17: 289-302

10. Suskind RR. 1985. Chloracne, the hallmark of dioxin intoxication. Scand.J.Work Environ.Health 11: 165-171

11. Schmidt JV and Bradfield CA. 1996. Ah receptor signaling pathways. Annu.Rev.Cell Dev.Biol. 12: 55-89

12. Flesch-Janys D, Berger J, Gurn P, Manz A, Nagel S et al. 1995. Exposure to polychlorinated dioxins and furans (PCDD/F) and mortality in a cohort of workers from a herbicide-producing plant in Hamburg, Federal Republic of Germany. Am.J.Epidemiol. 142: 1165-1175

13. Steenland K, Piacitelli L, Deddens J, Fingerhut M, and Chang LI. 1999. Cancer, heart disease, and diabetes in workers exposed to 2,3,7,8- tetrachlorodibenzo-p-dioxin [see comments]. J.Natl.Cancer Inst. 91: 779-786

14. Bertazzi PA, Consonni D, Bachetti S, Rubagotti M, Baccarelli A et al. 2001. Health effects of dioxin exposure: a 20-year mortality study. Am.J.Epidemiol. 153: 1031-1044

15. Anonymous. 1997. IARC Working Group on the Evaluation of Carcinogenic Risks to Humans: Polychlorinated Dibenzo-Para-Dioxins and Polychlorinated Dibenzofurans. Lyon, France, 4-11 February 1997. IARC Monogr Eval.Carcinog.Risks Hum. 69: 1-631

16. Bertazzi PA, Pesatori AC, Consonni D, Tironi A, Landi MT, and Zocchetti C. 1993. Cancer incidence in a population accidentally exposed to 2,3,7,8- tetrachlorodibenzo- para-dioxin [see comments]. Epidemiology. 4: 398-406

17. Manz A, Berger J, Dwyer JH, Flesch-Janys D, Nagel S, and Waltsgott H. 1991. Cancer mortality among workers in chemical plant contaminated with dioxin. Lancet 338: 959- 964

18. Fingerhut MA, Halperin WE, Marlow DA, Piacitelli LA, Honchar PA et al. 1991. Cancer mortality in workers exposed to 2,3,7,8- tetrachlorodibenzo-p-dioxin [see comments]. N.Engl.J.Med. 324: 212-218

19. Pirkle JL, Wolfe WH, Patterson DG, Needham LL, Michalek JE et al. 1989. Estimates of the half-life of 2,3,7,8-tetrachlorodibenzo-p-dioxin in Vietnam veterans of Operation Ranch. J.Toxicol.Environ.Health 27: 165-171

20. Safe S, Wang F, Porter W, Duan R, and McDougal A. 1998. Ah receptor agonists as endocrine disruptors: antiestrogenic activity and mechanisms. Toxicol.Lett. 102-103: 343- 347

21. Milstone LM and LaVigne JF. 1984. 2,3,7,8-tetrachlorodibenzo-p-dioxin induces hyperplasia in confluent cultures of human keratinocytes. J.Invest.Dermatol. 82: 532-534

22. Choi EJ, Toscano DG, Ryan JA, Riedel N, and Toscano WA, Jr. 1991. Dioxin induces transforming growth factor-alpha in human keratinocytes. J.Biol.Chem. 266: 9591-9597

23. Gaido KW and Maness SC. 1994. Regulation of gene expression and acceleration of differentiation in human keratinocytes by 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicol.Appl.Pharmacol. 127: 199-208

24. Moolgavkar S, Luebeck EG, Buchmann A, and Bock KW. 1996. Quantitative analysis of enzyme-altered liver foci in rats initiated with diethylnitrosamine and promoted with 2,3,7,8-tetrachlorodebenzo-p-dioxin or 1,2,3,4,6,7,8-heptachlorodibenzo-p-dioxin. Toxicol.Appl.Pharmacol. 138: 31-42

25. Wiebel FJ, Klose U, and Kiefer F. 1991. Toxicity of 2,3,7,8-tetrachlorodibenzo-p-dioxin in vitro: H4IIEC3-derived 5L hepatoma cells as a model system. Toxicol.Lett. 55: 161- 169

26. Wolfle D, Münzel P, Fischer G, and Bock KW. 1988. Altered growth control of rat hepatocytes after treatment with 3, 4,3',4'-tetrachlorobiphenyl in vivo and in vitro. Carcinogenesis 9: 919-924

27. Kamath AB, Xu H, Nagarkatti PS, and Nagarkatti M. 1997. Evidence for the induction of apoptosis in thymocytes by 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicol.Appl.Pharmacol. 142: 367-377

28. McConkey DJ, Hartzell P, Duddy SK, Hakansson H, and Orrenius S. 1988. 2,3,7,8- Tetrachlorodibenzo-p-dioxin kills immature thymocytes by Ca2+-mediated endonuclease activation. Science 242: 256-259

29. McConkey DJ and Orrenius S. 1989. 2,3,7,8-tetrachlorodibenzo-p-dioxin kills glucocorticoid- sensitive thymocytes in vivo. Biochem.Biophys.Res.Comm. 160: 1003- 1008

30. Worner W and Schrenk D. 1996. Influence of liver tumor promoters on apoptosis in rat hepatocytes induced by 2-acetylaminofluorene, ultraviolet light, or transforming growth factor beta 1. Cancer Res. 56: 1272-1278

31. Fernandez-Salguero P, Hilbert DM, Rudikoff S, Ward JM, and Gonzalez FJ. 1996. Aryl hydrocarbon receptor-deficient mice are resistant to 2,3,7,8-tetrachlorodibenzo-p-dioxin- induced toxicity. Toxicol.Appl.Pharmacol. 140: 173-179

32. Mimura J, Yamashita K, Nakamura K, Morita M, Takagi TN et al. 1997. Loss of teratogenic response to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in mice lacking the Ah (dioxin) receptor. Genes Cells 2: 645-654

33. Hoffman EC, Reyes H, Chu F-F, Sander F, Conley LH et al. 1991. Cloning of a factor required for activity of the Ah (dioxin) receptor. Science 252: 954-958

34. Hankinson O. 1995. The aryl hydrocarbon receptor complex. Annu.Rev.Pharmacol.Toxicol. 35: 307-340

35. Sutter TR and Greenlee WF. 1992. Classification of members of the Ah gene battery. Chemosphere 25: 223-226

36. Whitlock JPJ. 1993. Mechanistic aspects of dioxin action. Chem.Res.Toxicol. 6: 754-763

37. Gaido KW, Maness SC, Leonard LS, and Greenlee WF. 1992. 2,3,7,8- Tetrachlorodibenzo-p-dioxin-dependent regulation of transforming growth factors-alpha and -beta 2 expression in a human keratinocyte cell line involves both transcriptional and post-transcriptional control. J.Biol.Chem. 267: 24591-24595

38. Guo J, Sartor M, Karyala S, Medvedovic M, Kann S et al. 2004. Expression of genes in the TGF-beta signaling pathway is significantly deregulated in smooth muscle cells from aorta of aryl hydrocarbon receptor knockout mice. Toxicol.Appl.Pharmacol. 194: 79-89

39. Wang X, Harris PKW, Ulrich R, and Voorman RL. 1996. Identification of dioxin- responsive genes in HepG2 cells using differential mRNA display RT-PCR. Biochem.Biophys.Res.Comm. 220: 784-788

40. Strobeck MW, Fribourg AF, Puga A, and Knudsen ES. 2000. Restoration of retinoblastoma mediated signaling to Cdk2 results in cell cycle arrest. Oncogene 19: 1857-1867

41. Puga A, Maier A, and Medvedovic M. 2000. The transcriptional signature of dioxin in human hepatoma HepG2 cells. Biochem.Pharmacol. 60: 1129-1142

42. Wang F, Wang W, and Safe S. 1999. Regulation of constitutive gene expression through interactions of Sp1 protein with the nuclear aryl hydrocarbon receptor complex. Biochemistry 38: 11490-11500

43. Ge N-L and Elferink CJ. 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblatoma protein. J.Biol.Chem. 273: 22708-22713

44. Puga A, Barnes SJ, Dalton TP, Chang C, Knudsen ES, and Maier MA. 2000. Aromatic hydrocarbon receptor interaction with the retinoblastoma protein potentiates repression of E2F-dependent transcription and cell cycle arrest. J.Biol.Chem. 275: 2943-2950

45. Nguyen TA, Hoivik D, Lee JE, and Safe S. 1999. Interactions of nuclear receptor coactivator/corepressor proteins with the aryl hydrocarbon receptor complex. Arch.Biochem.Biophys. 367: 250-257

46. Wang S and Hankinson O. 2002. Functional involvement of the Brahma/SWI2-related gene 1 protein in cytochrome P4501A1 transcription mediated by the aryl hydrocarbon receptor complex. J.Biol.Chem. 277: 11821-11827

47. Beischlag TV, Wang S, Rose DW, Torchia J, Reisz-Porszasz S et al. 2002. Recruitment of the NCoA/SRC-1/p160 family of transcriptional coactivators by the aryl hydrocarbon receptor/aryl hydrocarbon receptor nuclear translocator complex. Mol.Cell Biol. 22: 4319-4333

48. Ma Q and Whitlock JPJ. 1996. The aromatic hydrocarbon receptor modulates the Hepa 1c1c7 cell cycle and differentiated state independently of dioxin. Mol.Cell Biol. 16: 2144- 2150

49. Weiss C, Kolluri SK, Kiefer F, and Gottlicher M. 1996. Complementation of Ah receptor deficiency in hepatoma cells: negative feedback regulation and cell cycle control by the Ah receptor. Exp.Cell Res. 226: 154-163

50. Trimarchi JM and Lees JA. 2002. Sibling rivalry in the E2F family. Nat.Rev.Mol.Cell Biol. 3: 11-20

51. Dyson N. 1998. The regulation of E2F by pRB-family proteins. Genes Dev. 12: 2245- 2262

52. Bernard HP, Darlington GJ, and Ruddle FH. 1973. Expression of liver phenotypes in cultured mouse hepatoma cells: synthesis and secretion of serum albumin. Dev.Biol. 35: 83-96

53. Sun W, Zhang J, and Hankinson O. 1997. A mutation in the aryl hydrocarbon receptor (AHR) in a cultured mammalian cell line identifies a novel region of AHR that affects DNA binding. J.Biol.Chem. 272: 31845-31854

54. Brooks SC, Locke ER, and Soule HD. 1973. Estrogen receptor in a human cell line (MCF-7) from breast carcinoma. J.Biol.Chem. 248: 6251-6253

55. Scheffner M, Munger K, Byrne JC, and Howley PM. 1991. The state of the p53 and retinoblastoma genes in human cervical carcinoma cell lines. Proc.Natl.Acad.Sci.U.S.A 88: 5523-5527

56. Chang C-Y and Puga A. 1998. Constitutive activation of the aromatic hydrocarbon receptor. Mol.Cell.Biol. 18: 525-535

57. Fukunaga BN, Probst MR, Reisz-Porszasz S, and Hankinson O. 1995. Identification of functional domains of the aryl hydrocarbon receptor. J.Biol.Chem. 270: 29270-29278

58. Whitaker LL, Su H, Baskaran R, Knudsen ES, and Wang JY. 1998. Growth suppression by an E2F-binding-defective retinoblastoma protein (RB): contribution from the RB C pocket. Mol.Cell Biol. 18: 4032-4042

59. Knudsen ES and Wang JYJ. 1996. Differential regulation of retinoblastoma protein function by specific cdk phosphorylation sites. J.Biol.Chem. 271: 8313-8320

60. Probst MR, Reisz-Porszasz S, Agbunag RV, Ong MS, and Hankinson O. 1993. Role of the aryl hydrocarbon receptor nuclear translocator protein in aryl hydrocarbon (dioxin) receptor action. Mol.Pharmacol. 44: 511-518

61. Weinmann AS and Farnham PJ. 2002. Identification of unknown target genes of human transcription factors using chromatin immunoprecipitation. Methods 26: 37-47

62. Wells J and Farnham PJ. 2002. Characterizing transcription factor binding sites using formaldehyde crosslinking and immunoprecipitation. Methods 26: 48-56

63. Wang W, Smith IR, and Safe S. 1998. Aryl hydrocarbon receptor-mediated antiestrogenicity in MCF-7 cells: modulation of hormone-induced cell cycle enzymes. Arch.Biochem.Biophys. 356: 239-248

64. Gottlicher M, Cikryt P, and Wiebel FJ. 1990. Inhibition of growth by 2,3,7,8- tetrachlorodibenzo-p-dioxin in 5L rat hepatoma cells is associated with the presence of Ah receptor. Carcinogenesis 11: 2205-2210

65. Göttlicher M and Wiebel FJ. 1991. 2,3,7,8-Tetrachlorodibenzo-p-dioxin causes unbalanced growth in 5L rat hepatoma cells. Toxicol.Appl.Pharmacol. 111: 496-503

66. Kolluri SK, Weiss C, Koff A, and Göttlicher M. 1999. p27kip1 induction and inhibition of proliferation by the intracellular Ah receptor in developing thymus and hepatoma cells. Genes Dev. 13: 1742-1753

67. Reisz-Porszasz S, Probst MR, Fukunaga BN, and Hankinson O. 1994. Identification of functional domains of the aryl hydrocarbon receptor nuclear translocator protein (ARNT). Mol.Cell.Biol. 14: 6075-6086

68. Ma Q, Dong L, and Whitlock JPJ. 1995. Transcriptional activation by the mouse Ah receptor. Interplay between multiple stimulatory and inhibitory functions. J.Biol.Chem. 270: 12697-12703

69. Harbour JW and Dean DC. 2000. The Rb/E2F pathway: expanding roles and emerging paradigms. Genes Dev. 14: 2393-2409

70. Trouche D, Cook A, and Kouzarides T. 1996. The CBP co-activator stimulates E2F1/DP1 activity. Nucleic Acids Res. 24: 4139-4145

71. Marzio G, Wagener C, Gutierrez MI, Cartwright P, Helin K, and Giacca M. 2000. E2F family members are differentially regulated by reversible acetylation. J.Biol.Chem. 275: 10887-10892

72. Kobayashi A, Numayama-Tsuruta K, Sogawa K, and Fujii-Kuriyama Y. 1997. CBP/p300 functions as a possible transcriptional coactivator of Ah receptor nuclear translocator (Arnt). J.Biochem.(Tokyo) 122: 703-710

73. Elferink CJ, Ge NL, and Levine A. 2001. Maximal Aryl Hydrocarbon Receptor Activity Depends on an Interaction with the Retinoblastoma Protein. Mol.Pharmacol. 59: 664-673

74. Chan HM, Smith L, and La Thangue NB. 2001. Role of LXCXE motif-dependent interactions in the activity of the retinoblastoma protein. Oncogene 20: 6152-6163

75. Koliopanos A, Kleeff J, Xiao Y, Safe S, Zimmermann A et al. 2002. Increased arylhydrocarbon receptor expression offers a potential therapeutic target for pancreatic cancer. Oncogene 21: 6059-6070

76. Hestermann EV, Stegeman JJ, and Hahn ME. 2002. Relationships among the cell cycle, cell proliferation, and aryl hydrocarbon receptor expression in PLHC-1 cells. Aquat.Toxicol. 58: 201-213

77. Abdelrahim M, Smith R, III, and Safe S. 2003. Aryl Hydrocarbon Receptor Gene Silencing with Small Inhibitory RNA Differentially Modulates Ah-Responsiveness in MCF-7 and HepG2 Cancer Cells. Mol.Pharmacol. 63: 1373-1381

78. Levine-Fridman A, Chen L, and Elferink CJ. 2004. Cytochrome P4501A1 promotes G1 phase cell cycle progression by controlling aryl hydrocarbon receptor activity. Mol.Pharmacol. 65: 461-469

79. Ait-Si-Ali S, Polesskaya A, Filleur S, Ferreira R, Duquet A et al. 2000. CBP/p300 histone acetyl-transferase activity is important for the G1/S transition. Oncogene 19: 2430-2437

Table 1. Gene-specific primer sets for real-time PCR analysis of re lative mRNA expression levels

Gene Name Accession Number Forward Primer (5'-3 ') Reverse Primer (5'-3') Product Size

β-actin M12481 CATCCGTAAAGACCTCTATGC C ACGCAGCTCAGTAACAGTCC 287 Cyp1a1 NM_009992 GTGTCTGGTTACTTTGACAAG TGG AACATGGACATGCAAGGACA 199 Cdk2 NM_183417 CATCTTTGCTGAAATGCACC GCTAAAATCTTGCCGAGCC 319

Cyclin E NM_007633 TCTTTCATCCCCACCCCTAAC ACACCTCCATCAGCCAATCC 253 Dhfr NM_010049 CAGGAGGAAAAAGGCATCAA ATCCCCAGGATCACAAAACA 203 DNA pol α NM_008892 TCTTCCCCTTCATTTCTCCC AACTGCTCTGCTATGTTCTTG 240 p27Kip1 NM_009875 GCAGCTTGCCCGAGTTCTAC TTCTTGGGCGTCTGCTCCAC 325

Table 2. Promoter-specific primer sets for chromatin immun oprecipitation analysis

Gene Name Forward Primer (5'-3') Reverse Primer (5'-3') Product Size

Cyp1a1 CTATCTCTTAAACCCCACCCCAA CTAAGTATGG TGGAGGAAAGGGTG 357 Cdk2 CAGCCCTTGACAATTTGTCC TCCGTAG ACCAGAAACACC 275

Cyclin E CCTGACATCTAGCCCCAC GGACATTTAAAAATCCCAGCG 213

Dhfr CGGCAATCCTAGCGTGAAGGCTGGT TCTCCGTTCTTGCCAATC 215 DNA pol α ACGCTCCAGACGCGCACTAC TTGGAAC GGGAAGCGGAAG 103

Chapter III Figure Legends

Fig. 1. TCDD inhibits S-phase progression in MCF-7 and Hepa-1 cell lines. A. MCF-7 breast carcinoma cells were serum-starved in minimal essential medium-α for 3 days. Cells were released into medium containing 5% fetal bovine serum and either DMSO vehicle or 10 nM TCDD for 0 to 30 hours. Cells were pulse-labeled with BrdU for 1 hour at the indicated times. Cells were fixed and processed for BrdU incorporation to detect DNA synthesis and stained with 7-Amino-actinomycin D to determine cell cycle phase by flow cytometry.

Percentage of S-phase cells was determined by gating for the BrdU-positive cell population.

Asterisks (*) indicate significant differences (P<0.05) from DMSO-treated controls using one- way ANOVA. B. Hepa-1c1c7 hepatoma cells were cultured to 80-90% confluence and treated with DMSO vehicle or 10 nM TCDD for 16 or 24 hours. BrdU labeling and flow cytometry analysis were performed as in A. Asterisks (*) indicate significant differences (P<0.05) from

DMSO-treated controls using one-way ANOVA. C. Hepa-1 cells were cultured and treated as in

B. Cell cycle distribution was then determined by flow cytometry. DNA content (7-AAD intensity) is plotted against cell number. The percentage of cells in G0-G1, S, and G2-M phases was determined by ModFit software. 7-AAD, 7-Amino-actinomycin D; DMSO, vehicle control.

Fig. 2. TCDD inhibits expression of E2F-dependent, S-phase specific genes. Hepa-1c1c7 cells were cultured in the presence of serum to 80-90% confluence and treated with vehicle,

10nM TCDD, 1 µg/mL actinomycin D, or actinomycin D plus 10 nM TCDD for the time periods indicated. Thereafter, cells were harvested for total RNA isolation and mRNA of the indicated genes was quantified by real-time PCR amplification. Products were confirmed by agarose gel electrophoresis. The ordinate represents threshold cycle (Ct) values normalized to β-actin. The

values shown represent the mean ± S.D. of at least two determinations from one representative

experiment. Cyp1a1, Cytochrome P4501A1; p27Kip1, Cyclin-dependent kinase inhibitor 1B;

Cdk2, Cyclin-dependent kinase 2; Cyclin E, Cyclin E1; Dhfr, Dihydrofolate reductase; Pol α,

DNA polymerase alpha 1; DMSO, vehicle control.

Fig. 3. AHR-mediated gene repression does not require transactivation. A. AHR deletion mutant series used for subsequent binding and transcription assays. Two domains previously implicated in RB binding, the LXCXE motif and the glutamine rich-domain, are shown.

Additional elements include the bHLH domain required for dimerization, DNA binding, and nuclear localization, and the PAS domain implicated in ARNT dimerization and ligand binding.

B. Full-length AHR is required for maximal binding to RB. Bacterially synthesized vector or

RB protein tagged with His6 at the amino terminus was incubated with nickel-agarose slurry, cleared through a column, and stored at 40C. Control and RB-bound columns were established

using 100 µL aliquots of the pre-adsorbed slurry. Full-length and truncation peptides of the

mouse AHR were in vitro translated and labeled with [35S]methionine. Equal volumes of AHR

or truncated peptides were recycled 10 times through control and RB-bound columns. Bound

proteins were eluted using 8 M urea, and eluted fractions were denatured by boiling and analyzed

by SDS-PAGE in 7.5% acrylamide gels. Detection of [35S]AHR was by autoradiography. The

graph represents quantitation of bound protein, normalized to control, by PhosphorImager

analysis. C. AHR-dependent transactivation is impaired with expression of AHR mutant

proteins. C33A cell cultures were transfected with the control plasmid pCMVβgal together with

the reporter pAhRDtkLuc3. Additional cotransfected plasmids were as follows: ---, vector;

wtAHR, pcDNAI/B6AHR; AHR mutants are as described in the Experimental Procedures. In

addition, the plasmid pcDNAIneo/mARNT was co-transfected with all AHR-expressing

plasmids. The ordinate represents luciferase activity normalized to β-galactosidase. The values shown represent the mean ± S.D. from one representative experiment of at least two independent trials. D. AHR-mediated transactivation is not required for repression of E2F-dependent transcription. C33A cell cultures were transfected with the control plasmid pCMVβgal together with the reporter p3×E2FLuc. Additional cotransfected plasmids were as follows: ---, vector;

RB, pCMVNeoRB; wtAHR, pcDNAI/B6AHR; AHR mutants are as described in the

Experimental Procedures. In addition, the plasmid pcDNAIneo/mARNT was co-transfected with all AHR-expressing plasmids. The ordinate represents luciferase activity normalized to β- galactosidase. The values shown represent the mean ± S.E. of at least twelve log-transformed determinations from four independent experiments. Asterisks (*) indicate significant differences

(P<0.05) from RB-transfected cells using one-way ANOVA. DMSO, vehicle control.

Fig. 4. ARNT dimerization is not required for AHR-mediated gene repression. A.

Formation of an AHR/ARNT complex is not required for repression of E2F-dependent transcription. c4 cell cultures were transfected with the control plasmid pCMVβgal together with the reporter plasmid p3×E2FLuc. Additional cotransfected plasmids were as follows: ---, vector; RB, pCMVNeoRB; AHR, pcDNAI/B6AHR; ARNT, pcDNAIneo/mARNT. The ordinate represents luciferase activity normalized to β-galactosidase. The values shown represent mean ± S.D. from one representative experiment of at least three independent trials. B. Hepa-1

cell cultures (left panel) or the c35 cell line harboring a mutant AHR defective in DNA-binding

(right panel) were transfected with the control plasmid pCMVβgal together with the reporter

p3×E2FLuc. Additional cotransfected plasmids were as follows: ---, vector; RB, pCMVNeoRB.

The ordinate represents luciferase activity normalized to β-galactosidase. Cells were treated

with DMSO vehicle or with 10 nM TCDD in DMSO for 24 h following transfection. The values

shown represent the mean ± S.D. from one representative experiment of at least two independent trials.

Fig. 5. AHR represses E2F-dependent gene expression by blocking p300 coactivator activity. Hepa-1 cell cultures were transfected with the control plasmid pCMVβgal together with the reporter p3×E2FLuc. Additional cotransfected plasmids were as follows: ---, vector;

RB, pCMVNeoRB; HDAC, pCMVhuHDAC1; BRG1, p300, and DNp300 are as described in the

Experimental Procedures. The ordinate represents luciferase activity normalized to β- galactosidase. Cells were treated with DMSO vehicle (open bars) or with 10 nM TCDD in

DMSO (hatched bars) for 24 h following transfection. The values shown represent the mean ± S.D. from one representative experiment of at least two independent trials. Asterisks (*) indicate significant differences (P<0.05) from DMSO-treated controls using one-way ANOVA.

DMSO, vehicle control.

Fig. 6. Recruitment of AHR to E2F-regulated promoters results in loss of p300. Hepa-1 cell were cultured as stated in the legend to Fig. 3 and treated with either DMSO vehicle or 10 nM TCDD for 1.5 h. Cells were harvested and native chromatin from nuclear lysates was prepared. Immunoprecipitations were carried out using antibodies to AHR, E2F1, HDAC-1, p300, and RB as indicated. A control (No Ab) using no antibody was included to confirm the specificity of the precipitations. Purified genomic DNA was subjected to PCR amplifications with gene-specific primers. Input lane represents promoter-specific amplification of 0.2% of the total chromatin sample. Products were separated and analyzed using 10% polyacrylamide gel electrophoresis. See Figure 2 legend for gene abbreviations.

Figure 1

C 1500

800 GO-G1: 43.7% GO-G1: 64.3% GO-G1: 53.2% GO-G1: 69.7% S: 39.8% S: 31.3% S: 34.8% 2500 S: 24.5% G2-M: 16.3% 1500 G2-M: 4.4% G2-M: 12.0% G2-M: 5.9% 600 1000 lls e 1500

c 1000 f 400

o # 500 500 200 500

0 0 0 0 200 400 600 200 400 600 200 400 600 200 400 600 7-AAD 7-AAD 7-AAD 7-AAD

DMSO, 16h TCDD, 16h DMSO, 24h TCDD, 24h

Figure 2

Figure 3

A LXCXE (325-329) bHLH PAS Q-rich AHR 1 121 374 805 ∆ 323-494

∆ 323-607 ∆495-805

Figure 4

100

Figure 5

101

Figure 6

DMSO TCDD

b C b C t 1 1 u A R A 0 R A 0 F 0 A F 0 p o H 2 3 B o H 2 D 3 B In N A E HD p R N A E H p R Cyp1a1

Cdk2

Cy clin E

Dhfr

Pol α

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Chapter IV

Modulation of E2F-dependent gene expression by an interaction between the AHR and E2F proteins

Jennifer L. Marlowe, Xiaoqing Chang and Alvaro Puga*

*Corresponding author:

Keywords: AHR, TCDD, E2F, cell cycle, chromatin immunoprecipitation

Abstract

Dioxin (TCDD) exposure results in a large number of cellular and molecular perturbations. One of the major effects of dioxin in cells is to disrupt normal cell cycle progression, either through impeding or promoting cell division, differentiation, or death, depending on the cell type under investigation. Most if not all of the effects of TCDD are mediated through its activation of the aryl hydrocarbon receptor (AHR) transcription factor.

Several laboratories have recently identified one potential mechanism by which activation of the

AHR by TCDD acts to inhibit cell cycle progression. The AHR can form protein complexes with the retinoblastoma (RB) protein and inhibit its normal inactivation during the G1/S transition of the cell cycle. The effect of the AHR-RB interaction is thus to preclude E2F- dependent gene expression required for cell cycle progression. In the current investigation, we

have identified an independent interaction of the AHR with E2F family proteins. The activation

of AHR by TCDD appears to act to inhibit E2F activity even in the absence of RB, likely

through the interaction of AHR with E2F. However, we have also discovered a novel role for

the AHR in regulating the constitutive activities of a number of E2F-regulated, cell cycle-

dependent genes. Cells lacking either AHR or E2F1 show a similar reduction in expression of

roughly 10 known E2F-regulated genes. The expression levels of at least half of these genes are

also decreased in TCDD-treated cells. Experiments using chromatin immunoprecipitation and

promoter tiling arrays indicate that the AHR constitutively targets these genes through canonical

AHR binding sites, and may only be recruited to sites of E2F binding upon TCDD treatment. In

all, these experiments reveal the context-specific transcriptional effects of the basal- and TCDD-

induced activities of the AHR.

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Introduction

TCDD is known for its potent activity as a tumor promoter, as well as for its ability to cause a plethora of seemingly unrelated biological responses in different tissues, organ systems, and species. The mechanisms by which TCDD acts to cause such a wide variety of effects are

still being discovered. It is widely accepted that the vast majority of these effects are mediated

by the aryl hydrocarbon receptor, as cells or organisms which lack AHR expression exhibit no

TCDD-induced toxicity. It is, however, becoming increasingly clear that the effects which do

occur as the result of AHR activation by TCDD are cell-type specific, at least in part due to the

variety of proteins and signal transduction pathways with which the AHR interacts. In terms of

its effects on cell cycle progression, the interaction of the AHR with RB-E2F proteins appears to

be an important determinate of the biological consequences of TCDD exposure.

E2F transcription factor family members are downstream targets of the retinoblastoma

protein family, and are important regulators of the G1/S phases of the cell cycle. However, the

E2Fs also have functions beyond the S-phase transition and impact cell proliferation in multiple different ways (1). The E2F family consists of 8 different members, the first 5 of which have been extensively characterized and the last 3 of which are less well-studied. Generally, E2F1,

E2F2 and E2F3 are potent activators of transcription, bind exclusively to the RB protein, and are cyclically expressed during the cell cycle. E2F4, which interacts with the RB-family members pRB, p107, and p130, and E2F5, which only binds p130, are poor transcriptional activators and function more as repressors through their recruitment of RB proteins to E2F-regulated promoters

(2;3). In general, the transcriptional activator E2Fs promote cell cycle progression, whereas the

E2Fs which repress transcription function in cell cycle exit and differentiation (1). E2F6 and

E2F7 are quite distinct from the other E2F members. Both lack the transactivation and RB-

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binding domains, and they repress transcription in an RB-independent manner (4). Little is known of E2F8, a newly discovered member of the E2F family.

As stated above, the most intensively studied and well-characterized function of E2F is its ability to regulate the G1/S-phase transition of the cell cycle. E2F proteins exert control over

the cell cycle through modulating the transcription of a variety of essential cell cycle control

genes, including cell cycle regulators, pRB and related pocket proteins, enzymes for nucleotide

biosynthesis, and proteins required for DNA replication (5). Interactions between E2F and RB

affect E2F-dependent transcription in two ways. First, the physical binding of RB to E2F

proteins inhibits the transcriptional transactivation activity of E2F. Second, repressor E2Fs

recruit RB proteins to gene promoters, and through the association of RB with chromatin-

modifying co-repressor proteins, mediate active repression of E2F-responsive genes (2;6;7).

Repressor E2F/RB complexes are prevalent in G0/early G1 phases, but these complexes disband

in late G1. This disruption is caused primarily by the phosphorylation of RB proteins by Cyclin

D-Cdk4/6 and Cyclin E-Cdk2 complexes. Late in G1, activator E2Fs turn on the transcription of genes required for entry into S-phase and DNA synthesis (5;8;9). However, the ability of a particular E2F protein to either induce S-phase, senescence, or some other outcome is dependent on the cellular context and on the specific sets of genes which are up- and down-regulated by

E2F (1).

AHR activation alters several cell cycle and signaling pathways, including those required for normal cell cycle regulation. The previous chapter focused on the ability of the activated

AHR to delay progression of the cell cycle during the G1 to S-phase transition. As this phase of the cell cycle is primarily controlled by the RB and E2F families of transcription factors, it is likely that the AHR would exert its effects on the cell cycle through some mechanism affecting

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these protein families. The presence of a canonical RB-binding motif in the AHR protein made it the most likely candidate, and several labs have focused on the characterization of this interaction as the principal mediator of AHR-dependent cell cycle delay (10-13). The ligand- activated AHR appears to cooperate with RB in its ability to repress E2F-dependent transcription and delay cell cycle progression (13;14). However, results obtained during our investigation of

the interactions of AHR with RB suggested that AHR directly interacts with and modulates the transcriptional activity of E2F as well. Published results in Hepa-1 cells as well as unpublished results in several other cell lines indicate that TCDD stimulation or ectopic expression of the

AHR inhibits expression of E2F-dependent reporter genes, and the extent of inhibition is equal to that exhibited by RB alone (13). The goal of this chapter is to characterize the E2F-specific interactions of the AHR, and to determine the impact of the AHR on E2F-dependent gene expression.

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Materials and Methods

Cell Lines, Growth Conditions, and Chemical Treatments

The mouse Hepa-1 hepatoma cell line (15), the human breast carcinoma cell line MCF-7

(16), and the adult African green monkey kidney CV-1 cell line (17) were maintained in minimal essential medium-α (Invitrogen) supplemented with 5% fetal bovine serum (Invitrogen), 1.0%

o 100X antibiotic/antimycotic solution (Sigma), and 26 mM NaHCO3 at 37 C in a humidified 5%

CO2 atmosphere. The human cervical cancer C33A cell line and the human osteosarcoma Saos-2 cell line (18) were similarly cultured in medium supplemented with 10% fetal bovine serum.

Wild type mouse embryonic fibroblasts (MEFs) from strain C57Bl6 were prepared by standard techniques from 14.5-day-old fetuses and grown in α-MEM medium as above. MEFs from RB-/-

and E2F1-/- mice were provided by E. Knudsen. AD-293 cells were obtained from Stratagene

(La Jolla, CA) and cultured in Dulbecco’s Modified Eagle Medium (Invitrogen) containing 10%

fetal bovine serum, 1.0% antibiotic/antimycotic solution, and 2 mM L-Glutamine (Invitrogen).

When indicated, cultures were treated with TCDD at a final concentration of 10 nM in Me2SO vehicle, and control cultures were treated with an equivalent volume of Me2SO.

C33A, Saos-2, and CV-1 cells do not express AHR, as determined from the lack of

expression of a transfected AHR/ARNT-responsive luciferase reporter plasmid and by

immunoblot analysis. Hepa-1 cells do not express RB, as determined by both immunoblot and

RT-PCR analysis. C33A cells harbor a mutation in the RB gene and express an unstable mutant

RB protein (19).

Transfections and Reporter Assays

Transient transfection experiments were performed using Lipofectamine Plus

(Invitrogen) on cells grown to 50-75% confluence in 24-well plates. All transfection mixtures

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were brought to the same amount of total DNA by the addition of the appropriate amount of empty vector. To control for variations in transfection efficiency, cells were also co-transfected with a pCMVβgal plasmid (Clontech) and transfections were normalized to β-galactosidase

expression. Cells were transfected in serum-free medium, and 3 h later one volume of medium

containing twice the final concentration of culture serum was added. After overnight incubation,

cells were washed once with phosphate-buffered saline and cultured in complete culture medium

with either TCDD or vehicle. Between 24 - 48 h after transfection, cells were washed twice with

PBS and lysed with 100 µl of reporter lysis buffer (Promega). Aliquots (50 µl) of cell lysate

were used to measure luciferase and β-galactosidase activities. Light units were determined

immediately upon the addition of 150 µl of luciferase assay buffer (20 mM tricine, 1.07 mM

MgCO2, 2.67 mM MgSO4, 33.3 mM dithiothreitol, 14.8 mg coenzyme A, 530 µM ATP, 0.1

mM EDTA, 10 mg luciferin), using a Wallac Victor2 1420 plate reader. β-galactosidase activity was determined by the addition of 50 uL of a 2X assay buffer (20mM PO4 buffer (pH 7.3), 2mM

MgCl2, 100mM β-mercaptoethanol, 1.33 mg/mL o-nitrophenyl-β-D-galactopyranoside).

Absorbance at 420 nm was measured following incubation of the samples for 30 minutes. Each

transfection was repeated a minimum of two times, and the results shown are the mean ± S.D. of

one representative set of results. Where noted, the results shown are the mean ± SE of log-

transformed results from multiple independent experiments.

Plasmid constructs

The plasmid pcDNAI/B6AHR, used to express the high affinity variant of murine AHR,

has been described (20). Mutant derivatives of the pcDNAI/B6AHR plasmid have also been

described (11). Briefly, plasmid p∆495-805 encodes a dominant-negative variant of the murine

AHR, expressing a protein capable of ligand binding and dimerization with ARNT but lacking

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transactivation activity (20). The plasmid p∆323-494 contains a deletion of the LXCXE RB- binding motif and the PAS B domain, and plasmid p∆323–607 lacks the LXCXE motif, the PAS

B domain, as well as part of the transactivation domain (21). Additional mutant derivatives of the pcDNAI/B6AHR plasmid, ∆376-805 and ∆1-425, have been described (13). Expression of

the peptides encoded by each AHR truncation mutant was confirmed by Western blot. The

reporter p3×E2FLuc, containing three E2F-responsive elements, has been previously described

(22). Other plasmids used directed the expression of human RB (pCMVNeoRB) (23). The

empty vectors pBlueScript II and pCMVScript (Stratagene) were used in transient transfection

experiments to bring all transfection groups to the same amount of total DNA.

The AdEasy XL Adenoviral Vector System (Stratagene) was used to generate

recombinant adenovirus for the expression of the high affinity variant of the murine AHR. The

pcDNAI/B6AHR vector was digested with HindIII, and the 2.6 kb fragment containing the AHR

gene was gel purified and cloned into the pShuttle-CMV vector digested with HindIII and

EcoRV. The pShuttle-CMV-B6AHR plasmid was then linearized using PmeI and transformed

into BJ5183-AD-1 cells, containing the adenovirus plasmid pAdEasy-1, to produce the

recombinant adenoviral plasmid pAdB6AHR. For control experiments, pShuttle-CMV-LacZ

was similarly processed. Finally, recombinants were transformed into XL10-Gold cells for

plasmid amplification.

In Vitro Transcription/Translation

One microgram of pcDNAI/B6AHR plasmid encoding the mouse high-affinity AHR

protein was used in 50-µl in vitro transcription/translation reactions with a TNT-coupled

reticulocyte lysate system (Promega). Reactions were prepared according to the manufacturer's

instructions and included 30 µCi of [35S]methionine (Amersham Pharmacia Biotech). Reactions

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were incubated at 30oC for 1.5 h. Translated proteins were used immediately for pull-down assays.

Pull-down Assays

Mutant RB and E2F1 GST fusion proteins were expressed in bacteria and 10 mL lysates were produced by suspension and sonication of cells in NET buffer containing protease inhibitors (20 mM Tris (pH 8.0), 100 mM NaCl, 1 mM EDTA, 10 mM NaF, 10 mM sodium pyrophosphate, 10 µg/mL aprotinin, 10 µg/mL leupeptin). Lysates were incubated overnight at

4oC with 50 µL of in vitro synthesized full-length [35S]AHR, and passed 5 times over columns

containing 500 µL of a glutathione-sepharose slurry (Amersham Pharmacia Biotech) pre-washed

with phosphate-buffered saline. Columns were washed thoroughly with PBS, and a final 500 µL

wash was collected to confirm the absence of protein in the flow through fraction by Western

blot analyses. Bound proteins were eluted with three 500 µL washes of a buffer containing 10

mM GSH. Elutions were combined and concentrated for analysis. Samples were denatured by

boiling and analyzed by SDS-PAGE in 7.5% acrylamide gels. Detection of [35S]AHR was by

autoradiography, and detection of GST fusion proteins was accomplished using an anti-GST

antibody.

Electrophoretic Mobility Shift Assays

Nuclear extracts were prepared essentially as described (24;25). Nuclear extracts of CV-

1 cells were prepared in a final volume of 100 µL buffer containing 0.1 mM EGTA, 1 mM

dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride, 420 mM NaCl, 1.5 mM MgCl2, 25%

glycerol, and 25 mM HEPES (pH 7.9), with addition of 10 mM NaF, 10 mM sodium

pyrophosphate, 10 µg/mL aprotinin, and 10 µg/mL leupeptin to inhibit phosphatase activity.

DNA binding reactions were performed in a 20 µL reaction volume with 10,000 dpm

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(approximately 0.1 ng) of double-stranded probe and 2–5 µg nuclear protein, in buffer containing

1 mM EGTA, 1 mM dithiothreitol, 6 mM MgCl2, 10% glycerol, 0.1% NP-40, 1 µg Salmon

Sperm DNA, 10 µg BSA, and 20 mM HEPES (pH 7.8). The double-stranded E2F-binding probe was synthesized using the oligonucleotide sequences 5’-AGCAATTTCGCGC-3’ and 5’-

CAAGTTTGGCGCGAAATTGCT-3’. Labeling was carried out in 15 µL reactions containing 4

µg annealed oligonucleotides, 50 mM NaCl, 10 mM Tris (pH 7.9), 10 mM MgCl2, 1 mM dithiothreitol, 5 units T4 DNA polymerase (New England Biolabs), 0.2 mM dATP dGTP dTTP, and 100 µCi [32P]dCTP (PerkinElmer). Binding reactions were incubated for 20 min at room temperature and samples were loaded onto non-denaturing 4% polyacrylamide gels. Following electrophoresis at 200 V for 2–3 hr in 0.5X Tris–borate buffer, gels were dehydrated and exposed to X-ray film. For antibody supershift assays, 1 µL of 1 µg/µL anti-E2F1, anti-DP1,

anti-AHR (Santa Cruz Technologies), anti-RB polyclonal serum (26), or affinity-purified rabbit

anti-mouse bHLH and YGOR antibodies prepared in our laboratory (made against the AHR

peptides ∆376-805 and ∆1-425, respectively), were added to the DNA binding reaction mixture

and incubated on ice for an additional 20 min prior to electrophoresis. The truncated AHR

peptides ∆376-805 and ∆1-425 (13) were purified from bacteria and increasing amounts were added to the reactions for an additional 20 minute incubation at room temperature.

Adenovirus Production and Infections

5 AD-293 cells (Stratagene) were plated at a density of 8 X 10 cells per 60-mm dish 24

hours prior to transfection. Primary virus stocks were established by transfection of the plasmids

pAdB6AHR and pAdLacZ using Lipofectamine Plus as described above. Cells were maintained

in normal growth medium for 7 to 10 days, or until 80-90% of cells were detached from the

plate. Medium and cells were harvested, cell pellets were resuspended in 1 mL of PBS, and cells

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were lysed by four rounds of freeze/thaw in a dry ice-methanol bath. Approximately 100 µL of the primary virus stock was then used to infect AD-293 cells grown to 70-80% confluence in a

10 cm dish. Purified virus from this first amplification was then used to infect 30 15-cm plates of AD-293 cells for production of the final virus stock. Cytopathic effect was established in these cells by 48 to 72 hours post-infection. Virus was isolated using 0.45 µM filtration followed by cesium chloride density gradient purification. Titration of the purified viral stocks was accomplished using the Adeno-X Rapid Titer Kit from Clontech (Mountain View, CA).

Recombinant adenovirus for the expression of RB and E2Fs (27) was provided by E. Knudsen.

Saos-2 cells were infected with purified adenovirus containing genes for the expression of LacZ, AHR, RB, and E2F1-4. Infections were carried out using 100 PFU per cell for 3 hours in normal growth medium. Following infections, cells were washed two times with PBS and placed into normal growth medium. Cells were harvested after 24 hours for co- immunoprecipitation experiments.

Co-immunoprecipitation and Western Blot

For co-immunoprecipitations with Hepa-1 and MEF cells, cells were grown in 10-cm plates, 4 plates per treatment, to 80-90% confluence and treated with 10 nM TCDD for 2 hours prior to harvesting for nuclear extracts (24;25). Nuclear extracts were obtained in buffer containing 2 mM EDTA, 2 mM dithiothreitol, 0.4 M KCl, 10% glycerol, and 25 mM HEPES

(pH 7.9). Approximately 300 µg of nuclear extract was incubated with 2 µg of antibody to AHR

(BIOMOL International) or E2F1 (Santa Cruz Biotechnology) in 1 mL of IP buffer containing

1% Triton X-100, 150 mM NaCl, 10 mM Tris (pH 7.4), 1 mM EDTA, 1mM EGTA (pH 8.0), 0.2 mM Na3VO4, 0.5% NP-40, 0.2 mM PMSF, 10 µg/mL aprotinin, and 10 µg/mL leupeptin.

Incubations were carried out using 25 µL protein A and G beads (Upstate Biotechnology)

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overnight at 40C. Beads were washed with 1 mL of IP buffer 6 times, suspended in 30 uL of

electrophoresis buffer containing 250 mM Tris (pH 6.8), 4% SDS, 10% glycerol, 0.006%

bromophenol blue, and 2% β-mercaptoethanol, and boiled for 5 minutes. Eluted proteins were analyzed in 7.5% SDS-PAGE, and AHR (BIOMOL International) and E2F1 (Lab Vision

Corporation) were detected by Western blot after semidry electrotransfer of the proteins to polyvinylidene difluoride membranes, as described (20).

For co-immunoprecipitations with Saos-2 cells, cultures were grown in 15-cm plates to

80-90% confluence, infected with adenovirus for 24 hours as described above, and treated with

10 nM TCDD for 2 hours prior to harvesting for whole cell extracts. Cells were washed twice with ice-cold PBS, harvested by scraping, and collected by centrifugation. Pelleted cells were washed one additional time with cold PBS and transferred to microcentrifuge tubes. Cell pellets were then resuspended in 300 µL NET-N lysis buffer (100 mM NaCl, 1 mM EDTA, 0.5% NP-

40, 20 mM Tris (pH 8.0)) containing 10 µg/mL aprotinin, 10 µg/mL leupeptin, 1 mM PMSF, 10 mM NaF, and 10 mM Na2P2O7. Co-immunoprecipitation reactions were carried out as described

above, using approximately 1 mg of whole cell extract incubated with 2 ug of antibody to AHR

in 1 mL IP buffer. Eluted proteins were analyzed in 7.5% SDS-PAGE, and RB, AHR, E2F1,

E2F3, and E2F4 (E2Fs from Santa Cruz Biotechnology) were detected by Western blot after

semidry electrotransfer of the proteins to polyvinylidene difluoride membranes.

Total RNA isolation, reverse transcription, and real-time PCR

MEF cells were cultured in 10-cm plates and treated under standard culture conditions

with DMSO vehicle or with 5 nM TCDD for 8 hours. Each treatment was performed in

duplicate for all cell lines analyzed. Total cellular RNA was isolated with the RNeasy Mini Kit

(Qiagen Inc.). Immediately following heating at 65°C for 10 min, first strand cDNAs were

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synthesized from 2-20 µg of total RNA using SuperScript II reverse transcriptase (Life

Technologies, Inc.) and random primers. cDNAs were diluted in equivalent volumes of ddH20 based upon starting RNA concentrations, and subjected to PCR amplifications with gene-specific primer sets for the various E2F-responsive and control genes (Table 1). Reaction volumes of 25

µL contained 13 µL QTaq DNA Polymerase Mix (Clontech), 1.75 uL SYBR green, 0.4 µM primers, and 2 µL cDNA template. Real-time quantitative PCR was performed using an MJ

Research Opticon 96-well rapid thermal cycler and fluorescence was measured after each of the repetitive cycles. A typical protocol included a 10 min denaturation step at 95oC followed by 40 cycles of 95oC denaturation for 30 sec, annealing for 30 sec at a primer-optimized temperature, and 72oC extension for 30 sec. Detection of the fluorescent product was carried out during the

o 72 C extension period, and emission data were quantified using threshold cycle (Ct) values. Ct values for all genes analyzed were determined 2 to 6 times, averaged, and means were determined from the average Ct values for each biological duplicate. All means were then normalized to values for β-actin. The relative or fold-change from control Ct values (MEF+/+

cells, DMSO treated) was determined for each sample, as determined by the following

calculation:

Fold change = 2-∆∆Ct

where ∆∆Ct = (CtTarget – CtActin)Time=t – (CtTarget – CtActin)Time=0. PCR product specificity from each

primer pair was confirmed using melting curve analysis and subsequent agarose gel

electrophoresis.

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Chromatin Immunoprecipitation

The ChIP protocol was performed with minor modifications as described previously by others (28;29). Hepa-1 cells were grown in 15-cm plates to 90-100% confluence (approximately

2 x 107 cells). One plate of cells was used per treatment, per antibody. Cells were treated with

Me2SO vehicle or 10 nM TCDD for 1.5 hours. Cross-linking was performed by adding

formaldehyde directly to the culture media to a final concentration of 1%, followed by 10

minutes of incubation. The formaldehyde reaction was quenched using glycine at a final

concentration of 0.125 M. Cells were rinsed twice with cold PBS, scraped from the dishes,

pelleted and washed again with PBS containing 0.5 mM PMSF. Cell pellets were resuspended in

cell lysis buffer (5 mM PIPES (pH 8.0), 85 mM KCl, 0.5% NP-40, 0.5 mM PMSF, 5 µg/ml

leupeptin, 5 µg/ml aprotinin) and incubated on ice for 10 min. Cells were then dounced with a B

pestle 10 to 20 times on ice to aid in nuclei release. Nuclei were pelleted and resuspended in

nuclei lysis buffer (50 mM Tris-HCl (pH 8.1), 10 mM EDTA, 1% SDS, 0.5 mM PMSF, 5 µg/ml

leupeptin, 5 µg/ml aprotinin) and incubated on ice for 10 min. Chromatin was sonicated to an

average length of 600 bp with 4 10-second pulses of 30 W, maintaining samples on ice for 30

seconds between pulses. Sonicated chromatin was then precleared for 15 min at 4°C with BSA-

and salmon sperm DNA-saturated protein A agarose (Upstate Biotechnology) in preparation for

immunoprecipitation. The supernatant was divided equally among all samples and incubated

overnight on a rotating platform at 4°C with 1 µg of AHR (BioMol) or E2F1 (Santa Cruz), antibodies. Protein A agarose slurry (20 µl) was added and incubated for 15 min at room temperature to allow for antibody binding. Agarose was pelleted and washed twice with 1X dialysis buffer (50 mM Tris-HCl (pH 8.0), 2 mM EDTA, 0.2% sarkosyl) and 4 times with IP wash buffer (100 mM Tris-HCl (pH 9.0), 500mM LiCl, 1% NP-40, 1% deoxycholic acid).

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Immune complexes were eluted from the beads with elution buffer (50 mM NaHCO3, 1% SDS).

Reversal of crosslinks was carried out by heating the eluates at 67°C for 4-5 h. The eluates were

then digested with proteinase K at 56°C for 1.5 h. DNA was extracted with

phenol/chloroform/isoamyl alcohol, precipitated with ethanol and dissolved in TE buffer.

Samples were diluted between 5 and 10 times and subjected to PCR amplifications with the

following promoter-specific primer sets: mouse Cyp1a1 (enhancer domain -1141 to -784 bp):

forward primer, 5’-CTATCTCTTAAACCCCACCCCAA-3’; reverse primer, 5’-

CTAAGTATGGTGGAGGAAAGGGTG-3’; amplification product, 357 bp. Mouse Dhfr:

forward primer, 5’-CGGCAATCCTAGCGTGAAGGCTGGT-3’; reverse primer, 5’-

TCTCCGTTCTTGCCAATC-3’; amplification product, 113 bp. PCR products were separated

for analysis in 10% polyacrylamide gels.

Promoter Microarrays

Custom microarray slides were generated by Invitrogen, incorporating tiles of roughly

5400 bp of the promoter regions of the genes of interest. For each promoter region, tiles were

arrayed approximately every 350 bp between +400 and -5,000 from the transcription initiation

0 0 site. Tiles were 60 bp in length and were chosen to have a Tm between 68 C and 75 C. Primers were designed to amplify DNA from ChIP experiments in order to generate amplicons that

0 0 would hybridize to the tiles. Primers were 18 - 22 bases long with a Tm between 55 C and 62 C and typical amplicon length between 100 and 120 bp. ChIP samples were generated from Hepa-

1c1c7 cells treated with Me2SO or 10 nM TCDD and immunoprecipitated with IgG control or anti-AHR antibodies.

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Results

AHR represses E2F-dependent transactivation in the absence of RB expression.

Previous results from our laboratory showed that AHR appears to cooperate with RB to enhance RB-mediated repression of E2F-dependent gene expression (13). In order to determine the effect of the AHR on E2F-dependent gene expression in the absence of RB, we utilized the

C33A cell line, which lacks RB expression (see Fig. 1A and Ref. (19)), in transient transfection assays. C33A cells were transfected with the luciferase reporter plasmid p3×E2FLuc, which responds to E2F-dependent transactivation, in addition to plasmids for the expression of RB or

AHR (Fig. 1B). As expected, transfection of the p3×E2FLuc reporter into C33A cells resulted in high levels of luciferase expression that were reduced to about 50% by expression of RB.

Ectopic expression of AHR in the absence of RB expression reduced E2F-dependent reporter activity to nearly the same extent as RB alone. The co-expression of RB and AHR resulted in an additive reduction in E2F-dependent reporter expression, as we have previously reported (11;13).

Interestingly, this effect of AHR on E2F-dependent reporter activity occurs in the absence of activation by exogenous ligand. It is well-documented that AHR overexpression results in its constitutive activation and nuclear localization (20), providing one potential explanation for this observation.

Through our investigation of the interaction of AHR with RB, it was determined that mutants of the AHR which are relatively poor binders of RB in vitro could still cooperate with

RB to repress E2F-dependent gene expression (11). This effect was attributed to the existence of two independent RB-binding domains within the AHR, both of which appeared to be required for maximal interaction between the two proteins (13), but do not appear to be required for AHR to affect RB activity. In order to determine the role that these mutants might play in mediating

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repression of E2F-dependent gene expression in the absence of RB, we used RB-negative C33A cells in transient transfection experiments using the p3×E2FLuc reporter plasmid. AHR and RB proteins corresponding to all transfected plasmids were expressed at comparable levels, as determined by Western immunoblotting of whole cell extracts of transfected C33A cells (data not shown). Figure 1C illustrates the ability of several AHR truncation proteins, including ∆494-

805, lacking the transactivation domain, and ∆323-607, lacking the ligand binding domain and a portion of the transactivation domain, to act in the absence of RB expression to repress E2F- dependent transcriptional activity as well as RB alone. Only ∆323-494, which lacks an important RB-binding LXCXE motif, was unable to repress reporter gene expression. These results mirror those reported for the ability of AHR to enhance RB-mediated repression of E2F activity (11). Significantly, transactivation is not required for the AHR to repress gene expression, suggesting that the AHR may be acting directly to block E2F activity in the absence of RB. Overall, the results show that RB is not required for AHR to suppress E2F-dependent gene expression, indicating that the AHR may actually act through a mechanism distinct from its ability to bind RB.

In addition to the ability of ectopically expressed AHR to repress E2F-dependent gene expression, we wanted to examine the ability of a TCDD-activated AHR to repress E2F activity in the absence of RB. In order to examine this possibility, we used an RB-deficient Hepa-1 cell line (Fig. 1A) transfected with the E2F-responsive luciferase reporter plasmid p3×E2FLuc. In agreement with previously published results in an RB-positive Hepa-1c1c7 cell line (11), activation of the AHR by TCDD treatment results in a reduction of E2F-dependent luciferase expression by about 50%, only slightly less than produced by RB expression alone (Fig. 1D).

Ectopic RB expression combined with AHR activation by TCDD results in enhanced repression

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of E2F-dependent luciferase expression beyond that seen with either RB or TCDD alone. These data indicate that the ligand-activated AHR is able to mediate repression of E2F-dependent gene expression through a mechanism which is independent of its direct interaction with the RB protein.

Previous work from our laboratory has shown that the AHR is recruited to native E2F- dependent promoters in cells expressing the RB protein (11). In order to determine if AHR is recruited to native E2F-driven promoters in the absence of RB, we used chromatin immunoprecipitation to analyze the Dhfr promoter in RB-negative Hepa-1 cells. Cells were

treated with Me2SO vehicle or 10 nM TCDD for 1.5 h, and chromatin was prepared from nuclear lysates. Immunoprecipitations were carried out using antibodies to AHR and E2F1. Controls containing no antibody (“mock”) were included to confirm the specificity of the precipitations.

Figure 1E shows the results of PCR from immunoprecipitated chromatin, analyzing AHR binding sequences from the Cyp1a1 promoter as well as sequences containing E2F-binding sites from the Dhfr promoter. AHR antibodies immunoprecipitated AHR-responsive enhancer sequences from the Cyp1a1 promoter in TCDD-treated cells, but not in vehicle-treated cells, as expected. There was little to no change observed between Me2SO- and TCDD-treated cells in the association of E2F1 with the Dhfr promoter. However, the AHR antibody did immunoprecipitate the Dhfr promoter in cells treated with TCDD, but not with Me2SO vehicle.

These results indicate that in the absence of RB protein expression, the AHR is recruited to E2F- bound regions of E2F-dependent promoters.

AHR interacts with E2F1 in vitro.

Nuclear translocation of the AHR, through its activation by exogenous ligand, appears to be required for repression of E2F-dependent gene expression, and that the AHR is directly

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recruited to E2F-dependent gene promoters (Fig. 1 and Ref. (11)). Published data (presented in

Chapter 3) implies that one mechanism by which the AHR is recruited to E2F-dependent

promoters is through its direct interaction with the retinoblastoma protein (11;13). However, the

recruitment of AHR to such promoters in the absence of expressed RB protein must occur

through some other mechanism. There are two possibilities for this to occur. First, the AHR is

recruited through the presence of XREs in the promoters of E2F-regulated genes. At least in the

case of the mouse Dhfr promoter, this is not the case, as a scan of the first 5000 bp upstream of

the transcription start site reveals just one canonical XRE sequence at position -4778. Second is

the possibility that the AHR is recruited through some unidentified protein interaction. The most

obvious candidate is a member of the E2F protein family. Indeed, several lines of evidence

indicate that AHR and E2F do form complexes both in vitro and in vivo.

In order to examine whether AHR interacts with E2F proteins in addition to RB, we used

an E2F1-GST fusion construct in pull-down assays. We also used a series of RB-GST fusion

proteins in the same experiment, both as positive and negative controls for AHR binding. Most

RB-binding proteins require the presence of the small pocket domain of RB (30), consisting of

the A and B domains shown in Figure 2A. In addition, the C-terminal domain of the large RB

pocket is required for high affinity interaction with other RB-binding proteins, including E2F

(31). Therefore, mutants of the large pocket domain of RB (23) were used in the in vitro pull down assay to determine if AHR-RB binding requires similar domain interactions as E2F-RB.

Mutant peptides (Fig. 2A) were expressed in bacteria as GST fusion proteins, and bound to glutathione-agarose columns. Full-length in vitro synthesized [35S]AHR was passed over the columns, and AHR bound to the columns was analyzed by electrophoresis in 7.5% SDS-

polyacrylamide gels (Fig. 2B). The results show that the large pocket of RB, consisting of the A,

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B, and C domains, is required for interaction between RB and the full-length AHR protein. Both the GST-C706F protein, containing a one amino acid mutant in the B-domain of full-length RB that produces a highly unstable protein (32), and the GST-A/B/C protein, containing only the amino acids spanning the large A/B pocket, bound the labeled AHR protein. AHR bound poorly to deletion mutants of the large pocket domain, including the GST-AE∆1 peptide, encompassing the A- and B-domains of RB, and the GST-SE peptide, consisting solely of the C-domain. Most significant to our current studies, the GST-E2F1 construct was able to bind AHR at least as well as RB, demonstrating that E2F1 also establishes direct protein interactions with the AHR in vitro.

AHR interacts independently with E2F and RB proteins in cells.

To test for AHR/E2F interactions under physiological conditions, we performed co- immunoprecipitation experiments using nuclear lysates of RB-negative (see Figure 1A) Hepa-1 cells treated with Me2SO vehicle or with 10 nM TCDD for 2h. Cell lysates were incubated with

anti-E2F1 or anti-AHR specific antibodies, and bound proteins were analyzed by SDS-PAGE,

blotted, and probed with anti-E2F1 or anti-AHR antibodies (Fig. 3A). Probing with anti-E2F1

showed that anti-AHR antibodies precipitated E2F1 from nuclear extracts of TCDD treated cells

but not from cells treated with Me2SO vehicle control. Conversely, probing with anti-AHR

showed that anti-E2F1 antibodies immunoprecipitated AHR from nuclear extracts of TCDD

treated cells but not from Me2SO-treated cells. These data suggest that in addition to the in vitro interaction between AHR and E2F1 identified in the previous experiment (Fig. 2B), AHR and

E2F1 associate in vivo in a manner dependent upon ligand-mediated nuclear translocation of the

AHR. In addition, the lack of RB expression in the Hepa-1 cell line utilized for this experiment

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suggests that the AHR-E2F1 interaction is an independent phenomenon which does not require the presence of the RB protein.

In order to confirm that RB is not required for AHR and E2F1 to interact, we performed co-immunoprecipitation experiments in MEF cells from wild type and RB-null mice. Reactions were performed as outlined above for experiments in Hepa-1 cells. For both Rb+/+ and Rb-/-

MEF cells, nuclear extracts were immunoprecipitated with anti-AHR antibodies and blots were subsequently probed with anti-AHR or anti-E2F1 antibodies (Fig. 3B). Equivalent amounts of total input protein were loaded to confirm the presence of the AHR in nuclei from TCDD treated samples and little to no AHR present in Me2SO control treated samples. In addition, TCDD treatment did not affect protein levels of E2F1 in either wild type or Rb-/- cells, for both cells

show the same amount of E2F1 expression. Confirming the results obtained in Hepa-1 cells,

probing with anti-E2F1 showed that anti-AHR antibodies precipitated E2F1 from nuclear

extracts of TCDD treated cells, but little to no E2F1 was precipitated from cells treated with

Me2SO vehicle control. In addition, the amount of E2F1 protein precipitated by anti-AHR

antibodies was similar in Rb+/+ and Rb-/- cells. Thus, AHR is able to complex with E2F1 in the

absence of RB.

Previous studies in our lab have shown that AHR and RB form complexes both in vitro

and in vivo (11;13). Given that we have now identified an interaction between the AHR and

E2F1 as well, we wanted to determine whether the interaction between the AHR and RB is

dependent on the presence of E2F1. In order to test this possibility, we utilized Saos-2 cells,

which are both RB- and AHR-negative, to express AHR, RB, and E2F1 using recombinant

adenoviruses. Immunoprecipitations were carried out using anti-AHR antibodies, and blots were

probed with anti-AHR, anti-RB, and anti-E2F1 antibodies (Fig. 3C). For cells in which AHR

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and RB are co-expressed, anti-AHR antibodies were able to precipitate RB from Saos-2 extracts.

In addition, anti-AHR antibodies also precipitated E2F1 from cell extracts where AHR and E2F1 were co-expressed. Finally, when all three proteins were co-expressed in the same cells, anti-

AHR antibodies precipitated both RB and E2F1. These data suggest that AHR can interact equally well with either RB or E2F1 independently of the presence of the other. In addition, given that anti-AHR immunoprecipitates RB equally well in the presence of E2F1 and vice versa, AHR appears to have no specific binding preference for either RB or E2F1.

RB is known to interact with multiple E2F family members. Of the 8 identified E2F family members, RB exhibits a preference for E2F1-4, while the RB family members p107 and p130 show a preference for E2F4 and E2F5 (33). We asked whether AHR has the same ability as RB to interact with multiple E2F family members, or if it has a specific preference for E2F1.

We utilized recombinant adenoviruses, as previously outlined, to express AHR, E2F1, E2F3, and

E2F4 in Saos-2 cells. Immunoprecipitations were carried out using anti-AHR antibodies, and blots were probed with anti-E2F1, anti-E2F3, and anti-E2F4 antibodies (Fig. 3D). As previously shown, anti-AHR antibodies precipitated AHR and E2F1 from Saos-2 extracts (see Fig. 3C). In addition, anti-AHR antibodies also precipitated E2F3 and E2F4 from cell extracts in which these

E2F members and AHR were co-expressed, thereby demonstrating that AHR can interact equally well with several other E2F family members.

AHR displaces RB from RB-E2F-DP1 complexes.

The previous co-immunoprecipitation experiments show that AHR interacts with both

RB and E2F1 independently (Fig. 3C), with no apparent preference for one protein over the other. It could be, however, that the AHR interacts with both RB and E2F proteins simultaneously, or that it forms independent complexes containing AHR-RB and AHR-E2F. To

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distinguish between these two possibilities, electrophoretic mobility shift assays (EMSA) were performed to determine whether AHR binds to RB when RB is complexed with E2F-DP1 heterodimers at E2F DNA-binding sites. Nuclear extracts of CV-1 cells, which express high levels of RB but lack expression of AHR, were used for all mobility shift assays. The specificity of the labeled E2F-binding probe was confirmed by competition assays with either excess unlabeled wild type or mutant oligonucleotides (data not shown). CV-1 nuclear extracts were incubated with labeled probe and with antibodies against RB, E2F1, DP1, AHR, and as a negative control, NF-κB, to identify the bands containing the RB, E2F, and DP1 protein-DNA complexes (arrows in Fig. 4A). Truncated AHR peptides were subsequently analyzed for their ability to shift or block the formation of RB-E2F complexes. Two truncated forms of the AHR were used, encoding the amino-terminal 375 amino acids (∆376-805) and the carboxy-terminal

379 amino acids (∆1-425). The ∆376-805 peptide contains the LXCXE sequence, while the ∆1-

425 peptide encompasses the entire glutamine-rich transactivation domain. These peptides were shown to form complexes with RB in vitro (13). Both protein fragments were tagged with a His6 peptide and purified from bacteria on nickel-agarose columns. Both truncated forms of the

AHR protein displaced RB from its DNA-binding complex (Fig. 4B), suggesting that AHR sequesters RB away from E2F at its DNA-binding site. In addition, a novel band of slower mobility than E2F-DP1 complexes appeared as a result of AHR inclusion, suggesting a direct interaction between the truncated AHR peptides and E2F (or possibly DP1), in agreement with the observation of AHR-E2F interactions in pull-down assays and co-immunoprecipitation experiments. Incubation of extracts with both the truncated peptides and antibodies specific for those peptides (anti-bHLH for ∆376-805 and anti-YGOR for ∆1-425) blocked formation of the

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novel complexes, indicating that the AHR peptides were indeed part of those complexes (Fig.

4C).

The AHR is required for constitutive expression of E2F-regulated genes.

In order to analyze the impact of the AHR on E2F-regulated genes in vivo, we used MEF

cells from wild type mice as well as mice lacking AHR and E2F1. In total, 12 cell cycle-related

and E2F-regulated genes were identified that had both E2F binding sites as well as canonical

AHR binding sites within 5000 bp of the transcription start site (Table 1). mRNA was isolated

from MEF+/+, AHR-/-, and E2F1-/- cells treated under standard culture conditions with Me2SO vehicle or with 5 nM TCDD for 8 hours, and cDNA was synthesized and amplified using the gene-specific primer sets listed in Table 2. Figure 5A shows the expression levels for Cyp1a1, relative to Me2SO-treated MEF+/+ cells. As expected, MEF+/+ and E2F1-/- cells show a robust increase in Cyp1a1 mRNA expression levels when cells are treated with TCDD for 8 hours. In contrast, Cyp1a1 expression does not change in treated AHR-/- cells. For most of the 12 E2F- regulated genes analyzed, expression levels were lower in E2F1-/- cells relative to MEF+/+ cells and were generally unaffected by TCDD (Figure 5B). Only two genes were increased in E2F1-/- cells, Apaf1 and Hmgn1, and these genes were also increased in AHR-/- cells. Interestingly, all of the genes which showed decreased expression in E2F1-/- cells were also decreased in AHR-/- cells, in most cases to a greater extent than in the E2F-/- cells. Finally, roughly half of the genes affected by constitutive activity of the AHR actually decreased in expression when the AHR was activated by TCDD treatment, consistent with data obtained using E2F-driven reporter genes

(Fig. 1), and this TCDD-mediated decrease is absent in cells lacking E2F1. Overall, these data indicate that the constitutive activity of the AHR is necessary for maintaining normal expression

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levels of a multitude of genes regulated by E2F, and that activation of the AHR by exogenous ligands perturbs this normal regulation.

The constitutive and TCDD-bound AHR binds differentially to E2F-regulated promoters.

As stated above, the E2F-regulated genes examined in Figure 5 were chosen for the presence of both E2F and AHR canonical binding sites. Hence, these genes have the potential to be modulated by the AHR either through direct binding of the AHR to the DNA of these promoters, or through binding of the AHR to E2F. In order to differentiate between these possibilities, we analyzed four of these genes (Apaf1, Cdt1, Dnmt1 and Mcm3) using chromatin immunoprecipitation studies combined with custom tiling microarrays. Microarrays were designed so as to incorporate 60 bp arrayed every 350 bp, spanning +400 to -5000 from the transcription start site of each gene. Hepa-1c1c7 cells treated with Me2SO vehicle or 10 nM

TCDD were used in ChIP experiments, and samples were immunoprecipitated with either IgG

control or anti-AHR antibodies. In order generate ample DNA for use in microarray studies, the

ChIP samples were amplified with primers to produce amplicons that could hybridize to the

tiling arrays. Both Me2SO and TCDD samples precipitated with anti-AHR antibody were hybridized to IgG-immunoprecipitated samples. Results for each specific tile on the arrays for the four genes of interest are presented in Figure 6. Presented on the abscissa are the promoter positions from -5000 bp from the transcription start site of each gene to +1000 bp, along with the associated positions of the AHR and E2F binding sites, and on the ordinate are the DMSO/IgG or TCDD/IgG ratio results from the microarray. The results show that there are one or more positions for each gene in which the DMSO/IgG ratio is greater than one, possibly explaining the constitutive expression levels of these genes when the AHR is expressed. In a few cases, it is also higher than the TCDD/IgG ratio, indicating a possible shift in the location of the AHR in its

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constitutive and TCDD-bound forms. In addition, TCDD treatment generally leads to higher binding ratios over IgG controls for all of the genes analyzed, with the higher ratios often correlating to positions of AHR binding sites or positions in which AHR and E2F binding sites are in close proximity to one another. Overall, the data show that there are multiple locations of

AHR binding in these E2F-regulated genes, and that the effect of the AHR on these genes is likely dependent on both AHR-DNA binding (particularly regarding the activity of the unliganded AHR) as well as the interaction of the AHR and E2F proteins. Notably, TCDD/IgG ratios are generally greatest in regions correlating with both AHR and E2F binding sites. Thus, the impact of the AHR-E2F interaction may be particularly important when the AHR is activated by ligand, as in its TCDD-bound state, the vast majority of the receptor is located in the nucleus, likely resulting in the observed increase in AHR-precipitated promoter sequences, as well as the resulting impact on gene expression.

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Discussion

Exposure to polycyclic aromatic hydrocarbons (PAHs) results in a plethora of toxic and carcinogenic responses in animals and in humans. TCDD, commonly studied as a model for outcomes of exposure to other polychlorinated hydrocarbons, is a potent tumor promoter in several rodent models, and has been labeled a human carcinogen (34;35). The specific mechanism(s) by which TCDD exhibits its promotional effects are speculative, but is believed to occur at the most fundamental level through AHR-mediated perturbations in the expression of literally hundreds of individual, cell type- and tissue-specific genes. The data presented here and in several other manuscripts indicates that one of these mechanisms is through the interaction of the AHR with the RB-E2F family of transcriptional regulators. E2F proteins are major regulators of cell cycle progression, and one of the major effects of dioxin exposure is to perturb cell cycle progression through enhancing or inhibiting cell division, differentiation, or death. It is interesting to note that as many as 7% of known genes, involved in mechanisms regulating differentiation, development, proliferation, and apoptosis, may be regulated by E2F (36). The

AHR appears to regulate expression of at least a subset of these genes through two mechanisms: direct AHR-DNA binding and AHR-E2F interactions.

Previous work from multiple laboratories found that the AHR and RB proteins form complexes both in vitro and in vivo (10;13). In the present studies, we show that the AHR can

also directly interact with proteins of the E2F family independently of RB. Pull-down assays

confirm that E2F1 and multiple RB mutant proteins interact equally well with an in vitro-

synthesized AHR (Fig. 2). Co-immunoprecipitation experiments show that AHR interacts in

vivo with multiple E2F family members, both in RB-positive and RB-negative cell lines (Fig. 3).

Despite the fact that AHR shows no preference for either RB or E2F in co-IP studies, gel shift

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assays indicate that the AHR may in fact displace RB in binding directly to E2F (Fig. 4). It is also possible, however, that both AHR-E2F-DP as well as AHR-RB-E2F-DP complexes exist, and that AHR binds directly to E2F when RB is phosphorylated and inactive, as is the case at the

G1/S transition. Certainly, there is precedent for the promiscuous nature of AHR in binding a wide variety of transcription factors, as has been demonstrated for several nuclear receptors (ER,

AR, TR), the orphan receptor COUP-TF1, NF-κB, and multiple coactivators and corepressor, including p300/CBP, SRC-1, NCoA-2, pCIP, ERAP140, and SMRT (37). Several of these interactions are clearly demonstrated to be cell-type specific, and only impinge upon AHR signaling under specific cellular contexts (38). It is unclear at this point whether AHR interacts with RB and/or E2F only under certain circumstances, or if these are ubiquitous protein-protein interactions which occur broadly and in all cell types.

In cells which express RB, the TCDD-activated AHR is recruited to E2F-regulated promoters, and coactivator recruitment to these promoters is prevented. The result is a repression of expression for these genes (11). Our current studies show that the AHR can repress an E2F-dependent reporter in the absence of RB, and can form complexes with E2F proteins.

Thus, it appears that the ligand-activated AHR can act by two mechanisms to repress gene expression, either through binding to RB and preventing its phosphorylation and release from

E2F, or through binding to E2F, preventing its ability to transactivate gene expression in a manner analogous to that of RB. In some contexts, AHR may actually mimic the activities of

RB, suppressing expression of genes required for cell cycle progression, and ultimately preventing cell division. Interestingly, RB is a potent tumor suppressor (39), and AHR has been shown to inhibit tumorigenesis under some conditions (40;41). However, the AHR is also known to mediate the tumor promoting activity of ligands such as TCDD (42). Many studies

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have demonstrated that interactions of the AHR with specific proteins can alter gene expression in a variety of ways, and the AHR is known to up- and down-regulate literally hundreds of different genes (43;44).

We have shown that the TCDD-activated AHR inhibits the transcription of a number of

important cell cycle regulatory genes in cells expressing the RB protein. However, it appears

that this activity of the AHR is not universal for all E2F-regulated genes in all RB-expressing

cells. For example, only about half of the genes examined in this study decrease in expression

following TCDD treatment in wild-type MEF cells. Surprisingly, the same results indicate that

the AHR is required for maintaining constitutive mRNA levels of a subset of E2F-regulated

genes in these cells, and that the AHR is at least as important in controlling their expression

levels as E2F itself (Fig. 5). While most research into transcriptional regulation by the AHR is

focused on the activities of the ligand-activated AHR, there is evidence for AHR-dependent gene

regulation in the absence of ligand-mediated nuclear translocation of the receptor (45-48). We

also show that the promoters of some of the genes analyzed here are constitutively bound by the

AHR (Fig. 6). Treatment of cells with TCDD results in the translocation of the bulk of

cytoplasmic AHR to the nucleus. In cells treated with TCDD, the proportion of AHR-bound

promoter sequences over IgG control increased for all 4 promoters examined. Interestingly,

mRNA expression levels were decreased for Cdt1, Dnmt1, and Mcm3 in TCDD-treated cells

(Fig. 5B), as was observed in assays of E2F-dependent reporter genes (Fig. 1). The same

decrease is not observed in E2F1-/- cells, indicating that this effect is dependent on the presence

of both AHR and E2F. The interaction of the AHR and E2F proteins may provide a mechanistic

explanation for this result. However, the relative importance of AHR-RB interaction versus

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AHR-E2F interaction in the recruitment of AHR to the promoters of many cell cycle-regulatory genes, and the resulting repression of E2F-dependent gene expression, must be clarified.

It is a common observation in studies of AHR-mediated functions that the basal activities of the receptor are quite distinct from its activities upon activation by exogenous ligand. One hypothesis is that the toxicities mediated by TCDD can be explained as deregulated physiological functions of the AHR (49), and it is often the case that the loss of AHR expression from a particular cell type produces the same physiological response as the aberrant and persistent activation of the receptor by dioxin. This seems to be true for the observed effects of

TCDD on cell cycle regulation. It is well known that the AHR is required for normal cell cycle progression (50;51), and cells lacking the AHR cycle more slowly than cells which express the

AHR (52-54). One potential explanation for this observation is the requirement of the AHR for basal expression of important cell cycle regulators, as our current results indicate. This basal regulation appears in some cases to be disrupted by the total nuclear translocation of the AHR by

TCDD activation. We believe this disruption occurs at least in part through the ability of the

AHR to interact and inhibit the activities of the E2F protein. E2F itself plays contrasting roles in different cellular contexts. In addition to being both an activator and repressor of transcription, it serves functions in cell proliferation and differentiation, apoptosis regulation, and acts to both suppress and promote tumorigenesis (1). The AHR functions in many of these same capacities, and it may be that TCDD exerts a portion of its toxic effects by targeting the AHR to E2F- regulated genes and disrupting their normal patterns of expression. Interestingly, it has recently been observed that the essential oil component eugenol, a potential ligand of the AHR, inhibits cell growth in an AHR-dependent manner in HaCat cells (55), and in an E2F1-dependent manner in melanoma cells (56).

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There exist multiple redundant pathways that couple proliferation with apoptosis in order to protect the cell when normal proliferation controls are lost (57). One striking example of this coupling is the activities of the E2F family. In addition to controlling gene expression for cell cycle progression, several E2F family members, in particular E2F1, are able to activate apoptosis. E2F1 is characterized as an oncogene based on evidence from cell transformation assays, and from transgenic mouse studies in which E2F1 overexpression increases rates of tumor formation (2;58). In contrast, E2F1 has been labeled a tumor suppressor based on the finding that E2F1-deficient mice develop a range of tumor types (59;60). Evidence suggests that the ability of E2F1 to act as a tumor suppressor lies in its ability to initiate apoptosis in cells that lose normal cell cycle control. Several mechanisms mediating E2F1-dependent apoptosis have been identified, including the inhibition of anti-apoptotic signaling pathways, transcriptional activation of specific genes that positively regulate apoptosis, and inhibition of genes that are negative regulators of apoptosis (61). Tumor promoters such as TCDD are generally believed to act by affecting the rate of division, terminal differentiation, or death of tumor precursor cells.

One widely accepted mechanism of tumor promotion/progression is inhibition of apoptosis (62).

The capacity of TCDD to act as a tumor promoter, particularly in rodent liver, has been attributed to its ability to inhibit the apoptotic elimination of initiated cells bearing genotoxic lesions (63). However, the precise mechanism(s) of this effect remains elusive, and seems to differ with the organism, tissue, or cell type examined. It is possible that the AHR may play a role in the inhibition of apoptosis through repression of genes required for E2F1-dependent apoptosis, as it does for genes controlling cell proliferation. For example, if normal proliferation controls are lost through the aberrant inactivation of RB, cells may be eliminated through the induction of E2F-dependent apoptotic activities. Activation of the AHR by TCDD could inhibit

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this activity by preventing the induction of pro-apoptotic gene targets of E2F, thus preventing the elimination of cells which have lost normal cell cycle control. Preliminary data relative to this hypothesis are presented in the Appendix.

It is clear from a number of studies that the AHR modulates the expression of a plethora of genes from a wide variety of molecular pathways, and that the sub-set of modulated genes may be completely different depending on the cell type examined (43;64-67). The same is true for the constitutive activity of the AHR, as assayed in cells lacking receptor expression (68;69).

For most of these genes, the mechanism by which the AHR exerts its effects remains undetermined, particularly for genes in which the AHR, a known transcriptional activator, downregulates their expression. In the current studies, we have identified yet another set of genes whose expression appears to be controlled by the AHR, as well as a potential mechanism by which this occurs. The significance of the AHR-E2F interaction in different cell types remains to be determined.

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References

1. Dimova DK and Dyson NJ. 2005. The E2F transcriptional network: old acquaintances with new faces. Oncogene 24: 2810-2826

2. Dyson N. 1998. The regulation of E2F by pRB-family proteins. Genes Dev. 12: 2245- 2262

3. Nevins JR. 1998. Toward an understanding of the functional complexity of the E2F and retinoblastoma families. Cell Growth Differ. 9: 585-593

4. Frolov MV and Dyson NJ. 2004. Molecular mechanisms of E2F-dependent activation and pRB-mediated repression. J.Cell Sci. 117: 2173-2181

5. Trimarchi JM and Lees JA. 2002. Sibling rivalry in the E2F family. Nat.Rev.Mol.Cell Biol. 3: 11-20

6. Harbour JW and Dean DC. 2000. The Rb/E2F pathway: expanding roles and emerging paradigms. Genes Dev. 14: 2393-2409

7. Ferreira R, Naguibneva I, Pritchard LL, Ait-Si-Ali S, and Harel-Bellan A. 2001. The Rb/chromatin connection and epigenetic control: opinion. Oncogene 20: 3128-3133

8. Stevaux O and Dyson NJ. 2002. A revised picture of the E2F transcriptional network and RB function. Curr.Opin.Cell Biol. 14: 684-691

9. DeGregori J. 2002. The genetics of the E2F family of transcription factors: shared functions and unique roles. Biochim.Biophys.Acta 1602: 131-150

10. Ge N-L and Elferink CJ. 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblatoma protein. J.Biol.Chem. 273: 22708-22713

11. Marlowe JL, Knudsen ES, Schwemberger S, and Puga A. 2004. The aryl hydrocarbon receptor displaces p300 from E2F-dependent promoters and represses S phase-specific gene expression. J.Biol.Chem. 279: 29013-29022

12. Huang G and Elferink CJ. 2005. Multiple mechanisms are involved in Ah receptor- mediated cell cycle arrest. Mol.Pharmacol. 67: 88-96

13. Puga A, Barnes SJ, Dalton TP, Chang C, Knudsen ES, and Maier MA. 2000. Aromatic hydrocarbon receptor interaction with the retinoblastoma protein potentiates repression of E2F-dependent transcription and cell cycle arrest. J.Biol.Chem. 275: 2943-2950

14. Strobeck MW, Fribourg AF, Puga A, and Knudsen ES. 2000. Restoration of retinoblastoma mediated signaling to Cdk2 results in cell cycle arrest. Oncogene 19: 1857-1867

135

15. Bernard HP, Darlington GJ, and Ruddle FH. 1973. Expression of liver phenotypes in cultured mouse hepatoma cells: synthesis and secretion of serum albumin. Dev.Biol. 35: 83-96

16. Brooks SC, Locke ER, and Soule HD. 1973. Estrogen receptor in a human cell line (MCF-7) from breast carcinoma. J.Biol.Chem. 248: 6251-6253

17. Jensen FC, Girardi AJ, Gilden RV, and Koprowski H. 1964. Infection of human and simian tissue cultures with Rous sarcoma virus. Proc.Natl.Acad.Sci.USA 52: 53-59

18. Fogh J, Fogh JM, and Orfeo T. 1977. One hundred and twenty-seven cultured human tumor cell lines producing tumors in nude mice. J.Natl.Cancer Inst. 59: 221-226

19. Scheffner M, Munger K, Byrne JC, and Howley PM. 1991. The state of the p53 and retinoblastoma genes in human cervical carcinoma cell lines. Proc.Natl.Acad.Sci.U.S.A 88: 5523-5527

20. Chang C-Y and Puga A. 1998. Constitutive activation of the aromatic hydrocarbon receptor. Mol.Cell.Biol. 18: 525-535

21. Fukunaga BN, Probst MR, Reisz-Porszasz S, and Hankinson O. 1995. Identification of functional domains of the aryl hydrocarbon receptor. J.Biol.Chem. 270: 29270-29278

22. Whitaker LL, Su H, Baskaran R, Knudsen ES, and Wang JY. 1998. Growth suppression by an E2F-binding-defective retinoblastoma protein (RB): contribution from the RB C pocket. Mol.Cell Biol. 18: 4032-4042

23. Knudsen ES and Wang JYJ. 1996. Differential regulation of retinoblastoma protein function by specific cdk phosphorylation sites. J.Biol.Chem. 271: 8313-8320

24. Dignam JD, Lebovitz RM, and Roeder RG. 1983. Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11: 1475-1489

25. Carrier F, Owens RA, Nebert DW, and Puga A. 1992. Dioxin-dependent activation of murine Cyp1a-1 transcription requires protein kinase C-dependent phosphorylation. Mol.Cell.Biol. 12: 1856-1863

26. Knudsen ES and Wang JY. 1997. Dual mechanisms for the inhibition of E2F binding to RB by cyclin- dependent kinase-mediated RB phosphorylation. Mol.Cell Biol. 17: 5771- 5783

27. Nevins JR, DeGregori J, Jakoi L, and Leone G. 1997. Functional analysis of E2F transcription factor. Methods Enzymol. 283: 205-219

28. Weinmann AS and Farnham PJ. 2002. Identification of unknown target genes of human transcription factors using chromatin immunoprecipitation. Methods 26: 37-47

136

29. Wells J and Farnham PJ. 2002. Characterizing transcription factor binding sites using formaldehyde crosslinking and immunoprecipitation. Methods 26: 48-56

30. Morris EJ and Dyson NJ. 2001. Retinoblastoma protein partners. Adv.Cancer Res. 82: 1- 54

31. Hiebert SW. 1993. Regions of the retinoblastoma gene product required for its interaction with the E2F transcription factor are necessary for E2 promoter repression and pRb- mediated growth suppression. Mol.Cell Biol. 13: 3384-3391

32. Lee JO, Russo AA, and Pavletich NP. 1998. Structure of the retinoblastoma tumour- suppressor pocket domain bound to a peptide from HPV E7. Nature 391: 859-865

33. Du W and Pogoriler J. 2006. Retinoblastoma family genes. Oncogene 25: 5190-5200

34. Bertazzi PA, Consonni D, Bachetti S, Rubagotti M, Baccarelli A et al. 2001. Health effects of dioxin exposure: a 20-year mortality study. Am.J.Epidemiol. 153: 1031-1044

35. . 1997. IARC Working Group on the Evaluation of Carcinogenic Risks to Humans: Polychlorinated Dibenzo-Para-Dioxins and Polychlorinated Dibenzofurans. Lyon, France, 4-11 February 1997. IARC Monogr Eval.Carcinog.Risks Hum. 69: 1-631

36. Muller H, Bracken AP, Vernell R, Moroni MC, Christians F et al. 2001. E2Fs regulate the expression of genes involved in differentiation, development, proliferation, and apoptosis. Genes Dev. 15: 267-285

37. Carlson DB and Perdew GH. 2002. A dynamic role for the Ah receptor in cell signaling? Insights from a diverse group of Ah receptor interacting proteins. J.Biochem.Mol.Toxicol. 16: 317-325

38. Beischlag TV and Perdew GH. 2005. ER alpha-AHR-ARNT protein-protein interactions mediate estradiol-dependent transrepression of dioxin-inducible gene transcription. J.Biol.Chem. 280: 21607-21611

39. Knudsen ES and Knudsen KE. 2006. Retinoblastoma tumor suppressor: where cancer meets the cell cycle. Exp.Biol.Med.(Maywood.) 231: 1271-1281

40. Kohle C and Bock KW. 2006. Activation of coupled Ah receptor and Nrf2 gene batteries by dietary phytochemicals in relation to chemoprevention. Biochem.Pharmacol. 72: 795- 805

41. Bradshaw TD, Trapani V, Vasselin DA, and Westwell AD. 2002. The aryl hydrocarbon receptor in anticancer drug discovery: friend or foe? Curr.Pharm.Des 8: 2475-2490

42. Bock KW and Kohle C. 2005. Ah receptor- and TCDD-mediated liver tumor promotion: clonal selection and expansion of cells evading growth arrest and apoptosis. Biochem.Pharmacol. 69: 1403-1408

137

43. Puga A, Maier A, and Medvedovic M. 2000. The transcriptional signature of dioxin in human hepatoma HepG2 cells. Biochem.Pharmacol. 60: 1129-1142

44. Ovando BJ, Vezina CM, McGarrigle B, and Olson JR. 2006. Hepatic Gene Down- Regulation Following Acute and Subchronic Exposure to 2,3,7,8-Tetrachlorodibenzo-p- dioxin. Toxicol.Sci.

45. Allan LL and Sherr DH. 2005. Constitutive activation and environmental chemical induction of the aryl hydrocarbon receptor/transcription factor in activated human B lymphocytes. Mol.Pharmacol. 67: 1740-1750

46. Pocar P, Augustin R, and Fischer B. 2004. Constitutive expression of CYP1A1 in bovine cumulus oocyte-complexes in vitro: mechanisms and biological implications. Endocrinology 145: 1594-1601

47. Wang F, Wang W, and Safe S. 1999. Regulation of constitutive gene expression through interactions of Sp1 protein with the nuclear aryl hydrocarbon receptor complex. Biochemistry 38: 11490-11500

48. Boesch JS, Miskimins R, Miskimins WK, and Lindahl R. 1999. The same xenobiotic response element is required for constitutive and inducible expression of the mammalian aldehyde dehydrogenase-3 gene. Arch.Biochem.Biophys. 361: 223-230

49. Bock KW and Kohle C. 2006. Ah receptor: dioxin-mediated toxic responses as hints to deregulated physiologic functions. Biochem.Pharmacol. 72: 393-404

50. Ma Q and Whitlock JPJ. 1996. The aromatic hydrocarbon receptor modulates the Hepa 1c1c7 cell cycle and differentiated state independently of dioxin. Mol.Cell Biol. 16: 2144- 2150

51. Weiss C, Kolluri SK, Kiefer F, and Gottlicher M. 1996. Complementation of Ah receptor deficiency in hepatoma cells: negative feedback regulation and cell cycle control by the Ah receptor. Exp.Cell Res. 226: 154-163

52. Elizondo G, Fernandez-Salguero P, Sheikh MS, Kim GY, Fornace AJ et al. 2000. Altered cell cycle control at the G(2)/M phases in aryl hydrocarbon receptor-null embryo fibroblast. Mol.Pharmacol. 57: 1056-1063

53. Tohkin M, Fukuhara M, Elizondo G, Tomita S, and Gonzalez FJ. 2000. Aryl hydrocarbon receptor is required for p300-mediated induction of DNA synthesis by adenovirus E1A. Mol.Pharmacol. 58: 845-851

54. Abdelrahim M, Smith R, III, and Safe S. 2003. Aryl hydrocarbon receptor gene silencing with small inhibitory RNA differentially modulates Ah-responsiveness in MCF-7 and HepG2 cancer cells. Mol.Pharmacol. 63: 1373-1381

138

55. Kalmes M, Neumeyer A, Rio P, Hanenberg H, Fritsche E, and Blomeke B. 2006. Impact of the arylhydrocarbon receptor on eugenol- and isoeugenol-induced cell cycle arrest in human immortalized keratinocytes (HaCaT). Biol.Chem. 387: 1201-1207

56. Ghosh R, Nadiminty N, Fitzpatrick JE, Alworth WL, Slaga TJ, and Kumar AP. 2005. Eugenol causes melanoma growth suppression through inhibition of E2F1 transcriptional activity. J.Biol.Chem. 280: 5812-5819

57. Tsantoulis PK and Gorgoulis VG. 2005. Involvement of E2F transcription factor family in cancer. Eur.J.Cancer 41: 2403-2414

58. Johnson DG. 2000. The paradox of E2F1: oncogene and tumor suppressor gene. Mol.Carcinog. 27: 151-157

59. Yamasaki L, Jacks T, Bronson R, Goillot E, Harlow E, and Dyson NJ. 1996. Tumor induction and tissue atrophy in mice lacking E2F-1. Cell 85: 537-548

60. Field SJ, Tsai FY, Kuo F, Zubiaga AM, Kaelin WG, Jr. et al. 1996. E2F-1 functions in mice to promote apoptosis and suppress proliferation. Cell 85: 549-561

61. Phillips AC and Vousden KH. 2001. E2F-1 induced apoptosis. Apoptosis. 6: 173-182

62. Roberts RA, Nebert DW, Hickman JA, Richburg JH, and Goldsworthy TL. 1997. Perturbation of the mitosis/apoptosis balance: a fundamental mechanism in toxicology. Fundam.Appl.Toxicol. 38: 107-115

63. Schwarz M, Buchmann A, Stinchcombe S, Kalkuhl A, and Bock K. 2000. Ah receptor ligands and tumor promotion: survival of neoplastic cells. Toxicol.Lett. 112-113: 69-77

64. Fletcher N, Wahlstrom D, Lundberg R, Nilsson CB, Nilsson KC et al. 2005. 2,3,7,8- Tetrachlorodibenzo-p-dioxin (TCDD) alters the mRNA expression of critical genes associated with cholesterol metabolism, bile acid biosynthesis, and bile transport in rat liver: a microarray study. Toxicol.Appl.Pharmacol. 207: 1-24

65. Hanlon PR, Zheng W, Ko AY, and Jefcoate CR. 2005. Identification of novel TCDD- regulated genes by microarray analysis. Toxicol.Appl.Pharmacol. 202: 215-228

66. Thackaberry EA, Jiang Z, Johnson CD, Ramos KS, and Walker MK. 2005. Toxicogenomic profile of 2,3,7,8-tetrachlorodibenzo-p-dioxin in the murine fetal heart: modulation of cell cycle and extracellular matrix genes. Toxicol.Sci. 88: 231-241

67. Fisher MT, Nagarkatti M, and Nagarkatti PS. 2004. Combined screening of thymocytes using apoptosis-specific cDNA array and promoter analysis yields novel gene targets mediating TCDD-induced toxicity. Toxicol.Sci. 78: 116-124

68. Yoon CY, Park M, Kim BH, Park JY, Park MS et al. 2006. Gene expression profile by 2,3,7,8-tetrachlorodibenzo-p-dioxin in the liver of wild-type (AhR+/+) and aryl hydrocarbon receptor-deficient (AhR-/-) mice. J.Vet.Med.Sci. 68: 663-668

139

69. Fong CJ, Burgoon LD, and Zacharewski TR. 2005. Comparative microarray analysis of basal gene expression in mouse Hepa-1c1c7 wild-type and mutant cell lines. Toxicol.Sci. 86: 342-353

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Table 1 Cell cycle-regulated genes and their canonical E2F and AHR binding sites. Sequences comprising the first 5000 bp upstream of the transcription start site of each gene were scanned for the invariant AHR binding nucelotides GCGTG or CACGC. The TFSEARCH transcription factor binding site program was used to identify E2F binding sites. Binding sites are listed by position from the first base of the promoter (-1) and actual sequence of the site. AHR binding site E2F Binding Site Gene Name Accession Number Function Position Sequence Position Sequence apoptosome; tumor -3574 GCGTG -2965 TTCGCCGC suppressor; -640 CACGC -1620 GCTGGAAA Apaf1 NM_009684 nervous/immune -346/-124 CACGC -509 TTTCCTGC system development -64 GCGTG -273 GCGGGAGA -3764 CACGC -2998 TTTCCCGC regulation of cell -800 CACGC -2378 GAGCCAAA proliferation; -372 CACGC -2274/-2136 TTTCGCCC Cdca7 NM_025866 neoplastic -340 GCGTG -1992 GCTCCAAA transformation -321 CACGC -280 TTTCGCGC +157 GCGTG -70 GCGGGAAA -3460/-2126 GCGTG -326 TTGGCGC -411/-279 CACGC/GCGTG -254 CGCGCGAA positive regulation of Cdca7-like NM_146040 -188/-96 CACGC/GCGTG -158 TTTGCGCG cell proliferation +5/+37 GCGTG +85 CACGC Cdk subunit; -366 CACGC -3136 GCGCGAAA Cks1 NM_016904 degradation of p27 13 GCGTG -102 ACGCCAAA methylation/gene -3554 CACGC -3065 TTTGGGGC Dnmt1 NM_010066 expression; integrity of -2960 GCGTG -89/+16 TTGCGCGC chromosomes -1513 GCGTG +24 GCGCGAAA histone methylation; -567 CACGC -549 GCGCCAA Ezh2 NM_007971 development; cell -512 CACGC -43 GGGCCAAA growth control -172 CACGC -7 TTTGGCGC DNA repair; -398/-296 CACGC -638 GCGCGAA Hmgn1 NM_008251 gene expression +93 CACGC -4546 CACGC -4341 GCTCCAAA cell cycle; DNA -4268 CACGC -2307/-1955 TTTGGCTC Mcm3 NM_008563 replication and -3638/-1889 GTGCG -77 GCGGGAAA repair; transcription -42 CACGC -60 TTTGGCGC Histone binding; -444 CACGC -3515 TTTAGCGC Nasp NM_016777 chromatin assemby -58 CACGC -130 G CGCGATA lipid -2246 GCGTG -2285 CGCGGAA Pltp NM_011125 binding/transport -392 GCGTG -709 TTTCGCCC HDL remodeling; -298 GCGTG -233 TTTCTCGC -4565 CACGC -1062 GGGCCAAA DNA replication; -3100 GCGTG -539 TTTGGAGC chromosome -2793 GCGTG -312 GCGCCAA Ris2 (Cdt1) NM_026014 stability; pre- -816/-645 GCGTG -268 CCGCGAAA replication complex -467 GCGTG 8 GCGCGGAA factor -258/-231 CACGC 32 TTTGGCGC -41/+71 GCGTG -4081/-4023 GTGCG -4018 GGGCCAAA E2F-binding partner; -3216/-2852 CACGC/GCGTG -1326 TTGGCGC regulation of -2780/-1547 GTGCG/CACGC -1297 GAGCCAAA -1462/-1319 GTGCG -966 GCGCGAAA Tfdp1 NM_009361 transcription; -1311/-1075 CACGC apoptosis; cell cycle; -1070/-893 CACGC development -616/-610 GTGCG -417/-350 CACGC/GCGTG

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Table 2 Gene-specific primer sets for real-time PCR analysis of relative mRNA expression levels

Gene Name Accession Number Forward Primer (5'-3') Reverse Primer (5'-3') Product Size

β-actin M12481 catccgtaaagacctctatgcc acgcagctcagtaacagtcc 287 Cyp1a1 NM_009992 gtgtctggttactttgacaagtgg aacatggacatgcaaggaca 199 Apaf1 NM_009684 tcagcaaacgagaggaaaag catagggggagaagtcacag 235 Cdca7 NM_025866 aagaggaggaggaagaggag ggtcggattatatgcggaag 142 Cdca7-like NM_146040 tgcttccgttccaaatacttc cctcctcttcatcatcttcctc 202 Cks1 NM_016904 caaatacgacgacgaggag tcagaaagatggcagggag 277 Dnmt1 NM_010066 ctgaccgcttctacttcctc tccctttccccttccctttc 111 Ezh2 NM_007971 aagacaccacctaaacgcc accactccactccacattc 273 Hmgn1 NM_008251 agataaaagggaagaggggag ttaaaaaatgggatgaggtggg 289 Mcm3 NM_008563 gcaggaagaatgaaaagaggg aggaagcaggaagtgagag 199 Nasp NM_016777 atgctcctgctccttctacc cccaacactccattctccattc 252 Pltp NM_011125 agaccatcaccatcccagac gcccccatcatataagaaccag 203 Ris2 (Cdt1) NM_026014 tccttgcctgtttctttcatc gccatccaacataccctactc 184 Tfdp1 NM_009361 ttagaggtggagaggcagag ttgacaatgatgaagggcaag 182

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Chapter IV Figure Legends

Fig. 1. AHR represses E2F-dependent transcription in multiple cell lines. A. Total protein equivalents from whole cell extracts of C33A, Hepa-1, MCF-7, CV-1, and Saos-2 cells were analyzed by Western blot for the expression of RB protein. B. C33A cells were grown in 24- well plates to a density of ~50% and transfected with transfected with the control plasmid pCMVβgal and the indicated expression plasmids using a cationic lipid-mediated method. The reporter plasmid is p3XE2Fluc, containing a luciferase gene driven by 3 copies of an E2F response element. Additional cotransfected plasmids were as follows: ---, vector; RB, pCMVNeoRB; AHR, pcDNAI/B6AHR. Cells were harvested for luciferase expression analysis

48 h after transfection. The ordinate represents luciferase activity normalized to β-galactosidase.

The values shown represent the mean ± S.D. from one representative experiment of at least two independent trials. Asterisks (*) indicate significant differences (P<0.05) from vector- transfected cells using one-way ANOVA, and exclamation points (!) represent significant differences from RB-transfected cells. C. AHR-mediated transactivation is not required for repression of E2F-dependent transcription in the absence of RB. C33A cell cultures were transfected as in B. Cotransfected plasmids were as follows: ---, vector; RB, pCMVNeoRB;

AHR, pcDNAI/B6AHR; AHR mutants are as described in the Experimental Procedures. The ordinate represents luciferase activity normalized to β-galactosidase. The values shown represent the mean ± S.E. of at least twelve log-transformed determinations from four independent experiments. Asterisks (*) indicate significant differences (P<0.05) from untransfected cells using one-way ANOVA. D. Hepa-1 cells were transfected with pCMVβgal, p3XE2Fluc, and pCMVNeoRB as above. In this case, cells were treated with 10 nM TCDD or

143

vehicle control 24 h before harvest. The ordinate represents luciferase activity normalized to β- galactosidase. The values shown represent the mean ± S.D. from one representative experiment of at least two independent trials. Asterisks (*) indicate significant differences (P<0.05) from vector-transfected, vehicle-treated cells using one-way ANOVA, and exclamation points (!) represent significant differences from vector-transfected, TCDD cells. E. AHR and E2F1 associate at common promoters. Hepa-1 cells from 10 150-mm plates were treated with 10 nM

TCDD or vehicle control for 1½ hours. Following formaldehyde crosslinking, chromatin was immunoprecipitated with αAHR, αE2F1, or no antibody (mock). DNA from the immunocomplex was extracted and purified, and polymerase chain reaction was used for detection of AHR or E2F1 binding sites in the promoters of the CYP1A1 and DHFR genes. The

PCR products were separated on a 10% acrylamide gel and visualized by ethidium bromide.

DMSO, vehicle control.

Fig. 2. AHR binds RB and E2F in vitro with equal affinities. A. Illustration of the full-length

RB protein expressed in vitro as a GST fusion protein, and domain deletions including GST-

A/B/C, which retains the large pocket domain, GST-SE, which consists solely of the C domain, and GST-AE∆1, consisting of the small pocket domain. GST-C706F is a one amino acid mutant in the B-domain of wild-type RB. B. AHR interacts with E2F1 as well as RB mutant proteins.

Truncated RB and full-length E2F1 GST fusion proteins were synthesized in bacteria, and proteins were captured by passing bacterial lysates over a glutathione sepharose slurry. [35S]-

AHR was synthesized using an in vitro transcription-translation protocol and passed several times over the glutathione columns. Bound proteins were eluted and analyzed by SDS-PAGE and autoradiography.

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Fig. 3. In vivo evidence for a direct interaction between the AHR and E2F1 transcription factors. A. AHR immunoprecipitates with E2F1 from Hepa-1 cell extracts. Cells grown in 150- mm plates were treated with 10 nM TCDD or vehicle control for 1 h. Nuclear extracts (300-500

µg total protein) were incubated with 10µL of a protein A/G bead mixture for 1 h at 40C.

Supernatants were then transferred to tubes containing protein A/G beads and 1µg of either an

AHR or E2F1 polyclonal antibody, followed by overnight rotation at 40C. After thorough washing, beads were boiled in an SDS loading buffer, and supernatants were analyzed by SDS-

PAGE and Western blot using the corresponding antibodies. B. AHR immunoprecipitates with

E2F1 from MEF cell extracts. TCDD treatment and experimental conditions were as in A. Cells were obtained from MEF+/+ or RB-/- C57BL6 mice. C. Saos-2 cells were infected at 100 PFU per cell with purified adenovirus containing genes for the expression of LacZ, AHR, RB, and

E2F1. After 24 hours, cells were treated with TCDD for 2 hours and harvested for whole cell extracts. Co-immunoprecipitations were carried out on 1 mg total protein extract with 2 ug of anti-AHR antibody, and eluted proteins were analyzed by Western blot. D. Saos-2 cells were infected at 100 PFU per cell with purified adenovirus containing genes for the expression of

LacZ, AHR, E2F1, E2F3, and E2F4. Co-immunoprecipitations and Western blots were carries out as in C.

Fig. 4. AHR interacts with E2F in gel shift assays and displaces RB from RB-E2F-DP1 complexes. A. Identification of DP, E2F, and RB-containing protein complexes in gel shift assays. Electrophoretic mobility shift assays were performed using 32P-labeled oligonucleotides of an E2F binding site incubated with CV-1 nuclear extracts. Samples were analyzed by acrylamide electrophoresis and autoradiography. Protein constituents of particular complexes

145

were identified by incubating samples with antibodies to the proteins of interest. B. AHR displaces RB from E2F binding sites occupied by RB/E2F/DP1 complexes, generating a novel complex containing E2F and AHR. Electrophoretic mobility shift assays were performed as above. AHR truncation mutants consisting of either the amino-terminal (∆376-805) or carboxy- terminal (∆1-425) half of the full-length protein were synthesized and purified from bacteria and incubated with samples following addition of the E2F-binding oligonucleotide. The double headed arrow denotes the novel complex. C. AHR is detected in complex with E2F and DP1.

Antibodies to ∆376-805 (αbHLH) and ∆1-425 (αYGOR) were incubated with CV-1 nuclear extracts following addition of the purified AHR peptides. Samples were subsequently analyzed by acrylamide elecrophoresis and autoradiography.

Fig. 5. AHR expression is required for basal expression levels of E2F-regulated genes. A.

AHR-mediated induction of Cyp1a1 gene expression is lost in AHR-/- MEF cells. MEF cells were cultured in the presence of serum in 10-cm plates and treated with Me2SO vehicle (DMSO) or 10 nM TCDD for 8 hours. Thereafter, cells were harvested for total RNA isolation, and mRNA of the indicated genes was quantified by real time PCR amplification. Products were

confirmed by agarose gel electrophoresis. The ordinate represents threshold cycle (Ct) values normalized to β-actin. The values shown represent the mean ± S.D. of two replicate experiments.

Cyp1a1, cytochrome P4501A1. B. AHR expression is required for basal expression of many

E2F-regulated genes. MEF cells were processed as in A. Tfdp1, transcription factor DP 1; Apaf1,

apoptotic peptidase activating factor 1; Cdt1, chromatin licensing and DNA replication factor 1;

Dnmt1, DNA methyltransferase (cytosine-5) 1; Mcm3, minichromosome maintenance deficient

3, Cdca7, cell division cycle associated 7; Cdca7-like, Cell division cycle associated 7-like;

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Cks1, cyclin-dependent kinase subunit 1; Ezh2, enhancer of zeste homolog 2; Hmgn1, high mobility group nucleosomal binding domain 1; Nasp, nuclear autoantigenic sperm protein; Pltp, phospholipid transfer protein. DMSO, vehicle control.

Fig. 6. The constitutive and TCDD-bound AHR binds differentially to E2F-regulated promoters. Samples were generated from the immunoprecipitation of chromatin isolated from

Hepa-1c1c7 cells treated with vehicle control or 10 nM TCDD. Immunoprecipitate DNA was isolated with IgG control or anti-AHR antibodies and used to probe custom microarray slides generated by Invitrogen, incorporating tiles of roughly 5400 bp of the promoter regions of the genes of interest. Each data point therefore represents tiles arrayed approximately every 350 bp between +400 and -5,000 from the transcription initiation site. Primers were designed to amplify DNA from ChIP experiments in order to generate amplicons that would hybridize to the tiles. The ordinate represents the ratios of experimental values to IgG controls, and the abscissa displays consensus DNA binding sites for AHR (blue) or E2F (pink). The values shown represent the mean ± S.D. of two replicate experiments. Apaf1, apoptotic peptidase activating factor 1; Cdt1, chromatin licensing and DNA replication factor 1; Dnmt1, DNA methyltransferase (cytosine-5) 1; Mcm3, minichromosome maintenance deficient 3. DMSO, vehicle control.

147

Figure 1

1 7 A. - - 2 - A a F 1 s p - 3 C o 3 e V a C H M C S RB

E. DMSO TCDD 1 1 r t R R e k F k F t u c H 2 c H 2 a o o np A E A E W I m α α m α α CYP1A1

DHFR

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Figure 2

A. A B C

1 379 572 772 928

A/B/C

C706F *

A/B

B. t C F u / 6 1 B 0 F p / 7 E /B 2 In A C S A E

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Figure 3

A. B.

αAHR αE2F1 TCDD - + - + - + - + D T D T AHR AHR

E2F1 E2F1 +/+ -/- +/+ -/- Extracts αAHR IP

1 F C. 2 E d 1 A F / B 2 B F F R E R D. 2 2 d d d E E d d A A A / / / A A 1 / / R R R R cZ F R R R R a H B H 2 H H cZ F cZ F H H H H L A R A E A A a 2 a 2 d d d d d d d L A E A L A E A d d d d d d d d A A A A A A A A A A A A A A A AHR E2F1

E2F1 E2F3

RB E2F4

IP: α-AHR Inputs IP: α-AHR

150

Figure 4

A. B. t

t c c B 1 1 a a - R k r F t B 2 P H F x R E D A N

E α α α α α Extr ∆376-805 ∆1-425

RB/E2F/DP1

E2F/DP1

H L R H O b G t α t c + c Y a a α r 5 5 r t 0 0 t R + x 8 H 8 x 5 5 e - L - e 2 O 2 6 6 l 4 4 ll 7 H 7 l - G - e 3 b 3 e 1 Y 1 C ∆ α ∆ C ∆ α ∆

151

Figure 5A

152

Figure 5B

153

Figure 6

4 4 DMSO/Ig/IgGG TCDD/IgG AHR DMSO/IgG E2F TCDD/IgG 3 3 AHR

3 l l l l l o

o E2F o o E2F o r r r r r t t t n n n o o o c cont c c cont

G G G G G g g 2 2 Ig Ig Ig to to to o o o i i i t t t a a a Ratio to I Ratio to I R R 1 R 1

0 0 -5000 -4000 -3000 -2000 -1000 +10 1000 -5000 -4000 -3000 -2000 -1000 +10 1000 Apaf1 5'-upstream regulatory region Cdt1 5'-upstream regulatory region 4 4 DMSO/IgG TCDD/IgG DMSO/IgG AHR TCDD/IgG E2F AHR 3 l l 3 l E2F o o o r r r ntrol ntrol ntrol o o o c c c G cont G cont G cont g g 2 g 2 to I to I to I Ratio Ratio Ratio Ratio to IgG Ratio to IgG Ratio to IgG 1 1

0 0 -6000 -5000 -4000 -3000 -2000 -1000 +10 1000 -6000 -5000 -4000 -3000 -2000 -1000 +10 1000 Dnmt1 5'-upstream regulatory region Mcm3 5'-upstream regulatory region

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Chapter V

ChIP-on-chip microarray analysis of AHR promoter binding sites in Hepa-1c1c7 cells

Jennifer L. Marlowe and Alvaro Puga*

*Corresponding author:

Keywords: TCDD, chromatin immunoprecipitation, ChIP-on-chip

Abstract

The aryl hydrocarbon receptor (AHR) is a ligand-inducible transcription factor of toxicological and physiological importance. The contribution of the AHR to toxicological outcomes has been extensively researched, and there are many environmental ligands of the

AHR that have been investigated for their effects in humans and ecological systems, including

TCDD and benzo[a]pyrene. The AHR is known to regulate gene expression primarily at the level of induction of cytochrome P450 drug-metabolizing enzymes. It has become evident that outside of this well-characterized role, the AHR also functions in many cellular signaling pathways. Mechanisms of transcriptional regulation by transcription factors, including the AHR, generally require binding of those factors to motifs in gene promoters or enhancers. This binding may be either direct at specific DNA motifs, or may be mediated by other proteins bound to the

DNA. In addition, transcription factors may bind DNA sequences that are totally unrelated to the canonical binding site for that factor. It is thus reasonable to presume that there exist currently unrecognized genes under direct regulation of the AHR. We have used chromatin immunoprecipitation (ChIP) to isolate DNA enriched for sequences that bind the AHR, either directly or via protein-protein interactions, from Hepa-1c1c7 cells treated with vehicle control or

TCDD. Immunoprecipitated DNA was amplified by ligation-mediated PCR, and this amplified

DNA was used to probe microarrays (Nimblegen Systems, Inc.) containing 5.0 kb of promoter

DNA for a comprehensive set of 30,000 genes within the mouse genome. Our results indicate that there are a subset of genes that are regulated by the AHR under physiologic conditions, and that TCDD treatment results in a significant alteration in the profile of AHR-regulated genes.

Hence, TCDD-mediated toxicity may be in part explained by the shift of AHR transcriptional activity from physiological target genes to primarily toxicant-induced target genes.

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Introduction

Although the AHR is required for dioxin-like compounds to exert their biological effects,

the role of this bHLH transcription factor extends beyond the activation of the best-known AHR

target genes, namely the Phase I (CYP1) and Phase II (NQO1, ALDH3A1, GSTA1-Ya,

UGT1A6) metabolism genes (1). The toxicological effects observed following TCDD treatment

in vivo and in a variety of cell cultures, including a disruption in liver homeostasis, immune

functions, and cell cycle perturbations in hepatoma and other cell lines, are simply not

adequately explained by activation of these target genes. Activation of the AHR by TCDD, for

example, confers expression changes in literally hundreds of genes (2). Notably, there have been

few functions described for the AHR outside of its activity as a transcription factor, or to

modulate the activities of other transcription factors (3). The basic mechanisms of

transcriptional regulation in general consist of a requirement for direct binding of transcription

factors to motifs in gene promoters or enhancers. This binding may be direct at specific DNA

motifs, or mediated by other proteins bound to DNA. This appears to be the case among some

E2F-regulated genes, in which AHR binding results in down-regulation of their expression

(Chapters 3 & 4, and Ref. (4)). In addition, transcription factors may bind DNA sequences that are totally unrelated to the canonical binding site for that factor (5). For example, several studies found that a significant number of promoters bound directly by E2F1 or E2F4 contain no identifiable E2F binding sites (6;7), and that RB is actually bound to a number of gene promoters both constitutively as well as during distinct cell cycle phases (8). Considering this, it is reasonable to presume that there exist currently unrecognized genes under direct regulation of the AHR, and that the resulting changes in expression of these genes following AHR activation may in part explain AHR-dependent toxicity.

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Chromatin immunoprecipitation (ChIP) techniques can be used to isolate DNA enriched

for promoter or enhancer sequences that bind transcription factors, either directly or via protein-

protein interactions (see Fig. 2). Typically, this technique is used to probe for particular

sequences using PCR analysis, and is thus limited to analyzing only pre-determined sequences.

Recently, researchers have begun using microarrays to analyze the DNA isolated from ChIP

experiments. Array experiments using DNA isolated from ChIP (commonly known as ChIP-on-

chip) offer researchers the ability to monitor chromatin structure as well as identify the target

genes of transcription factors en masse. Gene expression microarrays combined with ChIP-on-

chip analysis also offer the opportunity to identify broad patterns of regulated genes to give

novel insight into the biological processes regulated by a particular DNA-binding transcription

factor (9;10). Utilization of both types of microarrays has led to major advances in the

determination of the many ways in which transcription factors control normal cellular

physiology. ChIP-on-chip has been used to identify an array of novel gene targets for commonly

occurring factors such as p53, E2F1, estrogen receptor, NF-Y, and CTCF (11-15).

Developmental and transgenic mouse studies provide evidence that the AHR is more than

a sensor of xenobiotic exposure, but is also a key component of normal physiology (3). Thus it is

logical that as a transcription factor, the AHR would actively target genes for transcriptional

regulation in its “unliganded” state (possibly bound by endogenous ligands) in addition to those

it targets in its ligand-activated state. For example, it has been demonstrated that the AHR is

transiently activated in quiescent cells by the addition of serum, and this activation is followed

by a transient increase in Cyp1a1 protein levels (16). AHR has also been shown to be activated

by loss of cell contact in 10T1/2 fibroblasts (17). However, it is likely that any activities of the

“unliganded” AHR are actually mediated by a number of cell-type and tissue-specific ligands

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which have yet to be identified. There is also an abundance of evidence to suggest that the toxicological effects of ligands such as TCDD represent the deregulated physiology of the AHR

(3;18), and that clues as to the basal activities of the AHR may be found in examining the diversity of ligand-induced transcriptional targets of the AHR. We have therefore utilized the

DNA isolated from AHR-specific immunoprecipitations to probe promoter tiling microarrays from Nimblegen Systems, Inc. in order to identify as-yet-unidentified genes targeted by the AHR following TCDD treatment, as well as to analyze targets of the AHR in its unliganded or physiologic state.

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Materials and Methods

Chromatin Immunoprecipitation

The ChIP protocol was performed with minor modifications as described previously by

others (6;19). Figure 1 provides an illustration of the ChIP protocol, with additional steps for

sample amplification and array hybridization. Hepa-1c1c7 cells were seeded in 15-cm plates at a

density of 1x107 cells per plate in αMEM containing 5% FBS. The following day, cells were

treated with 10nM TCDD or the equivalent volume of DMSO vehicle for one hour. A total of

one plate of cells was used per treatment, per antibody, and three to five replicate sets were

performed for each treatment-antibody combination. Cross-linking was performed by adding

formaldehyde directly to the culture media to a final concentration of 1%, followed by

incubation for 10 minutes. The formaldehyde reaction was quenched using glycine at a final

concentration of 0.125 M. Cells were rinsed twice with cold PBS, scraped from the dishes,

pelleted and washed again with PBS containing 0.5 mM PMSF. Cell pellets were resuspended in

cell lysis buffer (5 mM PIPES (pH 8.0), 85 mM KCl, 0.5% NP-40, 0.5 mM PMSF, 5 µg/ml

leupeptin, 5 µg/ml aprotinin) and incubated on ice for 10 min. Nuclei were pelleted and

resuspended in nuclei lysis buffer (50 mM Tris-HCl (pH 8.1), 10 mM EDTA, 1% SDS, 0.5 mM

PMSF, 5 µg/ml leupeptin, 5 µg/ml aprotinin) and incubated on ice for 10 min. Chromatin was

sonicated to an average length of 600 bp with 4 10-second pulses of 30 W, maintaining samples

on ice for 30 seconds between pulses. Sonicated chromatin was then precleared for 1-2 hours at

4°C with BSA- and salmon sperm DNA-saturated protein A agarose (Upstate Biotechnology) in

preparation for immunoprecipitation. The supernatant was divided equally among all samples

and incubated overnight on a rotating platform at 4°C with 1 µg of AHR (BioMol) or E2F1

(Santa Cruz), antibodies. Protein A agarose slurry (20 µl) was added and incubated for 15 min at

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room temperature to allow for antibody binding. Agarose was pelleted and washed twice with

1X dialysis buffer (50 mM Tris-HCl (pH 8.0), 2 mM EDTA, 0.2% sarkosyl) and 4 times with IP

wash buffer (100 mM Tris-HCl (pH 9.0), 500mM LiCl, 1% NP-40, 1% deoxycholic acid).

Immune complexes were eluted from the beads with elution buffer (50 mM NaHCO3, 1% SDS).

Reversal of crosslinks was carried out by heating the eluates at 67°C for 4-5 h. The eluates were

then digested with proteinase K at 56°C for 1.5 h. DNA was then purified using the QIAquick

PCR purification kit (Qiagen) and eluted into 30-50 µL ddH20.

Real Time PCR

Samples were subjected to PCR amplifications with the following promoter-specific primer sets: mouse Cyp1a1 (enhancer domain -1141 to -784 bp): forward primer, 5’-

AAGCATCACCCTTTGTAGCC-3’; reverse primer, 5’-CAGGCAACACAGAGAAGTCG-3’; amplification product, 121 bp. Mouse Nqo1: forward primer, 5’-

TAAGAGCAGAACGCAGCAC-3’; reverse primer, 5’-ACCTGCCTACATAATCAGCC-3’; amplification product, 190 bp. Reaction volumes of 25 µL contained 12.5 µL Brilliant SYBR

Green QRT-PCR Master Mix (Stratagene), 0.4 µM primers, and 2 µL ChIP DNA template or 2

µL of a 50X dilution of total DNA. . Real-time quantitative PCR was performed using a Smart

Cycler rapid thermal cycler (Cepheid) and fluorescence was measured after each of the repetitive cycles. A typical protocol included a 2 min denaturation step at 95oC followed by 40 cycles of

95oC denaturation for 30 sec, annealing for 30 sec at a primer-optimized temperature, and 72oC extension for 30 sec. Detection of the fluorescent product was carried out during the 72oC extension period, and emission data were quantified using threshold cycle (Ct) values. Ct values for all genes analyzed were determined once, and means were calculated from the average Ct values for each biological duplicate. All means were then normalized to Ct values for total

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(“input”) DNA samples. The relative or fold-change from control Ct values (DMSO treated and immunoprecipitated with anti-rabbit IgG) was determined for each sample, as the following calculation illustrates:

Fold change = 2-∆∆Ct

where ∆∆Ct = (CtChIP– CtInput)anti-AHR – (CtChIP – CtInput)anti-IgG. PCR product specificity from each primer pair was confirmed using melting curve analysis and subsequent agarose gel electrophoresis.

Ligation-mediated PCR

Ligation-mediated PCR (LM-PCR) was performed essentially as described to create amplicons of ChIP DNA (20). Two to three replicate samples from each treatment group

(DMSO, anti-IgG; DMSO, anti-AHR; TCDD, anti-AHR) were combined for each LM-PCR reaction. In addition, 20 ng of total DNA from each group was amplified to use as a reference on the CpG microarray. First, the two unidirectional linkers JW102 (5’-

GCGGTGACCCGGGAGATCTGAATTC-3’) and JW103 (5’-GAATTCAGATC-3’) were annealed to one another. At the same time, the ChIP DNA was blunt-ended in order to eliminate the possibility of overhanging DNA ends in the samples, the reaction was purified using

QIAquick PCR purification columns, and eluted into 30 µL elution buffer. An overnight ligation reaction was performed to ligate the blunted ChIP DNA to the annealed linkers. Again, the reaction was purified with QIAquick columns and eluted in 30 µL elution buffer. Finally, the linker-ligated DNA was PCR-amplified using 25 µL chromatin, 10X Taq polymerase buffer, 140

µM dNTPs, 20 µM primer JW102, 1M betaine, and 1 µL Taq (Promega), and the reaction is purified as above and eluted into 30 µL elution buffer. The products of this step were used as

stocks for further PCR amplifications, until roughly 1-10 ug of DNA were produced. Finally,

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10ng of each stock were PCR-amplified as above to produce roughly 1-2 ug of DNA for each

group to be used in probing a promoter tiling array. In addition, PCR was performed on each

amplified sample to confirm the pre-amplification pattern of known AHR target genes (Cyp1a1,

Nqo1).

Promoter Tiling Array

Detailed information on promoter tiling arrays used in these experiments can be found

at http://www.nimblegen.com/. The promoter array design is a two array set from Nimblegen

Systems, Inc., containing 5.0 kb of promoter region for a comprehensive set of genes representing the mouse genome. All 5.0 kb regions are tiled at 110 bp intervals. For array hybridization, amplicons were analyzed and quantitated according to the instructions from

Nimblegen, and all labeling and hybridization reactions were carried out by Nimblegen. Each

ChIP sample was hybridized against the corresponding total (or “input”) DNA sample for that treatment-antibody combination. Arrays scans, data extraction, and preliminary analysis were also carried out by Nimblegen. Extracted data was visualized using the SignalMap software package.

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Results

Known AHR targets are enriched using ChIP

Hepa-1c1c7 cells are commonly used in studies of AHR-mediated gene regulation and of

prototypical AHR target genes, particularly those of the CYP1 family of cytochrome P450

enzymes, and are known to express high receptor levels (21). We used these cells in chromatin

immunoprecipitation experiments in order to isolate AHR-bound target DNA and to use this

DNA in probing promoter microarrays. Chromatin was immunoprecipitated using anti-AHR (for

samples treated with Me2SO and TCDD) or anti-rabbit IgG antibodies (for Me2SO-treated samples only). We confirmed the enrichment of known AHR-target genes in these samples using primers for the amplification of AHR-binding sequences in the enhancer region of the

Cyp1a1 gene (22) and the promoter of the Nqo1 gene (23). For TCDD-treated samples immunoprecipitated with anti-AHR antibodies, the proportion of the total sample containing

AHR target DNA should be greater than either the Me2SO-treated samples immunoprecipitated

with anti-AHR antibodies or those immunoprecipitated with anti-rabbit IgG antibodies. Figure

2A shows the PCR products after 40 cycles from replicate samples amplified with primers specific for the mouse Cyp1a1 enhancer region. Products shown correspond to the expected molecular weight, and are present in all total (“input”) DNA samples, as expected. The Cyp1a1 enhancer PCR product is not present in samples immunoprecipitated with IgG control antibodies, but is clearly amplified from either control-treated cells or TCDD-treated cell immunoprecipitated with anti-AHR. The mean cycle threshold (Ct) values from real-time PCR reactions of the biological replicate samples show that DNA from the TCDD-treated cells is enriched for both Cyp1a1 promoter DNA as well as Nqo1 enhancer DNA as compared to either

Me2SO (“DMSO”) or IgG controls (Fig. 2B). Interestingly, Cyp1a1 is enriched over IgG

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controls in Me2SO-treated cells immunoprecipitated with anti-AHR. This is not entirely surprising, as the AHR is known to exhibit basal activities, including regulation of the Cyp1a1 gene in the absence of exposure to exogenous ligands (16;24). Figure 2C shows the normalized

Ct values of ChIP DNA samples to the Ct values of their respective total input DNA. All values are calculated relative to the IgG control. Therefore, for vehicle-treated cells, the AHR immunoprecipitated approximately 3-fold more Cyp1a1 promoter DNA as IgG controls, and

350-fold more in TCDD-treated samples. For the Nqo1 promoter, the input-normalized values are 120-fold higher than IgG control, and there is no difference between vehicle and IgG for this gene.

AHR target promoter pattern is unchanged for LM-PCR-amplified samples

In order to use the ChIP DNA isolated from Hepa-1c1c7 cells to probe promoter microarrays, the samples must be amplified. We used a ligation-mediated PCR protocol in order to generate amplicons of ChIP and Input DNA samples for this purpose. The relative amounts of individual DNA sequences should be the same before and after amplification for each samples.

To confirm this, we used 10 ng of the final amplified product for each sample in a standard PCR reaction using the same Cyp1a1 and Nqo1 primers that were used to verify the success of the original ChIP experiment. The overall pattern of Ct values obtained by real-time PCR using these primer sets is unchanged as compared to the original samples (Fig. 3A vs. Fig. 2B), indicating that the relative enrichment of AHR target DNA sequences in the amplified ChIP samples is unchanged relative to the original samples. This is confirmed by normalizing the Ct values of the samples to their respective inputs (Fig. 3B). The enrichment relative to IgG control of both Cyp1a1 and Nqo1 promoter sequences in TCDD-treated samples is essentially the same as that observed for the original ChIP samples (Fig. 2C). Six-hundred nanograms each of total

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DNA and DNA isolated by ChIP was run on a DNA 7500 LabChip kit (Agilent) and analyzed

using the Agilent 2100 Bioanalyzer (Fig. 3C). Only samples with OD260/OD280 ratios greater

than 1.7 were used for labeling and microarray analysis.

The AHR targets different sets of genes in its physiological and TCDD-activated states

LM-PCR amplified ChIP samples were labeled and hybridized to promoter tiling

microarrays by Nimblegen Systems, Inc. Results were analyzed using the SignalMap software

package. SignalMap allows for the visualization of NimbleGen array data as peaks which correlate to specific annotated sequences, thereby enabling one to identify gene regions corresponding to those data points. High signal intensities or peaks in the data found using the

SignalMap platform should therefore indicate regions that are targeted by the transcription factor of interest, in this case the AHR. Figure 4 shows an example of the SignalMap analysis for chromosome two, in which data points are shown as peaks along the chromosome corresponding to the hybridization intensities of the samples at that particular location. Tables 1 and 2 list the

50 genes in the entire genome for which the signal intensities of the ChIP:input DNA was the

highest, indicating that the samples are enriched for DNA sequences targeted by the AHR. For

cells treated with vehicle control, in which the AHR is not activated by exogenous ligand but still

maintains poorly-understood basal or physiological functions, ChIP samples appear enriched in

sequences for genes functioning in development (Tsc22d1, Sema4g), transcription (Taf6, Taf9b,

Ixl, Rfxdc1, Falz), numerous physiological processes (Mpv17 (kidney), Npy1r and Falz (nervous system), Alox12e (arachidonic acid metabolism), Esr2 (sex organ functions), Ryr2 and NyrY1

(cardiovascular system), Fcrl5 (immune system)), cell cycle (Araf, Bcl2, Rbm7, Bcl2l2), cell adhesion/structure (Frmd6, Lrrc16, Omd, Sdc2), signaling cascades (Dusp14, Spag9, Rab18), and motor functions (Dnaic1, Bicd2). For cells treated with TCDD, the highest data peaks were

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found to correlate with genes functioning in similar pathways, including cell cycle (Kifap3,

Ccnl2, Scap2, Nek3, Tgfb3, Cetn2), nervous system function (Adam22, Pou6f2, Pex2), cell

adhesion/structure (Kifap3, Lad1, Fbn1, Bcar3, Hook3), immune system (Defb38, Cd209b,

Elmo1), and transcription (Ixl, Nap1l1, Paf1). Additional genes and pathways are also

implicated as TCDD-activated AHR targets, including lipid biosynthesis/metabolism (Nsdhl,

Pctp, Soat1, Dbi), retinoic acid production (Psip1), oxidative stress (Ero1l, Oxr1), and PKC activation (Dgkg). Interestingly, one gene, Ixl, was found in common between the vehicle- treated and TCDD-treated samples. The Ixl gene encodes a known component of the mammalian mediator complex, a major component of transcriptional activation and initiation

(25). Tables 3 and 4 list the same 50 genes matching the highest-intensity signals on each array, with the corresponding canonical AHR and E2F binding sites in their respective promoters.

AHR binding sites were identified by scanning the first 5000 base pairs upstream of the transcription start site for each gene, as identified by the Ensembl genome browser, for the invariate XRE-containing nucleotides 5’-GTGCG-3' or its complement. The same sequences

were analyzed by the TFSEARCH program (http://www.cbrc.jp/research/db/TFSEARCH.html) in order to identify potential E2F binding sites. Proximal sites are defined as those within 300 bp of the transcription start site, and distal binding site are up to 5000 bases upstream of this location. Finally, we scanned data from a majority of expression microarray studies that were conducted to assess gene expression changes mediated by the AHR, either through exogenous ligand activation or endogenous functions. Table 5 compares AHR target genes identified in our

ChIP-on-chip microarray experiment with genes found to be altered in expression microarrays.

Fourteen of the 38 annotated genes identified as AHR targets in DMSO-treated cells were altered in expression array studies, as well as eight of the 37 annotated genes in TCDD-treated cells.

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Discussion

The chromatin immunoprecipitation assay has been widely used for the identification of transcription factor binding sites and corresponding target genes. Applying the ChIP technique to newly available promoter or whole-genome tiling microarrays allows for an unbiased analysis of genome-wide transcription factor binding and the identification of as-yet unknown gene targets of that factor. In addition, ChIP-on-chip data can be combined with traditional expression microarray to broaden the analysis to entire transcriptional networks modified by a particular protein. Analysis of transcriptional pathways modified by AHR expression or activation by exogenous ligands has thus far not extended beyond the use of expression arrays. The results reported here represent the first ChIP-on-chip analysis of DNA sequences targeted for binding by the AHR, either through canonical AHR binding sites, protein-protein interactions, or through still uncharacterized mechanisms.

The gene lists presented in Tables 1 and 2 represent the most likely AHR targets in vehicle-treated and TCDD-treated cells, based upon analysis of the highest fifty signal ratios of the samples, an arbitrary cutoff, compared to their respective total input DNA. Strengthening the validity of this arbitrary cutoff is the fact that of all of the annotated genes identified as AHR targets in the ChIP-on-chip experiment, roughly 30% (22 of 75; see Table 5) have been reported to be altered in expression microarrays (2;26-37), several in multiple different reports. These expression microarrays were carried out in a variety of cell and organ systems in which TCDD is used as the AHR inducer, or in cell systems in which AHR expression is lost or the normal AHR activation pathway is altered in some way. For example, many genes identified in our promoter array were found to be differentially regulated in c1 cells, a Hepa-1c1c7 cell line derivative in which Cyp1a1 enzyme activity is lost. In these cells, it is thought that the AHR maintains some

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constitutive activity due to a lack of metabolism of endogenous AHR ligands by Cyp1a1 (28).

The differences in gene expression would therefore be attributable to increased nuclear localization and activity of the AHR, and as our data shows, a correlative increase in gene promoter targeting by the AHR. However, it should be noted that none of the best-characterized targets of the AHR were found within the cutoff point mentioned above. In the case of the

Cyp1a1 promoter, one of the genes used to verify the quality of the ChIP samples used to probe the microarrays, an incorrect annotation for the gene resulted in the absence of its promoter on the Nimblegen array.

Of the genes identified as AHR targets in our promoter array, nearly all of them have one or more canonical AHR binding sites in the regions 5000 bp upstream of the first exon of the annotated gene (see Tables 3 and 4 for AHR targets in Me2SO-treated and TCDD-treated cells, respectively). Many of these genes also contain typical E2F transcription factor binding sites, implicating multiple potential mechanisms for their regulation by the AHR. It is interesting to note that only one gene is shared by both the gene lists. The implication of this is unclear. It is possible that the AHR targets the same genes in both cases, but that differences in AHR recruitment to different binding sites, or interactions of the receptor with other proteins in its unliganded or liganded states, may result in relative increases in certain sequences over others under the different treatment conditions. In this case, the mechanisms of transcriptional regulation would be altered by the AHR being targeted to different binding sites of the same gene under different conditions, as was observed in Chapter 4 concerning certain E2F target genes, such that weak AHR binding under physiological conditions becomes strong AHR binding following induction by TCDD. However, it is also possible that the binding priority of the AHR shifts in response to exogenous activation by ligand, and the balance of AHR target

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genes is transferred from primarily physiological targets to primarily toxicological targets.

An emerging hypothesis of AHR-mediated toxicity concerns the ability of TCDD to

essentially deregulate normal functions of the AHR (3). In many cases, the defects observed

upon loss of AHR expression mimic those effects observed upon activation of the receptor by

persistent agonists like TCDD (18). It is notable that many of the gene promoters bound by

AHR in either its unliganded or TCDD-activated state are in similar pathways having been

previously implicated in AHR-mediated toxicity (38;39). These pathways include cell cycle

regulation (18), nervous system function and development (40-42), cell adhesion/structure

(17;43), immune system functions (44-46), and transcriptional regulation (47;48). Interestingly,

TCDD-induced AHR target genes are also involved in pathways specifically implicated in the

toxicity of TCDD, including oxidative stress response genes (49-51), PKC activation (52), and

genes involved in retinoic acid (53) and lipid biosynthesis/metabolism (54;55). Functions of the

AHR have also been associated with pathways implicated here as AHR targets under non-TCDD

induced conditions, including arachidonic acid metabolism (56), estrogen signaling (57), and

cardiovascular functions (58).

In most microarray experiments examining gene expression changes induced by TCDD,

literally hundreds of genes are found to be increased or decreased, the bulk of which are primary

responses to AHR activation (2). It seems likely that the mechanisms mediating up- or down-

regulation of genes by the AHR are quite different, and probably dependent not only on the particular promoter examined, but also on cell context. A variety of different mechanisms may account for differential binding of the AHR to promoter sequences. A few of those mechanisms are now known, including interactions or crosstalk of the receptor with other transcription factor proteins such as RB (59;60), E2F (4), NF-κB (61), ER (62), and AR (63), and binding of the

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receptor to non-canonical XRE sequences (64) as well as inhibitory XREs (65). Given that many of the genes identified here as direct AHR targets have also been found in analyses of AHR function in expression arrays, the genes are likely to be bonified AHR target genes. Further analyses are required in order to determine the mechanisms of AHR-dependent modulation of these genes, as well as their significance to AHR physiology and dioxin toxicology.

Considering the myriad of pathways and functions in which AHR function has been implicated, and the fact that many labs have used expression arrays to discern AHR involvement in transcriptional pathways, use of the ChIP-on-chip technique is a valuable addition to the continuing dissection of AHR gene targets. ChIP-on-chip may also be useful for determining differences in physiological versus toxicological targets of the AHR, and hence explaining

TCDD-mediated toxicity, as well as the toxicity of AHR ligands in general. While the physiological functions of the AHR remain ill-defined, it has been implicated in many of the signaling pathways in which the aforementioned genes are a part, particularly with regards to cell cycle control, nervous system function and development, and the vasculature. The data presented here suggest a definite shift in the transcriptional targets of the AHR under physiological and TCDD-induced conditions. It is our opinion that this shift in AHR targeting is a major key to understanding TCDD-mediated toxicity.

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References

1. Weiss C, Kolluri SK, Kiefer F, and Gottlicher M. 1996. Complementation of Ah receptor deficiency in hepatoma cells: negative feedback regulation and cell cycle control by the Ah receptor. Exp.Cell Res. 226: 154-163

2. Puga A, Maier A, and Medvedovic M. 2000. The transcriptional signature of dioxin in human hepatoma HepG2 cells. Biochem.Pharmacol. 60: 1129-1142

3. Bock KW and Kohle C. 2006. Ah receptor: dioxin-mediated toxic responses as hints to deregulated physiologic functions. Biochem.Pharmacol. 72: 393-404

4. Marlowe JL, Knudsen ES, Schwemberger S, and Puga A. 2004. The aryl hydrocarbon receptor displaces p300 from E2F-dependent promoters and represses S phase-specific gene expression. J.Biol.Chem. 279: 29013-29022

5. Wells J, Graveel CR, Bartley SM, Madore SJ, and Farnham PJ. 2002. The identification of E2F1-specific target genes. Proc.Natl.Acad.Sci.U.S.A 99: 3890-3895

6. Weinmann AS and Farnham PJ. 2002. Identification of unknown target genes of human transcription factors using chromatin immunoprecipitation. Methods 26: 37-47

7. Ren B, Cam H, Takahashi Y, Volkert T, Terragni J et al. 2002. E2F integrates cell cycle progression with DNA repair, replication, and G(2)/M checkpoints. Genes Dev. 16: 245- 256

8. Wells J, Yan PS, Cechvala M, Huang T, and Farnham PJ. 2003. Identification of novel pRb binding sites using CpG microarrays suggests that E2F recruits pRb to specific genomic sites during S phase. Oncogene 22: 1445-1460

9. Negre N, Lavrov S, Hennetin J, Bellis M, and Cavalli G. 2006. Mapping the distribution of chromatin proteins by ChIP on chip. Methods Enzymol. 410: 316-341

10. Blais A and Dynlacht BD. 2005. Constructing transcriptional regulatory networks. Genes Dev. 19: 1499-1511

11. Ceribelli M, Alcalay M, Vigano MA, and Mantovani R. 2006. Repression of new p53 targets revealed by ChIP on chip experiments. Cell Cycle 5: 1102-1110

12. Bieda M, Xu X, Singer MA, Green R, and Farnham PJ. 2006. Unbiased location analysis of E2F1-binding sites suggests a widespread role for E2F1 in the human genome. Genome Res. 16: 595-605

13. Testa A, Donati G, Yan P, Romani F, Huang TH et al. 2005. Chromatin immunoprecipitation (ChIP) on chip experiments uncover a widespread distribution of NF- Y binding CCAAT sites outside of core promoters. J.Biol.Chem. 280: 13606-13615

172

14. Jin VX, Leu YW, Liyanarachchi S, Sun H, Fan M et al. 2004. Identifying estrogen receptor alpha target genes using integrated computational genomics and chromatin immunoprecipitation microarray. Nucleic Acids Res. 32: 6627-6635

15. Mukhopadhyay R, Yu W, Whitehead J, Xu J, Lezcano M et al. 2004. The binding sites for the chromatin insulator protein CTCF map to DNA methylation-free domains genome- wide. Genome Res. 14: 1594-1602

16. Levine-Fridman A, Chen L, and Elferink CJ. 2004. Cytochrome P4501A1 Promotes G1 Phase Cell Cycle Progression by Controlling Aryl Hydrocarbon Receptor Activity. Mol.Pharmacol. 65: 461-469

17. Cho YC, Zheng W, and Jefcoate CR. 2004. Disruption of cell-cell contact maximally but transiently activates AhR-mediated transcription in 10T1/2 fibroblasts. Toxicol.Appl.Pharmacol. 199: 220-238

18. Marlowe JL and Puga A. 2005. Aryl hydrocarbon receptor, cell cycle regulation, toxicity, and tumorigenesis. J.Cell Biochem. 96: 1174-1184

19. Wells J and Farnham PJ. 2002. Characterizing transcription factor binding sites using formaldehyde crosslinking and immunoprecipitation. Methods 26: 48-56

20. Oberley MJ and Farnham PJ. 2003. Probing chromatin immunoprecipitates with CpG- island microarrays to identify genomic sites occupied by DNA-binding proteins. Methods Enzymol. 371: 577-596

21. Cuthill S, Poellinger L, and Gustaffson J-Å. 1987. The receptor for 2,3,7,8- tetrachlorodibenzo-p-dioxin in the mouse hepatoma cell line Hepa 1c1c7. J.Biol.Chem. 262: 3477-3481

22. Wu L and Whitlock JP, Jr. 1993. Mechanism of dioxin action: receptor - enhancer interactions in intact cells. Nucleic Acids Res. 21: 119-125

23. Nioi P and Hayes JD. 2004. Contribution of NAD(P)H:quinone oxidoreductase 1 to protection against carcinogenesis, and regulation of its gene by the Nrf2 basic-region leucine zipper and the arylhydrocarbon receptor basic helix-loop-helix transcription factors. Mutat.Res. 555: 149-171

24. Chang C-Y and Puga A. 1998. Constitutive activation of the aromatic hydrocarbon receptor. Mol.Cell.Biol. 18: 525-535

25. Lewis BA and Reinberg D. 2003. The mediator coactivator complex: functional and physical roles in transcriptional regulation. J.Cell Sci. 116: 3667-3675

26. Vezina CM, Walker NJ, and Olson JR. 2004. Subchronic exposure to TCDD, PeCDF, PCB126, and PCB153: effect on hepatic gene expression. Environ.Health Perspect. 112: 1636-1644

173

27. Boverhof DR, Burgoon LD, Tashiro C, Chittim B, Harkema JR et al. 2005. Temporal and dose-dependent hepatic gene expression patterns in mice provide new insights into TCDD- Mediated hepatotoxicity. Toxicol.Sci. 85: 1048-1063

28. Fong CJ, Burgoon LD, and Zacharewski TR. 2005. Comparative microarray analysis of basal gene expression in mouse Hepa-1c1c7 wild-type and mutant cell lines. Toxicol.Sci. 86: 342-353

29. Martinez JM, Afshari CA, Bushel PR, Masuda A, Takahashi T, and Walker NJ. 2002. Differential toxicogenomic responses to 2,3,7,8-tetrachlorodibenzo-p-dioxin in malignant and nonmalignant human airway epithelial cells. Toxicol.Sci. 69: 409-423

30. Hanlon PR, Zheng W, Ko AY, and Jefcoate CR. 2005. Identification of novel TCDD- regulated genes by microarray analysis. Toxicol.Appl.Pharmacol. 202: 215-228

31. Hanlon PR, Cimafranca MA, Liu X, Cho YC, and Jefcoate CR. 2005. Microarray analysis of early adipogenesis in C3H10T1/2 cells: cooperative inhibitory effects of growth factors and 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicol.Appl.Pharmacol. 207: 39-58

32. McHale CM, Zhang L, Hubbard AE, Zhao X, Baccarelli A et al. 2006. Microarray analysis of gene expression in peripheral blood mononuclear cells from dioxin-exposed human subjects. Toxicology

33. Jin B, Kim G, Park DW, and Ryu DY. 2004. Microarray analysis of gene regulation in the Hepa1c1c7 cell line following exposure to the DNA methylation inhibitor 5-aza-2'- deoxycytidine and 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicol.In Vitro 18: 659-664

34. Bemis JC, Alejandro NF, Nazarenko DA, Brooks AI, Baggs RB, and Gasiewicz TA. 2006. TCDD-Induced Alterations in Gene Expression Profiles of the Developing Mouse Paw Do Not Influence Morphological Differentiation of This Potential Target Tissue. Toxicol.Sci.

35. Fisher MT, Nagarkatti M, and Nagarkatti PS. 2004. Combined screening of thymocytes using apoptosis-specific cDNA array and promoter analysis yields novel gene targets mediating TCDD-induced toxicity. Toxicol.Sci. 78: 116-124

36. Fletcher N, Wahlstrom D, Lundberg R, Nilsson CB, Nilsson KC et al. 2005. 2,3,7,8- Tetrachlorodibenzo-p-dioxin (TCDD) alters the mRNA expression of critical genes associated with cholesterol metabolism, bile acid biosynthesis, and bile transport in rat liver: a microarray study. Toxicol.Appl.Pharmacol. 207: 1-24

37. Karyala S, Guo J, Sartor M, Medvedovic M, Kann S et al. 2004. Different global gene expression profiles in benzo[a]pyrene- and dioxin-treated vascular smooth muscle cells of AHR-knockout and wild-type mice. Cardiovasc.Toxicol. 4: 47-73

38. Fernandez-Salguero P, Pineau T, Hilbert DM, McPhail T, Lee SS et al. 1995. Immune system impairment and hepatic fibrosis in mice lacking the dioxin-binding Ah receptor. Science 268: 722-726

174

39. Brunnberg S, Andersson P, Lindstam M, Paulson I, Poellinger L, and Hanberg A. 2006. The constitutively active Ah receptor (CA-Ahr) mouse as a potential model for dioxin exposure--effects in vital organs. Toxicology 224: 191-201

40. Builee TL and Hatherill JR. 2004. The role of polyhalogenated aromatic hydrocarbons on thyroid hormone disruption and cognitive function: a review. Drug Chem.Toxicol. 27: 405- 424

41. Qin H and Powell-Coffman JA. 2004. The Caenorhabditis elegans aryl hydrocarbon receptor, AHR-1, regulates neuronal development. Dev.Biol. 270: 64-75

42. Williamson MA, Gasiewicz TA, and Opanashuk LA. 2005. Aryl hydrocarbon receptor expression and activity in cerebellar granule neuroblasts: implications for development and dioxin neurotoxicity. Toxicol.Sci. 83: 340-348

43. Waring JF, Gum R, Morfitt D, Jolly RA, Ciurlionis R et al. 2002. Identifying toxic mechanisms using DNA microarrays: evidence that an experimental inhibitor of cell adhesion molecule expression signals through the aryl hydrocarbon nuclear receptor. Toxicology 181-182: 537-550

44. Forawi HA, Tchounwou PB, and McMurray RW. 2004. Xenoestrogen modulation of the immune system: effects of dichlorodiphenyltrichloroethane (DDT) and 2,3,7,8- tetrachlorodibenzo-p-dioxin (TCDD). Rev.Environ.Health 19: 1-13

45. Birnbaum LS and Tuomisto J. 2000. Non-carcinogenic effects of TCDD in animals. Food Addit.Contam 17: 275-288

46. Nohara K, Pan X, Tsukumo S, Hida A, Ito T et al. 2005. Constitutively active aryl hydrocarbon receptor expressed specifically in T-lineage cells causes thymus involution and suppresses the immunization-induced increase in splenocytes. J.Immunol. 174: 2770- 2777

47. Wang S, Ge K, Roeder RG, and Hankinson O. 2004. Role of mediator in transcriptional activation by the aryl hydrocarbon receptor. J.Biol.Chem. 279: 13593-13600

48. Hankinson O. 2005. Role of coactivators in transcriptional activation by the aryl hydrocarbon receptor. Arch.Biochem.Biophys. 433: 379-386

49. Hassoun EA, Wilt SC, DeVito MJ, Van Birgelen A, Alsharif NZ et al. 1998. Induction of oxidative stress in brain tissues of mice after subchronic exposure to 2,3,7,8- tetrachlorodibenzo-p-dioxin. Toxicol.Sci. 42: 23-27

50. Slezak BP, Hatch GE, DeVito MJ, Diliberto JJ, Slade R et al. 2000. Oxidative stress in female B6C3F1 mice following acute and subchronic exposure to 2,3,7,8- tetrachlorodibenzo-p-dioxin (TCDD). Toxicol.Sci. 54: 390-398

175

51. Shertzer HG, Nebert DW, Puga A, Ary M, Sonntag D et al. 1998. TCDD causes a sustained oxidative stress response in C57BL/6J mice. Biochem.Biophys.Res.Comm. 253: 44-48

52. Long WP, Pray-Grant M, Tsai JC, and Perdew GH. 1998. Protein kinase C activity is required for aryl hydrocarbon receptor pathway-mediated signal transduction. Mol.Pharmacol. 53: 691-700

53. Fletcher N, Giese N, Schmidt C, Stern N, Lind PM et al. 2005. Altered retinoid metabolism in female Long-Evans and Han/Wistar rats following long-term 2,3,7,8-tetrachlorodibenzo- p-dioxin (TCDD)-treatment. Toxicol.Sci. 86: 264-272

54. Albro PW, Corbett JT, Harris M, and Lawson LD. 1978. Effects of 2,3,7,8- tetrachlorodibenzo-p-dioxin on lipid profiles in tissue of the Fischer rat. Chem.Biol.Interact. 23: 315-330

55. Schiller CM, Adcock CM, Moore RA, and Walden R. 1985. Effect of 2,3,7,8- tetrachlorodibenzo-p-dioxin (TCDD) and fasting on body weight and lipid parameters in rats. Toxicol.Appl.Pharmacol. 81: 356-361

56. Rifkind AB. 2006. CYP1A in TCDD toxicity and in physiology-with particular reference to CYP dependent arachidonic acid metabolism and other endogenous substrates. Drug Metab Rev. 38: 291-335

57. Dasmahapatra AK, Wimpee BA, Trewin AL, and Hutz RJ. 2001. 2,3,7,8- Tetrachlorodibenzo-p-dioxin increases steady-state estrogen receptor-beta mRNA levels after CYP1A1 and CYP1B1 induction in rat granulosa cells in vitro. Mol.Cell Endocrinol. 182: 39-48

58. Korashy HM and El Kadi AO. 2006. The role of aryl hydrocarbon receptor in the pathogenesis of cardiovascular diseases. Drug Metab Rev. 38: 411-450

59. Ge N-L and Elferink CJ. 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblatoma protein. J.Biol.Chem. 273: 22708-22713

60. Puga A, Barnes SJ, Dalton TP, Chang C, Knudsen ES, and Maier MA. 2000. Aromatic hydrocarbon receptor interaction with the retinoblastoma protein potentiates repression of E2F-dependent transcription and cell cycle arrest. J.Biol.Chem. 275: 2943-2950

61. Tian Y, Ke S, Denison MS, Rabson AB, and Gallo MA. 1999. Ah receptor and NF-kappaB interactions, a potential mechanism for dioxin toxicity. J.Biol.Chem. 274: 510-515

62. Klinge CM, Kaur K, and Swanson HI. 2000. The aryl hydrocarbon receptor interacts with estrogen receptor alpha and orphan receptors COUP-TFI and ERRalpha1. Arch.Biochem.Biophys. 373: 163-174

176

63. Barnes-Ellerbe S, Knudsen KE, and Puga A. 2004. 2,3,7,8-Tetrachlorodibenzo-p-dioxin blocks androgen-dependent cell proliferation of LNCaP cells through modulation of pRB phosphorylation. Mol.Pharmacol. 66: 502-511

64. Klinge CM, Bowers JL, Kulakosky PC, Kamboj KK, and Swanson HI. 1999. The aryl hydrocarbon receptor (AHR)/AHR nuclear translocator (ARNT) heterodimer interacts with naturally occurring estrogen response elements. Mol.Cell Endocrinol. 157: 105-119

65. Safe S, Wormke M, and Samudio I. 2000. Mechanisms of inhibitory aryl hydrocarbon receptor-estrogen receptor crosstalk in human breast cancer cells. J.Mammary.Gland.Biol.Neoplasia. 5: 295-306

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Table 1: 50 AHR gene target hits with the highest signal ratios from DMSO-treated cells.

RefSeq Gene Name Abbreviation Known or Putative Function

NM_026796 SET and MYND domain containing 2 Smyd2 metal ion binding NM_009741 B-cell leukemia/lymphoma 2 Bcl2 regulation of apoptosis XM_130308 Low density lipoprotein receptor-related 2 Lrp2 vesicle-mediated transport NM_146381 Olfactory receptor 1284 Olfr1284 G-protein coupled receptor activity AK035141 12 days embryo embryonic body AK081723 16 days embryo head cDNA NM_146919 Olfactory receptor 1188 Olfr1188 olfactory receptor activity NM_029963 Mitochondrial ribosomal protein S5 Mrps5 structural constituent of ribosome adenosine receptor activity, G- NM_027571 Purinergic Rec P2Y, G-protein coupled 12 P2ry12 protein coupled NM_183222 Fc receptor-like protein 5 Fcrl5 immunecomplex binding receptor component of molecular motors in NM_175138 Dynein, axonemal, intermediate chain 1 Dnaic1 cilium and flagellum TAF6 RNA polymerase II, TATA box BC058583 Taf6 RNA polymerase complex protein binding protein (TBP)-associated factor glomerulosclerosis; peroxisomal NM_008622 Mpv17 transgene, kidney disease mutant Mpv17 protein BC061486 PABP-dependent poly(A) nuclease 3 component of the mammalian NM_026042 Intersex-like (Drosophila) Ixl mediator complex histone binding; biogenesis of Fe/S NM_172746 HIRA interacting protein 3 Hirip3 proteins NM_147080 Olfactory receptor 615 Olfr615 olfactory receptor AK035883 16 days neonate cerebellum cDNA blood pressure regulation; feeding behavior; body size regulation; NM_010934 Neuropeptide Y receptor Y1 Npy1r glucose metabolism; sensory perception NM_144948 RNA binding motif protein 7 Rbm7 nucleic acid binding during meiosis BC086319 RIKEN cDNA 2310075C12 gene AK028335 12 days embryo embryonic body NM_027900 R3H domain containing 2 R3hdm2 unknown mediates inactive to active chromatin NM_177306 Regulatory factor X domain containing Rfxdc1 structure NM_024249 RIKEN cDNA 1810073N04 gene protein tyrosine/serine/threonine NM_019819 Dual specificity phosphatase 14 Dusp14 phosphatase activation of MAPK activity; protein NM_027569 Sperm associated antigen 9 Spag9 homooligomerization BC060715 Fetal Alzheimer antigen Falz brain development NM_147220 ATP-binding cassette transporter A9 Abca9 ATP-binding cassette gene

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Table 1: cont'd

RefSeq Gene Name Abbreviation Known or Putative Function

NM_145684 Arachidonate lipoxygenase, epidermal Alox12e leukotriene biosynthesis pathway prostate, ovary & mammary U81451 Estrogen receptor 2 (beta) Esr2 differentiation; ovulation U81451 Estrogen receptor 2 (beta) Esr2 NM_028127 FERM domain containing 6 Frmd6 cytoskeleton binding protein NM_026825 Leucine rich repeat containing 16 Lrrc16 cell adhesion and cellular trafficking NM_029791 Bicaudal D homolog 2 Bicd2 dynein-dynactin-interacting protein NM_023868 Ryanodine receptor 2, cardiac Ryr2 cardiac-specific calcium channel structural constituent of the NM_012050 Osteomodulin Omd extracellular matrix NM_029791 Bicaudal D homolog 2 Bicd2 dynein-dynactin-interacting protein NM_207652 Transforming growth factor beta 1 induced Tsc22d1 development anti-apoptosis factor associated with NM_007537 Bcl2-like 2 or Bcl-w Bcl2l2 mitochondria regulation of actin-cytoskeletal NM_008304 Syndecan 2 Sdc2 organization NM_183107 RIKEN cDNA 4930474M22 gene AK045711 Adult male corpora quadrigemina cDNA AK019744 Adult male testis cDNA small GTPase signal transduction; NM_181070 RAB18, member RAS oncogene family Rab18 membrane traffic/structure NM_146989 Olfactory receptor 1496 Olfr1496 olfactory receptor gene family nervous system development; axon NM_011976 Semaphorin Sema4g guidance; differentiation NM_146238 cDNA sequence BC023488 TAF9B RNA polymerase II, TATA box DNA-directed DNA polymerase NM_001001176 Taf9b binding protein (TBP)-associated factor activity; nucleotidyltransferase NM_009703 Raf-related oncogene Araf cell cycle regulation

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Table 2: 50 AHR gene target hits with the highest signal ratios from TCDD-treated cells.

RefSeq Gene Name Abbreviation Known or Putative Function

NM_009230 Sterol O-acyltransferase 1 Soat1 cholesterol, lipid, steroid metabolism negative regulation of cell proliferation; NM_010629 Kinesin-associated protein 3 Kifap3 positive regulation of calcium-dependent cell-cell adhesion NM_133664 Ladinin Lad1 basement membrane/ECM component acetyl-CoA/lipid binding; mitochondrial NM_007830 Diazepam binding inhibitor Dbi transport NM_007993 Fibrillin 1 Fbn1 ECM structural constituent NM_025626 RIKEN cDNA 3110001A13 gene NM_175113 RIKEN cDNA 3300001M20 gene NM_007993 Fibrillin 1 Fbn1 ECM structural constituent AK007018 Adult male testis cDNA AK041063 Adult male aorta and vein cDNA NM_175176 RIKEN cDNA 1700013E18 gene NM_013867 Breast cancer anti-estrogen resistance 3 Bcar3 regulation of cell adhesion and motility NM_013755 Glycogenin 1 Gyg1 carbohydrate/glycogen biosynthesis NM_153561 Nudix-type motif 6 Nudt6 hydrolase activity regulates membrane HCN channel NM_021483 Peroxin 2 Pex2 density; brain development cell cycle regulation; RNA polymerase II- NM_207678 Cyclin L2 Ccnl2 associated NM_133948 PC4 and SFRS1 interacting protein 1 Psip1 involved in retinoic acid production integral membrane component; NM_172880 Transmembrane protease, serine 11e Tmprss11e peptidase activity NM_001007220 Disintegrin/metallopeptidase domain 22 Adam22 myelination of peripheral nerves NM_018773 Src family associated phosphoprotein 2 Scap2 negative regulation of cell proliferation AK007017 Adult male testis cDNA BC083337 RNA polymerase II associated factor Paf1 NM_026042 Intersex-like (Drosophila) Ixl mammalian mediator complex protein NM_183036 Defensin β 38 Defb38 defense response to bacteria microtubule cytoskeleton organization NM_207659 hook homolog 3 (Drosophila) Hook3 and biogenesis positive regulation of phagocytosis; TNF- NM_026972 CD209b antigen Cd209b α biosynthesis protein kinase activity; cell cycle NM_011848 NIMA-related expressed kinase 3 Nek3 regulation NM_026972 CD209b antigen Cd209b see above NM_176935 RIKEN cDNA F730015K02 gene interact with Rab GTPases; membrane NM_138303 Yip1 domain family, member 2 Yipf2 traffic regulation

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Table 2: cont'd

RefSeq Gene Name Abbreviation Known or Putative Function protein phosphatase type 1 regulator NM_028755 Cyclic AMP-regulated phosphoprot., 21 Arpp21 activity NM_015781 Nucleosome assembly protein 1-like 1 Nap1l1 nucleosome assembly NM_015781 Nucleosome assembly protein 1-like 1 Nap1l1 nucleosome assembly NM_146933 Olfactory receptor 790 Olfr790 olfactory receptor activity NM_177614 RIKEN cDNA 4632413K17 gene NM_181577 RIKEN cDNA E030025D05 gene NM_008796 Phosphatidylcholine transfer protein Pctp lipid binding and transport negative regulation of cell cycle; organ NM_009368 Transforming growth factor, beta 3 Tgfb3 morphogenesis NM_198093 Engulfment and cell motility 1, ced-12 Elmo1 phagocytosis and cell migration transcriptional regulation in retinal NM_175006 POU domain, class 6, transcript. factor 2 Pou6f2 ganglion development integral ER membrane component; NM_015774 ERO1-like (S. cerevisiae) Ero1l redox state regulation NM_130885 Oxidation resistance 1 Oxr1 protection from oxidative stress AK054425 2 days pregnant adult female ovary cDNA diacylglycerol binding; protein kinase C NM_138650 Diacylglycerol kinase, gamma Dgkg activation NM_198418 Tudor domain containing 6 Tdrd6 arginine methylation NM_130868 Olfactory receptor 138 Olfr138 olfactory receptor activity BC016576 Isochorismatase domain containing 1 Doublesex and mab-3 related transcription transcription factor; development; sex NM_015826 Dmrt1 factor 1 differentiation NM_019405 Centrin 2 Cetn2 mitosis NM_010941 NAD(P)-dep. steroid dehydrogenase-like Nsdhl cholesterol, lipid, steroid biosynthesis

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Table 3: Number of AHR and E2F binding sites of the annotated genes in the top 50 AHR gene target hits from DMSO-treated cells. Proximal sites are within 300 bp of the transcription start site (as identified by the Ensembl genome browser), whereas distal sites are between 300 and 5000 bases upstream of the transcription start site.

Gene Proximal AHR Distal AHR Proximal E2F Distal E2F Smyd2 150 4 Bcl2 0 5 0 3 Lrp2 1 21 3 Olfr1284 000 1 Olfr1188 000 3 Mrps5 0 2 1 5 P2ry12 042 0 Fcrl5 000 3 Dnaic1 0 8 1 2 Taf6 1 3 2 3 Mpv17 011 1 Ixl 0 3 0 1 Hirip3 1 5 0 3 Olfr615 0 2 0 0 Npy1r 0 2 0 1 Rbm7 2 1 1 2 R3hdm2 0 0 0 2 Rfxdc1 1 2 0 2 Dusp14 1 3 1 5 Spag9 1 1 0 5 Falz 0 3 2 4 Abca9 0 3 0 6 Alox12e 1 5 0 4 Esr2 2 3 0 4 Frmd6 3 2 1 3 Lrrc16 0 2 0 5 Bicd2 0 4 1 3 Ryr2 0 3 0 2 Omd 0 0 0 1 Bicd2 0 4 1 3 Tsc22d1 1 2 4 3 Bcl2l2 2 1 0 0 Sdc2 0 1 0 1 Rab18 1 1 1 5 Olfr1496 0 2 0 3 Sema4g 2 1 0 1 Taf9b 1 7 0 5 Araf 1 2 1 1

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Table 4: Number of AHR and E2F binding sites of the annotated genes in the top 50 AHR gene target hits from TCDD-treated cells. Proximal sites are within 300 bp of the transcription start site (as identified by the Ensembl genome browser), whereas distal sites are between 300 and 5000 bases upstream of the transcription start site.

Gene Name Proximal AHR Distal AHR Proximal E2F Distal E2F Soat1 210 1 Kifap3 1 5 3 2 Lad1 2 10 5 Dbi 362 4 Fbn1 021 5 Bcar3 1 0 0 4 Gyg1 010 4 Nudt6 431 6 Pex2 0 1 0 3 Ccnl2 0 2 0 4 Psip1 210 2 Tmprss11e 0 0 0 1 Adam22 1 4 0 3 Scap2 2 1 0 2 Paf1 0 4 0 4 Ixl 0 3 0 1 Defb38 0 1 0 1 Hook3 3 3 1 3 Cd209b 0 0 0 3 Nek3 2 1 2 5 Cd209b 0 0 0 3 Yipf2 1 11 0 3 Arpp21 1 4 0 3 Nap1l1 0 2 0 3 Olfr790 0 1 0 2 Pctp 0 2 0 6 Tgfb3 0 4 0 2 Elmo1 2 0 3 2 Pou6f2 0 0 0 3 Ero1l 3 2 0 0 Oxr1 0 1 0 4 Dgkg 1 2 0 1 Tdrd6 1 0 1 3 Olfr138 0 3 0 3 Dmrt1 3 5 3 5 Cetn2 1 2 1 3 Nsdhl 2 2 0 1

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Table 5: Comparison of AHR target genes identified using ChIP-on-chip with available expression array data.

ChIP-on-chip Treatment ID Expression Array Expression Array Result Reference

DMSO TCDD Bcl-2 XBcl-2▼ TCDD-exposed female rats Vezina et al. 2004 Lrp2 XLrp2▲ liver of TCDD-treated mice Boverhof et al. 2005 Npy1r XNpy1r▼ TCDD-treated HepG2 cells Puga et al. 2000 Dnaic1 XDnalc4▼ c1 cells compared to Hepa-1c1c7 Fong et al. 2005 Dusp1 ▼ TCDD-treated HPL1A, A549 Martinez et al. 2002 Dusp1 ▼ TCDD in EGF-treated 10T1/2 cells Hanlon et al. 2005a Dusp14 X Dusp 4 ▲ dioxin-exposed humans McHale et al. 2006 Dusp5 ▼ TCDD-treated A549 cells Martinez et al. 2002 Spag9 XSpag9▼ c1 cells compared to Hepa-1c1c7 Fong et al. 2005 Alox12b ▼ TCDD-treated Hepa-1c1c7 cells Jin et al. 2004 Alox12e X Alox12b ▲ paw of TCDD-treated mice Bemis et al. 2006 Lrrc16 X Lrrc2 ▲ TCDD-treated Hepa-1c1c7 cells Jin et al. 2004 Bcl2l11 ▲ liver of TCDD-treated mice Boverhof et al. 2005 Bcl2l2 X Bcl2-w ▲ thymus of TCDD-treated mice Fisher et al. 2004 Sdc1 ▼ TCDD during adipocyte differentiation Hanlon et al. 2005a Sdc2 X Sdc1 ▼ c1 cells compared to Hepa-1c1c7 Fong et al. 2005 Syndecan Altered >2X by TCDD in rat liver Fletcher et al. 2005 Sema4g X Semaphorin ▼ TCDD-treated HepG2 cells Puga et al. 2000 Tsc22d1 ▲ c1 cells compared to Hepa-1c1c7 Fong et al. 2005 Tsc22d1 X Tsc22d1 Altered >2X by TCDD in rat liver Fletcher et al. 2005 TGFβ-inducible gene ▼ TCDD in EGF-treated 10T1/2 cells Hanlon et al. 2005a Rab18 X Rab11a ▲ dioxin-exposed humans McHale et al. 2006 Araf XAraf▲ c1 cells compared to Hepa-1c1c7 Fong et al. 2005 Kif2A ▼ c1 cells compared to Hepa-1c1c7 Fong et al. 2005 Kifap3 X Kif20A ▲ c4 cells compared to Hepa-1c1c7 Fong et al. 2005 Fbn1 XFbn2 ▲ TCDD-treated 10T1/2 cells Hanlon et al. 2005b Nudt1 ▲ TCDD-treated HPL1A cells Martinez et al. 2002 Nudt6 X Nudt4 ▼ livers of TCDD-treated rats Boverhof et al. 2006 Scap2 X Scap2 ▲ livers of TCDD-treated rat & mouse Boverhof et al. 2006 Nek2 ▼ TCDD-treated HepG2 cells Puga et al. 2000 Nek3 X Nek4 ▼ dioxin-exposed humans McHale et al. 2006 Tgfb3 X Tgf-b3 ▲ AHR-/- cells Karyala et al. 2004 Ero1l X Ero-1 ▼ TCDD during adipocyte differentiation Hanlon et al. 2005a Dgka ▼ livers of TCDD-treated rats Boverhof et al. 2006 Dgkg X Dgkz Altered >2X by TCDD in rat liver Fletcher et al. 2005

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Chapter V Figure Legends

Fig. 1. Basic components of the ChIP-on-chip methodology. Hepa-1c1c7 cells were treated

with 10nM TCDD or the equivalent volume of Me2SO vehicle for one hour. Cells were then

exposed to 1.0% formaldehyde in order to crosslink protein complexes to their DNA-binding

sites. Following cellular lysis, sonication was used to shear the DNA to an average length of

~500bp. Specific antibodies are used to immunoprecipitate proteins and their associated DNA.

For our experiments, anti-AHR (Biomol) was used to IP Me2SO or TCDD samples, and anti- rabbit IgG (on Me2SO-treated cells) was used as a negative control. Protein A beads were used to capture antibody-protein-DNA complexes. Heat-reversal of crosslinks was followed by DNA purification and PCR amplification with promoter-specific primer sets for confirmation of the presence of AHR targets. For each sample (IgG Control, Me2SO and TCDD), LM-PCR was

performed to amplify the immunoprecipitated DNA. In addition, 20 ng of total DNA was

similarly amplified. Total DNA was labeled (green) and competitively hybridized against the

amplified and labeled (red) ChIP DNA using a promoter tiling array from Nimblegen Systems,

Inc. DNA sequences that were bound by the protein of interest (AHR) are proportionately greater

in the ChIP samples compared to the total DNA samples.

Fig. 2. Verification of AHR target enrichment in ChIP samples. A. PCR analysis of samples

immunoprecipitated with anti-IgG (IgG, cell treatment is DMSO) or anti-AHR (DMSO and

TCDD) antibodies. Purified DNA samples were suspended in water and subjected to PCR using primers specific for the AHR-binding regions of the Cyp1a1 enhancer and Nqo1 promoter. Input samples represent 0.2% of total DNA from each individual immunoprecipitation. Replicates from each sample were combined for subsequent amplification reactions (Fig. 3). B. Real-time

185

quantitative PCR analysis of ChIP samples for the presence of the Cyp1a1 and Nqo1 promoter,

as outlined in A. Real-time PCR was performed using Brilliant SYBR Green QRT-PCR Master

Mix (Stratagene) and a Smart Cycler rapid thermal cycler (Cepheid). Fluorescence was measured

after each of the repetitive cycles. Cycle threshold (Ct) values are indicated on the ordinate. A

higher Ct value is indicative of less copies of a particular piece of DNA in that sample as compared to a sample showing a lower Ct value. Hence, a true target of AHR binding such as

Cyp1a1 should show a lower Ct value in samples precipitated with an anti-AHR antibody as

compared to IgG control, as well as in samples where the AHR has been activated by TCDD. C.

Real-time quantitative PCR results normalized to total input DNA each respective ChIP sample.

Ct values were manipulated as outlined in the Materials and Methods. Data are shown as normalized values relative to the IgG control, or as the fold-difference over IgG control. DMSO, vehicle control.

Fig. 3. Verification of AHR target enrichment in LM-PCR-amplified ChIP samples. A.

Real-time quantitative PCR analysis of LM-PCR amplified ChIP samples for the presence of the

CYP1a1 promoter. Purified DNA isolated from immunoprecipitations as outlined for Fig. 2 was subjected to a ligation-mediated PCR-based amplification method. Replicate ChIP samples (see

Fig. 2A) were combined for each sample in order to carry out LM-PCR. Twenty nanograms of total DNA were similarly amplified. All amplified samples were purified and quantified for subsequent PCR analysis. Real-time PCR was performed using Brilliant SYBR Green QRT-PCR

Master Mix (Stratagene) and a Smart Cycler rapid thermal cycler (Cepheid). Fluorescence was measured after each of the repetitive cycles. Cycle threshold (Ct) values for the Cyp1a1 and

Nqo1 promoters are indicated on the ordinate. B. Real-time quantitative PCR results normalized

to total input DNA each respective ChIP sample. Ct values were manipulated as outlined in the

186

Materials and Methods. Data are shown as normalized values relative to the IgG control, or as

the fold-difference over IgG control. C. Confirmation of size range of amplified ChIP samples.

Total DNA samples and DNA isolated by ChIP was amplified by LM-PCR, and 600 ug of each

sample was run on a DNA 7500 LabChip kit (Agilent) and analyzed using the Agilent 2100

bioanalyzer. DMSO, vehicle control.

Fig. 4. Map of the top 20 genes hits found on Chromosome 2 by Chip-on-chip. Amplified and quantified ChIP samples (Fig. 3) were sent to Nimblegen Systems, Inc. for labeling and hybridization to commercially available promoter tiling arrays. Data was extracted and analyzed at Nimblegen, and visualized using the SignalMap software package. SignalMap displays the sample hits in relation to the known state of sequence annotation, including genes, exons, and repetitive elements. Shown is a representative set of the highest 20 hybridization intensity signals from Chromosome 2 of the mouse genome, generated using the SignalMap software. At the top are shown approximate numerical position along the chromosome, and below are indicated the relative intensity signals (“score”) of the gene hits on that chromosome. One can immediately see the completely different pattern of signals generated by the Me2SO-treated (DMSO control) samples as compared to the TCDD-treated samples. Indicated are the gene identifications of the highest single hit for each treatment, details for which can be found in Tables 1 and 2.

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Figure 1

Step 1. Use formaldehyde to crosslink proteins to DNA, and isolate nuclei

Step 2. Sonicate nuclei to break DNA into short (~500bp) segments

Step 3. Immunoprecipitate protein-DNA Step 4. Formaldehyde crosslinking is complexes to enrich specific sequences reversed to isolate DNA from proteins

Step 5. ChIP-enriched DNA is purified, amplified by LM-PCR, and dyed with Cy5 (red). Total (input) DNA is also amplified by LM-PCR and dyed with Cy3 (green).

Step 6. Both IP-enriched and un-enriched DNA samples are hybridized to the same oligonucleotide array.

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Figure 2

A. IgG DMSO TCDD Input Cyp1a1 ChIP promoter

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Figure 3

G O D G O D g g I D I D MS C MS C D T D T Total DNA ChIP DNA

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Figure 4

7 7 7 7 7 0 0 0 0 1 10 1 1 1 x x x x x 2 5 0 3 6 9 1 1 chr2 Lrp2 375

0 DMSO Fbn1 423 0 TCDD

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Chapter VI

Conclusions

The exposure of human populations to AHR ligands has been associated with a number of toxicological outcomes and diseases, including cancer, chloracne, cardiovascular disease, diabetes, endometriosis, neurocognitive deficits, and developmental abnormalities (57;62).

While prevailing scientific evidence suggests that these outcomes are mediated primarily by aryl hydrocarbon receptor activation, the exact molecular mechanisms by which AHR ligands exert their effects are still ambiguous. It is clear, however, that AHR signaling impinges upon numerous molecular pathways both in its physiological state and in its ligand-activated form.

The relative importance of these pathways in mediating AHR-dependent toxicities is still being determined. Mapping these pathways in various systems for different ligands, cell types, and exposure conditions will aid in predicting safe exposure levels and identifying susceptible human populations. To this end, this dissertation attempts to shed some light on the complexity of AHR signaling, and to define specific mechanisms by which the AHR acts to perturb normal cellular homeostasis when it is activated by environmental toxicants. It is apparent from the work presented here that the molecular pathways in which the AHR interacts are dependent upon the particular cellular environment being examined. In other words, only some of the pathways upon which the AHR acts will be affected some of the time, depending at least in part on the particular preferences of the AHR for protein interactions and/or DNA binding, as well as the ultimate effect of these interactions on transcription.

One of the major effects of AHR activation is the perturbation of normal cell cycle progression. Activation of the AHR can result in the inhibition or progression of cell division, differentiation, or apoptosis. Again, the specific outcomes for a particular ligand depend upon

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cellular context. For example, AHR activation by TCDD activates apoptosis in thymocytes

(140), but inhibits apoptosis in transformed liver cells (141); it enhances cell proliferation in keratinocytes (142), but inhibits proliferation in hepatocytes (143); it inhibits differentiation of adipocytes (144), but accelerates differentiation in the skin (145). It is interesting to consider the

notion that a potent tumor promoter could both induce apoptosis and inhibit cell proliferation under some conditions. Both of these effects are contrary to the ultimate effect of a carcinogen, which is to enhance overall cell proliferation, resulting in tumor formation (146). The immediate response of cells to a carcinogen such as TCDD is likely an adaptive response to the presence of a foreign chemical, which causes such secondary effects as the generation of reactive oxygen species and general cellular damage. Thus, the cell is responding to an adverse environment by ceasing to proliferate until such cellular damage is cleared. Ultimately, the persistence of the carcinogen in inducing oxidative and genotoxic stress and cellular damage will overwhelm cell defense systems, and aberrant cell cycle control may ensue.

The AHR has been found to interact with and modulate the expression of literally

hundreds of factors. It is thus not surprising that its effects would vary greatly depending on the

cell type, and hence on the particular factors with which the AHR is able to interact at any given

time. One of the major themes of this dissertation is the interaction of the AHR with the major

cell cycle regulatory proteins RB and E2F (summarized in Figure 1). The interaction of the

TCDD-activated AHR with RB in particular appears to mediate cell cycle arrest in hepatocyte-

derived cell lines (34;114;147;148). Given that the RB protein is ubiquitously expressed in most

if not all cell types, it would seem logical to conclude that the AHR would mediate cell cycle

arrest in any cell type treated with TCDD. This is clearly not the case. While the AHR-RB

interaction may mediate important toxicological outcomes in some cell types, it is not the major

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Hepatocyte-derived cells A. Others? Cyclin E Mitogens P P P

RB Cyclin D/cdk E2F1 Cell cycle Cyclin E progression Pol α AHR Cdk2 RB AHR activation DHFR E2F1 Cyclin A

MCM7 G1/S arrest (?) AHR E2F1

Fibroblast-derived cells B. Others? AHR RB Dnmt1 (?) Cdt1 G /S arrest E2F1 Mcm3 1 Mitogens Cdca7

Cyclin D/cdk E2F1 Dnmt1 Cdt1 Cell cycle (?) Mcm3 progression R ARNT E2F1 Cdca7 AH

AHR AHR activation E2F1

Fig. 1. Multiple models for the interplay of AHR in cell cycle pathways. A. For the predominately hepatocyte-derived cell lines examined here, in quiescent or in normal cycling cells, RB/E2F interactions downregulate S-phase genes and mitogens activate cell cycle progression; under these conditions, activation of aryl hydrocarbon receptor (AHR) causes its translocation to the nucleus where it functions as an environmental checkpoint in cooperation with RB/E2F, inhibiting cyclin D,E/cdk-dependent RB phosphorylation, promoting repression of S-phase specific genes and causing cell cycle arrest. Evidence also suggest that the AHR may act directly on E2F to inhibit its normal function as a transcriptional regulator. B. In fibroblast cells, induction of the AHR is not sufficient to induce cell cycle arrest, although it is still able to interact with RB and E2F, and to cause the downregulation of a number of E2F-regulated genes. The mechanism of this downregulation may involve AHR/RB, AHR/E2F, or both. AHR is also required for constitutive expression of a number of E2F-regulated genes, and hence normal cell cycle progression, seemingly through direct binding of AHR to promoter sequences even when the AHR is not activated by TCDD.

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determinant of dioxin effects in all cell types. Furthermore, although the interaction of the AHR

with E2F family proteins may explain additional aspects of dioxin toxicity unrelated to the RB protein, the effect of AHR activation on E2F-dependent gene transcription is not universal for all

E2F-regulated genes.

At the most fundamental level, the toxicological effects mediated by the AHR appear to be the result of gene expression perturbations. The results presented in this dissertation illustrate the effect of the AHR on a variety of different genes by a number of different mechanisms (Fig.

1). The AHR is primarily viewed as a transcription factor that activates gene expression through its binding to cognate sequences in specific gene promoters. We now know that this is only one mechanism by which the AHR exerts its influence on gene transcription. The first part of this

dissertation focuses on an interaction of the AHR with the RB protein. One of the primary

functions of the RB protein is to prevent the expression of genes required for cell cycle

progression through inhibiting the transactivation function of E2F proteins. The AHR appears to

cooperate with RB in this function. It maintains RB in a hypophosphorylated state, inhibits the

release of RB from E2F proteins, and prevents the recruitment of coactivators to the promoters of

several genes involved in mediating S-phase progression. However, with respect to AHR-

mediated cell cycle perturbations, there are multiple mechanisms at work. In addition to

inhibiting the expression of E2F-regulated genes, the AHR appears to upregulate the expression

of the p27Kip1 cyclin dependent kinase inhibitor in a manner dependent upon transactivation by the AHR (149;150). It also appears that ARNT plays some as-yet uncharacterized role in AHR- mediated cell cycle arrest (150), although the data presented in Chapter 3 suggests that at least at the level of gene expression, AHR inhibits transcription even in the absence of ARNT.

Furthermore, RB is but one protein member of a family of proteins that includes the p107 and

195

p130 proteins. The studies presented here do not address the potential role of these proteins in

mediating the effects of AHR on the cell cycle. It is known that all three RB family proteins

interact with many of the E2F family proteins through conserved pocket domains (151), and in

the case of RB, this same domain mediates interaction with the AHR (128). Although the results

presented in Chapter 4 show unequivocally that the AHR and E2F form protein-protein complexes, and that the AHR affects E2F-dependent gene expression in the absence of RB, it may be that the observed effects are actually the result of p107 and p130 expression, and an interaction between the AHR and these proteins. Moreover, very little is known of the stoichiometric relationships among any of these proteins, and the relationship of the AHR to RB or E2F may be more or less important in different cell types as a result. Finally, it may be as well that the effect of the AHR on inhibiting cell cycle progression through RB is the result of its sustained activation by persistent ligands, and would not occur if AHR were transiently activated by a more metabolizable ligand (152). Valuable insight into AHR-dependent toxicity may result from the examination of these protein interactions under a number of different cellular contexts and with a number of different ligands.

The second part of this dissertation focuses on an interaction of the AHR with the E2F protein that is independent of RB binding. While this interaction certainly plays a role in the mechanisms by which the AHR is able to modify or perturb expression of specific genes, it is uncertain as to the significance of this interaction under different contexts. The results presented in Chapter 4 indicate that the AHR has a physiological role in maintaining normal expression levels of a large number of E2F-regulated, cell cycle-associated genes in MEF cells. Several interesting and complicating factors are worth noting in this regard. First, these same genes are not effected by AHR expression in Hepa-1 cells, as there was no difference in expression of

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these genes observed between the Hepa-1, c2, c4, and c35 cell lines (data not shown). The c2, c4, and c35 cell lines have reduced AHR expression levels, lack ARNT expression, and have a defect in AHR-dependent DNA binding, respectively, compared with the parent Hepa-1 cell line

(153;154). Notably, the effect on E2F-dependent gene expression in MEF cells was observed only in the complete absence of AHR expression. Clearly, none of the Hepa-1 derived cells referenced above exactly mimics the AHR status of the MEF cells examined here. Secondly, the effects of TCDD on gene expression levels varied depending on the gene examined, with about half of the genes decreasing in expression in response to TCDD treatment, and about half showing no effect at all. Also, two of the genes examined actually increased significantly in the absence of AHR expression, in contrast to the majority of other genes analyzed. However, in all cases, the pattern of gene expression in AHR-deficient cells mirrored that in E2F-deficient cells, indicating a potential co-regulatory mechanism of AHR and E2F for these genes. Finally, while

MEF cells are relatively refractory to cell cycle perturbations by TCDD treatment (data not shown), lack of AHR expression results in a significant inhibition of cell proliferation in this cell type. While this effect has been attributed to an increase of TGF-β proteins (55;155), it is also possible that the constitutive maintenance by the AHR of the expression of genes involved in cell cycle progression is a major determinate of proliferation in MEF cells, but likely not in all cells.

ChIP experiments indicate that the AHR constitutively binds to the promoters of at least a fraction of these important cell cycle genes, and each one has multiple canonical AHR binding sequences within 5000 bp of the transcription start site, implying a mechanism by which the

AHR could modulate transcription. However, the AHR-E2F interaction complicates this picture, and implies an alternative mechanism for AHR-dependent modulation of these genes through disruption of normal E2F-dependent transactivation. This appears to be the case for some of the

197

genes examined, but not all, and activation of the AHR by TCDD clearly targets additional receptor to certain promoter regions, as determined by promoter tiling arrays. Additional studies will be required in order to tease out the relative importance of these two opposing mechanisms.

It is clear from the aforementioned-results that the AHR modulates the expression of a number of genes in a manner depend at least in part on protein-protein interactions rather than direct DNA binding, and that the receptor can in fact repress gene expression in addition to being a transcriptional activator. These alternative regulatory mechanisms may apply broadly to the hundreds of genes which are modified following AHR activation by TCDD and other ligands. A large number of important cell cycle genes identified in Chapters 3 and 4 appear to be direct targets of the AHR, and these targets would not necessarily be identified simply by scanning promoters for AHR binding sites. Chapter 5 explores a technique for the unbiased detection of

AHR binding sites, namely ChIP-on-chip. While this technique has been used to identify novel promoter targets of a number of transcription factors, it has not been used for the AHR. The results of the promoter array presented here indicate several things. First, the potential AHR targets identified are genes residing in a number of pathways known to be modulated by AHR activity. A large percentage of the genes identified in the promoter array have also been identified as being affected by either TCDD treatment or by the AHR itself in cDNA expression microarrays. This is significant in that for most of these genes, a direct connection to AHR regulation at the transcriptional level has never been demonstrated. Finally, the promoter arrays indicate that the AHR preferentially targets different genes under physiological conditions as it does under toxicological conditions. This is significant in two ways. First, while it is assumed that the AHR is activated by endogenous ligands, and it is known that the AHR performs a variety of basal functions in normal cells, this is the first demonstration that the AHR actually

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binds to many promoter regions when it is not activated by exogenous ligands. In other words, it is likely that the primary physiological role of the AHR is to function through its constitutive activities as a transcription modulator for the expression of a large variety of genes. Second, it could be that the toxicity of AHR ligands is explained at least in part by a shift in the genes primarily targeted by the AHR in normal cells to a completely different set of primary targets in

TCDD-treated cells. Hence, the expression of genes that are induced or suppressed by AHR activity would be reversed in TCDD treated cells. The particular set of AHR target genes in specific cell types would thus impinge upon the AHR-mediated toxicity observed in that cell type.

This thesis implicates the AHR in a number of new pathways and transcriptional control mechanisms. The results presented throughout this work show the diversity of AHR functions related both to toxicological endpoints and normal cell physiology, and illustrate the ability of this important transcription factor to regulate the expression of a large number of genes by a variety of distinct mechanisms. Far from resolving any of the complexities surrounding AHR functions or the toxicities of its ligands, this thesis has illustrated the immense intricacies of this protein, and just how far we are from truly understanding its role in biology and toxicology.

Hopefully, this work also illustrates the necessity for specifically defining AHR interactions and targets in a cell-type specific manner, as it is not possible to generalize many if not all AHR- mediated effects among different systems. As functional genomic and proteomic techniques for dissecting molecular networks in individual systems improve, a more clear picture will emerge of the pathways, transcription factors, and genes effected by AHR signaling. Ultimately, this will improve our understanding of the impact and threat of environmental exposures on human and ecological systems.

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References for Introduction and Discussion

1. Rushmore TH and Kong AN. 2002. Pharmacogenomics, regulation and signaling pathways of phase I and II drug metabolizing enzymes. Curr.Drug Metab 3: 481-490

2. Nebert DW, Goujon FM, and Gielen JE. 1972. Aryl hydrocarbon hydroxylase induction by polycyclic hydrocarbons: simple autosomal dominant trait in the mouse. Nature New Biol. 236: 107-110

3. Poland A, Glover E, Robinson JR, and Nebert DW. 1974. Genetic expression of aryl hydrocarbon hydroxylase activity. Induction of monooxigenase activities and cytochrome P1-450 formation by 2,3,7,8-tetrachlorodibenzo-p-dioxin in mice genetically "non- responsive" to other aromatic hydrocarbons. J.Biol.Chem. 249: 5599-5606

4. Poland A, Glover E, and Kende AS. 1976. Stereospecific, high affinity binding of 2,3,7,8- tetrachlorodibenzo-p-dioxin by hepatic cytosol. Evidence that the binding species is receptor for induction of aryl hydrocarbon hydroxylase. J.Biol.Chem. 251: 4936-4946

5. Okey AB, Bondy GP, Mason ME, Kahl GF, Eisen HJ et al. 1979. Regulatory gene product of the Ah locus. Characterization of the cytosolic inducer-receptor complex and evidence for its nuclear translocation. J.Biol.Chem. 254: 11636-11648

6. Poland A and Knutson JC. 1982. 2,3,7,8,-tetrachlorodibenzo-p-dioxin and related halogenated aromatic hydrocarbons: examination of the mechanisms of toxicity. Annu.Rev.Pharmacol.Toxicol. 22: 517-554

7. Walisser JA, Bunger MK, Glover E, and Bradfield CA. 2004. Gestational exposure of Ahr and Arnt hypomorphs to dioxin rescues vascular development. Proc.Natl.Acad.Sci.U.S.A 101: 16677-16682

8. Walisser JA, Glover E, Pande K, Liss AL, and Bradfield CA. 2005. Aryl hydrocarbon receptor-dependent liver development and hepatotoxicity are mediated by different cell types. Proc.Natl.Acad.Sci.U.S.A 102: 17858-17863

9. Abbott BD, Birnbaum LS, and Perdew GH. 1995. Developmental expression of two members of a new class of transcription factors: I. Expression of aryl hydrocarbon receptor in the C57BL/6N mouse embryo. Dev.Dynamics 204: 133-143

10. Puga A, Tomlinson CR, and Xia Y. 2005. Ah receptor signals cross-talk with multiple developmental pathways. Biochem.Pharmacol. 69: 199-207

11. Gu YZ, Hogenesch JB, and Bradfield CA. 2000. The PAS superfamily: sensors of environmental and developmental signals. Annu.Rev.Pharmacol.Toxicol. 40: 519-561

12. Thackaberry EA, Gabaldon DM, Walker MK, and Smith SM. 2002. Aryl hydrocarbon receptor null mice develop cardiac hypertrophy and increased hypoxia-inducible factor- 1alpha in the absence of cardiac hypoxia. Cardiovasc.Toxicol. 2: 263-274

200

13. Baba T, Mimura J, Nakamura N, Harada N, Yamamoto M et al. 2005. Intrinsic function of the aryl hydrocarbon (dioxin) receptor as a key factor in female reproduction. Mol.Cell Biol. 25: 10040-10051

14. Hillegass JM, Murphy KA, Villano CM, and White LA. 2006. The impact of aryl hydrocarbon receptor signaling on matrix metabolism: implications for development and disease. Biol.Chem. 387: 1159-1173

15. Marlowe JL and Puga A. 2005. Aryl hydrocarbon receptor, cell cycle regulation, toxicity, and tumorigenesis. J.Cell Biochem. 96: 1174-1184

16. Kewley RJ, Whitelaw ML, and Chapman-Smith A. 2004. The mammalian basic helix- loop-helix/PAS family of transcriptional regulators. Int.J.Biochem.Cell Biol. 36: 189-204

17. Petrulis JR and Perdew GH. 2002. The role of chaperone proteins in the aryl hydrocarbon receptor core complex. Chem.Biol.Interact. 141: 25-40

18. Denison MS and Heath-Pagliuso S. 1998. The Ah receptor: a regulator of the biochemical and toxicological actions of structurally diverse chemicals. Bull.Environ.Contam Toxicol. 61: 557-568

19. Safe S. 1990. Polychlorinated biphenyls (PCBs), dibenzo-p-dioxins (PCDDs), dibenzofurans (PCDFs), and related compounds: environmental and mechanistic considerations which support the development of toxic equivalency factors (TEFs). Crit Rev.Toxicol. 21: 51-88

20. Denison MS and Nagy SR. 2003. Activation of the aryl hydrocarbon receptor by structurally diverse exogenous and endogenous chemicals. Annu.Rev.Pharmacol.Toxicol. 43: 309-334

21. U.S.Department of Health and Human Services. 2005. CERCLA Priority List of Hazardous Substances. Agency for Toxic Substances and Disease Registry, Atlanta, GA.

22. Faroon OM, Keith S, Jones D, and de Rosa C. 2001. Carcinogenic effects of polychlorinated biphenyls. Toxicol.Ind.Health 17: 41-62

23. Faroon OM, Keith S, Jones D, and de Rosa C. 2001. Effects of polychlorinated biphenyls on development and reproduction. Toxicol.Ind.Health 17: 63-93

24. Golden R, Doull J, Waddell W, and Mandel J. 2003. Potential human cancer risks from exposure to PCBs: a tale of two evaluations. Crit Rev.Toxicol. 33: 543-580

25. Ross G. 2004. The public health implications of polychlorinated biphenyls (PCBs) in the environment. Ecotoxicol.Environ.Saf 59: 275-291

26. Bostrom CE, Gerde P, Hanberg A, Jernstrom B, Johansson C et al. 2002. Cancer risk assessment, indicators, and guidelines for polycyclic aromatic hydrocarbons in the ambient air. Environ.Health Perspect. 110 Suppl 3: 451-488

201

27. Nebert DW, Eisen HJ, and Hankinson O. 1984. The Ah receptor: binding specificity only for foreign chemicals?. [Review]. Biochem.Pharmacol. 33: 917-924

28. Fernandez-Salguero P, Pineau T, Hilbert DM, McPhail T, Lee SS et al. 1995. Immune system impairment and hepatic fibrosis in mice lacking the dioxin-binding Ah receptor. Science 268: 722-726

29. Gonzalez FJ, Fernandez-Salguero P, Lee SS, Pineau T, and Ward JM. 1995. Xenobiotic receptor knockout mice. [Review]. Toxicology Letters 82-83: 117-121

30. Schmidt JV, Su GH-T, Reddy JK, Simon MC, and Bradfield CA. 1996. Characterization of a murine Ahr null allele: involvement of the Ah receptor in hepatic growth and development. Proc.Natl.Acad.Sci.USA 93: 6731-6736

31. Singh SS, Hord NG, and Perdew GH. 1996. Characterization of the activated form of the aryl hydrocarbon receptor in the nucleus of HeLa cells in the absence of exogenous ligand. Archives of Biochemistry & Biophysics 329: 47-55

32. Chang C-Y and Puga A. 1998. Constitutive activation of the aromatic hydrocarbon receptor. Mol.Cell.Biol. 18: 525-535

33. Ma Q and Whitlock JPJ. 1996. The aromatic hydrocarbon receptor modulates the Hepa 1c1c7 cell cycle and differentiated state independently of dioxin. Mol.Cell Biol. 16: 2144- 2150

34. Weiss C, Kolluri SK, Kiefer F, and Gottlicher M. 1996. Complementation of Ah receptor deficiency in hepatoma cells: negative feedback regulation and cell cycle control by the Ah receptor. Exp.Cell Res. 226: 154-163

35. Murray IA, Reen RK, Leathery N, Ramadoss P, Bonati L et al. 2005. Evidence that ligand binding is a key determinant of Ah receptor-mediated transcriptional activity. Arch.Biochem.Biophys. 442: 59-71

36. Rannug A, Rannug U, Rosenkranz HS, Winqvist L, Westerholm R et al. 1987. Certain photooxidized derivatives of tryptophan bind with very high affinity to the Ah receptor and are likely to be endogenous signal substances. J.Biol.Chem. 262: 15422-15427

37. Helferich WG and Denison MS. 1991. Ultraviolet photoproducts of tryptophan can act as dioxin agonists. Mol.Pharmacol. 40: 674-678

38. Paine AJ and Francis JE. 1980. The induction of benzo[a]pyrene-3-mono-oxygenase by singlet oxygen in liver cell culture is mediated by oxidation products of histidine. Chem.Biol.Interact. 30: 343-353

39. Adachi J, Mori Y, Matsui S, Takigami H, Fujino J et al. 2001. Indirubin and indigo are potent aryl hydrocarbon receptor ligands present in human urine. J.Biol.Chem. 276: 31475-31478

202

40. Seidel SD, Winters GM, Rogers WJ, Ziccardi MH, Li V et al. 2001. Activation of the Ah receptor signaling pathway by prostaglandins. J.Biochem.Mol.Toxicol. 15: 187-196

41. Phelan D, Winter GM, Rogers WJ, Lam JC, and Denison MS. 1998. Activation of the Ah receptor signal transduction pathway by bilirubin and biliverdin. Arch.Biochem.Biophys. 357: 155-163

42. Denison MS, Pandini A, Nagy SR, Baldwin EP, and Bonati L. 2002. Ligand binding and activation of the Ah receptor. Chem.Biol.Interact. 141: 3-24

43. Yannai S, Day AJ, Williamson G, and Rhodes MJ. 1998. Characterization of flavonoids as monofunctional or bifunctional inducers of quinone reductase in murine hepatoma cell lines. Food Chem.Toxicol. 36: 623-630

44. Ashida H. 2000. Suppressive effects of flavonoids on dioxin toxicity. Biofactors 12: 201- 206

45. Gradelet S, Astorg P, Leclerc J, Chevalier J, Vernevaut MF, and Siess MH. 1996. Effects of canthaxanthin, astaxanthin, lycopene and lutein on liver xenobiotic-metabolizing enzymes in the rat. Xenobiotica 26: 49-63

46. Bjeldanes LF, Kim JY, Grose KR, Bartholomew JC, and Bradfield CA. 1991. Aromatic hydrocarbon responsiveness-receptor agonists generated from indole-3-carbinol in vitro and in vivo: comparisons with 2,3, 7,8-tetrachlorodibenzo-p-dioxin. Proc.Natl.Acad.Sci.USA 88: 9543-9547

47. Wattenberg LW and Loub WD. 1978. Inhibition of polycyclic aromatic hydrocarbon- induced neoplasia by naturally occurring indoles. Cancer Res. 38: 1410-1413

48. Perdew GH and Babbs CF. 1991. Production of Ah receptor ligands in rat fecal suspensions containing tryptophan or indole-3-carbinol. Nutrition & Cancer 16: 209-218

49. Rogers JM and Denison MS. 2000. Recombinant cell bioassays for endocrine disruptors: development of a stably transfected human ovarian cell line for the detection of estrogenic and anti-estrogenic chemicals. In Vitr.Mol.Toxicol. 13: 67-82

50. Amakura Y, Tsutsumi T, Nakamura M, Kitagawa H, Fujino J et al. 2002. Preliminary screening of the inhibitory effect of food extracts on activation of the aryl hydrocarbon receptor induced by 2,3,7,8-tetrachlorodibenzo-p-dioxin. Biol.Pharm.Bull. 25: 272-274

51. Moon YJ, Wang X, and Morris ME. 2006. Dietary flavonoids: effects on xenobiotic and carcinogen metabolism. Toxicol.In Vitro 20: 187-210

52. Bradshaw TD, Trapani V, Vasselin DA, and Westwell AD. 2002. The aryl hydrocarbon receptor in anticancer drug discovery: friend or foe? Curr.Pharm.Des 8: 2475-2490

53. Mandal PK. 2005. Dioxin: a review of its environmental effects and its aryl hydrocarbon receptor biology. J.Comp Physiol [B] 175: 221-230

203

54. Pelclova D, Urban P, Preiss J, Lukas E, Fenclova Z et al. 2006. Adverse health effects in humans exposed to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD). Rev.Environ.Health 21: 119-138

55. Gonzalez FJ and Fernandez-Salguero P. 1998. The aryl hydrocarbon receptor: studies using the AHR-null mice. Drug Metab Dispos. 26: 1194-1198

56. Cole P, Trichopoulos D, Pastides H, Starr T, and Mandel JS. 2003. Dioxin and cancer: a critical review. Regul.Toxicol.Pharmacol. 38: 378-388

57. Steenland K, Bertazzi P, Baccarelli A, and Kogevinas M. 2004. Dioxin revisited: developments since the 1997 IARC classification of dioxin as a human carcinogen. Environ.Health Perspect. 112: 1265-1268

58. Anonymous. 1997. IARC Working Group on the Evaluation of Carcinogenic Risks to Humans: Polychlorinated Dibenzo-Para-Dioxins and Polychlorinated Dibenzofurans. Lyon, France, 4-11 February 1997. IARC Monogr Eval.Carcinog.Risks Hum. 69: 1-631

59. Larsen JC. 2006. Risk assessments of polychlorinated dibenzo- p-dioxins, polychlorinated dibenzofurans, and dioxin-like polychlorinated biphenyls in food. Mol.Nutr.Food Res. 50: 885-896

60. Liem AK, Furst P, and Rappe C. 2000. Exposure of populations to dioxins and related compounds. Food Addit.Contam 17: 241-259

61. Neuberger M, Landvoigt W, and Derntl F. 1991. Blood levels of 2,3,7,8- tetrachlorodibenzo-p-dioxin in chemical workers after chloracne and in comparison groups. Int.Arch.Occup.Environ.Health 63: 325-327

62. Sweeney MH and Mocarelli P. 2000. Human health effects after exposure to 2,3,7,8- TCDD. Food Addit.Contam 17: 303-316

63. Martin JV. 1984. Lipid abnormalities in workers exposed to dioxin. Br.J.Ind.Med. 41: 254-256

64. Dalton TP, Kerzee JK, Wang B, Miller M, Dieter MZ et al. 2001. Dioxin exposure is an environmental risk factor for ischemic heart disease. Cardiovasc.Toxicol. 1: 285-298

65. Vena J, Boffetta P, Becher H, Benn T, Bueno-de-Mesquita HB et al. 1998. Exposure to dioxin and nonneoplastic mortality in the expanded IARC international cohort study of phenoxy herbicide and chlorophenol production workers and sprayers. Environ.Health Perspect. 106 Suppl 2: 645-653

66. Henriksen GL, Ketchum NS, Michalek JE, and Swaby JA. 1997. Serum dioxin and diabetes mellitus in veterans of Operation Ranch Hand. Epidemiology 8: 252-258

67. Remillard RB and Bunce NJ. 2002. Linking dioxins to diabetes: epidemiology and biologic plausibility. Environ.Health Perspect. 110: 853-858

204

68. Johnson E, Shorter C, Bestervelt L, Patterson D, Needham L et al. 2001. Serum hormone levels in humans with low serum concentrations of 2,3,7,8-TCDD. Toxicol.Ind.Health 17: 105-112

69. Birnbaum LS and Cummings AM. 2002. Dioxins and endometriosis: a plausible hypothesis. Environ.Health Perspect. 110: 15-21

70. Birnbaum LS and Tuomisto J. 2000. Non-carcinogenic effects of TCDD in animals. Food Addit.Contam 17: 275-288

71. Dragan YP and Schrenk D. 2000. Animal studies addressing the carcinogenicity of TCDD (or related compounds) with an emphasis on tumour promotion. Food Addit.Contam 17: 289-302

72. Carpenter DO. 2006. Polychlorinated biphenyls (PCBs): routes of exposure and effects on human health. Rev.Environ.Health 21: 1-23

73. Connor KT and Aylward LL. 2006. Human response to dioxin: aryl hydrocarbon receptor (AhR) molecular structure, function, and dose-response data for enzyme induction indicate an impaired human AhR. J.Toxicol.Environ.Health B Crit Rev. 9: 147-171

74. Bock KW and Kohle C. 2006. Ah receptor: dioxin-mediated toxic responses as hints to deregulated physiologic functions. Biochem.Pharmacol. 72: 393-404

75. Powell-Coffman JA, Bradfield CA, and Wood WB. 1998. Caenorhabditis elegans orthologs of the aryl hydrocarbon receptor and its heterodimerization partner the aryl hydrocarbon receptor nuclear translocator. Proc.Natl.Acad.Sci.U.S.A 95: 2844-2849

76. Duncan DM, Burgess EA, and Duncan I. 1998. Control of distal antennal identity and tarsal development in Drosophila by spineless-aristapedia, a homolog of the mammalian dioxin receptor. Genes Dev. 12: 1290-1303

77. Dehal P, Satou Y, Campbell RK, Chapman J, Degnan B et al. 2002. The draft genome of Ciona intestinalis: insights into chordate and vertebrate origins. Science 298: 2157-2167

78. Butler RA, Kelley ML, Powell WH, Hahn ME, and Van Beneden RJ. 2001. An aryl hydrocarbon receptor (AHR) homologue from the soft-shell clam, Mya arenaria: evidence that invertebrate AHR homologues lack 2,3,7,8-tetrachlorodibenzo-p-dioxin and beta-naphthoflavone binding. Gene 278: 223-234

79. Hahn ME, Karchner SI, Evans BR, Franks DG, Merson RR, and Lapseritis JM. 2006. Unexpected diversity of aryl hydrocarbon receptors in non-mammalian vertebrates: insights from comparative genomics. J.Exp.Zoolog.A Comp Exp.Biol. 305: 693-706

80. Huang X, Powell-Coffman JA, and Jin Y. 2004. The AHR-1 aryl hydrocarbon receptor and its co-factor the AHA-1 aryl hydrocarbon receptor nuclear translocator specify GABAergic neuron cell fate in C. elegans. Development 131: 819-828

205

81. Hahn ME. 2002. Aryl hydrocarbon receptors: diversity and evolution. Chem.Biol.Interact. 141: 131-160

82. Mason ME and Okey AB. 1982. Cytosolic and nuclear binding of 2,3,7,8- tetrachlorodibenzo-p- dioxin to the Ah receptor in extrahepatic tissues of rats and mice. Eur.J.Biochem. 123: 209-215

83. Dolwick KM, Schmidt JV, Carver LA, Swanson HI, and Bradfield CA. 1993. Cloning and expression of a human Ah receptor cDNA. Mol.Pharmacol. 44: 911-917

84. Carver LA, Hogenesch JB, and Bradfield CA. 1994. Tissue specific expression of the rat Ah-receptor and ARNT mRNAs. Nucleic Acids Res. 22: 3038-3044

85. Manchester DK, Gordon SK, Golas CL, Roberts EA, and Okey AB. 1987. Ah receptor in human placenta: stabilization by molybdate and characterization of binding of 2,3,7,8- tetrachlorodibenzo-p- dioxin, 3-methylcholanthrene, and benzo(a)pyrene. Cancer Res. 47: 4861-4868

86. Lahvis GP, Lindell SL, Thomas RS, McCuskey RS, Murphy C et al. 2000. Portosystemic shunting and persistent fetal vascular structures in aryl hydrocarbon receptor-deficient mice. Proc.Natl.Acad.Sci.U.S.A 97: 10442-10447

87. Khorram O, Garthwaite M, and Golos T. 2002. Uterine and ovarian aryl hydrocarbon receptor (AHR) and aryl hydrocarbon receptor nuclear translocator (ARNT) mRNA expression in benign and malignant gynaecological conditions. Mol.Hum.Reprod. 8: 75- 80

88. Williamson MA, Gasiewicz TA, and Opanashuk LA. 2005. Aryl hydrocarbon receptor expression and activity in cerebellar granule neuroblasts: implications for development and dioxin neurotoxicity. Toxicol.Sci. 83: 340-348

89. Lahvis GP, Pyzalski RW, Glover E, Pitot HC, McElwee MK, and Bradfield CA. 2005. The aryl hydrocarbon receptor is required for developmental closure of the ductus venosus in the neonatal mouse. Mol.Pharmacol. 67: 714-720

90. Harstad EB, Guite CA, Thomae TL, and Bradfield CA. 2006. Liver deformation in Ahr- null mice: evidence for aberrant hepatic perfusion in early development. Mol.Pharmacol. 69: 1534-1541

91. Lund AK, Goens MB, Kanagy NL, and Walker MK. 2003. Cardiac hypertrophy in aryl hydrocarbon receptor null mice is correlated with elevated angiotensin II, endothelin-1, and mean arterial blood pressure. Toxicol.Appl.Pharmacol. 193: 177-187

92. Lin TM, Ko K, Moore RW, Simanainen U, Oberley TD, and Peterson RE. 2002. Effects of aryl hydrocarbon receptor null mutation and in utero and lactational 2,3,7,8- tetrachlorodibenzo-p-dioxin exposure on prostate and seminal vesicle development in C57BL/6 mice. Toxicol.Sci. 68: 479-487

206

93. Danilovich N and Ram SM. 2006. Recent female mouse models displaying advanced reproductive aging. Exp.Gerontol. 41: 117-122

94. Peters JM, Narotsky MG, Elizondo G, Fernandez-Salguero PM, Gonzalez FJ, and Abbott BD. 1999. Amelioration of TCDD-induced teratogenesis in aryl hydrocarbon receptor (AhR)-null mice. Toxicol.Sci. 47: 86-92

95. Shimizu Y, Nakatsuru Y, Ichinose M, Takahashi Y, Kume H et al. 2000. Benzo[a]pyrene carcinogenicity is lost in mice lacking the aryl hydrocarbon receptor. Proc.Natl.Acad.Sci.U.S.A 97: 779-782

96. Perdew GH. 1988. Association of the Ah receptor with the 90-kDa heat shock protein. J.Biol.Chem. 263: 13802-13805

97. Denis M, Cuthill S, Wikstrom AC, Poellinger L, and Gustafsson JA. 1988. Association of the dioxin receptor with the Mr 90,000 heat shock protein: a structural kinship with the glucocorticoid receptor. Biochem.Biophys.Res.Commun. 155: 801-807

98. Chen HS and Perdew GH. 1994. Subunit composition of the heteromeric cytosolic aryl hydrocarbon receptor complex. J.Biol.Chem. 269: 27554-27558

99. Kazlauskas A, Poellinger L, and Pongratz I. 1999. Evidence that the co-chaperone p23 regulates ligand responsiveness of the dioxin (Aryl hydrocarbon) receptor. J.Biol.Chem. 274: 13519-13524

100. Cox MB and Miller CA, III. 2004. Cooperation of heat shock protein 90 and p23 in aryl hydrocarbon receptor signaling. Cell Stress.Chaperones. 9: 4-20

101. Meyer BK, Pray-Grant MG, Vanden Heuvel JP, and Perdew GH. 1998. Hepatitis B virus X-associated protein 2 is a subunit of the unliganded aryl hydrocarbon receptor core complex and exhibits transcriptional enhancer activity. Mol.Cell Biol. 18: 978-988

102. Ma Q and Whitlock JP, Jr. 1997. A novel cytoplasmic protein that interacts with the Ah receptor, contains tetratricopeptide repeat motifs, and augments the transcriptional response to 2,3,7,8-tetrachlorodibenzo-p-dioxin. J.Biol.Chem. 272: 8878-8884

103. Carver LA and Bradfield CA. 1997. Ligand-dependent interaction of the aryl hydrocarbon receptor with a novel immunophilin homolog in vivo. J.Biol.Chem. 272: 11452-11456

104. Antonsson C, Whitelaw ML, McGuire J, Gustaffson J-Å, and Poellinger L. 1995. Distinct roles of the molecular chaperone hsp90 in modulating dioxin receptor function via the basic helix-loop-helix and PAS domains. Mol.Cell.Biol. 15: 756-765

105. Perdew GH and Bradfield CA. 1996. Mapping the 90 kDa heat shock protein binding region of the Ah receptor. Biochem.Mol.Biol.Int. 39: 589-593

207

106. Pollenz RS, Wilson SE, and Dougherty EJ. 2006. Role of endogenous XAP2 protein on the localization and nucleocytoplasmic shuttling of the endogenous mouse Ahb-1 receptor in the presence and absence of ligand. Mol.Pharmacol. 70: 1369-1379

107. Ramadoss P and Perdew GH. 2005. The transactivation domain of the Ah receptor is a key determinant of cellular localization and ligand-independent nucleocytoplasmic shuttling properties. Biochemistry 44: 11148-11159

108. Ikuta T, Eguchi H, Tachibana T, Yoneda Y, and Kawajiri K. 1998. Nuclear localization and export signals of the human aryl hydrocarbon receptor. J.Biol.Chem. 273: 2895-2904

109. Reyes H, Reiz-Porszasz S, and Hankinson O. 1992. Identification of the Ah receptor nuclear translocator protein (Arnt) as a component of the DNA binding form of the Ah receptor. Science 256: 1193-1195

110. McGuire J, Whitelaw ML, Pongratz I, Gustaffson J-Å, and Poellinger L. 1994. A cellular factor stimulates ligand-dependent release of hsp90 from the basic helix-loop-helix dioxin receptor. Mol.Cell.Biol. 14: 2438-2446

111. Swanson HI, Chan WK, and Bradfield CA. 1995. DNA binding specificities and pairing rules of the Ah receptor, ARNT, and SIM proteins. J.Biol.Chem. 270: 26292-26302

112. Mimura J and Fujii-Kuriyama Y. 2003. Functional role of AhR in the expression of toxic effects by TCDD. Biochim.Biophys.Acta 1619: 263-268

113. Nebert DW, Roe AL, Dieter MZ, Solis WA, Yang Y, and Dalton TP. 2000. Role of the aromatic hydrocarbon receptor and [Ah] gene battery in the oxidative stress response, cell cycle control, and apoptosis. Biochem.Pharmacol. 59: 65-85

114. Kolluri SK, Weiss C, Koff A, and Göttlicher M. 1999. p27kip1 induction and inhibition of proliferation by the intracellular Ah receptor in developing thymus and hepatoma cells. Genes Dev. 13: 1742-1753

115. Kolluri SK, Balduf C, Hofmann M, and Gottlicher M. 2001. Novel target genes of the Ah (dioxin) receptor: transcriptional induction of N-myristoyltransferase 2. Cancer Res. 61: 8534-8539

116. Matikainen T, Perez GI, Jurisicova A, Pru JK, Schlezinger JJ et al. 2001. Aromatic hydrocarbon receptor-driven Bax gene expression is required for premature ovarian failure caused by biohazardous environmental chemicals. Nat.Genet. 28: 355-360

117. Mimura J, Ema M, Sogawa K, and Fujii-Kuriyama Y. 1999. Identification of a novel mechanism of regulation of Ah (dioxin) receptor function. Genes Dev. 13: 20-25

118. Baba T, Mimura J, Gradin K, Kuroiwa A, Watanabe T et al. 2001. Structure and expression of the Ah receptor repressor gene. J.Biol.Chem. 276: 33101-33110

208

119. Pollenz RS. 2002. The mechanism of AH receptor protein down-regulation (degradation) and its impact on AH receptor-mediated gene regulation. Chem.Biol.Interact. 141: 41-61

120. Rowlands JC, McEwan I, and Gustaffson J-Å. 1996. Transactivation by the human aryl hydrocarbon receptor and aryl hydrocarbon nuclear translocator proteins: direct interactions with basal transcription factors. Mol.Pharmacol. 50: 538-548

121. Swanson HI and Yang JH. 1998. The aryl hydrocarbon receptor interacts with transcription factor IIB. Mol.Pharmacol. 54: 671-677

122. Hankinson O. 2005. Role of coactivators in transcriptional activation by the aryl hydrocarbon receptor. Arch.Biochem.Biophys. 433: 379-386

123. Wang S, Ge K, Roeder RG, and Hankinson O. 2004. Role of mediator in transcriptional activation by the aryl hydrocarbon receptor. J.Biol.Chem. 279: 13593-13600

124. Nguyen TA, Hoivik D, Lee JE, and Safe S. 1999. Interactions of nuclear receptor coactivator/corepressor proteins with the aryl hydrocarbon receptor complex. Arch.Biochem.Biophys. 367: 250-257

125. Klinge CM, Jernigan SC, Risinger KE, Lee JE, Tyulmenkov VV et al. 2001. Short heterodimer partner (SHP) orphan nuclear receptor inhibits the transcriptional activity of aryl hydrocarbon receptor (AHR)/AHR nuclear translocator (ARNT). Arch.Biochem.Biophys. 390: 64-70

126. Swanson HI. 2002. DNA binding and protein interactions of the AHR/ARNT heterodimer that facilitate gene activation. Chem.Biol.Interact. 141: 63-76

127. Gradin K, Toftgard R, Poellinger L, and Berghard A. 1999. Repression of dioxin signal transduction in fibroblasts. Identification Of a putative repressor associated with Arnt. J.Biol.Chem. 274: 13511-13518

128. Ge N-L and Elferink CJ. 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblatoma protein. J.Biol.Chem. 273: 22708-22713

129. Puga A, Barnes SJ, Dalton TP, Chang C, Knudsen ES, and Maier MA. 2000. Aromatic hydrocarbon receptor interaction with the retinoblastoma protein potentiates repression of E2F-dependent transcription and cell cycle arrest. J.Biol.Chem. 275: 2943-2950

130. Klinge CM, Kaur K, and Swanson HI. 2000. The aryl hydrocarbon receptor interacts with estrogen receptor alpha and orphan receptors COUP-TFI and ERRalpha1. Arch.Biochem.Biophys. 373: 163-174

131. Tian Y, Ke S, Denison MS, Rabson AB, and Gallo MA. 1999. Ah receptor and NF- kappaB interactions, a potential mechanism for dioxin toxicity. J.Biol.Chem. 274: 510- 515

209

132. Harper PA, Riddick DS, and Okey AB. 2006. Regulating the regulator: factors that control levels and activity of the aryl hydrocarbon receptor. Biochem.Pharmacol. 72: 267-279

133. Davarinos NA and Pollenz RS. 1999. Aryl hydrocarbon receptor imported into the nucleus following ligand binding is rapidly degraded via the cytosplasmic proteasome following nuclear export. J.Biol.Chem. 274: 28708-28715

134. Ma Q, Renzelli AJ, Baldwin KT, and Antonini JM. 2000. Superinduction of CYP1A1 gene expression. Regulation of 2,3,7, 8-tetrachlorodibenzo-p-dioxin-induced degradation of Ah receptor by cycloheximide. J.Biol.Chem. 275: 12676-12683

135. Andersson P, McGuire J, Rubio C, Gradin K, Whitelaw ML et al. 2002. A constitutively active dioxin/aryl hydrocarbon receptor induces stomach tumors. Proc.Natl.Acad.Sci.U.S.A 99: 9990-9995

136. Nohara K, Pan X, Tsukumo S, Hida A, Ito T et al. 2005. Constitutively active aryl hydrocarbon receptor expressed specifically in T-lineage cells causes thymus involution and suppresses the immunization-induced increase in splenocytes. J.Immunol. 174: 2770- 2777

137. Tauchi M, Hida A, Negishi T, Katsuoka F, Noda S et al. 2005. Constitutive expression of aryl hydrocarbon receptor in keratinocytes causes inflammatory skin lesions. Mol.Cell Biol. 25: 9360-9368

138. Moennikes O, Loeppen S, Buchmann A, Andersson P, Ittrich C et al. 2004. A constitutively active dioxin/aryl hydrocarbon receptor promotes hepatocarcinogenesis in mice. Cancer Res. 64: 4707-4710

139. Puga A, Marlowe J, Barnes S, Chang CY, Maier A et al. 2002. Role of the aryl hydrocarbon receptor in cell cycle regulation. Toxicology 181-182: 171-177

140. Kamath AB, Nagarkatti PS, and Nagarkatti M. 1998. Characterization of phenotypic alterations induced by 2,3,7,8-tetrachlorodibenzo-p-dioxin on thymocytes in vivo and its effect on apoptosis. Toxicol.Appl.Pharmacol. 150: 117-124

141. Schrenk D, Schmitz HJ, Bohnenberger S, Wagner B, and Worner W. 2004. Tumor promoters as inhibitors of apoptosis in rat hepatocytes. Toxicol.Lett. 149: 43-50

142. Ray SS and Swanson HI. 2003. Alteration of keratinocyte differentiation and senescence by the tumor promoter dioxin. Toxicol.Appl.Pharmacol. 192: 131-145

143. Bauman JW, Goldsworthy TL, Dunn CS, and Fox TR. 1995. Inhibitory effects of 2,3,7,8- tetrachlorodibenzo-p-dioxin on rat hepatocyte proliferation induced by 2/3 partial hepatectomy. Cell Prolif. 28: 437-451

210

144. Alexander DL, Ganem LG, Fernandez-Salguero P, Gonzalez F, and Jefcoate CR. 1998. Aryl-hydrocarbon receptor is an inhibitory regulator of lipid synthesis and of commitment to adipogenesis. J.Cell Sci. 111 ( Pt 22): 3311-3322

145. Loertscher JA, Lin TM, Peterson RE, and Allen-Hoffmann BL. 2002. In utero exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin causes accelerated terminal differentiation in fetal mouse skin. Toxicol.Sci. 68: 465-472

146. Roberts RA, Nebert DW, Hickman JA, Richburg JH, and Goldsworthy TL. 1997. Perturbation of the mitosis/apoptosis balance: a fundamental mechanism in toxicology. Fundam.Appl.Toxicol. 38: 107-115

147. Marlowe JL, Knudsen ES, Schwemberger S, and Puga A. 2004. The aryl hydrocarbon receptor displaces p300 from E2F-dependent promoters and represses S phase-specific gene expression. J.Biol.Chem. 279: 29013-29022

148. Mitchell KA, Lockhart CA, Huang G, and Elferink CJ. 2006. Sustained aryl hydrocarbon receptor activity attenuates liver regeneration. Mol.Pharmacol. 70: 163-170

149. Elferink CJ, Ge NL, and Levine A. 2001. Maximal Aryl Hydrocarbon Receptor Activity Depends on an Interaction with the Retinoblastoma Protein. Mol.Pharmacol. 59: 664-673

150. Huang G and Elferink CJ. 2005. Multiple mechanisms are involved in Ah receptor- mediated cell cycle arrest. Mol.Pharmacol. 67: 88-96

151. Giacinti C and Giordano A. 2006. RB and cell cycle progression. Oncogene 25: 5220- 5227

152. Levine-Fridman A, Chen L, and Elferink CJ. 2004. Cytochrome P4501A1 Promotes G1 Phase Cell Cycle Progression by Controlling Aryl Hydrocarbon Receptor Activity. Mol.Pharmacol. 65: 461-469

153. Fong CJ, Burgoon LD, and Zacharewski TR. 2005. Comparative microarray analysis of basal gene expression in mouse Hepa-1c1c7 wild-type and mutant cell lines. Toxicol.Sci. 86: 342-353

154. Sun W, Zhang J, and Hankinson O. 1997. A mutation in the aryl hydrocarbon receptor (AHR) in a cultured mammalian cell line identifies a novel region of AHR that affects DNA binding. J.Biol.Chem. 272: 31845-31854

155. Elizondo G, Fernandez-Salguero P, Sheikh MS, Kim GY, Fornace AJ et al. 2000. Altered cell cycle control at the G(2)/M phases in aryl hydrocarbon receptor-null embryo fibroblast. Mol.Pharmacol. 57: 1056-1063

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Appendix

Summary

The capacity to inhibit apoptosis, and therefore allow the survival of cells bearing

genotoxic lesions, may be the primary mechanism by which TCDD promotes tumor formation in

rat liver and other tissues (1). TCDD inhibits apoptosis in DEN-initiated rat liver tumors to about 10% of control levels (2), and inhibits apoptosis induced by treatment with DNA- damaging agents in liver cell cultures. There has been little investigation into the mechanism by which TCDD inhibits apoptosis. One study points to the abrogation of p53 activity in TCDD- treated cells as part of the mechanism of action (3), but little else has been noted. The fact that the AHR interacts with and possibly inhibits some transcriptional activities of E2F1 points to one possible mechanism by which TCDD may inhibit apoptosis. E2F1 is a potent activator of the apoptotic cell death pathway under physiologic conditions and under conditions of cell cycle deregulation. E2F1 is believed to mediate the apoptotic elimination of cells in which normal cell cycle control is lost due to a disruption in the pathway regulating RB activity. Much of the E2F- mediated apoptotic response is attributed to the transcriptional activation of pro-apoptotic genes.

The fact that RB and AHR have a similar capacity to inhibit E2F-dependent transcription (4),

and that RB can inhibit E2F1-induced apoptosis, indicates that exposure to TCDD may also

attenuate E2F1-induced apoptosis through activation of the AH receptor. The goal of this series of experiments was to test this hypothesis.

Background

Exposure to polycyclic aromatic hydrocarbons (PAHs) results in a plethora of toxic and carcinogenic responses in animals and in humans. TCDD (2,3,7,8-tetrachlorodibenzo-p-dioxin),

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commonly studied as a model for outcomes of exposure to other polychlorinated hydrocarbons,

is a potent tumor promoter in several rodent models, and has been labeled a human carcinogen

(5;6). The specific mechanisms by which TCDD exhibits its promotional effects are speculative.

Exposure to TCDD either in vivo or in vitro results in an array of seemingly contradictory

responses, including both increased and decreased rates of cell proliferation, inhibition and

promotion of apoptosis, and differentiation, depending on the cell or tissue type examined (7).

It is widely accepted that the majority of the effects of TCDD exposure are mediated by

the aromatic hydrocarbon receptor (AHR). The AHR is a ligand-activated transcription factor

that, upon ligand binding, translocates to the nucleus where it complexes with the aromatic

hydrocarbon nuclear translocator (ARNT). The AHR/ARNT heterodimer binds to specific

consensus sites in the regulatory domains of Phase I and Phase II drug-metabolizing enzymes to

activate their transcription (8). This represents an adaptive response required for the

detoxification of foreign compounds (i.e. PAHs), the effects of which do not adequately explain

the myriad of outcomes resulting from TCDD treatment in different systems, particularly

regarding gene repression effects (7). Many studies indicate that the AHR also functions in the

direct and indirect modulation of the transcriptional program, at least in part by associating with

additional transcription factors (9;10) and coactivators or corepressor (11;12). In doing so, the

ligand-activated receptor is able to alter expression of genes regulating proliferation, apoptosis, differentiation, and signal transduction pathways (7). TCDD itself is poorly metabolized, allowing it to accumulate and produce sustained effects via chronic receptor activation (13).

Tumor promoters are generally believed to act by affecting the rate of division, terminal differentiation, or death of tumor precursor cells. One widely accepted mechanism of tumor promotion/progression is the inhibition of apoptosis (14). The capacity of TCDD to act as a

213

tumor promoter, particularly in rodent liver, has been attributed to its ability to inhibit the

apoptotic elimination of initiated cells bearing genotoxic lesions (15). However, the precise

mechanism(s) of this effect remains elusive, and differs with the organism, tissue, or cell type

examined. In diethylnitrosamine (DEN)-initiated rats, both acute and chronic TCDD treatment

results in an approximate 10-fold decrease in the rate of apoptosis in preneoplastically

transformed liver foci, with no effect on the background rate of apoptosis in normal hepatocytes

(2). The overall effect of TCDD is thus to accelerate the rate at which DNA-damaged cells

convert to a neoplastic phenotype (16). As stimulation of cell division in similar assays is

negligible (17), the primary effect is the inhibition of apoptosis. TCDD treatment also inhibits

apoptosis in c-myc transgenic mice (15). TCDD promotes ovarian tumor formation in rats, an

outcome that is also suggested to result from an effect on apoptosis (18). Absence of tumor

promotion by TCDD treatment in rat strains lacking a functional AHR suggests that the receptor

is required for this effect (19). Effects of TCDD on apoptosis have also been documented in

cultured cells. TCDD inhibits apoptosis in cultured hepatocytes treated with UV light or 2-

acetylaminofluorene, an effect that was attributed in part to attenuation of p53 activity (1;3).

Apoptosis induced by growth factor withdrawal in human epithelial cells is inhibited by TCDD

treatment, in correlation with activation of the epidermal growth factor (EGF) signaling pathway

(20). Finally, studies of AHR-null mice confirm the importance of the AHR in tissue

homeostasis, as hepatocytes in these mice exhibit accelerated rates of apoptosis associated with

increased production of TGF-β (21). In vitro cell populations lacking the receptor also have higher rates of apoptotic death (22).

The mechanism by which TCDD inhibits apoptosis in preneoplastic liver cells, or in any system, is unknown. AHR activation alters several cell cycle and signaling pathways, including

214

those required for normal cell cycle regulation. Recent data indicate that the AHR directly

interacts with the retinoblastoma (RB) protein (4;10), which complexes with E2F family proteins

to control cell cycle progression. E2F complexes control the transcription of a variety of essential cell cycle control genes, including cell cycle regulators, RB and related pocket proteins, enzymes for nucleotide biosynthesis, and proteins required for DNA replication (23). Either RB or the RB-related proteins p107 and p130 directly bind E2F to inhibit transcriptional activation and mediate active repression of E2F-responsive genes, thereby inhibiting cell cycle progression from G1 into S-phase (24). The activated AHR cooperates with RB in this response (25), and it may also affect E2F transcriptional activity in the absence of RB binding (4).

In addition to controlling gene transcription for cell cycle progression, several E2F family members, in particular E2F1, are able to activate apoptosis. E2F1 is characterized as an oncogene based on evidence from cell transformation assays, and from transgenic mouse studies in which E2F1 overexpression increases rates of tumor formation (26). In contrast, E2F1 has been labeled a tumor suppressor based on the finding that E2F1-deficient mice develop a range of tumor types (27;28). Evidence suggests that the ability of E2F1 to act as a tumor suppressor lies in its ability to initiate apoptosis in cells that lose normal cell cycle control. The AHR may play a role in the inhibition of this response through transcriptional repression of genes required for E2F1-dependent apoptosis.

There is abundant in vivo and in vitro evidence suggesting that E2F1 acts as a positive regulator of apoptosis. E2F1-/- mice exhibit enlarged lymphoid organs as the result of a defect in

the normal apoptotic elimination of thymocytes (28). Deletion of E2F1 in TgT121 transgenic mice, which express a truncated SV40 T antigen that inactivates RB, results in an 80% decrease in the apoptotic index of targeted cells (29). This effect results from an inhibition of p53-

215

dependent apoptosis. Overexpression of E2F1 in epithelial tissues of transgenic mice inhibits

tumor promotion in correlation with the induction of apoptosis following topical TPA treatment,

suggesting that E2F1 exerts its tumor-suppressive effects during the promotion stage of tumor

development (30). Treatment of gliomas in nude mice with an E2F1-expressing adenovirus

arrests tumor growth (31), while expression of E2F1 in c-myc transgenic mice results in an

increase in the apoptotic rate and delayed malignant conversion during liver tumor progression

(32). Mice lacking RB die in utero, exhibiting massive apoptosis attributed in part to

unregulated E2F transcriptional activity (33). Finally, E2F1 overexpression and RB deficiency

lead to apoptosis in a number of cell types (34), including in primary rat hepatocytes (35), and

increased DNA-binding and transcriptional activities of E2F are correlated with induction of

apoptosis under specific conditions (36).

Several mechanisms mediating E2F1-dependent apoptosis have been identified, including

the inhibition of anti-apoptotic signaling pathways, transcriptional activation of specific genes

that positively regulate apoptosis, and inhibition of genes that are negative regulators of

apoptosis (33). The best characterized of these mechanisms is the transcriptional activation of

genes in the p53 pathway. The p53 family consists of key tumor suppressor proteins that

activate apoptosis to eliminate damaged or oncogenic cells. One member of the family that is

capable of inducing apoptosis is p73 (37). p73 is upregulated in response to ectopic or endogenous E2F1 expression (38;39), and may be required for E2F1-mediated apoptosis under some conditions (40). Other studies implicate p53 itself as a mediator of E2F1-dependent apoptosis (41;42). p53 is regulated through a process of ubiquitination and degradation by

Mdm-2, and inhibitors of this process allow for the stabilization of p53 and the subsequent activation of apoptosis signaling pathways (43). The ability of Mdm-2 to mediate p53

216

degradation is inhibited by direct binding to p19ARF (human p14ARF). The transcription of the

ARF gene is controlled in part by the action of E2F1. The upregulation of p19ARF by E2F1 occurs under many experimental conditions, both in cell culture and in animal models (44;45).

Additional but less well-characterized pro-apoptotic targets of E2F1 are TGF-β (46) and the

procaspase-activator APAF1 (47). It is probable that E2F1 activates multiple pathways in

concert to initiate apoptosis, although the importance of the different pathways seems to vary by

cell and tissue type.

Results obtained during investigation of the interactions of AHR with RB suggested that

AHR directly interacts with and modulates the transcriptional activity of E2F1. Published results

in Hepa-1 cells as well as unpublished results in several other cell lines indicate that TCDD

stimulation or ectopic expression of the AHR inhibits expression of E2F-dependent reporter genes, and the extent of inhibition is equal to that exhibited by RB alone (4). Data presented in

Chapter 4 reveal several lines of evidence indicating that AHR and E2F1 interact directly in cell culture. AHR binds as well to E2F1 as to RB in in vitro pull-down assays using bacterially synthesized E2F1 and radioactively labeled receptor. Gel shift assays using CV-1 nuclear extracts and purified AHR peptides show that AHR can displace RB from E2F binding sites occupied by RB/E2F/DP1 complexes, generating a novel complex containing E2F and AHR. In addition, the AHR can be immunoprecipitated using an anti-E2F1 antibody in Hepa-1 cells, which lack RB. In total, these data point to a direct interaction between AHR and E2F1 that potentially results in the repression of E2F-dependent transcription. The outcome of this interaction on the expression of specific genes, or the mechanism by which the AHR mediates transcriptional repression, has not been investigated. The following experiments address the

217

hypothesis that the AHR can repress E2F-mediated apoptosis through the repression of E2F-

induced, apoptosis-specific genes.

Discussion

The results presented in the following figures indicate that the AHR may actually inhibit

E2F-induced apoptosis when E2F1 is overexpressed by adenoviral vectors in Saos-2 cells.

Adenoviral-mediate expression of the AHR represses cell death induced by E2F1 in these cells at

least as well if not better than RB. In addition, the AHR is able to suppress the E2F1-mediated

induction of several pro-apoptotic genes in a manner analogous to RB. Considering the large

amount of data presented in Chapter 4 indicating an interaction between the AHR and E2F

proteins, it could be that AHR is acting in a similar manner as RB in the repression E2F-

dependent apoptosis, and RB binding to E2F is known to inhibit E2F transactivation activity.

The inhibition of apoptosis by the AHR plays a role in the progression of tumorigenesis observed

in TCDD-treated animals. The inhibition of E2F-transcriptional activity observed here is a

potential mechanism whereby the TCDD-activated AHR could act to promote carcinogenesis by

inhibiting the apoptotic elimination of cells that have lost normal cell cycle control. However, the results presented here could not be repeated using additional adenoviral stocks for expression of the same factors. The reason for this is unknown at this point, but may be the result of the sometimes-innate toxicity of adenoviral vector stocks, or an effect of extremely high expression

levels of these proteins. Although the results presented in the following figures are promising in

terms of identifying a mechanism of AHR-mediated tumor promotion, overall the results of these

experiments are inconclusive. The use of alternative methods or additional experimental

approaches is required to determine the impact of the AHR on E2F-dependent apoptosis.

218

References

1. Worner W and Schrenk D. 1996. Influence of liver tumor promoters on apoptosis in rat hepatocytes induced by 2-acetylaminofluorene, ultraviolet light, or transforming growth factor beta 1. Cancer Res. 56: 1272-1278

2. Stinchcombe S, Buchmann A, Bock KW, and Schwarz M. 1995. Inhibition of apoptosis during 2,3,7,8-tetrachlorodibenzo-p- dioxin-mediated tumour promotion in rat liver. Carcinogenesis 16: 1271-1275

3. Worner W and Schrenk D. 1998. 2,3,7,8-Tetrachlorodibenzo-p-dioxin suppresses apoptosis and leads to hyperphosphorylation of p53 in rat hepatocytes. Environ.Toxicol.Pharmacol. 6: 239-247

4. Puga A, Barnes SJ, Dalton TP, Chang C, Knudsen ES, and Maier MA. 2000. Aromatic hydrocarbon receptor interaction with the retinoblastoma protein potentiates repression of E2F-dependent transcription and cell cycle arrest. J.Biol.Chem. 275: 2943-2950

5. Bertazzi PA, Consonni D, Bachetti S, Rubagotti M, Baccarelli A et al. 2001. Health effects of dioxin exposure: a 20-year mortality study. Am.J.Epidemiol. 153: 1031-1044

6. Anonymous. 1997. IARC Working Group on the Evaluation of Carcinogenic Risks to Humans: Polychlorinated Dibenzo-Para-Dioxins and Polychlorinated Dibenzofurans. Lyon, France, 4-11 February 1997. IARC Monogr Eval.Carcinog.Risks Hum. 69: 1-631

7. Puga A, Maier A, and Medvedovic M. 2000. The transcriptional signature of dioxin in human hepatoma HepG2 cells. Biochem.Pharmacol. 60: 1129-1142

8. Hankinson O. 1995. The aryl hydrocarbon receptor complex. Annu.Rev.Pharmacol.Toxicol. 35: 307-340

9. Wang F, Wang W, and Safe S. 1999. Regulation of constitutive gene expression through interactions of Sp1 protein with the nuclear aryl hydrocarbon receptor complex. Biochemistry 38: 11490-11500

10. Ge N-L and Elferink CJ. 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblatoma protein. J.Biol.Chem. 273: 22708-22713

11. Nguyen TA, Hoivik D, Lee JE, and Safe S. 1999. Interactions of nuclear receptor coactivator/corepressor proteins with the aryl hydrocarbon receptor complex. Arch.Biochem.Biophys. 367: 250-257

12. Wang S and Hankinson O. 2002. Functional involvement of the Brahma/SWI2-related gene 1 protein in cytochrome P4501A1 transcription mediated by the aryl hydrocarbon receptor complex. J.Biol.Chem. 277: 11821-11827

219

13. Ma Q and Whitlock JPJ. 1996. The aromatic hydrocarbon receptor modulates the Hepa 1c1c7 cell cycle and differentiated state independently of dioxin. Mol.Cell Biol. 16: 2144- 2150

14. Roberts RA, Nebert DW, Hickman JA, Richburg JH, and Goldsworthy TL. 1997. Perturbation of the mitosis/apoptosis balance: a fundamental mechanism in toxicology. Fundam.Appl.Toxicol. 38: 107-115

15. Schwarz M, Buchmann A, Stinchcombe S, Kalkuhl A, and Bock K. 2000. Ah receptor ligands and tumor promotion: survival of neoplastic cells. Toxicol.Lett. 112-113: 69-77

16. Luebeck EG, Buchmann A, Stinchcombe S, Moolgavkar SH, and Schwarz M. 2000. Effects of 2,3,7,8-tetrachlorodibenzo-p-dioxin on initiation and promotion of GST-P- positive foci in rat liver: A quantitative analysis of experimental data using a stochastic model. Toxicol.Appl.Pharmacol. 167: 63-73

17. Buchmann A, Stinchcombe S, Korner W, Hagenmaier H, and Bock KW. 1994. Effects of 2,3,7,8-tetrachloro- and 1,2,3,4,6,7,8- heptachlorodibenzo-p-dioxin on the proliferation of preneoplastic liver cells in the rat. Carcinogenesis 15: 1143-1150

18. Davis BJ, Mccurdy EA, Miller BD, Lucier GW, and Tritscher AM. 2000. Ovarian tumors in rats induced by chronic 2,3,7,8-tetrachlorodibenzo-p-dioxin treatment. Cancer Res. 60: 5414-5419

19. Viluksela M, Bager Y, Tuomisto JT, Scheu G, Unkila M et al. 2000. Liver tumor- promoting activity of 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in TCDD-sensitive and TCDD-resistant rat strains. Cancer Res. 60: 6911-6920

20. Davis JW, Melendez K, Salas VM, Lauer FT, and Burchiel SW. 2000. 2,3,7,8- Tetrachlorodibenzo-p-dioxin (TCDD) inhibits growth factor withdrawal-induced apoptosis in the human mammary epithelial cell line, MCF-10A. Carcinogenesis 21: 881-886

21. Gonzalez FJ and Fernandez-Salguero P. 1998. The aryl hydrocarbon receptor: studies using the AHR-null mice. Drug Metab Dispos. 26: 1194-1198

22. Elizondo G, Fernandez-Salguero P, Sheikh MS, Kim GY, Fornace AJ et al. 2000. Altered cell cycle control at the G(2)/M phases in aryl hydrocarbon receptor-null embryo fibroblast. Mol.Pharmacol. 57: 1056-1063

23. Trimarchi JM and Lees JA. 2002. Sibling rivalry in the E2F family. Nat.Rev.Mol.Cell Biol. 3: 11-20

24. Dyson N. 1998. The regulation of E2F by pRB-family proteins. Genes Dev. 12: 2245-2262

25. Strobeck MW, Fribourg AF, Puga A, and Knudsen ES. 2000. Restoration of retinoblastoma mediated signaling to Cdk2 results in cell cycle arrest. Oncogene 19: 1857-1867

220

26. Johnson DG. 2000. The paradox of E2F1: oncogene and tumor suppressor gene. Mol.Carcinog. 27: 151-157

27. Yamasaki L, Jacks T, Bronson R, Goillot E, Harlow E, and Dyson NJ. 1996. Tumor induction and tissue atrophy in mice lacking E2F-1. Cell 85: 537-548

28. Field SJ, Tsai FY, Kuo F, Zubiaga AM, Kaelin WG, Jr. et al. 1996. E2F-1 functions in mice to promote apoptosis and suppress proliferation. Cell 85: 549-561

29. Pan H, Yin C, Dyson NJ, Harlow E, Yamasaki L, and Van Dyke T. 1998. Key roles for E2F1 in signaling p53-dependent apoptosis and in cell division within developing tumors. Mol.Cell 2: 283-292

30. Pierce AM, Schneider-Broussard R, Gimenez-Conti IB, Russell JL, Conti CJ, and Johnson DG. 1999. E2F1 has both oncogenic and tumor-suppressive properties in a transgenic model. Mol.Cell Biol. 19: 6408-6414

31. Fueyo J, Gomez-Manzano C, Yung WK, Liu TJ, Alemany R et al. 1998. Overexpression of E2F-1 in glioma triggers apoptosis and suppresses tumor growth in vitro and in vivo. Nat.Med. 4: 685-690

32. Conner EA, Lemmer ER, Omori M, Wirth PJ, Factor VM, and Thorgeirsson SS. 2000. Dual functions of E2F-1 in a transgenic mouse model of liver carcinogenesis. Oncogene 19: 5054-5062

33. Phillips AC and Vousden KH. 2001. E2F-1 induced apoptosis. Apoptosis. 6: 173-182

34. Adams PD and Kaelin WG, Jr. 1996. The cellular effects of E2F overexpression. Curr.Top.Microbiol.Immunol. 208: 79-93

35. Fan G, Ma X, Kren BT, and Steer CJ. 1996. The retinoblastoma gene product inhibits TGF-beta1 induced apoptosis in primary rat hepatocytes and human HuH-7 hepatoma cells. Oncogene 12: 1909-1919

36. Fan G, Ma X, Kren BT, and Steer CJ. 2002. Unbound E2F modulates TGF-beta1-induced apoptosis in HuH-7 cells. J.Cell Sci. 115: 3181-3191

37. Stiewe T and Putzer BM. 2001. p73 in apoptosis. Apoptosis. 6: 447-452

38. Irwin M, Marin MC, Phillips AC, Seelan RS, Smith DI et al. 2000. Role for the p53 homologue p73 in E2F-1-induced apoptosis. Nature 407: 645-648

39. Stiewe T and Putzer BM. 2000. Role of the p53-homologue p73 in E2F1-induced apoptosis. Nat.Genet. 26: 464-469

40. Lissy NA, Davis PK, Irwin M, Kaelin WG, and Dowdy SF. 2000. A common E2F-1 and p73 pathway mediates cell death induced by TCR activation. Nature 407: 642-645

221

41. Wu X and Levine AJ. 1994. p53 and E2F-1 cooperate to mediate apoptosis. Proc.Natl.Acad.Sci.USA 91: 3602-3606

42. Qin XQ, Livingston DM, Kaelin WG, Jr., and Adams PD. 1994. Deregulated transcription factor E2F-1 expression leads to S-phase entry and p53-mediated apoptosis. Proc.Natl.Acad.Sci.U.S.A 91: 10918-10922

43. Oren M. 1999. Regulation of the p53 tumor suppressor protein. J.Biol.Chem. 274: 36031- 36034

44. Parisi T, Pollice A, Di Cristofano A, Calabro V, and La Mantia G. 2002. Transcriptional regulation of the human tumor suppressor p14(ARF) by E2F1, E2F2, E2F3, and Sp1-like factors. Biochem.Biophys.Res.Commun. 291: 1138-1145

45. Zhu JW, DeRyckere D, Li FX, Wan YY, and DeGregori J. 1999. A role for E2F1 in the induction of ARF, p53, and apoptosis during thymic negative selection. Cell Growth Differ. 10: 829-838

46. Thatikunta P, Raj GV, Kundu M, Khalili K, and Amini S. 1997. The transcription factor E2F-1 modulates TGF-beta1 RNA expression in glial cells. Oncogene 14: 2959-2969

47. Moroni MC, Hickman ES, Lazzerini DE, Caprara G, Colli E et al. 2001. Apaf-1 is a transcriptional target for E2F and p53. Nat.Cell Biol. 3: 552-558

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Appendix Figure Legends

Fig. 1. AHR inhibits E2F1-induced apoptosis in transfected Saos-2 cells. Plasmids encoding

E2F1, AHR, and RB were transfected into Saos-2 cells grown to 50% confluence, along with a

plasmid for expression of GFP (green fluorescent protein). Cells were maintained in 5% FBS for

72 hours following transfection. Floating and adherent cells were then harvested and stained

with propidium iodide for flow cytometry analysis using a Beckman Coulter Epics XL flow

cytometer (Fullerton, CA). GFP-positive cells were analyzed for DNA content, and cells with

sub-G1 DNA content were counted as apoptotic. The results show that AHR can inhibit E2F1-

induced apoptosis at least as well as RB.

Fig. 2. Co-expression of the AHR inhibits E2F-1-overexpression-induced apoptosis in Soas-

2 cells. A. Adenovirus was generated for the expression of AHR, ARNT, RB, E2F1 and LacZ as

described in the Materials and Methods section of Chapter 4. Saos-2 cells were grown in 35-mm

dishes for 24 hours in normal serum media and for an additional 24 hours in 0.1% serum media.

Cells were then infected for 3 hours, washed 2 times with 1X PBS, and returned to 0.1% serum

media for an additional 72 hours. Infections were carried out at 100 PFU per cell for E2F1, and

200 PFU or 600 PFU per cell for LacZ. Following 72 hours, floating and attached cells were

collected and the percentage of dead cells for each group was counted using trypan blue

exclusion. B. Cells were infected as described in A with 100 PFU per cells for E2F1 and 300

PFU per cell for RB. Sevety-two hours after infections, dead cells were counted amongst the

floating and attached cell populations using trypan blue exclusion. C. Cells were infected as described in B, using 300 PFU per cell for both AHR and ARNT adenoviral stocks. Trypan blue

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exclusion was used to determined the percentage of viable cells at 72 hours post-infection. The

results show that expression of LacZ has no effect on E2F-dependent apoptosis in these cells,

while both RB and AHR show a similar ability of suppress apoptosis induced by E2F1

expression. In addition, co-expression of ARNT with AHR enhances the repression of E2F1-

induced apoptosis as compared to AHR alone.

Fig. 3. RB inhibits the expression of genes that are necessary for E2F-dependent apoptosis.

Saos-2 cells were infected with adenoviruses for the expression of E2F1 and RB as described in

Figure 2. At 24 hours post-infection, total cellular RNA was isolated and cDNAs were

synthesized as described in the Materials and Methods section of Chapters 3 and 4. Real-time

quantitative PCR was performed using a Smart Cycler rapid thermal cycler (Cepheid) and

fluorescence was measured after each of the repetitive cycles, and emission data were quantified

using threshold cycle (Ct) values. Ct values for all genes analyzed were averaged and normalized

to values for β-actin. The results show that RB inhibits the E2F-dependent induction of both

apoptosis genes (p73, Apaf1, p14Arf, Chk1) and cell cycle control genes (Cyclin E1).

Fig. 4. AHR inhibits the expression of genes that are necessary for E2F-dependent

apoptosis. Saos-2 cells were infected with adenoviruses for the expression of E2F1, AHR and

ARNT as described in Fig. 2. At 24 hours post-infection, total cellular RNA was isolated and

cDNAs were synthesized for analysis of E2F-dependent gene expression as described in Fig. 3.

The results show that the AHR inhibits E2F-dependent induction of pro-apoptotic gene expression at least as well as RB. The co-expression of E2F1 with ARNT in addition to AHR results in the reduction of expression of these genes to basal levels.

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Figure 1

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Figure 2

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Figure 3

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Figure 4

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