Modulation of Aryl Hydrocarbon Receptor-Dependent Transcription by Halogenated Compounds and Pharmaceuticals

Melanie Lynn Powis

A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Pharmacology and Toxicology University of Toronto

Melanie Lynn Powis

Master of Science

Department of Pharmacology and Toxicology University of Toronto

2011 Abstract

The aryl hydrocarbon receptor (AHR) mediates the toxic effects of halogenated aromatic hydrocarbons (HAHs), including 2,3,7,8-tetrachlorodibenzo-p-dioxin, 2,3,4,7,8- pentachlorodibenzofuran and 2,3,7,8-tetrachlorodibenzofuran. Y322 is believed to play a role in binding-independent activation of AHR by atypical inducers, such as omeprazole. I examined

AHR-mediated regulation of and coactivator recruitment to CYP1A1, CYP1B1, HES1 and

TiPARP in T-47D and HuH7 cells. All compounds induced expression of each gene in both cell lines, with some temporal differences between the HAHs and omeprazole. Chromatin immunoprecipitation assays demonstrated activator-, cell line- and gene-selectivity in AHR coactivator recruitment. Omeprazole induced AHR degradation which was prevented by MG-

132 pre-treatment. Y322 was found to be important for maximal AHR activation by 2,3,7,8-

TCDD and 2,3,4,7,8-PeCDF, but required for 2,3,7,8-TCDF and Omp in an AHR-deficient

MCF-7 cells. My findings provide further evidence for cell-, gene- and ligand-dependent differences in AHR-mediated gene expression and coactivator recruitment, and a role for Y322 in AHR activation.

Acknowledgments

I would like to acknowledge the following individuals for their contributions to both the completion of this thesis and my graduate experience as a whole:

• I would like to thank my supervisor, Dr. Jason Matthews, for the opportunity to be graduate student in his laboratory. I am truly grateful for his guidance and support, in addition to the opportunity to conduct interesting research while cultivating new laboratory skills.

• Secondly, I would like to thank my Master’s advisor, Dr. Peter McPherson, whose guidance and input were very helpful.

• I would like to thank the current and past members of the Matthews’ laboratory: Shaimaa Ahmed, Sarra Al-Saigh, Trine Celius, Raymond Lo and Laura MacPherson whose assistance and support were greatly appreciated. I truly enjoyed our discussions, both laboratory and otherwise.

• Next, I would like to thank the University of Toronto, Canadian Institute of Health Research, and Canadian Breast Cancer Foundation for financial support of this research.

• Finally, I would like to thank my family and friends for their continued encouragement and support during this experience.

iii

Table of Contents

Acknowledgments...... iii

Table of Contents...... iv

List of Tables ...... vii

List of Figures...... viii

List of Abbreviations ...... ix

1 Introduction...... 1

1.1 General Background...... 1

1.2 Structure of AHR ...... 1

1.3 AHR Signalling Pathway...... 3

1.4 Proposed Alternative AHR Signalling Pathway...... 5

1.5 Attenuation of AHR Signalling ...... 6

1.6 AHR Ligands...... 9

1.7 Naturally Occurring AHR Ligands...... 11

1.8 Atypical AHR Activators...... 11

1.9 AHR Coregulators...... 12

1.10 AHR Gene Battery...... 16

1.11 Physiological Role of AHR...... 17

1.12 AHR-mediated Adaptive and Toxic Response Pathways...... 19

1.13 Mutations to the LBD of AHR...... 21

2 Rationale and Objectives...... 23

2.1 Rationale ...... 23

2.2 Research Objectives...... 24

3 Materials...... 25

3.1 Chemicals...... 25

3.2 Plastic ware...... 26

3.3 Instruments...... 26

4 Methods...... 27

4.1 Cell Culture...... 27

4.2 RNA Extraction and qPCR ...... 28

4.3 Chromatin Immunoprecipitation (ChIP) Assays ...... 29

4.3.1 AHR Recruitment Experiments ...... 29

4.3.2 Coactivator Recruitment Experiments...... 31

4.4 Western Blot...... 31

4.4.1 Western Blot of Whole Cell Lysates ...... 31

4.4.2 Western Blot of Nuclear and Cytoplasmic Extracts ...... 32

4.5 Site-Directed Mutagenesis...... 33

4.6 Transformation...... 34

4.7 Transient Transfection...... 35

4.8 Statistical Analysis...... 35

5 Results...... 36

5.1 Ligand-, Cell Line- and Gene-Specific Differences in CYP1A1 and CYP1B1 mRNA Expression...... 36

5.2 Ligand- and Cell Line-Dependent AHR-Mediated HES1 and TiPARP mRNA Expression...... 37

5.3 Differential Recruitment of AHR to Response Elements in the Upstream Regulatory Regions of AHR Target Genes ...... 47

5.4 Ligand-induced Recruitment of Coactivators to the Enhancer AHREs of AHR Target Genes...... 52

5.5 Omeprazole Induced Nuclear AHR Degradation ...... 57

5.6 The Role of Y322 in Ligand-Induced AHR Signalling...... 60

6 Discussion, Conclusions, Limitations and Future Aims ...... 66

6.1 Discussion...... 66 v

6.1.1 Summary...... 66

6.1.2 Activator, Gene and Cell Line Differences in AHR-Mediated Expression of Target Genes ...... 67

6.1.3 Comparison of Activator-Induced AHR Recruitment to the Upstream Regulatory Regions of AHR Target Genes ...... 69

6.1.4 Activator-, Cell Line- and Gene-Dependent Differences in Coactivator Recruitment...... 70

6.1.5 Effect of Omp Treatment on AHR Degradation...... 72

6.1.6 Role of Y322 in Ligand-Mediated AHR Activation and Target Gene Expression...... 73

6.2 Conclusions...... 75

6.3 Limitations ...... 76

6.3.1 Time-points Chosen for mRNA and ChIP Experiments...... 76

6.3.2 ChIP Assay...... 76

6.3.3 Normalization of qPCR Data...... 77

6.3.4 Implications of Y322 Mutations on LBD Function...... 77

6.4 Further Aims ...... 78

References...... 79

List of Tables

Table 1. List of coregulators known to interact directly or indirectly with AHR and ARNT ...... 14

Table 2. qPCR Primers used for Analysis of cDNA...... 29

Table 3. qPCR Primers used for ChIP Assays...... 31

Table 4. Primers used for Site-Directed Mutagenesis...... 34

vii

List of Figures

Figure 1. Functional domains of the aryl hydrocarbon receptor (AHR)...... 3

Figure 2. Mechanism of activation of AHR by a ligand...... 7

Figure 3. Proposed Mechanism of AHR activation by atypical activators...... 8

Figure 4. Structures of selected AHR ligands and atypical AHR inducers...... 13

Figure 5. AHR-mediated CYP1A1 and CYP1B1 mRNA expression levels in T-47D cells...... 39

Figure 6. AHR-mediated CYP1A1 and CYP1B1 mRNA expression in HuH7 cells...... 41

Figure 7. AHR-mediated HES1 and TiPARP mRNA expression in T-47D cells...... 44

Figure 8. AHR-mediated HES1 and TiPARP mRNA expression in HuH7 cells...... 46

Figure 9. AHR recruitment to target gene AHREs is induced following 45min treatment...... 51

Figure 10. Recruitment of coactivators to AHREs of AHR-responsive genes following 45min treatment...... 56

Figure 11. Omeprazole treatment does not induce AHR degradation in whole cell lysates...... 58

Figure 12. Omeprazole treatment induces proteolytic degradation of AHR in nuclear extracts.. 59

Figure 13. Confirmation of variant expression in transiently transfected MCF-7 AHR100 cells.61

Figure 14. Y322 of AHR is required for maximal recruitment and CYP1A1 induction...... 62

Figure 15. Y322 of AHR is required for maximal recruitment and CYP1B1 induction...... 63

viii

List of Abbreviations

ACTR activator of thyroid and retinoic receptors

AHR aryl hydrocarbon receptor

AHRE AHR response elements

AHRR AHR repressor

AIB1 amplified in breast cancer 1

AIP AHR interacting protein

ALDH3A1 aldehyde dehydrogenase 3A1

AR androgen receptor

ARA9 aryl hydrocarbon receptor associated 9

ARA70 androgen receptor associated 70

ARNT aryl hydrocarbon receptor nuclear translocator

BAX Bcl-2-associated X protein bHLH/PAS basic-helix-loop-helix period ARNT single-minded transcriptional regulator

Brg-1 brahma related gene 1

BRAM-1 bone morphogenic protein receptor associated molecule 1

BRCA-1 breast cancer 1 protein, early onset

CARM-1 cofactor-associated arginine [R] methyltransferase

CBP CREB binding protein ix

CDK8 cyclin-dependent kinase 8

ChIP chromatin immunoprecipitation assay

CK2 casein kinase 2

CLOCK circadian locomotor output cycles kaput protein

CoCoA coiled-coil transcriptional coactivator

COUP-TF chicken ovalbumin upstream promoter-transcription factor

CREB cyclic adenosine monophosphate response element binding protein

CUL4B Cullin 4B protein

CYP1A1 cytochrome P450 1A1- human

CYP1A2 cytochrome P450 1A2- human

CYP1B1 cytochrome P450 1B1- human

DCC-FBS dextran-coated charcoal stripped fetal bovine serum

DIM 3,3’-diindolylmethane

DMEM Dulbecco’s Modified Eagle Media

DRE dioxin response elements

ER estrogen receptor

ERα estrogen receptor alpha

ERAP-140 estrogen receptor associated protein 140

FBS fetal bovine serum

GAC63 GRIP-1 associated coactivator 63

GRIP-1 glucocorticoid receptor interacting protein 1

HAH halogenated aromatic hydrocarbon hAHR human aryl hydrocarbon receptor

HES1 hairy and enhancer of split 1

HIF1α/2α hypoxia inducible factor 1α/2α

Hsp90 90kDa heat shock protein

IGFBP-1 insulin-like growth factor binding protein 1

I3C indole 3-carbinol

ICZ indolo[3,2-b]carbazole

IL-2 interleukin-2 mAHR mouse aryl hydrocarbon receptor

Med220 mediator protein 220

Mybbpla Myb binding protein 1a

NADPH nicotinamide adenine dinucleotide phosphate

NCoA1/2/3/4 nuclear receptor coactivator

Ned8 neural precursor cell developmentally down-regulated gene 8

NES nuclear export signal

NF1 nuclear factor 1

NFκB nuclear factor kappa B

NLS nuclear localization sequence

NQO1 NADPH quinine oxidoreductase I

Nrf-2 nuclear factor-like 2

Omp omeprazole ((RS)-6-methoxy-2-((4-methoxy-3,5-dimethylpyridin-2-yl) methylsulfinyl)-1H-benzo[d]imidazole) p160 160kDa family of nuclear receptor coactivator proteins p21 21kDa cyclin-dependent kinase 1 p23 23kDa heat shock protein 90-associated co-chaperone protein p27 27kDa cyclin-dependent kinase enzyme 1B p300 300kDa E1A binding protein; histone acetyl transferase activity

PAH polycyclic aromatic hydrocarbon

PCAF p300/ CREB binding protein associated factor

PCDD polycyclic dibenzo-p-dioxins

PCDF polycyclic dibenzofurans pCIP p300/CBP cointegrator proteins

2,3,4,7,8-PeCDF 2,3,4,7,8-pentachlorodibenzofuran

Per period circadian protein homolog1

PEST penicillin/ streptomycin

PKC protein kinase c

PML promyelocytic leukemia protein polII DNA polymerase II

PRMTI protein arginine methyltransferase I

xii

P-TEFb positive transcription elongation factor b

PXR pregnane X receptor rAHR rat aryl hydrocarbon receptor

RIP140 receptor interacting protein 140

Rb retinoblastoma protein

SHP small heterodimer partner

SIM1 single-minded homolog 1

SMRT silencing mediator for retinoid or thyroid-hormone receptors

SOB super optimal broth

SOC super optimal broth with catabolite repression

Sp1 specificity protein 1

SRC-1/2/3 steroid receptor coactivator 1/2/3

TAD transcriptional activation domains

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

2,3,7,8-TCDF 2,3,7,8-tetrachlorodibenzofuran

TAF2/4/6 transcription initiation factor TFIID subunit

TEF toxic equivalency factor

TEQ toxic equivalent

TFII B/D/F transcription factor II

TGF-β transforming growth factor beta

xiii

TIF2 transcriptional intermediary factor 2

TiPARP TCDD-inducible poly-(ADP-ribose) polymerase

TPR tetratricopeptide repeats

TR thyroid hormone receptor

TRAM1 thyroid hormone receptor molecule 1

TRAP220 thyroid hormone associated protein 220

TRegs regulatory T-cells

TRIP230 thyroid hormone receptor interactor protein 230

UBC9 ubiquitin-conjugated enzyme 9

UGT1A1/ A6 UDP-glucoronosyl transferase 1A1/ A6

XAP2 hepatitis B virus x-associated protein 2

xiv 1

1 Introduction 1.1 General Background

The aryl hydrocarbon receptor (AHR) is a ligand-activated member of the basic-helix-loop-helix Per (Period) ARNT (aryl hydrocarbon receptor nuclear translocator) SIM (single-minded) (bHLH/PAS) family of transcriptional regulators (Hankinson, 1995). Other members of the bHLH/PAS transcriptional regulator family include ARNT, hypoxia inducible factor 1α (HIF1α), single-minded homolog 1 protein (SIM1), circadian locomotor output cycles kaput protein (CLOCK), period circadian protein homolog1 (Per), and members of the p160 nuclear receptor coactivator family (Hankinson, 1995; Matthews et al., 2005; Soshilov and Denison, 2008). Treatment with aromatic hydrocarbons was first shown to induce a metabolic response in 1956, where benzo[a]pyrene treatment resulted in induction of aryl hydrocarbon hydroxylase activity (Conney et al., 1956; Schmidt and Bradfield, 1996; Snyder and Remmer, 1979). AHR was first identified in 1976 by Poland et al. who demonstrated binding of radio-labelled 2,3,7,8- tetrachlorodibenzo-para-dioxin (2,3,7,8-TCDD) to the receptor in mouse liver (Poland et al., 1976; Safe, 2001). AHR is able to bind many exogenous molecules, phytochemicals and pharmaceuticals; however, a definitive high-affinity endogenous ligand remains to be identified and, as such, AHR is considered to be an orphan receptor (Denison and Nagy, 2003). AHR mediates the toxic and carcinogenic effects of a variety of compounds, and environmental toxicants including halogenated aromatic hydrocarbons (HAHs), polychlorinated dibenzo-p- dioxins (PCDDs) and polychlorinated dibenzofurans (PCDFs). In addition to mediating toxicity, AHR has also been shown to play a significant role in both adaptive and developmental pathways (Schmidt and Bradfield, 1996; Stevens et al., 2009).

1.2 Structure of AHR

AHR contains several domains that are able to function independently (Gu et al., 2000; Hankinson, 1995, 2005). The amino-terminus of AHR contains a basic-helix-loop-helix (bHLH) motif (Figure 1.). This motif is well conserved across species and is characteristic of transcription factors that form homodimers or heterodimers, and are able to identify and bind to specific DNA sequences (Hankinson, 1995, 2005; Lees and Whitelaw, 1999; Nambu et al., 1991;

Soshilov and Denison, 2008). This domain is believed to be responsible for ligand-dependent nuclear localization of the receptor, as exposure of the nuclear localization sequence (NLS) within the bHLH domain results in translocation to the nucleus where the bHLH domain provides a platform for dimerization with ARNT (Ikuta et al., 1998; Lees and Whitelaw, 1999; Soshilov and Denison, 2008). AHR also possesses two Per ARNT Sim (PAS) domains, referred to as PAS A and PAS B which contain two copies of a 50-amino acid degenerate direct repeat sequence (Hankinson, 1995; Nambu et al., 1991; Soshilov and Denison, 2008). PAS A has been shown to be required for the formation of a stable AHR:ARNT heterodimer, while the bHLH and PAS B domains are the primary site of AHR dimerization for both the cytoplasmic chaperone 90kDa heat shock protein (Hsp90) and ARNT (Chapman-Smith et al., 2004; Coumailleau et al., 1995; Fukunaga et al., 1995). DNA binding activity occurs primarily through the basic domains of bHLH and PAS B (Hankinson, 2005). The PAS B domain of AHR contains the ligand binding domain (LBD), composed of several amino acids that are conserved across species (Coumailleau et al., 1995; Soshilov and Denison, 2008). PAS B has been shown to be essential for ligand-dependent activation of AHR, whereas previous studies have demonstrated that PAS B elicits an inhibitory effect on AHR activation and deletion of this domain resulted in a constitutively active AHR (Dolwick et al., 1993; Lindebro et al., 1995; McGuire et al., 2001). The carboxy-terminal of AHR contains the transcriptional activation domain (TAD) that is required for induction of target genes. The TAD includes three independently active sub- domains: the acidic sub-domain (a.a. 500-600), the glutamine (Q) rich sub-domain (a.a. 600-713) and the proline/ serine/ threonine (P/S/T) rich sub-domain (a.a. 713-848) (Hankinson, 2005; Ko et al., 1997; Kumar et al., 2001; Sogawa et al., 1995b). Based on other proteins containing multiple transactivating domains such as Sp1 and myogenin, the relative complexity of the TAD confers flexibility in transactivating potential (Beischlag et al., 2008). Each sub-domain was shown to possess low levels of transactivation individually; however, the additive activity of two or more of these domains resulted in initiation of reporter gene activity (Nguyen et al., 1999; Rowlands et al., 1996; Sogawa et al., 1995a). As is the case with AHR, the acidic and Q-rich domains of many transactivating proteins have been shown to directly interact with basal transcription factors (Gill et al., 1994; Weintraub et al., 1991). In a study by Kumar et al. (2001), it was demonstrated that the Q-rich sub-domain, particularly residues a.a. 663 to 688 were critical for transcriptional activation of AHR-responsive genes and required for in vitro recruitment of SRC-1 and RIP140 coactivator proteins (Kumar and Perdew, 1999; Kumar et al.,

2001; Kumar et al., 1999). While the P/S/T sub-domain was shown to have little function in transcriptional activation of AHR, previous studies have shown binding of coactivator proteins to both the acidic and P/S/T sub-domains (Kumar and Perdew, 1999; Kumar et al., 2001; Kumar et al., 1999). Jointly, these findings underscore the critical role of the Q-rich region of the AHR in mediating transactivation activity (Kumar et al., 1999).

1 848

bHLH Acidic Q-Rich P/S/T-Rich -COOH H2N- PAS A PAS B Domain Domain Domain Domain

27 39 230 397 500 600 713 848 DNA Binding Domain Ligand Binding Domain Transcriptional Activation Domain 40 79 121 289

ARNT Dimerization Domain

27 79 182 374

Hsp90 Binding Domain

Figure 1. Functional domains of the aryl hydrocarbon receptor (AHR). AHR contains an amino-terminal basic-helix-loop-helix (bHLH) domain, two Per-ARNT-SIM (PAS) domains referred to as PAS A and PAS B, and a carboxy-terminal transcriptional activation domain (TAD) composed of three sub-domains: the acidic domain, the glutamine (Q) rich domain, and the proline/ serine/ threonine (P/S/T) rich domain.

1.3 AHR Signalling Pathway

Prior to ligand binding, AHR resides in the cytoplasm bound to a chaperone protein complex, containing two molecules of Hsp90, AHR interacting protein (AIP also known as XAP2 and ARA9), and the co-chaperone protein, p23 (Carver et al., 1998; Denis et al., 1988; Perdew, 1988; Petrulis et al., 2003; Petrulis and Perdew, 2002). Hsp90 is a 90kDa protein which forms a hetero-complex with the transcriptionally inactive form of AHR. Displacement of Hsp90 is not

required for its translocation of the receptor to the nucleus but is required for AHR to bind to ARNT, since both Hsp90 and ARNT interact with the amino-terminal DNA binding HLH and PASB domains of AHR (Bradfield and Poland, 1988; Denis et al., 1988; Hankinson, 1995; Henry and Gasiewicz, 1993; Perdew, 1988). AIP is a 46kDa phosphorylated protein first identified in cross-linking studies by Perdew et al. (1992) to interact with AHR and Hsp90 through carboxy-terminal tetratricopeptide repeat (TPR) domains (Chen and Perdew, 1994; Coumailleau et al., 1995; Cox and Miller, 2002; Meyer and Perdew, 1999; Perdew, 1992; Whitelaw et al., 1995). Alanine substitution of the 4 most carboxy-terminal amino acids has been shown to prevent interaction with AHR but has no effect on Hsp90 binding to the receptor (Bell and Poland, 2000). To date, the functional role of AIP in AHR signalling remains unknown; however, studies suggest that AIP plays a role in cytoplasmic localization, repression of AHR transcriptional activity and reduction of proteasome-mediated degradation of the protein (Kazlauskas et al., 2000; Lees et al., 2003; Meyer and Perdew, 1999; Petrulis et al., 2003; Ramadoss et al., 2004). The phosphoprotein p23 is also referred to as the Hsp90-associated co- chaperone protein, and functions to interact with amino-terminal domains of Hsp90, stabilizing the ATP-bound conformation, preventing spontaneous dimerization with ARNT (Chadli et al., 2000; Felts and Toft, 2003; Kazlauskas et al., 1999; Kazlauskas et al., 2001). Presence of p23 is believed to enhance the release of ligand-bound AHR from Hsp90 and nuclear uptake of AHR by increasing the ability of the receptor to recognize the importin-β protein (Beischlag et al., 2008; Kazlauskas et al., 1999).

AHR is a promiscuous receptor as it exhibits profound ligand diversity, binding not only halogenated aromatic hydrocarbons (HAHs) and polycyclic aromatic hydrocarbons (PAHs), but phytochemicals, and clinically significant pharmacological agents; however, a definitive endogenous ligand has not been conclusively identified (Poland and Knutson, 1982; Prud'homme et al., 2010; Savouret et al., 2001; Song and Pollenz, 2002). Following binding within the LBD by a ligand, such as 2,3,7,8-tetrachlorodibenzo-p-dioxin (2,3,7,8-TCDD), AHR undergoes a conformational change to expose the NLS, resulting in binding of the importin-β protein and translocation of the complex to the nucleus (Figure 2.) (Hord and Perdew, 1994; Ikuta et al., 1998; Lees and Whitelaw, 1999; Okey et al., 1980; Pollenz et al., 1994; Safe, 2001). Once in the nucleus, AHR dissociates from the chaperone complex and binds to its obligatory dimerization partner, ARNT, turning it into its high affinity DNA-binding form (Denison and Nagy, 2003;

Hankinson, 1995, 2005; Probst et al., 1993). The AHR:ARNT heterodimer then binds to upstream genomic enhancer elements, referred to as aryl hydrocarbon or dioxin response elements (AHREs or DREs), containing the 5`-GCGTG-3` penta-nucleotide core (Denison et al., 1998; Safe, 2001; Whitlock, 1999). DNA binding induces the recruitment of coregulator proteins (coactivators and corepressors) resulting in changes in the expression of AHR- responsive genes (Hankinson, 2005). The phosphorylation of serine and tyrosine residues of the AHR is important for DNA-binding and maximal transactivation, whereas treatment with tyrosine kinase inhibitors reduce AHR-dependent regulation of CYP1A1 (Chen and Tukey, 1996; Pongratz et al., 1991). AHR-target genes include the drug-metabolizing enzymes cytochrome P4501A1 (CYP1A1) and CYP1B1, hairy and enhancer of split 1 (HES1), and the TCDD- inducible poly(ADP-ribose) polymerase (TiPARP) (Hankinson, 2005; Krusekopf et al., 2003).

1.4 Proposed Alternative AHR Signalling Pathway

Atypical activators, such as the substituted benzimidazole (RS)-6-methoxy-2-((4-methoxy-3,5- dimethylpyridin-2-yl) methylsulfinyl)-1H-benzo[d]imidazole (omeprazole, Omp), have been shown to induce AHR-dependent gene activation in a binding-independent manner (Denison and Nagy, 2003; Quattrochi and Tukey, 1993). A second mechanism, not completely distinct from the typical mechanism of AHR activation, is proposed to be mediated by phosphorylation of tyrosine within the LBD by a c-src tyrosine kinase independent of ligand binding (Figure 3.) (Backlund and Ingelman-Sundberg, 2005). Previous studies have demonstrated that, Omp- induced but not TCDD-induced AHR activation was inhibited in cells co-treated with TCDD or Omp and a tyrosine kinase inhibitor (Backlund and Ingelman-Sundberg, 2005; Lemaire et al., 2004). This supported the view that there is a mechanistic difference in AHR activation by typical and atypical ligands, such that tyrosine-kinase mediated activation is utilized by atypical AHR activators. The presence of a secondary mechanism of AHR activation remains disputed since atypical inducers activate the same AHR-dependent genes as binding-dependent ligands, but tend to be weak inducers of AHR that cannot out-compete the high affinity binding of [3H]- TCDD in binding assays (Backlund et al., 1999; Bradfield and Poland, 1988; Daujat et al., 1996; Daujat et al., 1992; Diliberto et al., 2001). Recent studies utilizing lower [3H]-TCDD concentrations and increased concentrations of the weak competitor have revealed that many

6 molecules that were previously reported to activate AHR indirectly, such as carbaryl, bind to the receptor (Denison and Nagy, 2003; Denison et al., 1998; Phelan et al., 1998b). As such, it is believed that many atypical binding-independent AHR inducers are actually weakly binding AHR ligands (Denison and Nagy, 2003; Phelan et al., 1998a; Phelan et al., 1998b)

1.5 Attenuation of AHR Signalling

Since the HAHs and PAHs that induce AHR signalling have long half-lives and are thus not readily degraded or eliminated by an organism, a mechanism for attenuation of AHR is crucial (Pollenz, 2002). Previous pulse-chase experiments by Ma and Baldwin (2000) have shown that exposure of mouse hepatoma cells to 2,3,7,8-TCDD results in a decrease in AHR half-life from 28h to 3h (Ma and Baldwin, 2000; Pollenz, 2002). Down-regulation of AHR signalling prevents chronic reactivation by stable ligands and prevents the generation of reactive oxygen species, and is achieved by either proteolytic degradation or transcriptional repression by the AHR repressor protein (AHRR). Following ligand-induced AHR signalling, AHR dissociates from the AHRE and ARNT exposing the nuclear export signal (NES) contained in the second helix domain between a.a. 62-73, resulting in the export of AHR from the nucleus to the cytoplasm (Burbach et al., 1992; Davarinos and Pollenz, 1999; Pollenz, 2002). Inhibition of nuclear export by co- treatment with 2,3,7,8-TCDD and leptomycin B has been shown to prevent AHR down- regulation (Davarinos and Pollenz, 1999; Ma and Baldwin, 2000). Once in the cytoplasm, AHR is tagged for degradation through the covalent addition of multiple 76 a.a. ubiquitin molecules, catalyzed by a multi-enzyme group containing the ubiquitin-activating enzyme (E1), the ubiquitin-conjugating enzyme (E2), and ubiquitin ligase (E3) (Ma and Baldwin, 2000). Following ubiquitination, AHR is degraded by the 26S proteasome (Davarinos and Pollenz, 1999; Ma and Baldwin, 2000; Pollenz et al., 2005; Roberts and Whitelaw, 1999). AHR degradation following treatment with prototypical AHR activators, such as 2,3,7,8-TCDD, is well-documented and may be inhibited by 1h pre-treatment with the selective 26S inhibitor MG- 132; however, degradation following treatment with atypical inducers remains unstudied (Ma and Baldwin, 2000; Pansoy et al., 2010; Pollenz, 2002, 2007). The second mechanism for attenuation of AHR signalling involves up-regulation of the AHRR in response to AHR activation (Haarmann-Stemmann et al., 2007; Mimura et al., 1999).

Typical Ligand

CYTOPLASM p23 Hsp90 AIP AHR Proteolytic Degradation (26S)

NUCLEUS AHR ARNT

p23 Hsp90 NCoA3 Target Genes AIP NCoA1 p300 AHR ARNT ie: CYP1A1

pol II AHRR AHRE

Figure 2. Mechanism of activation of AHR by a ligand. Prior to activation, AHR exists in the cytoplasm bound to its chaperone complex. AHR ligands, such as 2,3,7,8-TCDD, bind to the ligand binding domain of the receptor inducing a conformational change which exposes the nuclear localization sequence, resulting in the translocation of AHR to the nucleus. Once in the nucleus, AHR dissociates from its chaperone complex and heterodimerizes with ARNT. The heterodimer is then recruited to AHREs within the upstream enhancer regions of AHR- responsive genes, initiating coactivator recruitment and transcription. Signalling is attenuated either (1) by sequestration of available pools of ARNT by AHRR, or (2) by degradation of AHR through the 26S proteasome pathway.

Atypical Ligand p23 Hsp90 CYTOPLASM AIP AHR Kinase P

Proteolytic Degradation (26S) (?)

NUCLEUS AHR ARNT

p23 Hsp90 P NCoA3 AIP Target Genes NCoA1 p300 AHR ARNT ie: CYP1A1

pol II AHRR (?) AHRE

Figure 3. Proposed Mechanism of AHR activation by atypical activators. The inactive form of AHR exists within the cytoplasm bound to a chaperone complex. Treatment with an atypical AHR inducer, such as Omp, results in activation of a c-src tyrosine kinase. This results in phosphorylation of a tyrosine residue within the ligand binding domain of AHR, triggering a conformational change to expose the nuclear localization sequence. AHR then translocates to the nucleus where it dissociates from its chaperone complex and heterodimerizes with ARNT. The heterodimer then binds to AHREs in the enhancer regions of target genes, inducing recruitment of coactivators and transcriptional upregulation. While it is postulated that signalling through this alternative pathway would also be attenuated by AHRR and proteolytic degradation by 26S, these mechanisms have not been previously studied.

While AHRR is similar to AHR in that the bHLH and PAS A domains have a high level of identity, residues associated with ligand binding in the PAS B domain are extremely divergent and AHRR is not capable of binding to ligands. AHRR transcriptional upregulation occurs in response to treatment with AHR ligands, such as 3-methylchloanthrene (Baba et al., 2001; Mimura et al., 1999). AHRR acts as a negative regulator of AHR activity by out-competing it for dimerization with ARNT, blocking AHR mediated transcription initiation at AHREs (Hahn, 2002; Hosoya et al., 2008; Mimura et al., 1999).

1.6 AHR Ligands

AHR ligands include halogenated aromatic hydrocarbons (HAHs) such as poly-halogenated dibenzo-p-dioxins, dibenzofurans, and biphenyls, and polycyclic aromatic hydrocarbons (PAHs) including 3-methylcholanthrene, benzo(a)pyrene, benzanthracenes, and benzoflavones, phytochemicals, and clinically significant pharmacological agents (Denison et al., 1998; Poland and Knutson, 1982; Prud'homme et al., 2010; Savouret et al., 2001; Song and Pollenz, 2002). Since none of the identified ligands are normally occurring endogenous molecules or intermediate metabolites, a definitive endogenous ligand has yet to be determined (Hankinson, 1995). AHR mediates the toxic effects of its ligands which tend to be hydrophobic and planar or able to become coplanar, and include many environmental contaminants that are byproducts of the synthesis of various organochlorines, chlorine bleaching of wood pulp, during municipal, hospital and industrial waste incineration, metal production, and fossil fuel or wood burning (Denison and Nagy, 2003; DeVito et al., 1994; Hankinson, 1995; Kulkarni et al., 2008; Safe, 1990; Safe, 1986; Van den Berg et al., 1998). The most potent class of AHR activators is the most metabolically stable and are HAHs, including 2,3,7,8-TCDD, which are accommodated within the rectangular 3x10Ǻ LBD of AHR (Denison and Nagy, 2003; Hankinson, 1995; Poland and Knutson, 1982). 2,3,7,8-TCDD is known to accumulate in adipose tissue due to its high lipophilicity, and undergoes little hepatic metabolism (Diliberto et al., 2001; van der Molen et al., 1996). 2,3,7,8-TCDD binds to AHR with an affinity in to pM to nM range, while PCDFs, such as 2,3,4,7,8-pentachlorodibenzofuran (2,3,4,7,8-PeCDF) and 2,3,7,8- tetrachlorodibenzofuran (2,3,7,8-TCDF), exhibit lower binding affinities (Denison and Nagy, 2003). The structures of these ligands are provided in Figure 4.

It has been postulated that binding affinities to AHR have a direct correlation to electrochemical and thermodynamic properties of the ligands (Denison and Nagy, 2003; Waller and McKinney, 1995). The relative potency of a specific environmental toxicant is referred to as the toxic equivalency factor (TEF), while combined toxicity of mixtures of these compounds is referred to as the toxic equivalent (TEQ) (Van den Berg et al., 1998). TEFs were developed to assist in regulatory reviews and risk assessments of various compounds by comparing the toxic responses induced by exposure to these compounds, including dermal toxicity, immuno-toxicity, carcinogenicity, and adverse effects on development, reproduction and endocrine function (Van den Berg et al., 1998). Values are based on a compilation of both in vivo and in vitro data for compounds that bind to AHR and are expressed relative to the high-affinity prototypical AHR ligand TCDD (Van den Berg et al., 1998). In cases where the potency of a compound is determined from a single study, it is referred to as the relative potency value instead of a TEF (Van den Berg et al., 1998). In 2005, the World Health Organization reevaluated these values, based on new information from blood studies, and exposure data from studies examining household dust and soil contamination with dioxin-like compounds (Hong et al., 2009). For a compound to qualify for evaluation for a TEF value it must fulfill four basic requirements, (1) demonstrate a structural relationship to PCDDs or PCDFs, (2) be able to bind to AHR, (3) elicit AHR-mediated toxic and biochemical responses, and (4) be prevalent and accumulate in the food chain (Van den Berg et al., 1998). Based the established relative potencies of these compounds, 2,3,7,8-TCDD has a TEF of 1.0, while 2,3,4,7,8-PeCDF is approximately 1/3 as potent (TEF 0.3), and 2,3,7,8-TCDF is approximately 1/10 as potent (TEF 0.1) as 2,3,7,8-TCDD (Hong et al., 2009; Van den Berg et al., 1998). Since these TEF values are calculated based on data from a large number of studies, the predictive validity of the relative potencies in individual studies is unknown. A previous study by Zhang et al. (2008) demonstrated a correlation between binding affinity and induction of toxic response, whereby GAL4-coactivator assays comparing ligand treatments in Panc1, HEK293, and Hepa1c1c7 demonstrated that TEF may not be directly predictive of recruitment to and induction of AHR target genes in response to treatment with these ligands (Safe, 1986; Schecter et al., 2006; Stevens et al., 2009; Zhang et al., 2008).

1.7 Naturally Occurring AHR Ligands

There are a number of natural and synthetic dietary compounds that have also been shown to bind to AHR. Some plant compounds have been shown to be natural AHR inducers, such as curcumin and carotinoids (Ciolino et al., 1998; Denison and Nagy, 2003; Gradelet et al., 1996). Dietary indoles are generally derived from tryptophan and metabolized to more potent AHR inducers in the mammalian digestive tract, though most dietary ligands are only weak agonists (Denison and Nagy, 2003). The fact that tryptophan is a strong absorber of near-UV light led several laboratories to investigate the induction of CYP1A1 by tryptophan photo-products (Helferich and Denison, 1991; Rannug et al., 1987). These studies allowed researchers to identify several photo-products, particularly histidine oxidation products, that competitively bind to AHR and activate expression of AHR-dependent genes (Paine, 1976; Paine and Francis, 1980; Rannug et al., 1987). A specific example of this is activation of AHR by exposure to indole-3- carbinol (I3C), a compound found in cruciferous vegetables, that is hydrolyzed into indolo[3,2- b]carbazole (ICZ) and 3,3’-diindolylmethane (DIM) that can act as AHR agonists (Grose and Bjeldanes, 1992; Jellinck et al., 1993). ICZ is the most potent naturally occurring AHR activator and has been shown to induce AHR-mediated expression of target genes (Bjeldanes et al., 1991; Denison and Nagy, 2003). Flavanoids, such as flavones and flavanols, are present in vegetables, fruits and teas have been shown to be present in high enough circulating concentrations to antagonize AHR (Denison and Nagy, 2003). Resveratrol is another well-known AHR antagonist that is found in wine and grapes (Revel et al., 2001; Schneider et al., 2001).

1.8 Atypical AHR Activators

Unlike the HAHs, the proton pump inhibitor Omeprazole (Omp) is used as a pharmaceutical agent to treat acid reflux and is a member of the substituted benzimidazole family of chemicals (Howden et al., 1984; Quattrochi and Tukey, 1993). Omp is an AHR activator in that has been shown to induce AHR-mediated phase I metabolizing enzyme expression (Krusekopf et al., 2003; Quattrochi and Tukey, 1993; Yoshinari et al., 2008). However, it is considered to be an atypical AHR inducer as it is thought to activate AHR indirectly, and has been shown to be unable to competitively bind to AHR in [3H]-TCDD binding assays (Backlund and Ingelman- Sundberg, 2004; Denison and Nagy, 2003). Omp is believed to activate AHR by kinase

mediated tyrosine phosphorylation. Co-treatment with tyrosine kinase inhibitors was shown to have no effect on TCDD-induced AHR activation, but completely blocked the effects of Omp (Backlund and Ingelman-Sundberg, 2005; Lemaire et al., 2004). Co-treatment with tyrosine kinase inhibitors, lavendustin or genistein, was shown to have no effect, while co-treatment with the c-src specific tyrosine kinase inhibitor, herbimycin B inhibited Omp activation of AHR, indicating that it is a c-src tyrosine kinase-mediated effect. However, the indirect activation of AHR by Omp is somewhat controversial, since many atypical activators that were previously reported to activate AHR through this mechanism have actually been shown to weakly bind to the receptor in new [3H]-TCDD binding assays which utilize lower concentrations of TCDD and higher concentrations of the competitor (Backlund and Ingelman-Sundberg, 2005; Denison and Nagy, 2003).

1.9 AHR Coregulators

Binding of the AHR:ARNT heterodimer to AHREs initiates the recruitment of coactivators and basal transcriptional machinery to the regulatory regions of the AHR target genes. The acidic and Q-rich sub-domains of the carboxy-terminal TAD domain of AHR have been shown to interact directly with coregulators and transcription factors, while ARNT has also been shown to be involved in cofactor recruitment to the regulatory regions of target genes (Hestermann and Brown, 2003; Kumar et al., 1999; Matthews et al., 2005). Coactivators exist as components of multi-subunit complexes, and play various roles in transcriptional activity, including remodeling, relocating and dissociating nucleosomes to change chromatin structure to de-repress transcription, in addition to inducing the recruitment of RNA polymerase II and other basal transcription factors to the promoter region (Felsenfeld and Groudine, 2003; Goll and Bestor, 2002; Kraus and Wong, 2002). A list of coregulators that are known to interact with AHR or play a role in the transcription of AHR-responsive genes is provided in Table 1. The spectrum of coregulators required for the transcriptional activation differs greatly among genes (Hankinson, 2005). A previous study by Zhang et al. (2008) compared the ligand- and cell line- dependent effects on CYP1A1 induction, as well as activation and coactivator recruitment

AHR LIGANDS- Halogenated Aromatic Hydrocarbons (HAHs)

Cl O Cl Cl O Cl

Cl O Cl Cl Cl

2,3,7,8-tetrachlorodibenzo-para-dioxin 2,3,4,7,8- pentachlorodibenzofuran (2,3,7,8-TCDD) (2,3,4,7,8-PeCDF)

Cl O

Cl Cl

2,3,7,8- tetrachlorodibenzofuran (2,3,7,8-TCDF)

ATYPICAL AHR INDUCERS-

OCH3

H3C CH3

N N S O NH

CH3O

(RS)-6-methoxy-2-((4-methoxy-3,5-dimethylpyridin-2-yl) O(Opmethylsulfinyl)-1H-benzo[d]imidazole Omeprazole (Omp)

Figure 4. Structures of selected AHR ligands and atypical AHR inducers.

(NCoA1, CARM-1, TRAP220 and TIFII) in a Gal4 reporter system in Panc1, HEK 293, and Hepa1c1c7 cells (Zhang et al., 2008). They determined that each ligand induced CYP1A1 expression to a different extent in each cell line studied (Zhang et al., 2008). In addition, coactivator recruitment profiles differed greatly among ligands, as well as among cell lines (Zhang et al., 2008). There is a significant amount of functional redundancy between coactivators, whereby several different coactivators can be substituted for one another, and recruitment of these factors can be both cell line and gene specific (Hankinson, 2005; Kadonaga, 2004; Khorasanizadeh, 2004; Zhang et al., 2008). However, the expression of different coactivator recruitment profiles in different tissues supports the concept of selective aryl hydrocarbon receptor modulators where they are active, in certain tissues due to specific coactivator expression (Hankinson, 2005).

Table 1. List of coregulators known to interact directly or indirectly with AHR and ARNT

Name Function Reference CBP/p300 NCoA1/ SRC-1 NCoA2/ SRC-2/ (Beischlag et al., 2002; Hankinson, 2005; Harper GRIP1/ TIF2 Histone acetyl transferase (HAT) et al., 2006; Ma et al., NCoA3/SRC-3/ activity; transcriptional activation pCIP/ 2001; Zhang et al., 2008) AIB1/ACTR/TRAM1 NCoA4/ARA70 NCoA7/ERAP-140 Brg-1/ BRAM-1 Med220 (Wang et al., 2004a; Wang Chromatin remodelling activity; enhance PRMT1 et al., 2007; Wang et al., transcriptional activation of target genes CARM-1 2004b) CDK8 TRIP230 Enhance expression of AHR-responsive (Chen et al., 2006; Ge and CoCoA genes; specific function unknown Elferink, 1998; Harper et GAC63 al., 2006; Jones et al., 2002; Kim and Stallcup, NCoA4 2004; Kollara and Brown, BRCA1 2006) Rb Mybbp1a

PML Ned8 RIP140 SMRT (Carlson and Perdew, SHP Transcriptional repression; inhibits 2002; Gradin et al., 1999; AHRR AHR:ARNT induced gene expression Mimura et al., 1999; ARNT repressor Nguyen et al., 1999) P-TEFb Elongation factor (Tian et al., 2003) ER/ ERα AR (Carlson and Perdew, TR 2002; Chan et al., 1999; PKC AHR is believed to interact indirectly Chen and Tukey, 1996; Tyrosine kinases and with other steroid hormone receptor/ Gradin et al., 1996; Kolluri phosphatases signal transduction pathways through et al., 1999; Okino et al., TGF-β “cross-talk” 1992; Park et al., 2000; CK2 Porterfield, 2000; Reiners C2-ceramide and Clift, 1999; Safe et al., c-myc 1998; Zaher et al., 1998; p27 Zhang et al., 2008) TRAP220 HIF-1a Hsp90 (Carlson and Perdew, AHR complex proteins in the cytoplasm; p23 2002; Cox and Miller, help to modulate AHR activation by 2002; Meyer et al., 1998; ligands AIP Perdew, 1992) TFIIB TFIID TFIIF (Beischlag et al., 2008; Transcription factors known to be TAF4 Carlson and Perdew, 2002; recruited to promoter regions of AHR- TAF6 Hankinson, 2005; Ricci et responsive genes NF-1 al., 1999; Watt et al., 2005) Sp1 COUP-TF1 NF-κB UBC9 Sumo E3 Ligase activity (Beischlag et al., 2008) CUL4B

1.10 AHR Gene Battery

AHR signalling plays an important role in three distinct pathways: (1) development and differentiation, (2) adaptive metabolism, and (3) toxicity (Schmidt and Bradfield, 1996). It is responsible for inducing the transcriptional activation of a battery of physiological important genes, including enzymes responsible for metabolic transformation and excretion of xenobiotics (Hankinson, 2005; McMillan and Bradfield, 2007; Nebert et al., 2004; Puga et al., 2009; Uno et al., 2004; Xu et al., 2005). The most studied AHR-responsive gene is the phase I xenobiotic metabolizing enzyme cytochrome P450 1A1 (CYP1A1) (Androutsopoulos et al., 2009; Beischlag et al., 2008; Hankinson, 1995)Neuhold et al., 1989). Other AHR target genes include CYP1A2, CYP1B1, aldehyde dehydrogenase (ALDH3A1), NADPH quinine oxidoreductase (NQO1), UDP-glucoronosyl transferase (UGT1A6/ A1), nuclear factor-like 2 (Nrf2), Nrf2 nuclear factor (erythroid-derived 2)-like 2, and glutathione S-transferase A1 (GSTA1) (Bock and Kohle, 2006; Nebert et al., 2004; Okey et al., 2005). The hairy and enhancer of split 1 (HES1) is an oscillatory immediate early gene that has been shown to be a target of 2,3,7,8-TCDD-induced AHR-mediated activation (Thomsen, 2003). HES1 is a Notch effector protein that plays a role in neuronal differentiation and cell proliferation (Thomsen and Helboe, 2003; Thomsen et al., 2004; Yoshiura et al., 2007). TCDD-inducible poly-(ADP-ribose) polymerase (TiPARP) is an immediate early gene whose expression has been previously shown to be induced by 2,3,7,8- TCDD treatment (Frasor et al., 2004; Ma, 2002). TiPARP modulates the function of target proteins through the addition of ADP-ribose onto glutamic acid residues (Ma, 2002; Smith, 2001). Previous studies by Ma (2002) demonstrated that TiPARP is super-induced when proteosomal degradation of AHR is inhibited with MG-132 pre-treatment. Other genes that are known to be ligand-mediated AHR-responsive genes play a role in cell proliferation and differentiation, and include: cyclin-dependent kinase inhibitors (p21 and p27), transcription factors (c-jun and junD), interleukin-2 (IL-2), Bcl2-associated X protein (BAX), insulin-like growth factor binding protein 1 (IGFBP-1), Filaggrin, and ecto-ATPase (Bock and Kohle, 2006; Gao et al., 1998; Hoffer et al., 1996; Jeon and Esser, 2000; Kolluri et al., 1999; Thomsen et al., 2004).

1.11 Physiological Role of AHR

While an endogenous AHR ligand is yet to be definitively identified, AHR has been shown to fulfill many essential physiological roles, and is highly conserved among species. Mutants that are unable to bind ligand are still able to heterodimerize with ARNT and induce transcription, indicating that the ability of AHR to mediate aryl hydrocarbon exposure is a recent adaptation (Hahn, 2002). In an attempt to elucidate the physiological role of AHR, several studies have looked at the developmental implications of knocking out AHR expression (Fernandez-Salguero et al., 1995; Hushka et al., 1998; Peters et al., 1999; Pollenz, 2007; Schmidt et al., 1996). Analysis of mRNA expression has shown that AHR was expressed in the 8 different types of human tissues examined, with the highest levels of expression in placenta and lung, and the lowest levels in kidney, skeletal muscle and brain (Dolwick et al., 1993; Schmidt and Bradfield, 1996). AHR knockout studies in mice have demonstrated the implications of AHR deficiency in the closure of the ductus venosus, liver function, immune system development and ligand- dependent cellular activities (Fernandez-Salguero et al., 1995; Tijet et al., 2006; Walisser et al., 2004). In particular, studies of AHR null mice have revealed microvesicular steatosis in perinatal hepatocytes, prolonged extramedullary hematopoiesis and reduced liver size (Bunger et al., 2008). Studies demonstrating failure to close the ductus venosus, cardiac hypertrophy, hypertension, and elevated levels of vasoconstrictors in the blood of null mice suggest that AHR plays a role in vasculature and hematopoiesis in mammalian development (Bunger et al., 2008; Lund et al., 2003; Lundy et al., 2003; Stevens et al., 2009; Walisser et al., 2005). AHR is believed to play a role in the differentiation of skin since AHR-null mice develop follicular epidermal hyperplasia with hyperkeratosis and acanthosis, dermal fibrosis and anagenic hair follicles (Bock and Kohle, 2006; Fernandez-Salguero et al., 1995). Knockout mice also exhibited a myriad of phenotypic abnormalities, including growth defects, immune system impairment, cardiac hypertrophy, skin lesions, lower incidence of large infrontal bones, liver fibrosis and lipid accumulation, and hepatic defects (Abbott et al., 1999a; Abbott et al., 1999b; Baba et al., 2005; Fernandez-Salguero et al., 1995; Lund et al., 2006; Pollenz, 2007; Schmidt et al., 1996; Walisser et al., 2004; Walisser et al., 2005). Studies of AHR deficient mice have demonstrated a correlation between AHR and estrogen signalling whereby studies of female AHR null mice have not been shown to possess upstream changes in endocrine regulation of ovulatory-stimulating hormones, but instead exhibit insufficient follicular synthesis of estradiol

(Baba et al., 2005; Barnett et al., 2007; McMillan and Bradfield, 2007; Trewin et al., 2007). These female mice exhibit reproductive deficits including a reduction in the number of mature follicles, difficulties with conception, reduced litter number, reduced live birth size, poor fetal and offspring viability, and impaired mammary gland and ovary development (Abbott et al., 1999b; Baba et al., 2005).

AHR has been found to play a critical role in neuronal development, whereas it is expressed in a variety of brain regions during the crucial stages of development and over- expression results in neural differentiation (Abbott and Probst, 1995; Akahoshi et al., 2006; Huang et al., 2000; Kainu et al., 1995). Neuronal development deficiencies have also been reported in AHR null mice, whereby AHR deficiency resulted in abnormal granule neuron differentiation and undersized cerebellum, indicating that AHR plays a role in neural precursor cell proliferation (Collins et al., 2008; Latchney et al., 2011; Williamson et al., 2005). In addition, AHR has recently been shown to play a role the development of T-helper 17 and regulatory T-cells (Tregs) (Stevens et al., 2009). Naïve T-cells from AHR null mice have been shown to be ineffective at generating Tregs in vitro, which are responsible for reducing auto- immune and allergic diseases, and inhibiting anti-tumourigenic immune responses (Kimura et al., 2008; Mottet and Golshayan, 2007; Stevens et al., 2009). AHR is believed to modulate the differentiation of these Tregs by modulating expression of markers and through cross-talk with the TGF-β signalling (Stevens et al., 2009). AHR activation modifies the balance of Tregs to T- helper 17 cells by regulating the expression of the pro-inflammatory cytokines, since cross-talk with TGF-β induces Treg differentiation and T-helper 17 cell production (Larosa and Orange, 2008; Mottet and Golshayan, 2007; Stevens et al., 2009; Veldhoen et al., 2008). More recently, AHR has been shown to bind to the regulatory regions of Bach2, a protein that has been shown to repress B-cell differentiation. 2,3,7,8-TCDD treatment has been shown to suppress B-cell differentiation to immunoglobulin M secreting plasma cells (De Abrew et al., 2011; Holsapple et al., 1986; Morris et al., 1993; Sulentic et al., 1998).

Recent studies in AHR knock-out mice have demonstrated altered circulated red and white blood cells as well as a two-fold increase in the number of enriched hematopoietic stem cells in bone marrow (Singh et al., 2010). In agreement with these findings, another laboratory has demonstrated that inhibition of AHR results in CD34+ cell expansion, indicating that AHR plays a significant role in maintaining quiescence of hematopoietic precursors by negatively

regulating proliferation (Boitano et al., 2010; Singh et al., 2010). Taken together, these studies demonstrate that AHR may be advantageous in stem cell maintenance, as it functions to preserve long-term functionality of the stem cells, preventing premature exhaustion (Singh et al., 2010). Conversely, expression of constitutively active AHR in mice resulted in stomach tumours and a reduction in life span (Ma and Whitlock, 1996; Pollenz, 2007; Pollenz et al., 1998; Andersson et al., 2002; McGuire et al., 2001).

1.12 AHR-mediated Adaptive and Toxic Response Pathways

AHR is known to mediate most of the diverse and organ-specific toxic effects of environmental toxicants such as HAHs and PAHs (Bock and Kohle, 2006). It is postulated that the original role of AHR may have been to regulate a gene associated with differentiation or development; however, at some point it evolved the ability to regulate the transcription of biotransformation enzymes (Hahn, 2002). This system likely evolved to minimize the burden of extended polycyclic aromatic compounds on the body by increasing phase I xenobiotic metabolizing enzymes that hydroxylate the compound to increase its solubility in water and decrease its half- life (Conney, 1982; Schmidt and Bradfield, 1996; Stevens et al., 2009). This pathway is considered to be an example of adaptive metabolism since xenobiotics bind to cytosolic AHR to activate transcription of enzymes that lead to their eventual biotransformation as excretion (Schmidt and Bradfield, 1996; Stevens et al., 2009). Selective pressure for the evolution of this pathway is driven by the deleterious effects of these compounds on cellular processes (Schmidt and Bradfield, 1996). A major complication of this system exists, whereby PAHs can be metabolized to electrophilic metabolites which result in genotoxicity and disruption of cellular functions (Schmidt and Bradfield, 1996).

Due to its prevalence in the environment and high potency, the toxic effects of 2,3,7,8- TCDD are well-studied. Previous studies have shown that AHR-null mice are resistant to many of the manifestations of 2,3,7,8-TCDD-induced toxicity (Fernandez-Salguero et al., 1996; Lahvis and Bradfield, 1998; Mimura et al., 1999; Okey et al., 2005). In addition, deletions in the LBD and of 38 or 43 amino acids of the carboxy-terminal TAD have been shown to confer resistance to 2,3,7,8-TCDD-induced toxicity and lethality (Bradshaw and Bell, 2009; Moffat et al., 2010; Okey et al., 2005; Pohjanvirta et al., 1998). 2,3,7,8-TCDD binding to AHR has been shown to

induce phase I and II drug metabolizing enzymes, in addition to modulating the expression of various other genes that cause toxic responses. Binding affinity to AHR has been correlated to the magnitude of toxic response, whereby 2,3,7,8-TCDD induces 300-fold greater toxicity than its low-affinity dichloro cogeners (Safe, 1986; Schecter et al., 2006; Stevens et al., 2009). Mice expressing the high affinity AHR allele exhibited a greater 2,3,7,8-TCDD-induced toxic response and greater induction of xenobiotic metabolizing enzymes than their counterparts expressing a 10-fold lower affinity AHR allele (Poland and Glover, 1980; Poland et al., 1994; Stevens et al., 2009; Thomas et al., 2002).

In animal models, this toxic response has been shown to include porphyria, immunotoxicity, cleft palate, thymus involution, tooth abnormalities, altered offspring gender ratio, lymphoid involution, epithelial hyperplasia and metaplasia, impaired glucose tolerance and diabetes, reproductive and developmental toxicity, endocrine disruption, wasting syndrome, chloracne, tumour promotion, carcinogenesis, and death (Bock and Kohle, 2006; Bradshaw and Bell, 2009; Okey et al., 2005; Pohjanvirta, 2009; Poland and Knutson, 1982; Safe, 2001). IL-2 is known to be an AHR-responsive gene that plays a role in the homeostasis of T cells (Bock and Kohle, 2006). Previous studies suggest that 2,3,7,8-TCDD exposure induces immunosuppression through disregulation of co-regulator and co-stimulator molecules resulting in negative selection of T cells in the thymus (Bock and Kohle, 2006; Fisher et al., 2006). Maternal murine exposure to 2,3,7,8-TCDD has proven to have a tetratogenic response, whereby the epithelial to mesenchymal transformation of the palatal shelf does not occur during the embryonic and fetal stages of prenatal development, resulting in a cleft palate (Abbott and Birnbaum, 1991; Bock and Kohle, 2006). Endocrine disruption by 2,3,7,8-TCDD can lead to the development of endometriosis and breast cancer (Bock and Kohle, 2006; Warner et al., 2002). Chloracne was first identified in 1899 in workers involved in industrial production, and presents as hyperkeratotic skin disorder that affects sebaceous glands, the interfollicular epidermis and hair follicles causing persistent epidermal cysts (Bock and Kohle, 2006; Bradshaw and Bell, 2009). 2,3,7,8-TCDD is considered to be a tumour promoter, as treatment has been shown to initiate carcinogenesis by affecting cell surface plasticity, enhancing anti-apoptotic action, and affecting invasive cell growth (Stinchcombe et al., 1995). In addition, animal studies of cohorts exposed to industrial chemicals in the Netherlands, Germany and the United States exhibited increased prevalence of lung and gastrointestinal cancers, soft tissue sarcomas, and non-Hodgkin

lymphoma and death (Becher et al., 1995; Fingerhut et al., 1991; Hooiveld et al., 1998). The toxic and genotoxic effects that are mediated by AHR have been shown to be dependent on age, sex, and species (Poland and Knutson, 1982; Safe, 2001). For example, Kociba et al. (1978) demonstrated that 2 year dietary administration of 2,3,7,8-TCDD to Sprague-Dawley rats induced liver tumours in females but not males, which appears to be a hormone expression linked response (Kociba et al., 1978; Safe, 2001).

1.13 Mutations to the LBD of AHR

In recent years, mutations of several AHR receptor variants have been studied to elucidate the putative functional residues of the LBD. Phosphorylation has been shown to play an integral role in AHR activation and signalling. A previous study by Park et al. (2000) demonstrated that tyrosine phosphorylation is required for AHR to bind to DNA which is postulated to serve as a control for AHR-mediated target gene induction (Park et al., 2000). In addition, it has been proposed that activation of AHR by atypical inducers requires c-src kinase phosphorylation of a residue within the LBD (Backlund and Ingelman-Sundberg, 2004; Backlund et al., 1997; Kikuchi and Hossain, 1999; Lemaire et al., 2004). The effect of individual amino acid residues on 2,3,7,8-TCDD-responsiveness was previously demonstrated with C57BL/6J mice, relative to non-responsive DBA/2 mice, whereby the alanine to valine mutation of residue 375 conferred a 10-fold decrease in the binding affinity of AHR for 2,3,7,8-TCDD (Birnbaum et al., 1990; Celius and Matthews, 2010; Poland et al., 1994). More recently Pandini et al. (2009) examined the effect of individual amino acid mutations on 2,3,7,8-TCDD binding and 2,3,7,8-TCDD-induced DNA binding by AHR (Pandini et al., 2009). They determined that the following mutations to the mouse AHR receptor had a significant impact on AHR activation: T283M, H285A, F289A, Y316A, I319A/Y, F345A, and A375L (Pandini et al., 2009). Based on this evidence and the results of an earlier study by Backlund and Ingelman-Sundberg (2004), human Y322 residue is believed to be required for activation of binding-independent AHR activation; however, its role in binding-dependent activation is controversial. Tyrosine to phenylalanine mutation of rat AHR (rAHR-Y320F), which is believed to conserve the cyclic structure within the LBD but cannot be phosphorylated, resulted in attenuation of Omp-mediated AHR activation but had no effect on 2,3,7,8-TCDD-induced activation (Backlund and Ingelman-Sundberg, 2004). However,

expression of the same mutation of the equivalent residue in mouse AHR (mAHR-Y316F) resulted in a 55% reduction of 2,3,7,8-TCDD binding, and 16% reduction of 2,3,7,8-TCDD- induced AHRE binding relative to wt-mAHR (Pandini et al., 2009). Tyrosine to alanine mutation of the same residue (mAHR-Y316A), which abolishes the benzene ring of the residue and cannot be phosphorylated, was found to result in a 93% reduction of 2,3,7,8-TCDD specific binding and a 96% reduction of 2,3,7,8-TCDD-induced AHRE binding, relative to wt-mAHR (Pandini et al., 2009). The importance of Y322 in human AHR activation and its role in HAH- mediated activation remains to be elucidated.

2 Rationale and Objectives 2.1 Rationale

Despite numerous studies, many questions remained unanswered with respect to activation by both typical and atypical AHR inducers. The relative potencies of prototypical AHR ligands have been determined through a summation of available in vitro and in vivo data for each compound and are referred to as the toxic equivalency factors (TEF) values. However, previous studies in GAL-4 reporter assays have demonstrated that the TEF values were not necessarily predictive of the magnitude of ligand-induced AHR activation and CYP1A1 mRNA induction in some cell lines. In addition, previous studies suggest that AHR activation, target gene induction, and coactivator recruitment to AHREs are ligand- and cell line-dependent. CYP1A1 is the most studied AHR-responsive gene, but ligand-induced AHR signalling has been shown to mediate expression of other genes, including CYP1B1, HES1 and TiPARP. However, previous studies have not extended their investigations of ligand and cell line differences to include these lesser studied target genes. Since HAH compounds and Omp are believed to activate AHR through two distinct pathways, it is postulated that there should be some fundamental differences in how they mediate AHR signalling. As such, these differences should be reflected in AHR and coactivator recruitment to the regulatory regions of AHR-responsive genes, as well as target gene expression. It is well documented that typical AHR ligands induce proteolytic degradation of AHR through the 26S proteasome pathway to attenuate AHR signalling, but it remains unclear whether treatment with Omp also induces degradation.

While the atypical inducer Omp is believed to activate AHR through phsophorylation of residue Y322 of the LBD, the implication of this mechanistic difference on AHR signalling has not been extensively studied. Since little difference in the effects of atypical inducers relative to AHR ligands has been previously reported suggesting that, perhaps, Omp is actually a low affinity ligand of AHR. The mouse and rat equivalents of hAHR-Y322 of the LBD have previously been studied using reporter gene assays, and demonstrated that this residue is required for AHR activation by Omp. However, the role of Y322 in binding-dependent activation remains controversial. The impact of Y322 mutation on hAHR function in the presence of HAHs and Omp is yet to be studied. In addition, mutations to this residue are yet to be studied in a cellular system, which would provide a more realistic approach to assessing its role.

2.2 Research Objectives

Based on the above rationale, the objectives of my thesis research were as follows:

1. Compare inducer-, gene-, and cell line-dependent differences in HAH- and Omp- mediated AHR signalling.

To do so, I will compare the effects of ligands to atypical inducers, as well as to their relative potencies by first examining differences in AHR-mediated expression of CYP1A1, CYP1B1, HES1 and TiPARP mRNA by quantitative PCR (qPCR) in human breast cancer T-47D and human Hepatoma HuH7 cells. I will then look examine AHR and coactivator recruitment to the upstream regulatory regions of genes using chromatin immunoprecipitation (ChIP) assays. To examine whether Omp-mediated AHR signalling is attenuated by proteolytic degradation of AHR through the 26S proteasome pathway, I will perform Western blots of whole cell lysates and nuclear extracts with or without pre-treatment with the 26S proteasome inhibitor, MG132.

2. Examine the role of Y322 in ligand-mediated AHR activation.

To study the implications of Y322 on AHR signalling, I will overexpress phenylalanine and alanine variants in transiently transfected human breast cancer MCF-7 AHR100 cells. I will then examine the effects of these mutations on mRNA expression levels of CYP1A1 and CYP1B1, and AHR recruitment to the enhancer regions of these genes.

3 Materials 3.1 Chemicals

All sterile reagents for cell culture were purchased from Wisent (St. Brunno, QC), and included: 1:1 mixture of Dulbecco’s Modified Eagle Medium (DMEM) and Ham’s F12, high-glucose DMEM, and low-glucose DMEM media, fetal bovine serum (FBS), antibiotic (penicillin/ streptomycin, PEST), DNase/RNase-Free Sterile water and trypsin-EDTA. 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF, and 2,3,7,8-TCDF were purchased from Wellington Laboratories (Guelph, ON), while Omp was purchased from Calbiochem (San Diego, CA), and dimethyl sulfoxide (DMSO) from Sigma (St Louis, MO). Total RNA was isolated using RNAspin Mini Columns obtained from GE Healthcare (Mississauga, ON). Protein A Agarose Fast Flow 50% v/v used for ChIP assay were obtained from Sigma. Antibodies for ChIP assays were p300 (N-15), NCoA1 (M-341), NCoA3 (M-397), AHR (H-211), and rabbit immunoglobulin (IgG, sc-2027), and were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). DNA for ChIP assays was purified using the EZ-10 Spin PCR Purification Columns (BioBasic Inc.; Markham, ON). Primary antibodies used for Western Blots from both whole cell extract and nuclear extracts were obtained from Santa Cruz Biotechnology, and included: β-actin, AHR (H-211), and p53 (FF-393). ECL Anti-rabbit IgG horseradish peroxidase (HRP) linked whole secondary antibody (NA934V) was obtained from GE Healthcare, while donkey anti-mouse IgG HRP secondary antibody (sc-2314) was obtained from Santa Cruz Biotechnology. Western blots were visualized using the ECL-Advanced Western Blot Detection system (GE Healthcare). For site-directed mutagenesis, pfu Turbo was used (Stratagene; La Jolla, CA), and for reverse transcription, SuperScript II reverse transcriptase was used (Invitrogen; Carlsbad, CA). SYBR Green, obtained from KappaBiosystems (Woburn, MA), was used for all real-time PCR reactions. All primers were synthesized by Integrated DNA Technologies (IDT, Coralville, IA).

3.2 Plastic ware

All T25, T75 and T175 flasks for cell culture, as well as 6-well plates and 10cm cell culture dishes, were obtained from Starstedt (Montreal, QC). All 1.5ml and 1.7ml microcentrifuge tubes and 96-well PCR plates were manufactured by Progene and purchased from Ultident Scientific (St. Laurent, QC).

3.3 Instruments

HERAcell 150 incubators obtained from Kendro (Langenselbold, Germany) were used to maintain cells. The water bath was obtained from VWR International (Mississauga, ON; Sheldon manufacturing, Cornelius, OR). A Bright-Line Hemocytometer (Hausser Scientific, Horsham, PA) and VistaVision Light Microscope, purchased from VWR International, were used to quantify cells. The Locator JR Cryo Biological Storage System (Thermolyne), purchased from VWR International, was used to store cells in liquid nitrogen. Cells were centrifuged with Centrifuge 5702 (Eppendorf International; Hamburg, Germany). PCR DNA Purification and RNA extraction processes requiring centrifugation were performed using a Centrifuge 5415D (Eppendorf International). The concentration of RNA was measured using a Ultrospec 2100 pro Spectrophotometer obtained from Biochrom (Cambridge, UK). For ChIP assay, a Branson Digital Sonifier 450 (Danbury, CT) was utilized for sonification. A Mini LabRoller (Labnet Inc., Edison, NJ) rotator was used for both ChIP assays and protein extracts from whole cell lysates. Real-time PCR amplification of ChIP DNA and cDNA was performed on Chromo4 Real-Time PCR detector purchased from Bio-Rad Laboratories Inc. (Hercules, CA). The DNAEngine Peltier Thermal Cycler (BioRad) was used for site-directed mutagenesis and cDNA synthesis of extracted RNA. Protein concentrations were determined using 96-well MultiScan EX plate reader (Thermo Scientific; Rockford, IL).

4 Methods 4.1 Cell Culture

Prior to use, cells were stored in liquid nitrogen. Cells were grown to approximately 80-90% confluency then trypsinized and neutralized with fresh media. To pellet cells, they were centrifuged for 2 min at 4000rpm in a 50mL conical then resuspended in 90% FBS and 10% DMSO (v/v). The cells were then aliquoted into pre-labelled cryovials and placed in the -80ºC freezer overnight before being transferred to liquid nitrogen for long-term storage.

To prepare frozen T-47D cells for usage, a cryovial was thawed at room temperature and transferred to a T-25 tissue culture flask containing 1:1 mixture of Dulbecco’s Modified Eagle Medium (DMEM) and Ham’s F12 media supplemented with 10% (v/v) FBS and 1%

penicillin/streptomycin (PEST) and maintained at 37°C and 5% CO2. Once cells were approximately 80% confluent, the media was aspirated and cells were washed with PBS then trypsinized with 1mL of trypsin-EDTA and transferred to a T-75 flask. This was repeated until cells were transferred to a T-175 flask. Once in a T-175 flask, the medium was aspirated from culture flask and cells were washed once with 5 ml of sterile PBS. The PBS was aspirated and 4 ml of pre-warmed trypsin-EDTA was added to the flask and incubated for 2 min or until the cells detached completely. A volume of 8 ml of pre-warmed complete medium was added to the cell suspension in the flask to neutralize the tryspin. The suspension was then vigorously pipetted up and down until homogenous and 4 ml of the cell solution was transferred to a T-175 flask containing 25 ml of completed medium. Cells were then sub-cultured 1:2 to 1:3 every 2-3 days when they reached approximately 80% confluency.

HuH7 cells and AHR deficient human breast cancer MCF-7 AHR100 cells were prepared for use in the same manner as described above and maintained at 37°C and 5% CO2. HuH7 cells were cultured in high-glucose DMEM with 10% (v/v) FBS and 1% PEST and subcultured 1:5 to 1:10 every 2-3 days at approximately 80% confluency. MCF-7 AHR 100 cells were cultured in low-glucose DMEM supplemented with 10% FBS and 1% PEST and subcultured 1:10 every 2-3 days at approximately 80% confluency.

4.2 RNA Extraction and qPCR

T-47D cells were seeded in 6-well plates (300,000 cells/ well) in phenol red-free 1:1 DMEM: Ham’s F12 with 5% (v/v) dextran-coated charcoal stripped FBS (DCC) and 1% PEST. HuH7 cells were seeded in 6-well plates (200,000 cells/ well) in phenol red free high-glucose DMEM with 5% DCC and 1% PEST. Cells were incubated for 72h in media to minimize the compounding effects caused by esposure to estrogen and estrogenic compounds contained in the serum and media. Cells were treated with DMSO (solvent control), 0.01nM- 10nM concentrations of 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF, or 2,3,7,8-TCDF, or 10- 104nM Omp for 1.5, 6, and 24h. Total RNA was isolated using RNAspin Mini Columns (GE Healthcare) whereby cells were washed with phosphate buffered saline (PBS) twice, then 350μL of RA1 buffer containing 1% β-mercaptoethanol (GE Healthcare). Cells were then harvested by scraping and transferred to a 1.5mL microcentrifuge tube (Progene). To bind RNA to the column, 70% EtOH was added to the sample and mixed gently by pipetting, then loaded onto the RNA spin column and centrifuged for 1min at 9100rpm to bind the RNA to the column. The column was washed with 350μL of membrane desalting buffer (MDB; GE Healthcare) and centrifuged for 1min at 10,700g. DNA was digested with an on-column DNase digest utilizing 95μL DNase reaction mixture in reaction buffer, which was incubated at room temperature for 15min. Following incubation, the column was washed sequentially with 200μL RA2 buffer and 600μL RA3 buffer (GE Healthcare) and centrifuged for 1 min at 10,700g. The column was washed a third time with 250μL RA3 buffer (GE Healthcare) and centrifuged for 2min at 11,000g. Purified total RNA was eluted in 40μL of RNase-free water by centrifugation for 1min at 11,000g.

Following purification, the concentration of isolated RNA was determined by spectrophotometry (Biochrom) and the concentration of each sample was adjusted to 50 ng/ μL by adding the appropriate volume of DNase and RNase-Free Sterile water. 500ng of total RNA was reverse transcribed with SuperScript II reverse transcriptase (Invitrogen) in a total volume of 20 μL containing 1μM random hexamers, and 1μM dNTPs. cDNA was synthesized by PCR using the following protocol: incubated at 42°C for 1h followed by 15min incubation at 70°C. Following reverse transcription, CYP1A1, CYP1B1, HES1 and TiPARP expression was determined by quantitative PCR (QPCR) with SYBR Green (KapaBiosystems; Woburn, MA) (Primers listed in Table 2). Data was analyzed using the Opticon Monitor3 software (Bio-rad). All transcripts were run in triplicate and expression of each AHR target gene transcript was

analyzed using the comparative CT (ΔΔCT) method, expressed as a fold induction relative to time matched DMSO control and normalized against ribosomal 18s content.

Table 2. qPCR Primers used for Analysis of cDNA. Primer Name Sequence (5'-3') 18s mRNA Forward GCTTAATTTGACTCAACACGGGA 18s mRNA Reverse AGCTATCAATCTGTCAATCCTGTC hCYP1A1 mRNA Forward TGGTCTCCCTTCTCTACACTCTTGT hCYP1A1 mRNA Reverse ATTTTCCCTATTACATTAAATCAATGGTTCT hCYP1B1 mRNA Forward CCTATGTCCTGGCCTTCCTTT hCYP1B1 mRNA Reverse TGAGGAATAGTGACAGGCACAAA hHES1 mRNA Forward AGGCCACCCCTCCTCCTA hHES1 mRNA Reverse GTTTAGAGTCCGGAGGGAAGAGA hTiPARP mRNA Forward GGCAGATTTGAATGCCATGA hTIPARP mRNA Reverse TGGACAGCCTTCGTAGTTGGT

4.3 Chromatin Immunoprecipitation (ChIP) Assays

4.3.1 AHR Recruitment Experiments

T-47D cells were seeded in 10cm dishes (3 million cells/ 10cm dish) in phenol red-free 1:1 DMEM: Ham’s F12 with 5% (v/v) DCC and 1% PEST. Following 72 h incubation, cells were treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10 μM Omp for 45min. Following treatment, protein was cross-linked to DNA with 1% formaldehyde added directly into the media for 10min with mild rocking. After incubation with formaldehyde, cross- links were quenched with 0.125M glycine and incubated for an additional 5 min with rocking. Cells were washed twice with PBS and then 750μL of PBS+ 0.1% Tween20 was added to each 10cm dish. Cells were collected by scraping them into a 1.5mL microcentrifuge tube and pelleted by centrifugation for 3min at 10,000rpm and 4°C. Supernatant was aspirated and the pellet was resuspended in 400μL of TSEI buffer containing 1% protease inhibitor cocktail (PIC). Cells were sonicated on ice to shear DNA to 500-800bp fragments at 30% intensity for 80 pulses. Cell debris was pelleted by centrifugation for 10min at 13,000rpm and 4°C and the

supernatant was transferred to a new 1.5mL microcentrifuge tube. The extract was pre- immunocleared with 25μL of 1:1 Protein A Agarose Fast Flow beads (Sigma): TSE I buffer by rotation at 4°C for 2 hours. Beads were collected at the bottom of the tube by centrifugation for 1min at 5,000rpm at room temperature. 80μL of each sample was aliquoted into new microcentrifuge tubes containing a mixture of 100 μL of TSE I buffer containing 0.05μg/μL salmon sperm DNA (ssDNA), and 10mg/mL bovine serum albumin (BSA), and 0.5-1μg of antibody against IgG, AHR and ARNT then incubated with rotation overnight at 4°C. The following morning, 25μL of a slurry of 1:1 Protein A Agarose Fast Flow beads (Sigma): TSE I with BSA and ssDNA was added to each sample and incubated with rotation at 4°C for 1h. Following incubation, beads were pelleted for 1min at 5,000rpm at room temperature and supernatant was removed by aspiration. Beads were then washed sequentially 3 times with 1mL TSE I buffer, twice with 1mL TSE II buffer, twice with LiCl buffer and twice with TE buffer. During each wash, samples were incubated with the wash buffer for 5min then spun down and aspirated. After the first TE buffer wash, approximately 50μL of supernatant was left behind and aspirated using a syringe with a 30 ½ gauge needle. 500μL of TE was then added and samples were transferred to a new tube were the syringe aspiration step was repeated. After the final TE wash, 110μL of TE containing 1% SDS was added to each sample. Samples were incubated with rotation at room temperature for 1 hr before transferring them to incubate overnight at 60°C to reverse the cross-links. The following day, samples were centrifuged at 5,000rom for 1min to collect beads at the bottom of the tube and 100μL of the supernatant was transferred to a new 1.5mL microcentrifuge tube. 500μL of PB Binding Buffer was added to the tube and pipetted up and down until the mixture was homogenous. The mixture was then applied to an EZ-10 Spin PCR Purification Column (BioBasic Inc.) and centrifuged at 13,200rpm for 1min. The column was washed sequentially with 500 and 200μL of Wash Buffer (BioBasic Inc.) and centrifuged at 13,200rpm and 1min following each wash step. The columns were then dried by centrifugation at 13,2000rpm for 1min, and DNA was eluted into a new microcentrifuge tube with 100μL Elution Buffer (BioBasic Inc.). Recruitment of AHR to the regulatory regions of CYP1A1, CYP1B1, HES1 and an AHRE cluster was analyzed by qPCR (Primers listed in Table 3). Data

was expressed relative to a 100% total input chromatin and analyzed using the CT (ΔΔCT) method.

Table 3. qPCR Primers used for ChIP Assays. Primer Name Sequence (5'-3') hCYP1A1 ChIP Forward AGGCGTGGACCGAAAATG hCYP1A1 ChIP Reverse CTAGGTCTGCGTGTGGCTTCT hCYP1B1 ChIP Forward ATATGACTGGAGCCGACTTTCC hCYP1B1 ChIP Reverse GGCGAACTTTATCGGGTTGA hHES1 ChIP Forward GCGTGCAGTCCCAGATATATATAGAG hHES1 ChIP Reverse CCAGCTCCGGATCCTGTGT hAHRE Cluster ChIP Forward GGGAGGGCAGTCACGCTAT hAHRE Cluster ChIP Reverse GTCCTCCCCCGGTGAACT

4.3.2 Coactivator Recruitment Experiments

T-47D cells were seeded in 10cm dishes (3 million cells/ 10cm dish) in phenol red-free 1:1 DMEM: Ham’s F12 with 5% (v/v) DCC and 1% PEST. HuH7 cells were seeded in 10cm dishes (2 million cells/ 10cm dish) in phenol red-free high-glucose DMEM with 5% DCC and 1% PEST. Following 72 h incubation, cells were treated with DMSO, 10 nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF, or 2,3,7,8-TCDF, or 10μM Omp for 45min. After treatment, a ChIP assay was performed as described above with 0.5-1μg of antibodies against IgG, p300, NCoA1, NCoA3, polII, RIP140, and AHR. Recruitment of AHR to the regulatory regions of CYP1A1, CYP1B1, HES1 and an AHRE cluster was analyzed by qPCR (Primers listed in Table 3). Data were expressed relative to a 100% total input chromatin and analyzed using the CT (ΔΔCT) method.

4.4 Western Blot

4.4.1 Western Blot of Whole Cell Lysates

T-47D cells were seeded in 10cm dishes as described above. Following 72h incubation, cells were treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp for 0, 1, 3, and 24h. Cells were then washed twice with PBS, scraped with 500μL of PBS+ Tweeen20 and transferred to a 1.5mL microcentrifuge tube. Cells were collected by centrifugation for 3min at 10,000rpm at 4°C and the supernatant was aspirated. Pellets were

32 placed in the -20°C freezer for 10min to lyse cells. Pellets were resuspended in 200μL TENG buffer with 0.5% NP40, 1X PIC and 1mM DTT. Following resuspension, cells were frozen in the -80°C freezer for 10min then thawed and sonicated once for 10sec at 25% amplitude on ice. Samples were incubated on a rotator for 20min at 4°C. Cell debris was pelleted by centrifugation for 10min at 13,000rpm and 4°C. Protein concentration in the whole cell extract was determine by measuring absorbance of 10μL of 5X diluted sample 200μL of Bradford Reagent at 595nm with a 96-well plate reader relative to a standard curve of 1, 0.75, 0.5, 0.25, 0.125, and 0 mg/mL BSA in Bradford reagent. Samples were prepared in 4X Loading buffer with 100mM DTT and heated to 95°C for 5min to denature the proteins, then resolved using an 8% SDS-polyacrylamide gel electrophoresis run at 125V constant volts for 2h alongside Kaleidoscope Protein Ladder. Following electrophoresis, proteins were transferred to a nitrocellulose blotting membrane in 25mM Tris base (pH 8.3) containing 19.2nM glycine and 20% (v/v) methanol. The membrane was blocked with 2% (w/v) ECL-Advanced blocking agent (GE Healthcare) for 4h at room temperature with constant rocking, and incubated overnight at 4°C with 1:4000 anti-AHR (SA211) antibody in 2% blocking solution with constant rocking. The following morning, the blot was washed 4X with PBS+ 0.1% Tween20 and incubated with 1:200,000 horseradish peroxidase (HRP)-conjugated anti-rabbit secondary antibody in 2% blocking solution for 1h at room temperature with constant rocking. Following incubation, the blot was washed with PBS+ 0.1% Tween20 4X and the immunoblot was visualized using ECL- Advanced chemiluminescent substrate (GE Healthcare). Remaining developing solution was removed by washing with PBS+ 0.1% Tween20 for 15min. The blot was incubated with 1:500,000 β-actin antibody in 2% blocking solution for 20 min, washed with 4X with PBS+ 0.1% Tween20, and incubated for 20 min with 1:200,000 anti-mouse antibody in 2% blocking solution at room temperature with constant rocking. The blot was then visualized as described above.

4.4.2 Western Blot of Nuclear and Cytoplasmic Extracts

T-47D cells were seeded in 10cm dishes in phenol red-free media as described above. Following 72h incubation, cells were treated with DMSO, 10nM 2,3,7,8-TCDD or 10μM Omp for 0, 1, and 3h with and without 1h pre-treatment with 25μM MG132. After treatment, media

was aspirated and cells were washed once with PBS. To each dish, 500μL of hypotonic lysis

buffer (10mM HEPES, 1.5mM MgCl2, 10mM KCl, 0.05% NP40, 0.5mM DTT, 1X PIC) then cells were scraped and transferred to a 1.5mL microcentrifuge tube. Samples were incubated on ice for 10min to aid in lysis, the nuclei were pelleted by centrifugation for 3min at 3000rpm at 4°C. The supernatant (cytoplasmic fraction) was transferred to a new 1.5mL tube and frozen immediately at -20°C. Nuclear pelletes were washed once with 1mL nuclei wash buffer (10mM

HEPES, 1.5mM MgCl2, 10mM KCl) and pellets were resuspended in 200μL of high salt nuclear

extraction buffer (20mM HEPES, 400mM NaCl, 0.2mM EDTA, 1.5mM MgCl2, 25% v/v Glycerol, 1X PIC). Samples were incubated on ice for 30min with periodic vigorous vortexing. Debris was pelleted with centrifugation for 10min at 13,200rpm at 4°C. The supernatant (nuclear protein extract) was transferred to a new 1.5mL tube and the pellet was discarded. Protein concentrations in both the cytoplasmic and nuclear fractions were determined using a Bradford assay as described above. Samples were prepared in 4X Loading buffer with 100mM DTT and heated to 65°C for 10min to denature the proteins prior to loading. Protein from nuclear and cytoplasmic extracts was resolved by 8% SDS-polyacrylamide denaturing gel electrophoresis and transferred to nitrocellulose blotting membrane as previously described. The membrane was then blocked with 2% (w/v) ECL-Advanced blocking agent (GE Healthcare) and probed for AHR (primary: 1:4000; secondary anti-rabbit: 1:200,000) and β-actin (primary: 1:500,000; secondary anti-mouse: 1:200,000) as described previously. The nitrocellulose membrane was then stripped with 10mL of Restore Western Blot Stripping Buffer (Thermo Scientific) for 15min. After stripping, the blot was washed 5X for 15min in PBS+ 0.1% Tween20 then blocked in 2% ECL-Advanced blocking agent (GE Healthcare) for 1h. The membrane was then probed for p53 with primary antibody (1:2000) overnight at 4°C with constant rocking. The following morning, the blot was washed 4X in PBS+ 0.1% Tween20 and secondary anti-rabbit antibody was applied (1:200,000) for 1h at room temperature with constant rocking. The blot was washed 4X with PBS+ 0.1% Tween20 and developed as described above.

4.5 Site-Directed Mutagenesis

Primers for the generation of the Y322A and Y322F hAHR variants were designed and purchased from IDT (primers listed in Table 4); mutations are as underlined. The desired point-

mutations were introduced using site-directed mutagenesis with 1μL of 50ng/μL pRC-CMV2- hAHR, sterile water, 1X pfu Buffer, 0.2mM dNTPs, 0.2mM both 5’ and 3’ primers, and 2.5U pfu Turbo. The Y322A variant was introduced using PCR with the following protocol: 95°C for 5min, the 25 cycles of 95°C for 15sec, 60°C for 15sec, and 72°C for 10min, then 72°C for 7min, and 4°C for 10min. The Y322F mutation was introduced under the following PCR conditions: 95°C for 5min, the 25 cycles of 95°C for 1min, 50°C for 1min, and 72°C for 10min, then 72°C for 7min, and 4°C for 10min. PCR product was digested for 1h with 1μL DpnI (New England Biolabs; Pickering, ON) restriction enzyme at 37°C.

Table 4. Primers used for Site-Directed Mutagenesis. Primer Name Sequence (5'- 3') hAHR-Y322F Forward GAGAGGCTCAGGTTTTCAGTTTATTCATG

hAHR-Y322F Reverse CATGAATAAACTGAAAACCTGAGCCTCTC

hAHR-Y322A Forward AGCTGTGCACGAGAGGCTCAGGTGCTCAGTTTAT TCATGCAGCTGATAT hAHR-Y322A Reverse ATATCAGCTGCATGAATAAACTGAGCACCTGAGC CTCTCGTGCACAGCT

4.6 Transformation

Chemically competent DH5α E. coli cells were removed from the -80°C freezer and thawed on ice. A total of 100μL of cells was transferred to each 1.5mL microcentrifuge tube then 10μL of the site-directed mutagenesis PCR product was added to the tube. The reactions were incubated on ice for 20min, then for 5min at 37°C. A 500μL aliquot of SOC (SOB+ 1M glucose) was added to each reaction then samples were incubated at 37°C for 20min with constant shaking. Following incubation, samples were centrifuged for 1min at 10,000rpm and aspirated such that the residual volume was approximately 100μL. Cells were distributed using aseptic technique on LB-Ampicillin 10cm plates pre-dried at 37°C for 20min. Plates were inverted and incubated overnight at 37°C; the following day 4 colonies were chosen from each variant plate and grown

overnight in 3mL of LB broth+ Ampicillin at 37°C with constant shaking. The following morning, plasmids were purified using the Genelute Genomic DNA Miniprep kit (Sigma) and screened for inclusion of the Y322F and Y322A mutations at the DNA Sequencing and Synthesis Facility (Medical and Related Sciences (MaRS); Toronto, ON). Plasmids found to contain the desired mutations were then maxi-prepped using Qiagen Plasmid Maxi Prep kit (Valencia, CA) as per the manufacturer’s protocol.

4.7 Transient Transfection

MCF-7 AHR 100 cells were seeded in 6-well plates in high-glucose DMEM with 10% FBS and 1% PEST (300,000 cells/ well), and transiently transfected the following day with 500ng of empty vector, wt-hAHR, Y322A or Y322F variant in Opti-MEM and 2μL of Lipofectamine2000 (Invitrogen) according to manufacturer’s instructions. In addition, 1ng/μL of pEGFP was co- transfected into cells so that the efficiency of the transfection could be assessed the following morning by examining expression of the green fluorescent protein with fluorescence microscopy. Following 24h transfection, CYP1A1 mRNA was quantified in cells treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF or 10μM Omp for 6h as described previously. ChIP assays were performed as described above for each transfection by pooling three wells of a 6-well plate following 45min exposure to one of each treatments.

4.8 Statistical Analysis

All qPCR results were analyzed using Opticon Monitor 3 and Microsoft Excel, and expressed as the mean + standard deviation of representative data derived from at least two independent biological replicates. The statistical significant of results was determined using the GraphPad Prism 5 software. Statistical significance was evaluated using a one-way analysis of variance (ANOVA) with Tukey post-hoc test.

5 Results 5.1 Ligand-, Cell Line- and Gene-Specific Differences in CYP1A1 and CYP1B1 mRNA Expression

To examine ligand and temporal differences in the ability of these compounds to induce AHR regulated gene expression, T-47D and HuH7 cells were treated with DMSO (solvent control), or increasing concentrations of 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF, 2,3,7,8-TCDF, or Omp for 1.5, 6 and 24h. Use of full serum and phenol red containing media is associated with high constitutive estrogen receptor (ER) activity, which has been shown to influence AHR signalling due to cross- talk between AHR and the ER pathways (Pansoy et al., 2010). As such, both dextran-coated charcoal stripped FBS (DCC-FBS) and phenol red-free media were utilized for all experiments to avoid the compounding effects of estrogen, estrogenic compounds and other steroid or lipophilic compounds (Rogers and Denison, 2000). Following incubation, cells were harvested and RNA was isolated then reverse transcribed to examine the mRNA expression levels of CYP1A1, CYP1B1, HES1 and TiPARP. Following reverse transcription, expression was determined by qPCR and analyzed using the Opticon Monitor3 software. All transcripts were

analyzed in triplicate and expression of each gene was analyzed using the comparative CT

(ΔΔCT) method, expressed as the mean fold relative to time matched DMSO solvent control and normalized against ribosomal 18s content.

As expected, all ligands induced expression of CYP1A1, CYP1B1, HES1 and TiPARP mRNA for all time-points studied in a dose-dependent manner in both T-47D and HuH7 cells. In T-47D cells, a similar expression pattern of CYP1A1 and CYP1B1 mRNA was observed all three time-points for 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF and 2,3,7,8-TCDF, whereby mRNA levels continued to increase to the 24h time-point (Figure 5A-F). Following 1.5h treatment, Omp induced greater expression of CYP1A1 than all other ligands in T-47D cells (Figure 5A). Conversely, all treatments induced relatively similar levels of CYP1B1 mRNA expression following 1.5h treatment (Figure 5D). Following 6h treatment, only the highest (10μM) concentration of Omp significantly induced CYP1A1 and CYP1B1 mRNA levels. While expression of both CYP1A1 and CYP1B1 continued to increase 24h following treatments with 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, Omp-induced CYP1A1 mRNA expression levels did not further increase following 6h treatment (Figure 5B and C). The increase in Omp-

induced CYP1B1 between 6 and 24h was negligible (Figure 5E and F). In agreement with the reported relative potencies of the HAHs studied, comparison of the 0.1 nM treatments revealed that 2,3,7,8-TCDF induced significantly less CYP1A1 and CYP1B1 mRNA expression than 2,3,7,8-TCDD or 2,3,4,7,8-PeCDF following 6 and 24h treatments (Figure 5A-F).

In HuH7 cells, all ligands induced CYP1A1 and CYP1B1 mRNA expression in a dose- and time-dependent manner, with similar induction patterns for 6h and 24h treatments to T-47D cells (Figure 6A-F). Unlike T-47D cells, the HAHs induced greater CYP1A1 and CYP1B1 mRNA expression following 1.5h treatment than any dose of Omp (Figure 6A and D). In addition, significant induction of CYP1A1 and CYP1B1 mRNA in HuH7 cells was observed with both 1μM and 10μM Omp after 6h treatment of HuH7 cells (Figure 6B and E). As was observed for T-47D cells, 2,3,7,8-TCDD-, 2,3,4,7,8-PeCDF-, and 2,3,7,8-TCDF-induced CYP1A1 and CYP1B1 mRNA expression continued in increase up to 24h. Unlike T-47D cells, both the 1μM and 10μM doses of Omp induced CYP1A1 and CYP1B1 mRNA expression after 24h exposure in HuH7 cells. However, as demonstrated previously, no further increases in CYP1A1 or CYP1B1 mRNA levels were observed from 6 to 24h in cells treated with Omp (Figure 6C and F). Unlike T-47D cells, the relative potencies of the HAH ligands were not predictive of the level of CYP1A1 mRNA expression in HuH7 cells. TEF values were not predictive of ligand-induced CYP1B1 expression for all doses and time-points studied. Temporal differences in activator-mediated target gene induction are likely due to differences in the relative stability of these compounds in cell lines studied.

5.2 Ligand- and Cell Line-Dependent AHR-Mediated HES1 and TiPARP mRNA Expression

To further investigate the cell line and temporal differences in ligand-mediated AHR signalling, I examined the expression of HES1 and TiPARP mRNA following 1.5, 6, and 24h treatments. As described previously, cells were harvested and RNA was reverse transcribed to examine the mRNA expression levels of HES1 and TiPARP. Expression was determined by qPCR and analyzed using the Opticon Monitor3 software. All transcripts were analyzed in triplicate and expression of each gene transcript was analyzed using the comparative CT (ΔΔCT) method,

CYP1A1 A 6 * † 5 4

3 * * * † † 2 * † † 1 * Fold Induction- 1.5h * * * † 0

DMSO TCDD PeCDF TCDF Omp

B 16 * *† ** 12 ** * * 8 * *† 4 *† Fold Induction- 6h † †

DMSO TCDD PeCDF TCDF Omp

C 250 * * * 200 * * 150 * * 100 * *†

Fold Induction- 24h 50 *† *† †† * * 0

DMSO TCDD PeCDF TCDF Omp

CYP1B1 D 5 * * * 4 * *† 3 * *† 2 * *

1 Fold Induction- 1.5h

DMSO TCDD PeCDF TCDF Omp

E 9 *†

* 7 * * * * * 5 * * * 3

Fold Induction- 6h Fold * † † 1 0

DMSO TCDD PeCDF TCDF Omp

F 25 * * * * 20 * * 15 * * * 10 *† * Fold Induction- 24h Fold 5 * †† 0

DMSO TCDD PeCDF TCDF Omp

Figure 5. AHR-mediated CYP1A1 and CYP1B1 mRNA expression levels in T-47D cells. Cells were treated with the DMSO or 0.01, 0.1, 1 and 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF, or 2,3,7,8-TCDF, or 10, 100, 103, and 104nM Omp for 1.5, 6 and 24h. Changes in mRNA expression levels were determined by reverse transcription and qPCR and are expressed as the mean fold induction of CYP1A1 (A-C) and CYP1B1 (D-F) relative to time-matched DMSO + SEM normalized relative ribosomal 18s. Data presented is representative data of at least two individual biological replicates. Significant differences (p<0.05) relative to time-matched DMSO (*) and concentration-matched 2,3,7,8-TCDD (†) were determined by one-way ANOVA with Tukey post-hoc test.

CYP1A1 A 5 * * * 4 * * * *† 3 * * *† 2 * †

Fold Induction- 1.5h 1

DMSO TCDD PeCDF TCDF Omp B 12 * 10 * 8 * * * *† * *† 6 * *† 4

Fold Induction- 6h *† †† 2

DMSO TCDD PeCDF TCDF Omp

C 80 * * * * 60 * * * 40 * *† 20 Fold Induction- 24h *† *† † *† * †† 0

DMSO TCDD PeCDF TCDF Omp

CYP1B1 D 20 *†

16 * 12 * * 8 * * * * 4 †

Fold Induction- 1.5h 0

DMSO TCDD PeCDF TCDF Omp

E 35 * 30 * 25 *† 20 *† 15 * *† 10 *† *†

Fold Induction- 6h *† 5 0

DMSO TCDD PeCDF TCDF Omp

F *† 60

50 * * 40 * * 30

20 *† *†

Fold Induction- 24h 10 * *† † *† *† 0

DMSO TCDD PeCDF TCDF Omp

Figure 6. AHR-mediated CYP1A1 and CYP1B1 mRNA expression in HuH7 cells. HuH7 cells were treated with the DMSO or 0.01, 0.1, 1 and 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF, or 2,3,7,8-TCDF, or 10, 100, 103, and 104nM Omp for 1.5, 6 and 24h. Changes in mRNA expression levels were analyzed by qPCR and expressed as the mean fold induction relative to time-matched DMSO + SEM normalized relative and ribosomal 18s of CYP1A1 (A-C) and CYP1B1 (D-F). Data presented is representative data of at least two individual biological replicates. Significant differences (p<0.05) relative to time-matched DMSO (*) and concentration-matched 2,3,7,8-TCDD (†) were determined by one-way ANOVA with Tukey post-hoc test.

expressed as a the mean fold induction relative to time matched DMSO solvent control and normalized against ribosomal 18s content. In T-47D cells, all treatments induced expression of HES1 and TiPARP mRNA at the 1 and 10nM doses for 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF and 2,3,7,8-TCDF, and 1 and 10μM doses of Omp following 1.5h treatment (Figure 7A and D). Induction of TiPARP at this time-point appeared to be dose-dependent, while the level of induced mRNA expression of HES1 was very similar among doses. Following 6h, HES1 mRNA induction returned to being indistinguishable from that of the solvent control for most treatments (Figure 7B). This was also observed following 24h treatment, whereas only the 1 and 10μM significantly induced HES1 mRNA at this time-point (Figure 7C). Between 6 and 24h treatments, TiPARP mRNA expression levels decreased in a temporally dependent manner for all treatments, whereas by 24h TiPARP mRNA levels decreased to solvent control levels for all treatments except 10μM Omp (Figure 7E and F).

Similar to results in T-47D cells, all treatments induced HES1 and TiPARP mRNA expression following 1.5h treatment in HuH7 cells (Figure 8A and D). At 6h a similar expression pattern to T-47D cells was observed, whereby mRNA expression levels of both HES1 and TiPARP decreased for all treatments (Figure 8B and E). HES1 expression appeared to rebound slightly from being comparable to solvent controls at 6h to being slightly increased at 24h in all HAH treated samples, albeit this increase is relatively modest and was found to be statistically insignificant (Figure 8C). These results correlate well with previous evidence that suggests HES1 functions to repress transcription of bHLH transcription factor-dependent genes, and has been shown to be cyclically expressed (Hirata et al., 2002; Yoshiura et al., 2007). Contrary to findings in T-47D cells, TiPARP expression levels were also shown to slightly increase from 6 to 24h in HuH7 cells (Figure 8F). These results, coupled with the mRNA expression data for CYP1A1 and CYP1B1, demonstrate that HES1 and TiPARP are differentially regulated by AHR compared to CYP1A1 and CYP1B1.

HES1 A 3.5 *† 3 2.5 * ***† *† 2 *† 1.5 1 Fold Induction- 1.5h 0.5 0

DMSO TCDD PeCDF TCDF Omp

B 3 *† 2.5 * 2 *† * 1.5

1 Fold Induction- 6h 0.5

DMSO TCDD PeCDF TCDF Omp

C 5 *† 4

3 *† 2

Fold Induction- 24h 1

DMSO TCDD PeCDF TCDF Omp

TiPARP

D 7 * 6 * * 5 4 * * * * 3 † *† 2 Fold Induction- 1.5h 1 0

DMSO TCDD PeCDF TCDF Omp

E 2 * * 1.6 * † 1.2 * * * * 0.8 * *† Fold Induction- 6h 0.4

DMSO TCDD PeCDF TCDF Omp

F 3.5 3 2.5 2 * † 1.5 1

Fold Induction- 24h * * * 0.5 0

DMSO TCDD PeCDF TCDF Omp

Figure 7. AHR-mediated HES1 and TiPARP mRNA expression in T-47D cells. Cells were treated with the DMSO or 0.01, 0.1, 1 and 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8- TCDF, or 10, 100, 103, and 104nM Omp for 1.5, 6 and 24h. Expression analyzed by qPCR and expressed as the mean fold induction relative to time-matched DMSO + SEM, normalized relative to ribosomal 18s for HES1 (A-C), and TiPARP (D-F). Representative data from at least two individual biological replicates are presented in these graphs. Significant differences (p<0.05) relative to time-matched DMSO (*) and concentration-matched 2,3,7,8-TCDD (†) were determined by one-way ANOVA with Tukey post-hoc test.

HES1 A * 4 ** 3 * *

2 * *†

1 Fold Induction- 1.5h

DMSO TCDD PeCDF TCDF Omp

B 1.4

1 * * 0.6

Fold Induction- 6h

0.2

DMSO TCDD PeCDF TCDF Omp

C 1.6

1.2 † † † *† 0.8

0.4 Induction-Fold 24h 0

DMSO TCDD PeCDF TCDF Omp

TiPARP D 9 * * * 7 * * * 5 * *† * *† 3 * *

Fold Induction- 1.5h 1 0

DMSO TCDD PeCDF TCDF Omp E 4.5 * * * * * 3.5 * *† *† 2.5 * † * 1.5 Fold Induction- 6h

0.5 0

DMSO TCDD PeCDF TCDF Omp

F * 7 * 6 * 5 * 4 3 * *† * † 2 † † Fold Induction- 24h 1 0

DMSO TCDD PeCDF TCDF Omp

Figure 8. AHR-mediated HES1 and TiPARP mRNA expression in HuH7 cells. HuH7 cells were treated with the DMSO or 0.01, 0.1, 1 and 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10, 100, 103, and 104nM Omp for 1.5, 6 and 24h. Results were analyzed by qPCR and expressed as the mean fold induction relative to time-matched DMSO + SEM, normalized relative to ribosomal 18s HES1 (A-C) and TiPARP (D-F). Representative data from at least two individual biological replicates are presented. Significant differences (p<0.05) relative to time-matched DMSO (*) and concentration-matched 2,3,7,8-TCDD (†) were determined by one-way ANOVA with Tukey post-hoc test.

5.3 Differential Recruitment of AHR to Response Elements in the Upstream Regulatory Regions of AHR Target Genes

I then wanted to determine if target gene mRNA expression data correlated with AHR recruitment to the upstream regulatory regions of the above mentioned target genes by utilizing ChIP assays. In addition, I wished to examine potential ligand- and cell line-dependent differences in the recruitment of AHR to CYP1A1, CYP1B1, HES1 and a cluster of AHREs 100kb from TiPARP (AHRE cluster), and if this correlates to the pre-established relative potencies of each of these ligands. T-47D or HuH7 cells were treated with the solvent control (DMSO), 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp for 45min. The 45min time-point was chosen based on previous kinetic ChIP studies in these cell lines by Pansoy et al. (2010). Based on the results of dose-response experiments, the high 10nM HAH and 10μM Omp dose was examined for all further experiments. Recruitment of AHR to these enhancer regions is determined by cross-linking proteins to DNA following treatment with an AHR inducer. After quenching the cross-links, the cells were sonicated to increase the efficiency of the assay by aiding in cell lysis and generating approximately 500bp DNA fragments. DNA that was bound to AHR was immuno-purified with protein A beads bound to antibody against AHR. Following immuno-purification, the cross-links were reversed by denaturing the proteins with buffer containing SDS and heating the samples to 65ºC. Recruitment was then quantified by qPCR, analyzed with Opticon Monitor3, and expressed as the mean percent recruitment + SEM of at least two biological replicates relative to 100% total input chromatin. Following 45min treatment, recruitment of AHR to CYP1A1, CYP1B1, HES1 and the AHRE cluster was induced by all treatments. In T-47D cells, treatment with 10 nM 2,3,7,8-TCDD and 2,3,4,7,8- PeCDF resulted in equivalent recruitment of AHR to CYP1A1, CYP1B1, HES1 and the AHRE cluster whereas lower AHR recruitment levels were observed for 10 nM 2,3,7,8-TCDF and 10 μM Omp (Figure 9A, C, E and G). A similar AHR recruitment pattern was also observed in HuH7 cells, whereby 2,3,7,8-2,3,7,8-TCDD induced the highest level of AHR occupancy at CYP1A1, CYP1B1, HES1 and the AHRE cluster while 2,3,7,8-TCDF and Omp treatments induced lower levels of AHR recruitment at these regions (Figure 9B, D, F and H). Although Omp treatment did induce AHR recruitment to levels above that of the solvent control in HuH7 cells, this increase was not statistically significant (Figure 9F).

CYP1A1

A 1.6 * * *† 1.2

0.8 *† 0.4

0 IgG AHR AHR % AHR Recruitment-% AHR T-47D AHR AHR AHR DMSO TCDD PeCDF TCDF Omp

B 1.2 * *† 1 *† *† 0.8

0.6

0.4 * 0.2

AHR Recruitment-% HuH7 0 IgG AHR AHR AHR AHR AHR DMSO TCDD PeCDF TCDF Omp

-3721 +1 CYP1A1 AHRE -3651

CYP1B1

C 7 * * *† 5 *†

1 % AHR Recruitment- T-47D 0 IgG AHR AHR AHR AHR AHR

DMSO TCDD PeCDF TCDF Omp

D * 4 *

*† *† 2

1 * % AHR Recruitment- AHR % HuH7 0 IgG AHR AHR AHR AHR AHR DMSO TCDD PeCDF TCDF Omp

-1679 +1 CYP1B1 AHRE -1580

HES1

E 0.8 * * 0.6

0.4 *† *† 0.2

% AHR Recruitment- T-47D 0

IgG AHR AHR AHR AHR AHR DMSO TCDD PeCDF TCDF Omp

F 0.4 * *

0.3 * † 0.2

0.1

Recruitment- AHR % HuH7 0 IgG AHR AHR AHR AHR AHR

DMSO TCDD PeCDF TCDF Omp

-279 +1 HES1 AHRE -215

AHRE CLUSTER

G 6 *

* * 4 *†

% AHR Recruitment- AHR % T-47D 0 IgG AHR AHR AHR AHR AHR

DMSO TCDD PeCDF TCDF Omp H 4 * * 3 * *† 2

1 % AHR Recruitment- AHR % HuH7 0 IgG AHR AHR AHR AHR AHR DMSO TCDD PeCDF TCDF Omp

-139559 +1 TiPARP AHRE

-139504

Figure 9. AHR recruitment to target gene AHREs is induced following 45min treatment. T-47D and HuH7 cells were treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp for 45min. Recruitment of AHR to CYP1A1 (A, B), CYP1B1 (C, D), HES1 (E, F), and the AHRE Cluster (G, H) was determined by ChIP assay and analyzed by qPCR. Values represent the mean + SEM relative to 100% total sample. Representative data from at least two individual biological replicates are presented. Significant differences of p<0.05 were determined by one-way ANOVA with Tukey post-hoc test relative to antibody-matched DMSO (*) and concentration-matched 2,3,7,8 -TCDD (†).

5.4 Ligand-induced Recruitment of Coactivators to the Enhancer AHREs of AHR Target Genes

To identify any ligand and/or cell-line specific differences in coactivator recruitment we used ChIP assays, as described previously, to examine recruitment profiles of p300, NCoA1 and NCoA3, which are known to be involved in AHR-dependent transcription, in extracts from T- 47D and HuH7 cells following 45 min, (Matthews et al., 2005). In addition to these coregulators, I also examined the recruitment of ERα, polII, CBP, RIP140, and PCAF; however, only p300, NCoA1 and NCoA3 gave consistent results with ChIP assay and demonstrated ligand-dependent differences. In T-47D cells, 45min treatment with all ligands induced recruitment of p300 to CYP1A1 or CYP1B1, although only Omp was found to be statistically different then 2,3,7,8-TCDD (Figure 10A and C). Conversely, none of the ligand treatments induced p300 recruitment to the HES1 enhancer region, while all treatments induced p300 recruitment to the AHRE cluster but only 2,3,4,7,8-PeCDF was significantly different than 2,3,7,8-TCDD (Figure 10E and G). 2,3,7,8-TCDF and Omp were the only treatments to induce NCoA1 recruitment to CYP1A1 with 2,3,7,8-TCDD and 2,3,4,7,8-PeCDF preferentially inducing recruitment of NCoA3 to this gene. All treatments induced recruitment of NCoA1 to CYP1B1 and the AHRE cluster, whereby no ligand preferences were observed. However, notably lower levels of NCoA3 occupancy were observed at CYP1B1 and the AHRE cluster following Omp treatment. In contrast to the other genes, only Omp treatment resulted in NCoA1 recruitment to HES1. While there appeared to be a preference for recruitment of NCoA3 to HES1 following treatment with the HAHs, only PeCDF-induced recruitment was statistically greater than that of the solvent control. In HuH7 cells, significant increases in p300 levels were observe with all ligand following 45min treatment to CYP1A1, CYP1B1 and the AHRE cluster, but not HES1 (Figure 10B, D, F and H). In contrast to T-47D cells, none of the treatments induced NCoA1 recruitment to any of the genes studied in HuH7 cells. Omp was the only treatment to induce recruitment of NCoA3 to CYP1A1, while 2,3,4,7,8-PeCDF was the only treatment to induce NCoA3 to HES1. Preferential recruitment of NCoA3 over NCoA1 to CYP1B1 and the AHRE cluster was evident for all treatments in HuH7 cells.

CYP1A1

A 0.7 * DMSO

TCDD * PeCDF 0.5 TCDF Omp *† *† 0.3 * *† * * * *

0.1 % Coactivator% Recruitment- T-47D 0 IgG p300 NCoA1 NCoA3 B 0.7 *†

0.5 *† * 0.3

* 0.1 % Coactivator% Recruitment- HuH7 0 IgG p300 NCoA1 NCoA3

-3721 +1 CYP1A1 AHRE -3651

CYP1B1

C 2.5 DMSO * TCDD * 2 PeCDF *† TCDF Omp 1.5 *† ** 1 * *†

0.5 * * **† % Coactivator% Recruitment- T-47D 0 IgG p300 NCoA1 NCoA3

D 2.5

1.5

1 ** * * * ** * 0.5 % Coactivator% Recruitment- HuH7 0 IgG p300 NCoA1 NCoA3

-1679 +1 CYP1B1 AHRE -1580

HES1

E 0.14 DMSO * TCDD PeCDF 0.1 TCDF Omp

* 0.06

0.02

% Coactivator% Recruitment- T-47D 0 IgG p300 NCoA1 NCoA3

F 0.16 *

0.12

* * 0.08

* 0.04

Coactivator% Recruitment- HuH7 0 IgG p300 NCoA1 NCoA3

-279 +1 HES1 AHRE -215

AHRE Cluster G

3.5 DMSO TCDD * PeCDF 2.5 TCDF * * Omp *† 1.5

* * *† * *† 0.5 * * *

Coactivator% Recruitment- T-47D 0 IgG p300 NCoA1 NCoA3

H 0.9 * * *

0.7 * * * * *† 0.5

0.3

0.1 % Coactivator% Recruitment- HuH7 0 IgG p300 NCoA1 NCoA3

-139559 +1 TiPARP AHRE

-139504 Figure 10. Recruitment of coactivators to AHREs of AHR-responsive genes following 45min treatment. T-47D and HuH7 cells were treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp for 45min. Recruitment of selected coactivators to the AHREs of CYP1A1 (A, B), CYP1B1 (C, D), HES1 (E, F), and the AHRE Cluster (G, H) was determined by ChIP assay and analyzed by qPCR. Values are expressed as the mean + SEM, relative to 100% total sample; representative data from at least two individual biological replicates is presented. Significant differences (p<0.05) were determined by one-way ANOVA with Tukey post-hoc test relative to antibody-matched DMSO (*) and 2,3,7,8-TCDD (†).

5.5 Omeprazole Induced Nuclear AHR Degradation

Previous studies have demonstrated that following AHR activation by HAHs such as 2,3,7,8- TCDD, 2,3,4,7,8-PeCDF and 2,3,7,8-TCDF, AHR signalling is attenuated by degradation through the 26S proteasome pathway (Ma and Baldwin, 2000; Pansoy et al., 2010; Pollenz, 2002). While degradation has been well-documented to occur following activation of AHR by HAHs, the effect of atypical inducers, such as Omp, on AHR degradation remains to be investigated. As such, I wished to examine whether Omp treatment would induce AHR degradation. To determine differences in ligand-induced proteolytic degradation of AHR, I treated T-47D cells with the solvent (DMSO), 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF and 2,3,7,8-TCDF, or 10μM Omp for 1, 3 and 24h, then collected and lysed cells. These time-points were chosen based on a previous study of AHR degradation following ligand treatment by our laboratory (Pansoy et al., 2010). I examined AHR degradation in whole cell lysates using Western Blots incubated with antibody against AHR, and β-actin as a control for protein loading. In agreement with previous studies, treatments with 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8- TCDF resulted in proteolytic degradation of AHR following 3h treatment, whereas no Omp- induced AHR degradation was observed at any of the time-points studied (Figure 11). These results indicated that either (1) Omp-treatment was unable to induce proteolytic degradation of AHR and AHR signalling is attenuated via an alternative mechanism, or (2) Omp activated less AHR molecules thus making it more difficult to detect subtle changes in concentration. As such, I examined AHR degradation in both cytoplasmic and nuclear extracts to attempt to elucidate this mechanism. Since all HAHs induced relatively similar levels of AHR turnover in whole cell lysate studies, I only examine the effects of DMSO (solvent control), Omp and 2,3,7,8-TCDD in nuclear and cytoplasmic extracts.

To test whether any perceived changes in AHR concentration were associated with 26S proteasome activity, I examined the effects of 1h pre-treatment with MG-132, which has been shown to exhibit specific 26S proteasome inhibition (Anwar-Mohamed et al., 2008; Davarinos and Pollenz, 1999). Previous experiments by our laboratory have demonstrated that greater than 5h total exposure to MG-132 following treatment results in cell death (unpublished data). As such, I only examine the 1 and 3h time-points for these studies. T-47D cells were treated with DMSO, 10nM 2,3,7,8-TCDD, or 10μM Omp for 1 or 3h then harvested cells and fractionated them into nuclear and cytoplasmic extracts. I visualized differences in AHR by Western blot

58 with antibody against AHR, p53 and β-actin. I chose to immuno-blot for p53 as a control for both protein loading and efficiency of nuclear extraction, since p53 exists solely in the nucleus. Conversely, β-actin was used as a control for cytoplasmic contamination, and as a control for loading of the cytoplasmic fraction. These immuno-blots revealed that both 2,3,7,8-TCDD and Omp induced AHR turn-over in nuclear fractions of T-47D cells (Figure 12). Treatment with the solvent control (DMSO) or co-treatment with MG-132, was shown to have no effect on nuclear and cytoplasmic AHR concentrations (Figure 12A). In addition, both 2,3,7,8-TCDD- and Omp-mediated AHR degradation could be inhibited by 1h pre-treatment with MG-132, indicating that Omp induces AHR degradation through the 26S proteasome pathway (Figure 12B and C).

DMSO 0h 1h 3h 24h

AHR TCDD

β-Actin

AHR PeCDF β-Actin

AHR TCDF β-Actin

AHR Omp β-Actin

Figure 11. Omeprazole treatment does not induce AHR degradation in whole cell lysates. T-47D cells were treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp for 0, 1, 3, and 24h. Whole cell lysates were analyzed by Western blot probed with antibody against AHR and β-actin as a loading control. Data is representative of at least 2 biological replicates.

59 A - DMSO 3 3 MG132 - - + AHR p53 Nuclear

Β-actin

AHR

p53 Cytoplasmic

Β-actin

B TCDD 0 1 3 0 1 3 MG132 - - - + + + AHR

p53 Nuclear Β-actin

AHR Cytoplasmic p53 Β-actin

C Omp 0 1 3 0 1 3 MG132 - - - + + + AHR

p53 Nuclear Β-actin

AHR

p53 Cytoplasmic Β-actin

Figure 12. Omeprazole treatment induces proteolytic degradation of AHR in nuclear extracts. T-47D cells were treated with DMSO (A), 10nM 2,3,7,8-TCDD (B) or 10μM Omp (C) for 0, 1, and 3h with or without 1h pre-treatment with 26S proteasome inhibitor MG-132. Cells were fractionated and a Western blot of nuclear and cytoplasmic extracts was probed for AHR, p53 as a control for efficiency of nuclear extract and loading, and β-actin as a control for cytoplasmic contamination of nuclear samples and cytoplasmic loading.

5.6 The Role of Y322 in Ligand-Induced AHR Signalling

There is some controversy as to the role of residue Y322 in binding-dependent activation of AHR signalling. Previous studies have postulated that activation of AHR by atypical inducers requires phosphorylation of a residue within the LBD of AHR, whereas co-treatment with a c-src tyrosine kinase inhibitor abolished Omp-induced AHR signalling but had no effect on 2,3,7,8- TCDD-induced AHR activation (Backlund and Ingelman-Sundberg, 2005; Blankenship and Matsumura, 1997; Enan and Matsumura, 1996; Lemaire et al., 2004). To examine the role of residues within the LBD in AHR activation, Backlund and Ingelman-Sundberg (2004) investigated the effect of substitution of selected residues within the LBD on AHR activation by both 2,3,7,8-TCDD and Omp using a Gal4-AHR reporter gene system. In this study, it was demonstrated that residue Y320 of rAHR (equivalent to human Y322) was required for Omp- induced AHR activation but had no effect on 2,3,7,8-TCDD-induced AHR activation (Backlund and Ingelman-Sundberg, 2004). Contrary to these findings, Pandini et al (2009) identified the equivalent mAHR residue (Y316) as one of the conserved amino acids necessary for 2,3,7,8- TCDD-binding and DNA-binding of AHR, whereby mutation to either Y316F or Y316A significantly impaired AHR ligand- and DNA-binding activity (Pandini et al., 2009). Despite there being previous studies conducted in both mouse and rat, the functional relevance of the human equivalent residue (hAHR-Y322) has yet to be established. To examine the role of hAHR-Y322 in ligand-mediated activation of human AHR signalling, I transiently transfected MCF-7 AHR100 human breast cancer cells, that have been chronically exposed to benzo[a]pyrene to express very low levels of AHR (Ciolino et al., 2002), with empty pRC- CMV2, wt-hAHR, Y322F, or Y322A variants. The Y322F variant was studied for its ability to prevent phosphorylation while conserving the cyclic ring structure of tyrosine at this site. This mutation has been reported to have a minimal effect on 2,3,7,8-TCDD-dependent activation of CYP1A1-mediated reporter gene activity while it has been shown to impair the ability of 2,3,7,8- TCDD to bind to the receptor (Backlund and Ingelman-Sundberg, 2004; Pandini et al., 2009). The Y322A was studied since an equivalent mutation in mAHR abolishes AHR function (Pandini et al., 2009).

To confirm that the variants were expressed in transfected AHR100 cells in relatively equivalent amounts to that of the pRC-CMV2-wt-hAHR vector, I performed a Western blot of

A Em wt-hAHR Y322F Y322A

AHR

B-Actin

B Em wt-hAHR Y322F Y322A

AHR

Β-Actin

Figure 13. Confirmation of variant expression in transiently transfected MCF-7 AHR100 cells. MCF-7 AHR100 breast cancer cells were transfected with empty pRC-CMV2 plasmid or one of wt-hAHR, Y322F, and Y322A a Western blot of whole cell extracts was probed with antibodies against AHR and β-actin (loading control) to confirm equivalent protein expression of the variants. A, Immuno-blots against AHR and β-actin (loading control) are shown following 1min exposure, respectively. B, 5min and exposure revealed a band corresponding to endogenous AHR in cells expressing the empty vector.

CYP1A1

A 120 Empty *† wt-hAHR *† 100 *† *† Y322F 80 Y322A *† 60 *† 40 Treated wt-AHR

20 % Induction Relative to TCDD TCDD to Relative Induction % 0

B DMSO TCDD PeCDF TCDF Omp 0.35 *† 0.3 *† 0.25 *†

0.2 † *† 0.15 † *

0.1

0.05

AHR Recruitment-% CYP1A1 0 IgG AHR AHR AHR AHR AHR

DMSO TCDD PeCDF TCDF Omp

-3721 +1 CYP1A1 AHRE -3651

Figure 14. Y322 of AHR is required for maximal recruitment and CYP1A1 induction. AHR100 cells were transfected with empty pRC-CMV2 plasmid or wt-hAHR, Y322F and Y322A, and treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp and CYP1A1 mRNA expression was analyzed by qPCR and expressed as percentage relative to wt-AHR 2,3,7,8-TCDD-treated samples, normalized relative to ribosomal 18s (A). ChIP assay of AHR recruitment was analyzed by qPCR and results were expressed relative to 100% total input (B). Values represent the mean + SEM values; representative data from at least two individual biological replicates are shown. Significant differences of p<0.05 were determined by one-way ANOVA with Tukey post-hoc test relative to plasmid-matched DMSO (*) and treatment matched empty plasmid (†).

CYP1B1

A 120 Empty *† *† *† Wt-hAHR 100 Y322F *† 80 Y322A *† *† † 60

† † * 40 * *

Treated wt-AHR *

20 % Induction% Relative to TCDD 0 DMSO TCDD PeCDF TCDF Omp B 2 *† *† *†

1.6 † † 1.2

0.8 * * * ** *

% AHR Recruitment AHR % 0.4

0 IgG AHR AHR AHR AHR AHR DMSO TCDD PeCDF TCDF Omp

-1679 +1 CYP1B1 AHRE -1580 Figure 15. Y322 of AHR is required for maximal recruitment and CYP1B1 induction. Cells were treated with DMSO, 10nM 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF or 2,3,7,8-TCDF, or 10μM Omp and CYP1A1 mRNA expression was analyzed by qPCR and is presented as a percentage relative to wt-AHR samples treated with 2,3,7,8-TCDD and normalized relative to ribosomal 18s (A). ChIP assay of AHR recruitment to CYP1B1 was analyzed by qPCR and results were expressed relative to 100% total input (B). Values represent the mean + SEM; representative data from at least two individual biological replicates are shown. Significant differences of p<0.05 were determined by one-way ANOVA with Tukey post-hoc test relative to plasmid-matched DMSO (*) and treatment matched empty plasmid (†).

whole cell extracts from untreated but transiently transfected cells (Figure 13A). The immuno- blot indicated that the wt-hAHR, Y322F and Y322A variants were expressed at equivalent levels. Overexposure revealed a faint band corresponding to low levels of remaining endogenous AHR in cells transfected with the empty plasmid, which was evident in my results for CYP1A1 expression and AHR recruitment (Figure 13B). To examine the role of hAHR- Y322 in AHR signalling, I investigated the ability of each variant to induce expression of CYP1A1 and CYP1B1 mRNA following 6h treatment. In addition, I examined recruitment of AHR to the enhancer regions of these genes following 45min treatment. I found induction of HES1 and TiPARP mRNA to be too low in AHR100 cells expressing wt-hAHR to properly quantify differences due to Y322F and Y322A expression. Over-expression of the empty plasmid resulted in a small increase in CYP1A1 mRNA expression levels relative to DMSO for 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF and 2,3,7,8-TCDF (Figure 14A). A similar pattern of expression was observed for CYP1B1 expression, whereby all ligand treatments resulted in an increase in expression relative to DMSO (Figure 15A). It is likely that this increase in cells transfected with the empty plasmid is due to the presence of endogenous AHR. Transfection of AHR into AHR deficient cell-lines has been previous shown by both our laboratory and others to result in high background levels in solvent control (DMSO) (Celius and Matthews, 2010; Pandini et al., 2009; Roblin et al., 2004). However, the high background (DMSO) of CYP1A1 mRNA expression was eliminated in cells transfected with the mutant AHRs compared to wild-type. Pandini et al. (2009) previously demonstrated that mAHR-Y322A mutation resulted in a 93% reduction in binding of 2,3,7,8-TCDD to AHR, and a 96% reduction in AHRE binding by AHR (Pandini et al., 2009). Consistent with previous findings for the mAHR-Y316A, the equivalent hAHR-Y322A mutation was incapable of inducing AHR-dependent regulation of CYP1A1 mRNA levels. This pattern was also observed for CYP1B1 mRNA induction, whereas none of the ligands were able to induce expression of CYP1B1 in cells transiently transfected with the Y322A variant. In previous studies by Pandini et al. (2009), the Y316F mutation of mAHR drastically reduces 2,3,7,8-TCDD-inducible binding of AHR to AHREs (Pandini et al., 2009). Consistent with the findings of this study, AHRs containing Y322F mutation supported induction of CYP1A1 by the higher-affinity ligands 2,3,7,8-TCDD- and 2,3,4,7,8-PeCDF, albeit expression was diminished by approximately 50% relative to wt-hAHR. This mutation did, however, prevent 2,3,7,8-TCDF- and Omp-induced CYP1A1 and CYP1B1 mRNA expression.

To further elucidate the role of the Y322 residue in AHR signalling, I examined the effects of mutations to this residue on AHR recruitment to AHREs in the enhancer regions of both CYP1A1 and CYP1B1. To examine differences in the recruitment of these mutants to CYP1A1 and CYP1B1 relative to wild-type, I performed ChIP assays as previously described. No significant increase in AHR recruitment was observed for any ligand relative to DMSO at the CYP1A1 enhancer in cells transfected with the empty plasmid (Figure 14B). Consistent with mRNA findings, in cells transfected with the empty plasmid all treatments induced recruitment of AHR to the CYP1B1 enhancer relative to DMSO (Figure 15B). This provided further evidence for the presence of low levels of endogenous AHR in this AHR-deficient cell line. Consistent with the observed increases CYP1A1 mRNA expression, an increase in wt-hAHR recruitment to the CYP1A1 enhancer was observed in cells treated with the solvent control (DMSO). The is consistent with previous work by our laboratory as well as others (Celius and Matthews, 2010; Pandini et al., 2009; Roblin et al., 2004). All ligand treatments resulted in increased recruitment of wt-hAHR to the CYP1A1 and CYP1B1 AHREs. Consistent with mRNA findings, hAHR-Y322F expression resulted in approximately a 50% reduction in 2,3,7,8-TCDD- and 2,3,4,7,8-PeCDF-induced AHR recruitment to CYP1A1. Similarly, 2,3,7,8-TCDF- and Omp-induced AHR recruitment was attenuated by this mutation. In support of these findings for both mRNA expression and CYP1A1 binding, hAHR-Y322F resulted in approximately 60% reduction in AHR recruitment to CYP1B1 relative to wt-hAHR, while this mutation prevented 2,3,7,8-TCDF-and Omp-induced AHR recruitment.

6 Discussion, Conclusions, Limitations and Future Aims 6.1 Discussion

6.1.1 Summary

AHR is a ligand-dependent transcription factor that mediates the actions of many HAHs, PAHs, phytochemicals and pharmaceuticals, and is involved in the expression of many target genes including the most studied AHR-responsive gene CYP1A1. As such, I compared the ability of various AHR activators to regulate the expression of CYP1A1, CYP1B1, HES1 and TiPARP expression. I focused on these genes because they represent different classes of genes that have AHR recruited to their regulatory regions and whose transcriptional activation is mediated by AHR. HAH-induced CYP1A1 and CYP1B1 expression continued to increase up to 24h, while HES1 and TiPARP increased after 1.5h, but was reduced at 6h and modestly increased at 24h. No increase in Omp-induced CYP1A1 and CYP1B1 mRNA expression was observed following 6h treatment, which is likely due to the stability of this compound and the antagonistic effects of its metabolic byproduct on AHR (Gerbal-Chaloin et al., 2006; Nagaoka et al., 2006; Sivertsson et al., 2010). At sub-maximal concentrations, mRNA expression data for CYP1A1, CYP1B1 and TiPARP reflected the relative potencies of the HAH compounds. Few differences were observed at the maximal dose, indicating that the receptor is likely saturated at this concentration. To examine cell line-, gene-, and ligand-dependent differences, I performed ChIP assays to investigate AHR and coactivator recruitment to the upstream regulatory regions of CYP1A1, CYP1B1, HES1 and an AHRE cluster. Following 45min treatment, patterns for AHR recruitment to the upstream enhancer regions of CYP1A1, CYP1B1 and HES1 closely mimicked the relative potencies of the HAHs in T-47D and HuH7 cells, as well as the expression profiles for target genes. Recruitment to an AHRE cluster 100kb from TiPARP reflected the relative potency of the HAH ligands in HuH7 cells, but not T-47D cells. This region was examined since previous studies by our laboratory indicated that it was the highest ranked AHR-binding regions in promoter-focused microarrays (Ahmed et al., 2009; Pansoy et al., 2010). Omp induced AHR recruitment to the AHREs of all genes studied, albeit lower levels than all ligands studied which is likely because it is a relatively weak AHR activator (Backlund and Ingelman-Sundberg, 2004). Gene-, cell line- and ligand-dependent differences were observed for the recruitment of coactivators with similar function following 45min treatment. My findings demonstrate that

there is little correlation between the relative potency of ligands and coactivator recruitment. This is perhaps because recruitment of coactivators is not dependent on the magnitude of AHR response but, rather, whether AHR signalling has been activated or not. While it is well- documented that HAH treatment results in attenuation of AHR signalling by AHR degradation, I wished to examine whether or not Omp treatment would induce AHR degradation. A Western blot of whole cell lysates demonstrated that 2,3,7,8-TCDD, 2,3,4,7,8,-PeCDF and 2,3,7,8-TCDF induced AHR degradation following 3h treatment, while Omp did not induce degradation at any of the time-points studied. However, a Western blot of nuclear extracts demonstrated that both 2,3,7,8-TCDD and Omp treatments induced AHR degradations that could be inhibited by 1h pre- treatment with the 26S proteasome inhibitor MG-132. These findings suggest that changes in AHR concentration resulting from Omp treatment are too subtle to be detected by immuno-blot of whole cell lysates. There have been conflicting studies on the role of the mouse and rat equivalents of hAHR-Y322 in ligand-mediated AHR activation. As such, I wished to investigate the role of this residue in AHR signalling by examining the effects of Y322F and Y322A mutations on the regulation of and the recruitment of AHR to CYP1A1 and CYP1B1. Y322F mutation reduced gene induction and AHR recruitment by 2,3,7,8-TCDD and 2,3,4,7,8-PeCDF, while ablating the effects of 2,3,7,8-TCDF and Omp treatment. Y322A mutation attenuated the effects of all ligands on AHR signalling, indicating that Y322 is required for maximal AHR induction.

6.1.2 Activator, Gene and Cell Line Differences in AHR-Mediated Expression of Target Genes

TEF values represent the relative potencies of compounds capable of binding to AHR, and are established by analyzing all available in vivo and in vitro toxicity data. Zhang et al. (2008) previously showed that HAH- and PAH-induced CYP1A1 mRNA expression to TEF values and found that potency was not predictive of target gene expression in Panc1, HEK293 and Hepa1c1c7 cells. To determine if the predictive validity of TEF values is cell line-or gene- dependent, I examined temporal differences in expression of CYP1A1, CYP1B1, HES1, and TiPARP mRNA following 1.5, 6, and 24h treatments in T-47D and HuH7 cells. All genes exhibited a different level of expression following ligand treatments in both T-47D and HuH7, indicating that AHR-mediated expression is both gene- and cell line-dependent. I found that

each ligand induced CYP1A1 and CYP1B1 mRNA expression that continued to increase from 1.5 to 24h. Following 24h treatment, sub-maximal doses of 2,3,4,7,8-PeCDF induced CYP1A1 mRNA expression was approximately 85% that of 2,3,7,8-TCDD-induced expression, while 2,3,7,8-TCDF was approximately 37%. These results correlate well with the pattern of ligand- induced mRNA expression that is predicted by the TEF values, whereby 2,3,7,8-TCDD has a TEF of 1.0, 2,3,4,7,8-PeCDF of 0.3, and 2,3,7,8-TCDF of 0.1. There is no TEF value for Omp since it does not fulfill most of the basic requirements for assignment of a TEF, whereas it is structurally dissimilar to PCDDs and PCDFs, it is believed to be unable to bind to AHR, and it does not accumulate in the food chain. In T-47D cells, treatment resulted in greater induction of CYP1A1 mRNA than CYP1B1, whereas in HuH7 cells CYP1B1 mRNA expression was substantially greater than CYP1A1, indicating cell line-selectivity. Treatment with HAHs resulted in increasing CYP1A1 and CYP1B1 expression to 24h, while expression in cells treated with Omp leveled off after 6h treatment, with only the high dose inducing expression at this time-point in both cell lines studied. In addition, Omp induced lower levels of expression of TiPARP mRNA in T-47D and HuH7 cells, and HES1 in HuH7 cells. These temporal differences in HAHs and Omp-induced AHR-responsive gene expression may be attributed to the relative stability of the compounds. HAHs are known to be extremely stable and, due to the high lipophilicity of these compounds, they are known to accumulate in adipose tissue and undergo very little hepatic metabolism, whereas Omp is readily metabolized by CYP3A4 and CYP2C19 which are expressed in both of the cell lines studied, and possesses a half-life of approximately 2.17h when administered clinically to humans (Howden et al., 1984). Additionally, this plateau may be attributed to the presence of Omp’s degradation metabolite, omeprazole sulphide. Omeprazole has been shown to undergo biotransformation to omeprazole sulphide which acts as an antagonist of AHR and can inhibit nuclear translocation of AHR (Gerbal-Chaloin et al., 2006; Nagaoka et al., 2006; Sivertsson et al., 2010). To examine whether temporal differences between HAHs and Omp in target gene expression are due to metabolism, the concentrations of metabolites of omeprazole in the media could be tested. In addition, by targeting the pregnane X receptor (PXR) to control induction, CYP3A4 expression could be either decreased or super- induced to look at its effect on Omp-mediated signalling. Decreasing CYP3A4 expression should prolong the half-life of Omp and the plateau in CYP1A1 and CYP1B1 expression should disappear, while super-inducing CYP3A4 should prevent Omp-mediated activation of AHR signalling.

While CYP1A1 and CYP1B1 mRNA induction continually increased in a temporally- dependent manner, HES1 and TiPARP mRNA expression appeared to be oscillatory, whereby all ligands induced expression of both genes at 1.5h, then by 6h expression returned to comparable levels to that of the solvent control. At 24h, mRNA expression levels appeared to rebound slightly for all treatments in both T-47D and HuH7 cells for TiPARP, and HuH7 cells for HES1. These findings demonstrate distinct regulation of HES1 and TiPARP relative to CYP1A1 and CYP1B1. In addition, these findings correlate well with previous studies demonstrating maximal HES1 expression following 2h 2,3,7,8-TCDD treatment and oscillatory expression of HES1 (Thomsen et al., 2004). Oscillatory expression of HES1 has been previously documented by both our lab and others, whereby temporal changes in HES1 mRNA expression were observed in the 4.5h following treatment, and oscillation was shown to occur every 2h, correlating well with 2h half-life of HES1 protein (Hirata et al., 2002; Pansoy et al., 2010; Yoshiura et al., 2007). This pattern of expression may be attributed to auto-regulation of HES1, since expression of the HES1 protein has been shown to repress its transcriptional activation and decrease half-life which modulates cell-cycle progression and proliferation (Castella et al., 2000; Muller et al., 2002; Strom et al., 2000; Thomsen et al., 2004; Yoshiura et al., 2007). Like HES1, TiPARP is postulated to be under auto-regulatory control and to repress AHR activity (our laboratory’s unpublished data). My findings agree with previous reports by our laboratory demonstrating oscillatory expression of TiPARP (Ahmed et al., 2009).

6.1.3 Comparison of Activator-Induced AHR Recruitment to the Upstream Regulatory Regions of AHR Target Genes

To correlate mRNA expression with AHR and coactivator recruitment, I performed ChIP assays. In addition, I compared my findings for target gene induction and AHR recruitment by HAHs studied to their relative potencies to determine whether these values are reflective of the magnitude of AHR response in T-47D and HuH7 cells. A good correlation between TEF values and AHR recruitment to the regulatory regions of CYP1A1, CYP1B1 and HES1 was observed at the 10nM dose following 45min treatment in both T-47D and HuH7 cells, whereby high affinity ligands such as 2,3,7,8-TCDD and 2,3,5,7,8-PeCDF induced greater AHR recruitment than the low affinity ligand 2,3,7,8-TCDF and Omp. In turn, this pattern was mirrored in the mRNA expression of CYP1A1 and CYP1B1 but not HES1. Taken with the above mRNA expression

data, this suggests that ligand affinity affects AHR recruitment to the regulatory regions of genes. In addition, this demonstrates that TEF values are sometimes predictive of AHR recruitment and target gene induction in T-47D and HuH7 cells for these genes. Conversely, recruitment of AHR to the AHRE cluster closely followed the predicted potencies of these ligands in HuH7 cells but not T-47D cells. As such, it is likely that the predictive validity of TEF values for individual studies is both cell line- and context-dependent. While it is possible that there could be a mutation in AHR that confers the cell line-specific differences, the likelihood that these small nucleotide polymorphisms (SNPs) have an effect on binding affinity is very low (Celius and Matthews, 2010).

6.1.4 Activator-, Cell Line- and Gene-Dependent Differences in Coactivator Recruitment

Coregulators (coactivators and corepressors) are recruited to AHREs within the upstream regulatory regions of AHR-responsive genes following AHR activation, and function to enhance or repress gene expression (Hankinson, 2005). There is a certain degree of functional redundancy in coactivators that are recruited in response to AHR activation, which may be attributed to cell-line and gene differences (Hankinson, 2005). In particular, I examined differences in the recruitment of p300, NCoA1 and NCoA3, which have been shown to illicit histone acetyltransferase (HAT) activity (Beischlag et al., 2002; Kobayashi et al., 1997). Transcriptionally inactive or silenced genes exist tightly packed into nucleosomes composed of 147 bp of DNA wrapped around an octomer of four core histone proteins (H2A, H2B, H3, H4) and a linker histone H1, and whose N-terminal histone tail possesses a large number of amino acid residues that can be modified covalently (Kouzarides, 2007). The tight packing of nucleosomes prevents transcription factors from binding to their DNA recognition sequences. Chromatin remodeling is required for transcriptional activation of these genes, and is achieved through modifications to core histones, including acetylation and methylation, by such coactivators as HATs (Hankinson, 2005; Kouzarides, 2007). In addition, a previous study by Zhang et al. (2008) demonstrated cell line- and ligand-dependent differences coactivator activation in mammalian two-hybrid assays. As such, I wished to examine coactivator recruitment to the AHREs of CYP1A1, CYP1B1, HES1 and the AHRE cluster to compare the

effects of ligand, cell-line, and genes on recruitment following 45min treatment in a cellular system.

No correlation between the relative potencies of the HAH compounds and coactivator recruitment to AHREs of the target gene enhancer regions was observed. This is likely since coactivator recruitment occurs as a result of transcriptional activation of the gene, so the magnitude of AHR activation does not affect recruitment. As such, coactivator recruitment is likely an “on or off,” rather than a potency-dependent effect. Few differences were observed in p300 recruitment to any of the genes studied, following 45min treatment. In T-47D cells, there appears to be a preference for recruitment of NCoA3 by higher affinity ligands, 2,3,7,8-TCDD and 2,3,4,7,8-PeCDF, to the CYP1A1 enhancer, while treatment with lower affinity ligand 2,3,7,8-TCDF, and Omp induced recruitment of NCoA1. This pattern was not observed for recruitment to CYP1B1, the AHRE cluster or in HuH7 cells; while there appeared to be preference for recruitment of NCoA3 to HES1 following treatment with the HAHs, though only 2,3,4,7,8-PeCDF-induced recruitment statistically greater than the solvent control. These cell- line differences in NCoA1 recruitment have been previously documented in mammalian two- hybrid studies by Zhang et al. (2008), whereby NCoA1 was activated by 2,3,7,8-TCDD and 2,3,4,7,8-PeCDF treatment in HEK293 but not Panc1 and Hepa1c1c7 cells, while 2,3,7,8-TCDF did not induce recruitment of NCoA1 in any of these cell-lines (Zhang et al., 2008). These findings support previous evidence that recruitment of NCoA1 and NCoA3 is both cell line- and gene-dependent (Hankinson, 2005; Kadonaga, 2004; Khorasanizadeh, 2004; Zhang et al., 2008). Overall, this demonstrates that ligand-, cell line-, and gene-dependent preferences exist for recruitment of functionally redundant coactivators. Cell line-dependent preferences in expression of one coactivator over another may also be attributed to differences in the level of expression of these factors in the cell lines studied. Since I examined selected coactivators with HAT activity, gene-dependent differences in recruitment may be due to the level of transcriptional activity of the gene. Genes that highly are transcriptionally active, such as the oscillatory protein HES1, are likely to exhibit a more relaxed chromatin structure and remain transcriptionally derepressed. In turn, these genes are likely to require lower levels of chromatin remodelling HAT coactivators. Differences in the overall recruitment levels among coactivators may be due to efficacy of the antibodies used for these assays, whereby the antibody against one

coactivator may be better able to recognize the epitope of that protein, then another antibody against a different coactivator.

6.1.5 Effect of Omp Treatment on AHR Degradation

Previous studies have demonstrated that activation of AHR by halogenated aromatic compounds results AHR down-regulation through the ubiquitin-26S proteasomal degradation pathway. These studies did not, however, examine whether activation of AHR by an atypical inducer also results in AHR degradation (Pollenz, 2002). To investigate whether Omp treatment resulted in AHR degradation, I performed a Western blot of whole cell lysates following 1, 3, and 24h exposures. This immuno-blot demonstrated that 2,3,7,8-TCDD, 2,3,4,7,8-PeCDF and 2,3,7,8- TCDF induced comparable levels of AHR degradation following 3h treatment, while Omp did not result in any observable level of AHR degradation at any of the time-points studied. These results indicated that either (1) Omp-treatment was unable to induce proteolytic degradation of AHR and AHR signalling is attenuated by the AHRR or an alternative mechanism potentially involving dephosphorylation of AHR by a phosphatase, or (2) since Omp induced much less AHR recruitment to the enhancer regions and lower levels of target gene induction, it is likely that the assay is not sensitive enough to detect subtle changes in AHR concentration. To further investigate whether Omp treatment was capable of inducing AHR degradation, I examined degradation in nuclear extracts treated for 1 and 3h with and without MG-132 pre-treatment by performing a Western blot. A previous study by Ma and Baldwin (2000) demonstrated that inhibition of the 26S proteasome with MG-132 pre-treatment induced an increase in 2,3,7,8- TCDD-mediated AHR: ARNT nuclear accumulation resulting in a super induction of CYP1A1 expression (Ma and Baldwin, 2000). Consistent with previous findings, I observed substantial AHR degradation following 3h treatment with 2,3,7,8-TCDD, whereby MG-132 pre-treatment resulted in an accumulation of AHR in nuclear fractions. Omp also induced AHR degradation in nuclear extracts following 3h treatment but to a seemingly lesser degree then 2,3,7,8-TCDD. In addition, MG-132 pre-treatment induced nuclear accumulation of AHR, once again to a lesser degree than 2,3,7,8-TCDD. This suggests that, Omp-induced AHR activation results in degradation of the receptor through the 26S proteasome pathway. Omp treatment induced less nuclear localization of AHR and subsequent degradation than the high-affinity 2,3,7,8-TCDD

which supports mRNA and ChIP data for AHR recruitment, whereby Omp induced lower levels of target gene induction and AHR recruitment than high affinity ligands. Diminished AHR activation over-time can also be associated with the observed pattern of AHR degradation, suggesting that the relative potency of the ligand for the receptor plays a role in the extent of AHR activation and degradation. These findings indicate that Omp-induces AHR degradation through the 26S proteasome pathway, but that changes in the overall concentration of AHR are too subtle to be detected in whole cell lysates. In addition, this further demonstrates that there was little difference between the affects of Omp, an atypical inducer, and a low affinity ligand.

6.1.6 Role of Y322 in Ligand-Mediated AHR Activation and Target Gene Expression

While the role of residue Y322 in AHR activation is controversial, the phosphorylation status of tyrosine residues in AHR has been shown to affect receptor function, including activation and DNA binding activity (Park et al., 2000). While studies by Backlund and Ingel-Sundberg (2004) investigating the Y320F mutation suggest that the equivalent residue in the rat AHR LBD is required for activation by the Omp in a Gal-4 reporter system, it had no effect on 2,3,7,8-TCDD- induced activation. Contrary to these findings, Pandini et al. (2009) demonstrated that the equivalent residue of mouse AHR was necessary for maximal activation by 2,3,7,8-TCDD, whereby Y316F mutation of mAHR resulted in a 55% reduction of TCDD binding, and 16% reduction in 2,3,7,8-TCDD-induced AHRE binding. Pandini et al. (2009) also demonstrated that alanine substitution at the equivalent residue of mouse AHR resulted in a 93% reduction of 2,3,7,8-TCDD specific binding and a 96% reduction in TCDD-inducible AHRE binding, relative to wt-mAHR. Pandini and collegues did not, however, investigate the effects of this mutation on Omp-mediated activation, and neither study examined 2,3,7,8-TCDD-binding or AHRE binding in a cellular system. As such, I sought to elucidate the role of Y322 in activation of human AHR by examining the effects of Y322F and Y322A mutations on AHR recruitment and target gene expression. In MCF-7 AHR100 cells engineered to be AHR-deficient due to chronic exposure to benzo[a]pyrene, over-expressing wt-hAHR exhibited significant basal binding of AHR to the CYP1A1 and CYP1B1 enhancers which correlates with a high basal CYP1A1 and CYP1B1 mRNA expression. Interestingly, this basal binding to either CYP1A1 or CYP1B1 could be abolished by the introduction of either Y322F or Y322A variants. This effect has been

previously documented by both our laboratory and others (Celius and Matthews, 2010; Pandini et al., 2009; Roblin et al., 2004).

In agreement with the latter study by Pandini et al, my findings revealed that Y322 of the hAHR LBD is required for maximal activation by both binding-dependent and atypical AHR inducers. Mutation of tyrosine to phenylalanine is believed to conserve the cyclic structure of the residue within the LBD without allowing for phosphorylation, although the crystal structure of the hAHR LBD has not been published to confirm this. In agreement with the findings of Pandini et al. (2009), my study demonstrates that over-expression of Y322F results in an approximately 50% decrease in 2,3,7,8-TCDD- and 2,3,4,7,8-PeCDF-dependent CYP1A1 mRNA induction and AHR recruitment relative to wt-hAHR, while the effects of the lower- affinity compounds 2,3,7,8-TCDF and Omp were ablated. Similar results were also observed for CYP1B1, whereby over-expression of the Y322F variant resulted in an approximately 60% reduction in 2,3,7,8-TCDD- and 2,3,4,7,8-PeCDF-induced mRNA expression and AHR recruitment, while this mutation abolished the effects of 2,3,7,8-TCDF and Omp. Consistent with these findings, over-expression of the hAHR-Y322A variant abolished the effects of all ligands on CYP1A1 and CYP1B1 mRNA expression levels and AHR recruitment. This suggests that Y322 is required for maximal ligand-induced AHR activation and that this residue plays a role in ligand-affinity for the receptor. A great degree of similarity in the effects of these mutations was observed for the higher affinity ligands 2,3,7,8-TCDD and 2,3,4,7,8-PeCDF, while similarly little difference in CYP1A1 or CYP1B1 mRNA expression and AHR recruitment was observed between low affinity ligand 2,3,7,8-TCDF and Omp. In addition to similarities between the low affinity ligand TCDF and the atypical inducer Omp for mRNA induction, AHR recruitment, coactivator recruitment, and attenuation of AHR signalling, little difference was observed in CYP1A1 and CYP1B1 mRNA expression levels and AHR recruitment in response to over-expression of AHR variants. Overall, these results lend support to the theory that Omp is actually a weakly-bound AHR ligand, although further experiments are required to confirm this interaction (Denison and Nagy, 2003). However, to further elucidate the mechanism of AHR activation by Omp, competitive binding assays with radio-labelled ligand are required, in addition to re-examining the role of tyrosine kinases in Omp-mediated AHR activation.

6.2 Conclusions

The findings of my study demonstrate ligand-, gene-, and cell line-dependent differences, and provide new insights into the role of hAHR-Y322 in AHR signalling. Examination of mRNA levels revealed continued expression of CYP1A1 and CYP1B1 mRNA to 24h in a pattern consistent with the relative potencies of the HAH ligands in T-47D and HuH7 cells. HES1 expression did not correlate well to compound potencies, while TiPARP expression only correlated well with relative potencies in T-47D and not HuH7 cells. Unlike CYP1A1 and CYP1B1 expression which continually increased to 24h, HES1 and TiPARP expression were shown to be oscillatory, indicating that there is differential regulation of AHR-responsive genes. A good correlation between TEF values for HAHs and AHR recruitment to the enhancer regions of CYP1A1, CYP1B1 and HES1 was observed at the 10nM dose following 45min treatment in both T-47D and HuH7 cells. This pattern was mirrored in the mRNA expression data for CYP1A1 and CYP1B1 but not HES1. I concluded that potency of ligands is not necessarily reflected in the magnitude of target gene induction and AHR recruitment. Coactivator recruitment did not correlate to the relative potencies of the ligands; however, functional redundancy in coactivator function exists between p300, NCoA1 and NCoA3 and gene-, cell line- and ligand-selective recruitment was observed. Little difference was observed in either expression of or AHR recruitment to target genes by the low-affinity ligand 2,3,7,8-TCDF and Omp of both wild-type and mutant AHR, lending evidence to the theory that Omp is actually a low-affinity ligand for AHR rather than an atypical inducer. Previous studies had demonstrated HAHs induce AHR degradation, the effects of Omp treatment on degradation remained previously untested (Ma and Baldwin, 2000; Pollenz, 2007). Here I show that Omp induces AHR degradation in nuclear extracts, which is prevented by MG-132 pre-treatment indicating that Omp treatment does lead to attenuation of AHR signalling by 26S proteasomal degradation. Finally, mutation of hAHR-Y322 of the LBD to Y322F or Y322A was shown to impair and abolish AHR signalling, respectively, by both high and low affinity ligands. As such, Y322 is required for maximal ligand affinity and AHR activation. While the results of this study provide several interesting insights into gene-, cell line-, and ligand-dependent differences and the role of Y322, there are several inherent experimental limitations in this study.

6.3 Limitations

6.3.1 Time-points Chosen for mRNA and ChIP Experiments

Time-points chosen for mRNA and ChIP assay experiments represent an area of potential limitations for my study. I chose to examine 1.5, 6 and 24h time-points of mRNA induction for CYP1A1, CYP1B1, HES1 and TiPARP based on the time-points of chosen arbitrarily for previous experiments by our laboratory in T-47D cells. Due to the oscillatory nature of expression that was observed for both HES1 and TiPARP, additional time-points should have been examined to more clearly observe this effect. Perhaps a greater number of time-points should also have been examined, since previous studies suggest a 2h oscillation period for the expression of HES1 (Hirata et al., 2002). In addition, the time-points chosen were relatively arbitrary, so observed results may not be representative of data at ideal time-points for comparison. Previous studies by our laboratory have demonstrated that maximal AHR recruitment was observed following 30min treatment. I examined the effects of 45min treatment in attempt to better correlate mRNA expression with AHR binding. However, recruitment may have been slightly lower than maximal at this time-point. As such, results may have been slightly different if I had examined the 30 min time-point instead. This could particularly affect genes with low recruitment, such as HES1, where recruitment is already analyzed at the upper decipherable threshold of qPCR.

6.3.2 ChIP Assay

While ChIP assay is an excellent technique for studying protein/DNA interactions, it is not without limitations. Use of a specific antibody for immuno-precipitation is a critical step in performing a ChIP assay. While “ChIP-Grade” antibodies are required for specific immuno- precipitations, these very specific antibodies are not necessarily readily available (Deblois and Giguere, 2008). An additional limitation of this technique is inherent with the use of formaldehyde for cross-linking, since formaldehyde causes cross-linking not only between protein and DNA but also of induces protein-protein cross-links (Hanlon and Lieb, 2004; Haring et al., 2007). As such, formation of large protein complexes cross-linked to DNA may obstruct binding of the antibody to the epitope on the protein of interest, which is required for efficient

immuno-precipitation. In addition, the formation of these complexes can also mask the true recruitment patterns of a protein to a regulatory region of a gene if it interacts indirectly with the DNA at this site. It has been previously demonstrated that proteins directly bound to DNA in these regions exhibit greater recruitment in ChIP assays then their indirectly bound counter-parts, which may not accurately reflect the level of recruitment (Aparicio et al., 2005).

6.3.3 Normalization of qPCR Data

The method by which I normalized data may also introduce limitations into my study. For ChIP assays, I analyzed the results of qPCR by normalizing data relative to the % input chromatin. Input chromatin is taken from early on in the ChIP procedure and is pre-immuno-cleared prior to aliquoting the sample for immuno-precipitation. Normalization of ChIP data serves to correct for technical variation by accounting for differences in input chromatin amount, efficiency of immuno-precipitation, and DNA recovery but should not mask biologically significant differences (Haring et al., 2007). A major disadvantage to using the % input normalization method is the effect that differences in the handling methods of samples and % totals may have on the final results (Haring et al., 2007). Despite these limitations, expressing results relative to the percent total input chromatin remains the best choice for normalization of qPCR data for these experiments.

6.3.4 Implications of Y322 Mutations on LBD Function

The mouse equivalent of Y322 was identified previously as a well conserved amino acid, believed to be a putative residue for ligand binding based on nuclear magnetic resonance structures of the PAS B domain of human hypoxia-inducible factor 2α (HIF2α) which has a high degree of identity to AHR (Pandini et al., 2009). Using this projected structure of the AHR LBD, Pandini et al. (2009) elucidated a “TCDD- binding fingerprint” of residues believed to be required for 2,3,7,8-TCDD-mediated AHR activation. While my findings indicate that Y322 is required for maximal activation, it is still unknown whether they directly interact with ligands, or play a crucial role in proper structure and protein folding within the pocket. The effect of the equivalent mutations in mice was elucidated through molecular docking simulations with the

structure of HIF2α as a model; however, solving the 3D-structure of hAHR would be required to confirm the role of these residues in the binding pocket. Further experiments are required to examine the role of other human equivalent residues identified by Pandini et al. (2009) in the “TCDD-binding fingerprint,” so that the role they play in mediating target gene mRNA expression and AHR recruitment can be determined in a cellular system, rather than by reporter gene assays.

6.4 Further Aims

My results suggest a strong correlation in data between the low affinity AHR ligand and Omp, and are in support of Omp acting as a low affinity ligand, capable of binding to AHR (Denison and Nagy, 2003). To further test the mechanism of Omp-induced AHR activation, [3H]-TCDD binding assays would have to be performed, which, contrary to traditional binding assay protocols, utilize much lower concentrations of 2,3,7,8-TCDD and higher concentrations of the competititor (Denison and Nagy, 2003). To increase the ability of such a low potency inducer to competitively bind to AHR, a low affinity radiolabelled competitor, such as [3H]-TCDF could be used. In addition to binding analysis, a re-examination of the effects of tyrosine kinase inhibition on Omp-mediated AHR signalling would be required. Phosphorylation status has been implicated in such activities as nuclear translocation of the AHR complex and DNA binding activity. Likewise, tyrosine kinases play a significant role in the modulation of a variety of cellular activities. As such, treatment with tyrosine kinase inhibitors can have many off-target and wide-ranging effects. Previous investigations of the role of c-src tyrosine kinase inhibitors in Omp-mediated activation have been examined in a reporter gene system. Experiments involving tyrosine kinases should be repeated in a cellular system and ligand binding, AHR recruitment, and target gene expression should be assessed to determine what level of AHR regulation tyrosine kinases are acting upon.

References

Abbott, B.D., Birnbaum, L.S., 1991. TCDD exposure of human embryonic palatal shelves in organ culture alters the differentiation of medial epithelial cells. Teratology 43, 119-132. Abbott, B.D., Held, G.A., Wood, C.R., Buckalew, A.R., Brown, J.G., Schmid, J., 1999a. AhR, ARNT, and CYP1A1 mRNA quantitation in cultured human embryonic palates exposed to TCDD and comparison with mouse palate in vivo and in culture. Toxicol Sci 47, 62-75. Abbott, B.D., Probst, M.R., 1995. Developmental expression of two members of a new class of transcription factors: II. Expression of aryl hydrocarbon receptor nuclear translocator in the C57BL/6N mouse embryo. Dev Dyn 204, 144-155. Abbott, B.D., Schmid, J.E., Pitt, J.A., Buckalew, A.R., Wood, C.R., Held, G.A., Diliberto, J.J., 1999b. Adverse reproductive outcomes in the transgenic Ah receptor-deficient mouse. Toxicology and applied pharmacology 155, 62-70. Akahoshi, E., Yoshimura, S., Ishihara-Sugano, M., 2006. Over-expression of AhR (aryl hydrocarbon receptor) induces neural differentiation of Neuro2a cells: neurotoxicology study. Environ Health 5, 24. Androutsopoulos, V.P., Tsatsakis, A.M., Spandidos, D.A., 2009. Cytochrome P450 CYP1A1: wider roles in cancer progression and prevention. BMC Cancer 9, 187. Anwar-Mohamed, A., Elbekai, R.H., El-Kadi, A.O., 2008. MG-132 inhibits the TCDD-mediated induction of Cyp1a1 at the catalytic activity but not the mRNA or protein levels in Hepa 1c1c7 cells. Toxicology letters 182, 121-126. Aparicio, O., Geisberg, J.V., Sekinger, E., Yang, A., Moqtaderi, Z., Struhl, K., 2005. Chromatin immunoprecipitation for determining the association of proteins with specific genomic sequences in vivo. Current protocols in molecular biology / edited by Frederick M Ausubel [et al Chapter 21, Unit 21 23. Baba, T., Mimura, J., Gradin, K., Kuroiwa, A., Watanabe, T., Matsuda, Y., Inazawa, J., Sogawa, K., Fujii-Kuriyama, Y., 2001. Structure and expression of the Ah receptor repressor gene. The Journal of biological chemistry 276, 33101-33110. Baba, T., Mimura, J., Nakamura, N., Harada, N., Yamamoto, M., Morohashi, K., Fujii- Kuriyama, Y., 2005. Intrinsic function of the aryl hydrocarbon (dioxin) receptor as a key factor in female reproduction. Molecular and cellular biology 25, 10040-10051. Backlund, M., Ingelman-Sundberg, M., 2004. Different structural requirements of the ligand binding domain of the aryl hydrocarbon receptor for high- and low-affinity ligand binding and receptor activation. Molecular pharmacology 65, 416-425. Backlund, M., Ingelman-Sundberg, M., 2005. Regulation of aryl hydrocarbon receptor signal transduction by protein tyrosine kinases. Cell Signal 17, 39-48. Backlund, M., Johansson, I., Mkrtchian, S., Ingelman-Sundberg, M., 1997. Signal transduction- mediated activation of the aryl hydrocarbon receptor in rat hepatoma H4IIE cells. The Journal of biological chemistry 272, 31755-31763.

Backlund, M., Weidolf, L., Ingelman-Sundberg, M., 1999. Structural and mechanistic aspects of transcriptional induction of cytochrome P450 1A1 by benzimidazole derivatives in rat hepatoma H4IIE cells. European journal of biochemistry / FEBS 261, 66-71. Barnett, K.R., Tomic, D., Gupta, R.K., Babus, J.K., Roby, K.F., Terranova, P.F., Flaws, J.A., 2007. The aryl hydrocarbon receptor is required for normal gonadotropin responsiveness in the mouse ovary. Toxicology and applied pharmacology 223, 66-72. Becher, G., Skaare, J.U., Polder, A., Sletten, B., Rossland, O.J., Hansen, H.K., Ptashekas, J., 1995. PCDDs, PCDFs, and PCBs in human milk from different parts of Norway and Lithuania. Journal of toxicology and environmental health 46, 133-148. Beischlag, T.V., Luis Morales, J., Hollingshead, B.D., Perdew, G.H., 2008. The aryl hydrocarbon receptor complex and the control of gene expression. Crit Rev Eukaryot Gene Expr 18, 207-250. Beischlag, T.V., Wang, S., Rose, D.W., Torchia, J., Reisz-Porszasz, S., Muhammad, K., Nelson, W.E., Probst, M.R., Rosenfeld, M.G., Hankinson, O., 2002. Recruitment of the NCoA/SRC- 1/p160 family of transcriptional coactivators by the aryl hydrocarbon receptor/aryl hydrocarbon receptor nuclear translocator complex. Molecular and cellular biology 22, 4319-4333. Bell, D.R., Poland, A., 2000. Binding of aryl hydrocarbon receptor (AhR) to AhR-interacting protein. The role of hsp90. The Journal of biological chemistry 275, 36407-36414. Birnbaum, L.S., McDonald, M.M., Blair, P.C., Clark, A.M., Harris, M.W., 1990. Differential toxicity of 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in C57BL/6J mice congenic at the Ah Locus. Fundam Appl Toxicol 15, 186-200. Bjeldanes, L.F., Kim, J.Y., Grose, K.R., Bartholomew, J.C., Bradfield, C.A., 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. Proceedings of the National Academy of Sciences of the United States of America 88, 9543-9547. Blankenship, A., Matsumura, F., 1997. 2,3,7,8-Tetrachlorodibenzo-p-dioxin-induced activation of a protein tyrosine kinase, pp60src, in murine hepatic cytosol using a cell-free system. Molecular pharmacology 52, 667-675. Bock, K.W., Kohle, C., 2006. Ah receptor: dioxin-mediated toxic responses as hints to deregulated physiologic functions. Biochemical pharmacology 72, 393-404. Boitano, A.E., Wang, J., Romeo, R., Bouchez, L.C., Parker, A.E., Sutton, S.E., Walker, J.R., Flaveny, C.A., Perdew, G.H., Denison, M.S., Schultz, P.G., Cooke, M.P., 2010. Aryl hydrocarbon receptor antagonists promote the expansion of human hematopoietic stem cells. Science (New York, NY 329, 1345-1348. Bradfield, C.A., Poland, A., 1988. A competitive binding assay for 2,3,7,8-tetrachlorodibenzo-p- dioxin and related ligands of the Ah receptor. Molecular pharmacology 34, 682-688. Bradshaw, T.D., Bell, D.R., 2009. Relevance of the aryl hydrocarbon receptor (AhR) for clinical toxicology. Clinical toxicology (Philadelphia, Pa 47, 632-642. Bunger, M.K., Glover, E., Moran, S.M., Walisser, J.A., Lahvis, G.P., Hsu, E.L., Bradfield, C.A., 2008. Abnormal liver development and resistance to 2,3,7,8-tetrachlorodibenzo-p-dioxin toxicity in mice carrying a mutation in the DNA-binding domain of the aryl hydrocarbon receptor. Toxicol Sci 106, 83-92.

Burbach, K.M., Poland, A., Bradfield, C.A., 1992. Cloning of the Ah-receptor cDNA reveals a distinctive ligand-activated transcription factor. Proceedings of the National Academy of Sciences of the United States of America 89, 8185-8189. Carlson, D.B., Perdew, G.H., 2002. A dynamic role for the Ah receptor in cell signalling? Insights from a diverse group of Ah receptor interacting proteins. Journal of biochemical and molecular toxicology 16, 317-325. Carver, L.A., LaPres, J.J., Jain, S., Dunham, E.E., Bradfield, C.A., 1998. Characterization of the Ah receptor-associated protein, ARA9. The Journal of biological chemistry 273, 33580-33587. Castella, P., Sawai, S., Nakao, K., Wagner, J.A., Caudy, M., 2000. HES-1 repression of differentiation and proliferation in PC12 cells: role for the helix 3-helix 4 domain in transcription repression. Molecular and cellular biology 20, 6170-6183. Celius, T., Matthews, J., 2010. Functional analysis of six human aryl hydrocarbon receptor variants in human breast cancer and mouse hepatoma cell lines. Toxicology. Chadli, A., Bouhouche, I., Sullivan, W., Stensgard, B., McMahon, N., Catelli, M.G., Toft, D.O., 2000. Dimerization and N-terminal domain proximity underlie the function of the molecular chaperone heat shock protein 90. Proceedings of the National Academy of Sciences of the United States of America 97, 12524-12529. Chan, W.K., Yao, G., Gu, Y.Z., Bradfield, C.A., 1999. Cross-talk between the aryl hydrocarbon receptor and hypoxia inducible factor signalling pathways. Demonstration of competition and compensation. The Journal of biological chemistry 274, 12115-12123. Chapman-Smith, A., Lutwyche, J.K., Whitelaw, M.L., 2004. Contribution of the Per/Arnt/Sim (PAS) domains to DNA binding by the basic helix-loop-helix PAS transcriptional regulators. The Journal of biological chemistry 279, 5353-5362. Chen, H.S., Perdew, G.H., 1994. Subunit composition of the heteromeric cytosolic aryl hydrocarbon receptor complex. The Journal of biological chemistry 269, 27554-27558. Chen, Y.H., Beischlag, T.V., Kim, J.H., Perdew, G.H., Stallcup, M.R., 2006. Role of GAC63 in transcriptional activation mediated by the aryl hydrocarbon receptor. The Journal of biological chemistry 281, 12242-12247. Chen, Y.H., Tukey, R.H., 1996. Protein kinase C modulates regulation of the CYP1A1 gene by the aryl hydrocarbon receptor. The Journal of biological chemistry 271, 26261-26266. Ciolino, H.P., Dankwah, M., Yeh, G.C., 2002. Resistance of MCF-7 cells to dimethylbenz(a)anthracene-induced apoptosis is due to reduced CYP1A1 expression. Int J Oncol 21, 385-391. Ciolino, H.P., Daschner, P.J., Yeh, G.C., 1998. Resveratrol inhibits transcription of CYP1A1 in vitro by preventing activation of the aryl hydrocarbon receptor. Cancer research 58, 5707-5712. Collins, L.L., Williamson, M.A., Thompson, B.D., Dever, D.P., Gasiewicz, T.A., Opanashuk, L.A., 2008. 2,3,7,8-Tetracholorodibenzo-p-dioxin exposure disrupts granule neuron precursor maturation in the developing mouse cerebellum. Toxicol Sci 103, 125-136. Conney, A.H., 1982. Induction of microsomal enzymes by foreign chemicals and carcinogenesis by polycyclic aromatic hydrocarbons: G. H. A. Clowes Memorial Lecture. Cancer research 42, 4875-4917.

Conney, A.H., Miller, E.C., Miller, J.A., 1956. The metabolism of methylated aminoazo dyes. V. Evidence for induction of enzyme synthesis in the rat by 3-methylcholanthrene. Cancer research 16, 450-459. Coumailleau, P., Poellinger, L., Gustafsson, J.A., Whitelaw, M.L., 1995. Definition of a minimal domain of the dioxin receptor that is associated with Hsp90 and maintains wild type ligand binding affinity and specificity. The Journal of biological chemistry 270, 25291-25300. Cox, M.B., Miller, C.A., 3rd, 2002. The p23 co-chaperone facilitates dioxin receptor signalling in a yeast model system. Toxicology letters 129, 13-21. Daujat, M., Charrasse, S., Fabre, I., Lesca, P., Jounaidi, Y., Larroque, C., Poellinger, L., Maurel, P., 1996. Induction of CYP1A1 gene by benzimidazole derivatives during Caco-2 cell differentiation. Evidence for an aryl-hydrocarbon receptor-mediated mechanism. European journal of biochemistry / FEBS 237, 642-652. Daujat, M., Peryt, B., Lesca, P., Fourtanier, G., Domergue, J., Maurel, P., 1992. Omeprazole, an inducer of human CYP1A1 and 1A2, is not a ligand for the Ah receptor. Biochemical and biophysical research communications 188, 820-825. Davarinos, N.A., Pollenz, R.S., 1999. Aryl hydrocarbon receptor imported into the nucleus following ligand binding is rapidly degraded via the cytosplasmic proteasome following nuclear export. The Journal of biological chemistry 274, 28708-28715. De Abrew, K.N., Phadnis, A.S., Crawford, R.B., Kaminski, N.E., Thomas, R.S., 2011. Regulation of Bach2 by the aryl hydrocarbon receptor as a mechanism for suppression of B-cell differentiation by 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicology and applied pharmacology. Denis, M., Cuthill, S., Wikstrom, A.C., Poellinger, L., Gustafsson, J.A., 1988. Association of the dioxin receptor with the Mr 90,000 heat shock protein: a structural kinship with the glucocorticoid receptor. Biochemical and biophysical research communications 155, 801-807. Denison, M.S., Nagy, S.R., 2003. Activation of the aryl hydrocarbon receptor by structurally diverse exogenous and endogenous chemicals. Annual review of pharmacology and toxicology 43, 309-334. Denison, M.S., Phelan, D., Winter, G.M., Ziccardi, M.H., 1998. Carbaryl, a carbamate insecticide, is a ligand for the hepatic Ah (dioxin) receptor. Toxicology and applied pharmacology 152, 406-414. DeVito, M.J., Ma, X., Babish, J.G., Menache, M., Birnbaum, L.S., 1994. Dose-response relationships in mice following subchronic exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin: CYP1A1, CYP1A2, estrogen receptor, and protein tyrosine phosphorylation. Toxicology and applied pharmacology 124, 82-90. Diliberto, J.J., DeVito, M.J., Ross, D.G., Birnbaum, L.S., 2001. Subchronic Exposure of [3H]- 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in female B6C3F1 mice: relationship of steady- state levels to disposition and metabolism. Toxicol Sci 61, 241-255. Dolwick, K.M., Swanson, H.I., Bradfield, C.A., 1993. In vitro analysis of Ah receptor domains involved in ligand-activated DNA recognition. Proceedings of the National Academy of Sciences of the United States of America 90, 8566-8570.

Enan, E., Matsumura, F., 1996. Identification of c-Src as the integral component of the cytosolic Ah receptor complex, transducing the signal of 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) through the protein phosphorylation pathway. Biochemical pharmacology 52, 1599-1612. Felsenfeld, G., Groudine, M., 2003. Controlling the double helix. Nature 421, 448-453. Felts, S.J., Toft, D.O., 2003. p23, a simple protein with complex activities. Cell stress & chaperones 8, 108-113. Fernandez-Salguero, P., Pineau, T., Hilbert, D.M., McPhail, T., Lee, S.S., Kimura, S., Nebert, D.W., Rudikoff, S., Ward, J.M., Gonzalez, F.J., 1995. Immune system impairment and hepatic fibrosis in mice lacking the dioxin-binding Ah receptor. Science (New York, NY 268, 722-726. Fernandez-Salguero, P.M., Hilbert, D.M., Rudikoff, S., Ward, J.M., Gonzalez, F.J., 1996. Aryl- hydrocarbon receptor-deficient mice are resistant to 2,3,7,8-tetrachlorodibenzo-p-dioxin-induced toxicity. Toxicology and applied pharmacology 140, 173-179. Fingerhut, M.A., Halperin, W.E., Marlow, D.A., Piacitelli, L.A., Honchar, P.A., Sweeney, M.H., Greife, A.L., Dill, P.A., Steenland, K., Suruda, A.J., 1991. Cancer mortality in workers exposed to 2,3,7,8-tetrachlorodibenzo-p-dioxin. The New England journal of medicine 324, 212-218. Fisher, J.W., Campbell, J., Muralidhara, S., Bruckner, J.V., Ferguson, D., Mumtaz, M., Harmon, B., Hedge, J.M., Crofton, K.M., Kim, H., Almekinder, T.L., 2006. Effect of PCB 126 on hepatic metabolism of thyroxine and perturbations in the hypothalamic-pituitary-thyroid axis in the rat. Toxicol Sci 90, 87-95. Frasor, J., Stossi, F., Danes, J.M., Komm, B., Lyttle, C.R., Katzenellenbogen, B.S., 2004. Selective estrogen receptor modulators: discrimination of agonistic versus antagonistic activities by gene expression profiling in breast cancer cells. Cancer research 64, 1522-1533. Fukunaga, B.N., Probst, M.R., Reisz-Porszasz, S., Hankinson, O., 1995. Identification of functional domains of the aryl hydrocarbon receptor. The Journal of biological chemistry 270, 29270-29278. Gao, L., Dong, L., Whitlock, J.P., Jr., 1998. A novel response to dioxin. Induction of ecto- ATPase gene expression. The Journal of biological chemistry 273, 15358-15365. Ge, N.L., Elferink, C.J., 1998. A direct interaction between the aryl hydrocarbon receptor and retinoblastoma protein. Linking dioxin signalling to the cell cycle. The Journal of biological chemistry 273, 22708-22713. Gerbal-Chaloin, S., Pichard-Garcia, L., Fabre, J.M., Sa-Cunha, A., Poellinger, L., Maurel, P., Daujat-Chavanieu, M., 2006. Role of CYP3A4 in the regulation of the aryl hydrocarbon receptor by omeprazole sulphide. Cell Signal 18, 740-750. Gill, G., Pascal, E., Tseng, Z.H., Tjian, R., 1994. A glutamine-rich hydrophobic patch in transcription factor Sp1 contacts the dTAFII110 component of the Drosophila TFIID complex and mediates transcriptional activation. Proceedings of the National Academy of Sciences of the United States of America 91, 192-196. Goll, M.G., Bestor, T.H., 2002. Histone modification and replacement in chromatin activation. Genes & development 16, 1739-1742.

Gradelet, S., Leclerc, J., Siess, M.H., Astorg, P.O., 1996. beta-Apo-8'-carotenal, but not beta- carotene, is a strong inducer of liver cytochromes P4501A1 and 1A2 in rat. Xenobiotica; the fate of foreign compounds in biological systems 26, 909-919. Gradin, K., McGuire, J., Wenger, R.H., Kvietikova, I., fhitelaw, M.L., Toftgard, R., Tora, L., Gassmann, M., Poellinger, L., 1996. Functional interference between hypoxia and dioxin signal transduction pathways: competition for recruitment of the Arnt transcription factor. Molecular and cellular biology 16, 5221-5231. Gradin, K., Toftgard, R., Poellinger, L., Berghard, A., 1999. Repression of dioxin signal transduction in fibroblasts. Identification Of a putative repressor associated with Arnt. The Journal of biological chemistry 274, 13511-13518. Grose, K.R., Bjeldanes, L.F., 1992. Oligomerization of indole-3-carbinol in aqueous acid. Chemical research in toxicology 5, 188-193. Gu, Y.Z., Hogenesch, J.B., Bradfield, C.A., 2000. The PAS superfamily: sensors of environmental and developmental signals. Annual review of pharmacology and toxicology 40, 519-561. Haarmann-Stemmann, T., Bothe, H., Kohli, A., Sydlik, U., Abel, J., Fritsche, E., 2007. Analysis of the transcriptional regulation and molecular function of the aryl hydrocarbon receptor repressor in human cell lines. Drug metabolism and disposition: the biological fate of chemicals 35, 2262-2269. Hahn, M.E., 2002. Aryl hydrocarbon receptors: diversity and evolution. Chemico-biological interactions 141, 131-160. Hankinson, O., 1995. The aryl hydrocarbon receptor complex. Annual review of pharmacology and toxicology 35, 307-340. Hankinson, O., 2005. Role of coactivators in transcriptional activation by the aryl hydrocarbon receptor. Archives of biochemistry and biophysics 433, 379-386. Hanlon, S.E., Lieb, J.D., 2004. Progress and challenges in profiling the dynamics of chromatin and transcription factor binding with DNA microarrays. Current opinion in genetics & development 14, 697-705. Haring, M., Offermann, S., Danker, T., Horst, I., Peterhansel, C., Stam, M., 2007. Chromatin immunoprecipitation: optimization, quantitative analysis and data normalization. Plant methods 3, 11. Harper, P.A., Riddick, D.S., Okey, A.B., 2006. Regulating the regulator: factors that control levels and activity of the aryl hydrocarbon receptor. Biochemical pharmacology 72, 267-279. Helferich, W.G., Denison, M.S., 1991. Ultraviolet photoproducts of tryptophan can act as dioxin agonists. Molecular pharmacology 40, 674-678. Henry, E.C., Gasiewicz, T.A., 1993. Transformation of the aryl hydrocarbon receptor to a DNA- binding form is accompanied by release of the 90 kDa heat-shock protein and increased affinity for 2,3,7,8-tetrachlorodibenzo-p-dioxin. The Biochemical journal 294 ( Pt 1), 95-101. Hestermann, E.V., Brown, M., 2003. Agonist and chemopreventative ligands induce differential transcriptional cofactor recruitment by aryl hydrocarbon receptor. Molecular and cellular biology 23, 7920-7925.

Hirata, H., Yoshiura, S., Ohtsuka, T., Bessho, Y., Harada, T., Yoshikawa, K., Kageyama, R., 2002. Oscillatory expression of the bHLH factor Hes1 regulated by a negative feedback loop. Science (New York, NY 298, 840-843. Hoffer, A., Chang, C.Y., Puga, A., 1996. Dioxin induces transcription of fos and jun genes by Ah receptor-dependent and -independent pathways. Toxicology and applied pharmacology 141, 238-247. Holsapple, M.P., McCay, J.A., Barnes, D.W., 1986. Immunosuppression without liver induction by subchronic exposure to 2,7-dichlorodibenzo-p-dioxin in adult female B6C3F1 mice. Toxicology and applied pharmacology 83, 445-455. Hong, B., Garabrant, D., Hedgeman, E., Demond, A., Gillespie, B., Chen, Q., Chang, C.W., Towey, T., Knutson, K., Franzblau, A., Lepkowski, J., Adriaens, P., 2009. Impact of WHO 2005 revised toxic equivalency factors for dioxins on the TEQs in serum, household dust and soil. Chemosphere 76, 727-733. Hooiveld, M., Heederik, D.J., Kogevinas, M., Boffetta, P., Needham, L.L., Patterson, D.G., Jr., Bueno-de-Mesquita, H.B., 1998. Second follow-up of a Dutch cohort occupationally exposed to phenoxy herbicides, chlorophenols, and contaminants. American journal of epidemiology 147, 891-901. Hord, N.G., Perdew, G.H., 1994. Physicochemical and immunocytochemical analysis of the aryl hydrocarbon receptor nuclear translocator: characterization of two monoclonal antibodies to the aryl hydrocarbon receptor nuclear translocator. Molecular pharmacology 46, 618-626. Hosoya, T., Harada, N., Mimura, J., Motohashi, H., Takahashi, S., Nakajima, O., Morita, M., Kawauchi, S., Yamamoto, M., Fujii-Kuriyama, Y., 2008. Inducibility of cytochrome P450 1A1 and chemical carcinogenesis by benzo[a]pyrene in AhR repressor-deficient mice. Biochemical and biophysical research communications 365, 562-567. Howden, C.W., Meredith, P.A., Forrest, J.A., Reid, J.L., 1984. Oral pharmacokinetics of omeprazole. Eur J Clin Pharmacol 26, 641-643. Huang, P., Rannug, A., Ahlbom, E., Hakansson, H., Ceccatelli, S., 2000. Effect of 2,3,7,8- tetrachlorodibenzo-p-dioxin on the expression of cytochrome P450 1A1, the aryl hydrocarbon receptor, and the aryl hydrocarbon receptor nuclear translocator in rat brain and pituitary. Toxicology and applied pharmacology 169, 159-167. Hushka, L.J., Williams, J.S., Greenlee, W.F., 1998. Characterization of 2,3,7,8- tetrachlorodibenzofuran-dependent suppression and AH receptor pathway gene expression in the developing mouse mammary gland. Toxicology and applied pharmacology 152, 200-210. Ikuta, T., Eguchi, H., Tachibana, T., Yoneda, Y., Kawajiri, K., 1998. Nuclear localization and export signals of the human aryl hydrocarbon receptor. The Journal of biological chemistry 273, 2895-2904. Jellinck, P.H., Forkert, P.G., Riddick, D.S., Okey, A.B., Michnovicz, J.J., Bradlow, H.L., 1993. Ah receptor binding properties of indole carbinols and induction of hepatic estradiol hydroxylation. Biochemical pharmacology 45, 1129-1136. Jeon, M.S., Esser, C., 2000. The murine IL-2 promoter contains distal regulatory elements responsive to the Ah receptor, a member of the evolutionarily conserved bHLH-PAS transcription factor family. J Immunol 165, 6975-6983.

Jones, L.C., Okino, S.T., Gonda, T.J., Whitlock, J.P., Jr., 2002. Myb-binding protein 1a augments AhR-dependent gene expression. The Journal of biological chemistry 277, 22515- 22519. Kadonaga, J.T., 2004. Regulation of RNA polymerase II transcription by sequence-specific DNA binding factors. Cell 116, 247-257. Kainu, T., Gustafsson, J.A., Pelto-Huikko, M., 1995. The dioxin receptor and its nuclear translocator (Arnt) in the rat brain. Neuroreport 6, 2557-2560. Kazlauskas, A., Poellinger, L., Pongratz, I., 1999. Evidence that the co-chaperone p23 regulates ligand responsiveness of the dioxin (Aryl hydrocarbon) receptor. The Journal of biological chemistry 274, 13519-13524. Kazlauskas, A., Poellinger, L., Pongratz, I., 2000. The immunophilin-like protein XAP2 regulates ubiquitination and subcellular localization of the dioxin receptor. The Journal of biological chemistry 275, 41317-41324. Kazlauskas, A., Sundstrom, S., Poellinger, L., Pongratz, I., 2001. The hsp90 chaperone complex regulates intracellular localization of the dioxin receptor. Molecular and cellular biology 21, 2594-2607. Khorasanizadeh, S., 2004. The nucleosome: from genomic organization to genomic regulation. Cell 116, 259-272. Kikuchi, H., Hossain, A., 1999. Signal transduction-mediated CYP1A1 induction by omeprazole in human HepG2 cells. Exp Toxicol Pathol 51, 342-346. Kim, J.H., Stallcup, M.R., 2004. Role of the coiled-coil coactivator (CoCoA) in aryl hydrocarbon receptor-mediated transcription. The Journal of biological chemistry 279, 49842-49848. Kimura, A., Naka, T., Nohara, K., Fujii-Kuriyama, Y., Kishimoto, T., 2008. Aryl hydrocarbon receptor regulates Stat1 activation and participates in the development of Th17 cells. Proceedings of the National Academy of Sciences of the United States of America 105, 9721- 9726. Ko, H.P., Okino, S.T., Ma, Q., Whitlock, J.P., Jr., 1997. Transactivation domains facilitate promoter occupancy for the dioxin-inducible CYP1A1 gene in vivo. Molecular and cellular biology 17, 3497-3507. Kobayashi, A., Numayama-Tsuruta, K., Sogawa, K., Fujii-Kuriyama, Y., 1997. CBP/p300 functions as a possible transcriptional coactivator of Ah receptor nuclear translocator (Arnt). Journal of biochemistry 122, 703-710. Kociba, R.J., Keyes, D.G., Beyer, J.E., Carreon, R.M., Wade, C.E., Dittenber, D.A., Kalnins, R.P., Frauson, L.E., Park, C.N., Barnard, S.D., Hummel, R.A., Humiston, C.G., 1978. Results of a two-year chronic toxicity and oncogenicity study of 2,3,7,8-tetrachlorodibenzo-p-dioxin in rats. Toxicology and applied pharmacology 46, 279-303. Kollara, A., Brown, T.J., 2006. Functional interaction of nuclear receptor coactivator 4 with aryl hydrocarbon receptor. Biochemical and biophysical research communications 346, 526-534. Kolluri, S.K., Weiss, C., Koff, A., Gottlicher, M., 1999. p27(Kip1) induction and inhibition of proliferation by the intracellular Ah receptor in developing thymus and hepatoma cells. Genes & development 13, 1742-1753.

Kouzarides, T., 2007. Chromatin modifications and their function. Cell 128, 693-705. Kraus, W.L., Wong, J., 2002. Nuclear receptor-dependent transcription with chromatin. Is it all about enzymes? European journal of biochemistry / FEBS 269, 2275-2283. Krusekopf, S., Roots, I., Hildebrandt, A.G., Kleeberg, U., 2003. Time-dependent transcriptional induction of CYP1A1, CYP1A2 and CYP1B1 mRNAs by H+/K+ -ATPase inhibitors and other xenobiotics. Xenobiotica; the fate of foreign compounds in biological systems 33, 107-118. Kulkarni, P.S., Branco, L.C., Crespo, J.G., Afonso, C.A., 2008. Capture of dioxins by ionic liquids. Environmental science & technology 42, 2570-2574. Kumar, M.B., Perdew, G.H., 1999. Nuclear receptor coactivator SRC-1 interacts with the Q-rich subdomain of the AhR and modulates its transactivation potential. Gene expression 8, 273-286. Kumar, M.B., Ramadoss, P., Reen, R.K., Vanden Heuvel, J.P., Perdew, G.H., 2001. The Q-rich subdomain of the human Ah receptor transactivation domain is required for dioxin-mediated transcriptional activity. The Journal of biological chemistry 276, 42302-42310. Kumar, M.B., Tarpey, R.W., Perdew, G.H., 1999. Differential recruitment of coactivator RIP140 by Ah and estrogen receptors. Absence of a role for LXXLL motifs. The Journal of biological chemistry 274, 22155-22164. Lahvis, G.P., Bradfield, C.A., 1998. Ahr null alleles: distinctive or different? Biochemical pharmacology 56, 781-787. Larosa, D.F., Orange, J.S., 2008. 1. Lymphocytes. The Journal of allergy and clinical immunology 121, S364-369; quiz S412. Latchney, S.E., Lioy, D.T., Henry, E.C., Gasiewicz, T.A., Strathmann, F.G., Mayer-Proschel, M., Opanashuk, L.A., 2011. Neural precursor cell proliferation is disrupted through activation of the aryl hydrocarbon receptor by 2,3,7,8-tetrachlorodibenzo-p-dioxin. Stem cells and development 20, 313-326. Lees, M.J., Peet, D.J., Whitelaw, M.L., 2003. Defining the role for XAP2 in stabilization of the dioxin receptor. The Journal of biological chemistry 278, 35878-35888. Lees, M.J., Whitelaw, M.L., 1999. Multiple roles of ligand in transforming the dioxin receptor to an active basic helix-loop-helix/PAS transcription factor complex with the nuclear protein Arnt. Molecular and cellular biology 19, 5811-5822. Lemaire, G., Delescluse, C., Pralavorio, M., Ledirac, N., Lesca, P., Rahmani, R., 2004. The role of protein tyrosine kinases in CYP1A1 induction by omeprazole and thiabendazole in rat hepatocytes. Life Sci 74, 2265-2278. Lindebro, M.C., Poellinger, L., Whitelaw, M.L., 1995. Protein-protein interaction via PAS domains: role of the PAS domain in positive and negative regulation of the bHLH/PAS dioxin receptor-Arnt transcription factor complex. The EMBO journal 14, 3528-3539. Lund, A.K., Goens, M.B., Kanagy, N.L., Walker, M.K., 2003. Cardiac hypertrophy in aryl hydrocarbon receptor null mice is correlated with elevated angiotensin II, endothelin-1, and mean arterial blood pressure. Toxicology and applied pharmacology 193, 177-187. Lund, A.K., Goens, M.B., Nunez, B.A., Walker, M.K., 2006. Characterizing the role of endothelin-1 in the progression of cardiac hypertrophy in aryl hydrocarbon receptor (AhR) null mice. Toxicology and applied pharmacology 212, 127-135.

Lundy, S.K., Berlin, A.A., Lukacs, N.W., 2003. Interleukin-12-independent down-modulation of cockroach antigen-induced asthma in mice by intranasal exposure to bacterial lipopolysaccharide. The American journal of pathology 163, 1961-1968. Ma, H., Baumann, C.T., Li, H., Strahl, B.D., Rice, R., Jelinek, M.A., Aswad, D.W., Allis, C.D., Hager, G.L., Stallcup, M.R., 2001. Hormone-dependent, CARM1-directed, arginine-specific methylation of histone H3 on a steroid-regulated promoter. Curr Biol 11, 1981-1985. Ma, Q., 2002. Induction and superinduction of 2,3,7,8-tetrachlorodibenzo-rho-dioxin-inducible poly(ADP-ribose) polymerase: role of the aryl hydrocarbon receptor/aryl hydrocarbon receptor nuclear translocator transcription activation domains and a labile transcription repressor. Archives of biochemistry and biophysics 404, 309-316. Ma, Q., Baldwin, K.T., 2000. 2,3,7,8-tetrachlorodibenzo-p-dioxin-induced degradation of aryl hydrocarbon receptor (AhR) by the ubiquitin-proteasome pathway. Role of the transcription activaton and DNA binding of AhR. The Journal of biological chemistry 275, 8432-8438. Ma, Q., Whitlock, J.P., Jr., 1996. The aromatic hydrocarbon receptor modulates the Hepa 1c1c7 cell cycle and differentiated state independently of dioxin. Molecular and cellular biology 16, 2144-2150. Matthews, J., Wihlen, B., Thomsen, J., Gustafsson, J.A., 2005. Aryl hydrocarbon receptor- mediated transcription: ligand-dependent recruitment of estrogen receptor alpha to 2,3,7,8- tetrachlorodibenzo-p-dioxin-responsive promoters. Molecular and cellular biology 25, 5317- 5328. McGuire, J., Okamoto, K., Whitelaw, M.L., Tanaka, H., Poellinger, L., 2001. Definition of a dioxin receptor mutant that is a constitutive activator of transcription: delineation of overlapping repression and ligand binding functions within the PAS domain. The Journal of biological chemistry 276, 41841-41849. McMillan, B.J., Bradfield, C.A., 2007. The aryl hydrocarbon receptor sans xenobiotics: endogenous function in genetic model systems. Molecular pharmacology 72, 487-498. Meyer, B.K., Perdew, G.H., 1999. Characterization of the AhR-hsp90-XAP2 core complex and the role of the immunophilin-related protein XAP2 in AhR stabilization. Biochemistry 38, 8907- 8917. Meyer, B.K., Pray-Grant, M.G., Vanden Heuvel, J.P., Perdew, G.H., 1998. Hepatitis B virus X- associated protein 2 is a subunit of the unliganded aryl hydrocarbon receptor core complex and exhibits transcriptional enhancer activity. Molecular and cellular biology 18, 978-988. Mimura, J., Ema, M., Sogawa, K., Fujii-Kuriyama, Y., 1999. Identification of a novel mechanism of regulation of Ah (dioxin) receptor function. Genes & development 13, 20-25. Moffat, I.D., Boutros, P.C., Chen, H., Okey, A.B., Pohjanvirta, R., 2010. Aryl hydrocarbon receptor (AHR)-regulated transcriptomic changes in rats sensitive or resistant to major dioxin toxicities. BMC genomics 11, 263. Morris, D.L., Karras, J.G., Holsapple, M.P., 1993. Direct effects of 2,3,7,8-tetrachlorodibenzo-p- dioxin (TCDD) on responses to lipopolysaccharide (LPS) by isolated murine B-cells. Immunopharmacology 26, 105-112. Mottet, C., Golshayan, D., 2007. CD4+CD25+Foxp3+ regulatory T cells: from basic research to potential therapeutic use. Swiss medical weekly 137, 625-634.

Muller, P., Kietz, S., Gustafsson, J.A., Strom, A., 2002. The anti-estrogenic effect of all-trans- retinoic acid on the breast cancer cell line MCF-7 is dependent on HES-1 expression. The Journal of biological chemistry 277, 28376-28379. Nagaoka, R., Iwasaki, T., Rokutanda, N., Takeshita, A., Koibuchi, Y., Horiguchi, J., Shimokawa, N., Iino, Y., Morishita, Y., Koibuchi, N., 2006. Tamoxifen activates CYP3A4 and MDR1 genes through steroid and xenobiotic receptor in breast cancer cells. Endocrine 30, 261-268. Nambu, J.R., Lewis, J.O., Wharton, K.A., Jr., Crews, S.T., 1991. The Drosophila single-minded gene encodes a helix-loop-helix protein that acts as a master regulator of CNS midline development. Cell 67, 1157-1167. Nebert, D.W., Dalton, T.P., Okey, A.B., Gonzalez, F.J., 2004. Role of aryl hydrocarbon receptor- mediated induction of the CYP1 enzymes in environmental toxicity and cancer. The Journal of biological chemistry 279, 23847-23850. Nguyen, T.A., Hoivik, D., Lee, J.E., Safe, S., 1999. Interactions of nuclear receptor coactivator/corepressor proteins with the aryl hydrocarbon receptor complex. Archives of biochemistry and biophysics 367, 250-257. Okey, A.B., Bondy, G.P., Mason, M.E., Nebert, D.W., Forster-Gibson, C.J., Muncan, J., Dufresne, M.J., 1980. Temperature-dependent cytosol-to-nucleus translocation of the Ah receptor for 2,3,7,8-tetrachlorodibenzo-p-dioxin in continuous cell culture lines. The Journal of biological chemistry 255, 11415-11422. Okey, A.B., Franc, M.A., Moffat, I.D., Tijet, N., Boutros, P.C., Korkalainen, M., Tuomisto, J., Pohjanvirta, R., 2005. Toxicological implications of polymorphisms in receptors for xenobiotic chemicals: the case of the aryl hydrocarbon receptor. Toxicology and applied pharmacology 207, 43-51. Okino, S.T., Pendurthi, U.R., Tukey, R.H., 1992. Phorbol esters inhibit the dioxin receptor- mediated transcriptional activation of the mouse Cyp1a-1 and Cyp1a-2 genes by 2,3,7,8- tetrachlorodibenzo-p-dioxin. The Journal of biological chemistry 267, 6991-6998. Paine, A.J., 1976. Induction of benzo[a]pyrene Mono-oxygenase in liver cell culture by the photochemical generation of active oxygen species. Evidence for the involvement of singlet oxygen and the formation of a stable inducing intermediate. The Biochemical journal 158, 109- 117. Paine, A.J., Francis, J.E., 1980. The induction of benzo[a]pyrene-3-mono-oxygenase by singlet oxygen in liver cell culture is mediated by oxidation products of histidine. Chemico-biological interactions 30, 343-353. Pandini, A., Soshilov, A.A., Song, Y., Zhao, J., Bonati, L., Denison, M.S., 2009. Detection of the TCDD binding-fingerprint within the Ah receptor ligand binding domain by structurally driven mutagenesis and functional analysis. Biochemistry 48, 5972-5983. Pansoy, A., Ahmed, S., Valen, E., Sandelin, A., Matthews, J., 2010. 3-methylcholanthrene induces differential recruitment of aryl hydrocarbon receptor to human promoters. Toxicol Sci 117, 90-100. Park, S., Henry, E.C., Gasiewicz, T.A., 2000. Regulation of DNA binding activity of the ligand- activated aryl hydrocarbon receptor by tyrosine phosphorylation. Archives of biochemistry and biophysics 381, 302-312.

Perdew, G.H., 1988. Association of the Ah receptor with the 90-kDa heat shock protein. The Journal of biological chemistry 263, 13802-13805. Perdew, G.H., 1992. Chemical cross-linking of the cytosolic and nuclear forms of the Ah receptor in hepatoma cell line 1c1c7. Biochemical and biophysical research communications 182, 55-62. Peters, J.M., Narotsky, M.G., Elizondo, G., Fernandez-Salguero, P.M., Gonzalez, F.J., Abbott, B.D., 1999. Amelioration of TCDD-induced teratogenesis in aryl hydrocarbon receptor (AhR)- null mice. Toxicol Sci 47, 86-92. Petrulis, J.R., Kusnadi, A., Ramadoss, P., Hollingshead, B., Perdew, G.H., 2003. The hsp90 Co- chaperone XAP2 alters importin beta recognition of the bipartite nuclear localization signal of the Ah receptor and represses transcriptional activity. The Journal of biological chemistry 278, 2677-2685. Petrulis, J.R., Perdew, G.H., 2002. The role of chaperone proteins in the aryl hydrocarbon receptor core complex. Chemico-biological interactions 141, 25-40. Phelan, D., Winter, G.M., Rogers, W.J., Lam, J.C., Denison, M.S., 1998a. Activation of the Ah receptor signal transduction pathway by bilirubin and biliverdin. Archives of biochemistry and biophysics 357, 155-163. Phelan, D.M., Brackney, W.R., Denison, M.S., 1998b. The Ah receptor can bind ligand in the absence of receptor-associated heat-shock protein 90. Archives of biochemistry and biophysics 353, 47-54. Pohjanvirta, R., 2009. Transgenic mouse lines expressing rat AH receptor variants--a new animal model for research on AH receptor function and dioxin toxicity mechanisms. Toxicology and applied pharmacology 236, 166-182. Pohjanvirta, R., Wong, J.M., Li, W., Harper, P.A., Tuomisto, J., Okey, A.B., 1998. Point mutation in intron sequence causes altered carboxyl-terminal structure in the aryl hydrocarbon receptor of the most 2,3,7,8-tetrachlorodibenzo-p-dioxin-resistant rat strain. Molecular pharmacology 54, 86-93. Poland, A., Glover, E., 1980. 2,3,7,8,-Tetrachlorodibenzo-p-dioxin: segregation of toxocity with the Ah locus. Molecular pharmacology 17, 86-94. Poland, A., Glover, E., Kende, A.S., 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. The Journal of biological chemistry 251, 4936-4946. Poland, A., Knutson, J.C., 1982. 2,3,7,8-tetrachlorodibenzo-p-dioxin and related halogenated aromatic hydrocarbons: examination of the mechanism of toxicity. Annual review of pharmacology and toxicology 22, 517-554. Poland, A., Palen, D., Glover, E., 1994. Analysis of the four alleles of the murine aryl hydrocarbon receptor. Molecular pharmacology 46, 915-921. Pollenz, R.S., 2002. The mechanism of AH receptor protein down-regulation (degradation) and its impact on AH receptor-mediated gene regulation. Chemico-biological interactions 141, 41- 61.

Pollenz, R.S., 2007. Specific blockage of ligand-induced degradation of the Ah receptor by proteasome but not calpain inhibitors in cell culture lines from different species. Biochemical pharmacology 74, 131-143. Pollenz, R.S., Popat, J., Dougherty, E.J., 2005. Role of the carboxy-terminal transactivation domain and active transcription in the ligand-induced and ligand-independent degradation of the mouse Ahb-1 receptor. Biochemical pharmacology 70, 1623-1633. Pollenz, R.S., Santostefano, M.J., Klett, E., Richardson, V.M., Necela, B., Birnbaum, L.S., 1998. Female Sprague-Dawley rats exposed to a single oral dose of 2,3,7,8-tetrachlorodibenzo-p- dioxin exhibit sustained depletion of aryl hydrocarbon receptor protein in liver, spleen, thymus, and lung. Toxicol Sci 42, 117-128. Pollenz, R.S., Sattler, C.A., Poland, A., 1994. The aryl hydrocarbon receptor and aryl hydrocarbon receptor nuclear translocator protein show distinct subcellular localizations in Hepa 1c1c7 cells by immunofluorescence microscopy. Molecular pharmacology 45, 428-438. Pongratz, I., Stromstedt, P.E., Mason, G.G., Poellinger, L., 1991. Inhibition of the specific DNA binding activity of the dioxin receptor by phosphatase treatment. The Journal of biological chemistry 266, 16813-16817. Porterfield, S.P., 2000. Thyroidal dysfunction and environmental chemicals--potential impact on brain development. Environmental health perspectives 108 Suppl 3, 433-438. Probst, M.R., Reisz-Porszasz, S., Agbunag, R.V., Ong, M.S., Hankinson, O., 1993. Role of the aryl hydrocarbon receptor nuclear translocator protein in aryl hydrocarbon (dioxin) receptor action. Molecular pharmacology 44, 511-518. Prud'homme, G.J., Glinka, Y., Toulina, A., Ace, O., Subramaniam, V., Jothy, S., 2010. Breast cancer stem-like cells are inhibited by a non-toxic aryl hydrocarbon receptor agonist. PloS one 5, e13831. Puga, A., Ma, C., Marlowe, J.L., 2009. The aryl hydrocarbon receptor cross-talks with multiple signal transduction pathways. Biochemical pharmacology 77, 713-722. Quattrochi, L.C., Tukey, R.H., 1993. Nuclear uptake of the Ah (dioxin) receptor in response to omeprazole: transcriptional activation of the human CYP1A1 gene. Molecular pharmacology 43, 504-508. Ramadoss, P., Petrulis, J.R., Hollingshead, B.D., Kusnadi, A., Perdew, G.H., 2004. Divergent roles of hepatitis B virus X-associated protein 2 (XAP2) in human versus mouse Ah receptor complexes. Biochemistry 43, 700-709. Rannug, A., Rannug, U., Rosenkranz, H.S., Winqvist, L., Westerholm, R., Agurell, E., Grafstrom, A.K., 1987. Certain photooxidized derivatives of tryptophan bind with very high affinity to the Ah receptor and are likely to be endogenous signal substances. The Journal of biological chemistry 262, 15422-15427. Reiners, J.J., Jr., Clift, R.E., 1999. Aryl hydrocarbon receptor regulation of ceramide-induced apoptosis in murine hepatoma 1c1c7 cells. A function independent of aryl hydrocarbon receptor nuclear translocator. The Journal of biological chemistry 274, 2502-2510. Revel, A., Raanani, H., Younglai, E., Xu, J., Han, R., Savouret, J.F., Casper, R.F., 2001. Resveratrol, a natural aryl hydrocarbon receptor antagonist, protects sperm from DNA damage and apoptosis caused by benzo(a)pyrene. Reproductive toxicology (Elmsford, NY 15, 479-486.

Ricci, M.S., Toscano, D.G., Mattingly, C.J., Toscano, W.A., Jr., 1999. Estrogen receptor reduces CYP1A1 induction in cultured human endometrial cells. The Journal of biological chemistry 274, 3430-3438. Roberts, B.J., Whitelaw, M.L., 1999. Degradation of the basic helix-loop-helix/Per-ARNT-Sim homology domain dioxin receptor via the ubiquitin/proteasome pathway. The Journal of biological chemistry 274, 36351-36356. Roblin, S., Okey, A.B., Harper, P.A., 2004. AH receptor antagonist inhibits constitutive CYP1A1 and CYP1B1 expression in rat BP8 cells. Biochemical and biophysical research communications 317, 142-148. Rowlands, J.C., McEwan, I.J., Gustafsson, J.A., 1996. Trans-activation by the human aryl hydrocarbon receptor and aryl hydrocarbon receptor nuclear translocator proteins: direct interactions with basal transcription factors. Molecular pharmacology 50, 538-548. 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). Critical reviews in toxicology 21, 51-88. Safe, S., 2001. Molecular biology of the Ah receptor and its role in carcinogenesis. Toxicology letters 120, 1-7. Safe, S., Wang, F., Porter, W., Duan, R., McDougal, A., 1998. Ah receptor agonists as endocrine disruptors: antiestrogenic activity and mechanisms. Toxicology letters 102-103, 343-347. Safe, S.H., 1986. Comparative toxicology and mechanism of action of polychlorinated dibenzo- p-dioxins and dibenzofurans. Annual review of pharmacology and toxicology 26, 371-399. Savouret, J.F., Antenos, M., Quesne, M., Xu, J., Milgrom, E., Casper, R.F., 2001. 7- ketocholesterol is an endogenous modulator for the arylhydrocarbon receptor. The Journal of biological chemistry 276, 3054-3059. Schecter, A., Birnbaum, L., Ryan, J.J., Constable, J.D., 2006. Dioxins: an overview. Environmental research 101, 419-428. Schmidt, J.V., Bradfield, C.A., 1996. Ah receptor signalling pathways. Annual review of cell and developmental biology 12, 55-89. Schmidt, J.V., Su, G.H., Reddy, J.K., Simon, M.C., Bradfield, C.A., 1996. Characterization of a murine Ahr null allele: involvement of the Ah receptor in hepatic growth and development. Proceedings of the National Academy of Sciences of the United States of America 93, 6731- 6736. Schneider, Y., Duranton, B., Gosse, F., Schleiffer, R., Seiler, N., Raul, F., 2001. Resveratrol inhibits intestinal tumorigenesis and modulates host-defense-related gene expression in an animal model of human familial adenomatous polyposis. Nutrition and cancer 39, 102-107. Singh, K.P., Garrett, R.W., Casado, F.L., Gasiewicz, T.A., 2010. Aryl Hydrocarbon Receptor- Null Allele Mice Have Hematopoietic Stem/Progenitor Cells with Abnormal Characteristics and Functions. Stem cells and development. Sivertsson, L., Ek, M., Darnell, M., Edebert, I., Ingelman-Sundberg, M., Neve, E.P., 2010. CYP3A4 catalytic activity is induced in confluent Huh7 hepatoma cells. Drug metabolism and disposition: the biological fate of chemicals 38, 995-1002.

Smith, S., 2001. The world according to PARP. Trends in biochemical sciences 26, 174-179. Snyder, R., Remmer, H., 1979. Classes of hepatic microsomal mixed function oxidase inducers. Pharmacology & therapeutics 7, 203-244. Sogawa, K., Iwabuchi, K., Abe, H., Fujii-Kuriyama, Y., 1995a. Transcriptional activation domains of the Ah receptor and Ah receptor nuclear translocator. Journal of cancer research and clinical oncology 121, 612-620. Sogawa, K., Nakano, R., Kobayashi, A., Kikuchi, Y., Ohe, N., Matsushita, N., Fujii-Kuriyama, Y., 1995b. Possible function of Ah receptor nuclear translocator (Arnt) homodimer in transcriptional regulation. Proceedings of the National Academy of Sciences of the United States of America 92, 1936-1940. Song, Z., Pollenz, R.S., 2002. Ligand-dependent and independent modulation of aryl hydrocarbon receptor localization, degradation, and gene regulation. Molecular pharmacology 62, 806-816. Soshilov, A., Denison, M.S., 2008. Role of the Per/Arnt/Sim domains in ligand-dependent transformation of the aryl hydrocarbon receptor. The Journal of biological chemistry 283, 32995- 33005. Stevens, E.A., Mezrich, J.D., Bradfield, C.A., 2009. The aryl hydrocarbon receptor: a perspective on potential roles in the immune system. Immunology 127, 299-311. Stinchcombe, S., Buchmann, A., Bock, K.W., Schwarz, M., 1995. Inhibition of apoptosis during 2,3,7,8-tetrachlorodibenzo-p-dioxin-mediated tumour promotion in rat liver. Carcinogenesis 16, 1271-1275. Strom, A., Arai, N., Leers, J., Gustafsson, J.A., 2000. The Hairy and Enhancer of Split homologue-1 (HES-1) mediates the proliferative effect of 17beta-estradiol on breast cancer cell lines. Oncogene 19, 5951-5953. Sulentic, C.E., Holsapple, M.P., Kaminski, N.E., 1998. Aryl hydrocarbon receptor-dependent suppression by 2,3,7, 8-tetrachlorodibenzo-p-dioxin of IgM secretion in activated B cells. Molecular pharmacology 53, 623-629. Thomas, R.S., Penn, S.G., Holden, K., Bradfield, C.A., Rank, D.R., 2002. Sequence variation and phylogenetic history of the mouse Ahr gene. Pharmacogenetics 12, 151-163. Thomsen, C., Helboe, L., 2003. Regional pattern of binding and c-Fos induction by (R)- and (S)- citalopram in rat brain. Neuroreport 14, 2411-2414. Thomsen, J.S., Kietz, S., Strom, A., Gustafsson, J.A., 2004. HES-1, a novel target gene for the aryl hydrocarbon receptor. Molecular pharmacology 65, 165-171. Tian, Y., Ke, S., Chen, M., Sheng, T., 2003. Interactions between the aryl hydrocarbon receptor and P-TEFb. Sequential recruitment of transcription factors and differential phosphorylation of C-terminal domain of RNA polymerase II at cyp1a1 promoter. The Journal of biological chemistry 278, 44041-44048. Tijet, N., Boutros, P.C., Moffat, I.D., Okey, A.B., Tuomisto, J., Pohjanvirta, R., 2006. Aryl hydrocarbon receptor regulates distinct dioxin-dependent and dioxin-independent gene batteries. Molecular pharmacology 69, 140-153.

Trewin, A.L., Woller, M.J., Wimpee, B.A., Conley, L.K., Baldridge, M.G., Hutz, R.J., 2007. Short-term hormone release from adult female rat hypothalamic and pituitary explants is not altered by 2,3,7,8-tetrachlorodibenzo-p-dioxin. The Journal of reproduction and development 53, 765-775. Uno, S., Dalton, T.P., Sinclair, P.R., Gorman, N., Wang, B., Smith, A.G., Miller, M.L., Shertzer, H.G., Nebert, D.W., 2004. Cyp1a1(-/-) male mice: protection against high-dose TCDD-induced lethality and wasting syndrome, and resistance to intrahepatocyte lipid accumulation and uroporphyria. Toxicology and applied pharmacology 196, 410-421. Van den Berg, M., Birnbaum, L., Bosveld, A.T., Brunstrom, B., Cook, P., Feeley, M., Giesy, J.P., Hanberg, A., Hasegawa, R., Kennedy, S.W., Kubiak, T., Larsen, J.C., van Leeuwen, F.X., Liem, A.K., Nolt, C., Peterson, R.E., Poellinger, L., Safe, S., Schrenk, D., Tillitt, D., Tysklind, M., Younes, M., Waern, F., Zacharewski, T., 1998. Toxic equivalency factors (TEFs) for PCBs, PCDDs, PCDFs for humans and wildlife. Environmental health perspectives 106, 775-792. van der Molen, G.W., Kooijman, S.A., Slob, W., 1996. A generic toxicokinetic model for persistent lipophilic compounds in humans: an application to TCDD. Fundam Appl Toxicol 31, 83-94. Veldhoen, M., Hirota, K., Westendorf, A.M., Buer, J., Dumoutier, L., Renauld, J.C., Stockinger, B., 2008. The aryl hydrocarbon receptor links TH17-cell-mediated autoimmunity to environmental toxins. Nature 453, 106-109. Walisser, J.A., Bunger, M.K., Glover, E., Bradfield, C.A., 2004. Gestational exposure of Ahr and Arnt hypomorphs to dioxin rescues vascular development. Proceedings of the National Academy of Sciences of the United States of America 101, 16677-16682. Walisser, J.A., Glover, E., Pande, K., Liss, A.L., Bradfield, C.A., 2005. Aryl hydrocarbon receptor-dependent liver development and hepatotoxicity are mediated by different cell types. Proceedings of the National Academy of Sciences of the United States of America 102, 17858- 17863. Waller, C.L., McKinney, J.D., 1995. Three-dimensional quantitative structure-activity relationships of dioxins and dioxin-like compounds: model validation and Ah receptor characterization. Chemical research in toxicology 8, 847-858. Wang, F., Zhang, R., Beischlag, T.V., Muchardt, C., Yaniv, M., Hankinson, O., 2004a. Roles of Brahma and Brahma/SWI2-related gene 1 in hypoxic induction of the erythropoietin gene. The Journal of biological chemistry 279, 46733-46741. Wang, F., Zhang, R., Shi, S., Hankinson, O., 2007. The effect of aromatic hydrocarbon receptor on the phenotype of the Hepa 1c1c7 murine hepatoma cells in the absence of dioxin. Gene Regul Syst Bio 1, 49-56. Wang, S., Ge, K., Roeder, R.G., Hankinson, O., 2004b. Role of mediator in transcriptional activation by the aryl hydrocarbon receptor. The Journal of biological chemistry 279, 13593- 13600. Warner, M., Eskenazi, B., Mocarelli, P., Gerthoux, P.M., Samuels, S., Needham, L., Patterson, D., Brambilla, P., 2002. Serum dioxin concentrations and breast cancer risk in the Seveso Women's Health Study. Environmental health perspectives 110, 625-628.

Watt, K., Jess, T.J., Kelly, S.M., Price, N.C., McEwan, I.J., 2005. Induced alpha-helix structure in the aryl hydrocarbon receptor transactivation domain modulates protein-protein interactions. Biochemistry 44, 734-743. Weintraub, H., Davis, R., Tapscott, S., Thayer, M., Krause, M., Benezra, R., Blackwell, T.K., Turner, D., Rupp, R., Hollenberg, S., et al., 1991. The myoD gene family: nodal point during specification of the muscle cell lineage. Science (New York, NY 251, 761-766. Whitelaw, M.L., McGuire, J., Picard, D., Gustafsson, J.A., Poellinger, L., 1995. Heat shock protein hsp90 regulates dioxin receptor function in vivo. Proceedings of the National Academy of Sciences of the United States of America 92, 4437-4441. Whitlock, J.P., Jr., 1999. Induction of cytochrome P4501A1. Annual review of pharmacology and toxicology 39, 103-125. Williamson, M.A., Gasiewicz, T.A., Opanashuk, L.A., 2005. Aryl hydrocarbon receptor expression and activity in cerebellar granule neuroblasts: implications for development and dioxin neurotoxicity. Toxicol Sci 83, 340-348. Xu, C., Li, C.Y., Kong, A.N., 2005. Induction of phase I, II and III drug metabolism/transport by xenobiotics. Archives of pharmacal research 28, 249-268. Yoshinari, K., Ueda, R., Kusano, K., Yoshimura, T., Nagata, K., Yamazoe, Y., 2008. Omeprazole transactivates human CYP1A1 and CYP1A2 expression through the common regulatory region containing multiple xenobiotic-responsive elements. Biochemical pharmacology 76, 139-145. Yoshiura, S., Ohtsuka, T., Takenaka, Y., Nagahara, H., Yoshikawa, K., Kageyama, R., 2007. Ultradian oscillations of Stat, Smad, and Hes1 expression in response to serum. Proceedings of the National Academy of Sciences of the United States of America 104, 11292-11297. Zaher, H., Fernandez-Salguero, P.M., Letterio, J., Sheikh, M.S., Fornace, A.J., Jr., Roberts, A.B., Gonzalez, F.J., 1998. The involvement of aryl hydrocarbon receptor in the activation of transforming growth factor-beta and apoptosis. Molecular pharmacology 54, 313-321. Zhang, S., Rowlands, C., Safe, S., 2008. Ligand-dependent interactions of the Ah receptor with coactivators in a mammalian two-hybrid assay. Toxicology and applied pharmacology 227, 196- 206.