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2,3,7,8-TETRACHLORODIBENZO-p-DIOXIN-INDUCIBLE POLY(ADP-RIBOSE) POLYMERASE IS A MONO-ADP-RIBOSYLTRANSFERASE AND A LIGAND-INDUCED
REPRESSOR OF AHR TRANSACTIVATION
by
Laura Mariko MacPherson
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy
Department of Pharmacology and Toxicology University of Toronto
© Copyright by Laura Mariko MacPherson (2014)
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2,3,7,8-tetrachlorodibenzo-p-dioxin-inducible poly(ADP-ribose)
polymerase is a mono-ADP-ribosyltransferase and a ligand-
induced repressor of AHR transactivation
Laura Mariko MacPherson
Doctor of Philosophy
Department of Pharmacology and Toxicology University of Toronto
2014
Abstract
2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD)-inducible poly(ADP-ribose) polymerase
(TiPARP/ARTD14) is a member of the ARTD family and is regulated by the aryl hydrocarbon
receptor (AHR); however, little is known about TiPARP function. In this study we examined the
catalytic function of TiPARP and determined its role in AHR transactivation. We observed that
TiPARP exhibited auto-mono-ADP-ribosyltransferase activity and ribosylated core histones.
RNAi-mediated knockdown of TiPARP in T47D breast cancer cells increased TCDD-dependent
cytochrome P450 1A1 (CYP1A1) and CYP1B1 mRNA expression and recruitment of AHR to
both genes. Overexpression of TiPARP reduced AHR-dependent increases in CYP1A1-reporter
gene activity, which was restored by overexpression of AHR, but not ARNT. Deletion and
mutagenesis studies showed that TiPARP-mediated inhibition of AHR required the zinc finger
and catalytic domains. TiPARP and AHR co-localized in the nucleus, directly interacted and
both were recruited to CYP1A1 in response to TCDD. Overexpression of TiPARP enhanced
whereas RNAi-mediated knockdown of TiPARP reduced TCDD-dependent AHR proteolytic
degradation. TCDD-dependent induction of AHR target genes was also enhanced in Tiparp-/-
mouse embryonic fibroblasts compared to wildtype controls. Moreover, livers excised from
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TCDD-treated Tiparp-/- mice displayed significantly greater AHR target gene expression
compared with wildtype or heterozygous mice. Comparison of TiPARP to a known negative
regulator of AHR, AHR repressor (AHRR), revealed that TiPARP and AHRR had some notable
similarities but also differences between their mechanisms of repression. Similar to TiPARP,
AHRR was recruited to AHR target regulatory regions in response to TCDD and its
overexpression repressed reporter gene activity. However unlike TiPARP, knockdown of the
AHRR did not affect AHR transactivation or its proteasomal degradation. Despite some
mechanistic similarities, our data suggest that TiPARP and AHRR independently repress AHR
transactivation. Overall, our findings show that TiPARP is a mono-ADP-ribosyltransferase and a
transcriptional repressor of AHR, revealing a novel negative feedback loop controlling AHR
function.
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Acknowledgments There are many people who deserve recognition for their contributions to this thesis.
I would like to express my sincere gratitude to my supervisor, Dr. Jason Matthews for giving me
the opportunity to be a graduate student in his laboratory once again. It has been a pleasure to
work under his guidance and mentorship throughout the years. I am thankful for all that I have
learned from him and all the opportunities he has provided me during this experience.
I would also like to express my gratitude to my supervisory committee: Dr. Denis Grant, Dr.
Peter McPherson and Dr. Ali Salahpour for sharing their expertise in this work. All of their
suggestions were most helpful and I value their input tremendously.
Next, I would like to mention all of my wonderful Matthews’ lab members: Dr. Shaimaa Ahmed,
Sharanya Rajendra, Alvin Gomez, Debbie Bott and Laura Tamblyn for all their technical support
and insights. An extra special thanks to Shaimaa for always being an excellent resource in the
lab and the best coffee break buddy.
I would like to acknowledge the Canadian Institutes of Health Research, Ontario Graduate
Scholarship Program and the University of Toronto for financial support of this research.
Finally, last but certainly not least, I would like to thank my parents; Douglas and Patricia
MacPherson, my brothers; Gordon and Cameron and my lovely husband Tom for their
unconditional support and encouragment during this entire experience.
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Table of Contents Acknowledgments.......................................................................................................................... iv
Table of Contents............................................................................................................................ v
List of Tables .................................................................................................................................. x
List of Figures ................................................................................................................................ xi
List of Abbreviations ................................................................................................................... xiv
Chapter 1: Introduction ................................................................................................................... 1
1 Statement of Research Problem ................................................................................................. 1
2 ADP-ribosylation Reactions....................................................................................................... 2
2.1 Historical Overview............................................................................................................ 2
2.2 NAD+ Metabolism .............................................................................................................. 3
2.3 Mono-ADP-ribosylation ..................................................................................................... 7
2.3.1 Mono-ADP-ribosyltransferase families .................................................................. 7
2.3.2 ADP-ribosylhydrolases ........................................................................................... 9
2.4 Poly-ADP-ribosylation ..................................................................................................... 10
2.4.1 ADP-ribosyltransferase diphtheria toxin-like (ARTD) family ............................. 10
2.4.2 Poly(ADP-ribose) Catabolism .............................................................................. 22
2.5 Regulation of chromatin structure and transcription by ARTDs ...................................... 24
2.5.1 Modulation of chromatin structure by ARTD1 and poly(ADP-ribose)................ 24
2.5.2 Enhancer-binding of actions of ARTD1 ............................................................... 25
2.5.3 Promoter-binding and transcriptional co-regulation by ARTD1 .......................... 26
2.5.4 Transcriptional regulation by other ARTDs ......................................................... 27
3 Aryl hydrocarbon receptor ....................................................................................................... 28
3.1 Structure............................................................................................................................ 29
3.2 AHR ligands...................................................................................................................... 30
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3.2.1 Exogenous AHR ligands....................................................................................... 31
3.2.2 Naturally occurring AHR ligands ......................................................................... 32
3.3 AHR-mediated toxicity..................................................................................................... 33
3.4 AHR transactivation.......................................................................................................... 34
3.5 Negative regulation of AHR transactivation..................................................................... 36
3.5.1 Proteolytic Degradation of AHR .......................................................................... 37
3.5.2 Aryl hydrocarbon receptor repressor (AHRR) ..................................................... 38
3.5.3 Induction of a labile AHR degradation promoting factor (ADPF) ....................... 40
3.5.4 In vivo models used to study negative regulation of AHR ................................... 41
3.6 AHR target genes.............................................................................................................. 43
3.6.1 TCDD-inducible poly(ADP-ribose) polymerase (TiPARP): a novel AHR target ..................................................................................................................... 44
Chapter 2: Aims of the present study............................................................................................ 48
Chapter 3: Materials and Methods................................................................................................ 49
4 Materials................................................................................................................................... 49
4.1 Chemicals and biological reagents.................................................................................... 49
4.2 Plasticware ........................................................................................................................ 50
4.3 Instruments........................................................................................................................ 50
5 Methods.................................................................................................................................... 51
5.1 Maintenance of HuH7, COS7, TREx-FLP-IN 293, T47D, NCI-N87 and MCF7 cell lines ................................................................................................................................... 51
5.2 Generation of Expression Constructs................................................................................ 52
5.3 Site-directed mutagenesis ................................................................................................. 54
5.4 Transient transfection of HuH7 cells for RNA expression, Western blot and ChIP assays ................................................................................................................................ 56
5.5 Transient transfection of T47D cells for Western blot ..................................................... 56
5.6 Transient transfection of HuH7 cells and reporter gene assays........................................ 57
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5.7 Transient transfection and RNAi ...................................................................................... 57
5.8 mRNA time-course ........................................................................................................... 58
5.9 Tiparp-null mice and generation of mouse embryonic fibroblasts (MEF) ....................... 58
5.10 RNA isolation and cDNA synthesis................................................................................. 59
5.10.1 Mammalian cells ................................................................................................... 59
5.10.2 Mouse liver ........................................................................................................... 60
5.10.3 cDNA synthesis .................................................................................................... 60
5.11 Chromatin Immunoprecipitation (ChIP)........................................................................... 61
5.12 Western blot ...................................................................................................................... 63
5.13 Co-immunoprecipitation ................................................................................................... 64
5.14 In vitro translation and 35S PARP assays ......................................................................... 65
5.15 Indirect immunofluorescence........................................................................................... 65
5.16 GST purification............................................................................................................... 66
5.17 32P-NAD+ auto-ADP-ribosylation and hetero-ADP-ribosylation assays ......................... 67
5.18 Statistical Analysis ........................................................................................................... 67
Chapter 4: Results ......................................................................................................................... 69
6 TiPARP is a mono-ADP-ribosyltransferase............................................................................. 69
6.1 Auto-ADP-ribosylation..................................................................................................... 69
6.2 Hetero-ADP-ribosylation.................................................................................................. 76
7 Identification of TiPARP as a novel negative regulator of AHR............................................. 79
7.1 Temporal induction of TiPARP mRNA by TCDD........................................................... 79
7.2 TiPARP knockdown increased TCDD-induced AHR transactivation ............................. 81
7.3 Overexpression of TiPARP repressed AHR-mediated transactivation ............................ 88
7.4 Tiparp knockout increased TCDD-dependent AHR target transactivation and AHR protein levels ..................................................................................................................... 93
7.5 Tiparp knockout increased AHR-mediated gene expression in vivo................................ 96
viii
8 Understanding the mechanism of TiPARP-mediated repression of AHR-mediated transactivation .......................................................................................................................... 99
8.1 TiPARP selective repression of AHR activity.................................................................. 99
8.2 Ectopic AHR but not ARNT prevented TiPARP-mediated inhibition of AHR transactivation ................................................................................................................. 101
8.3 Identification of a putative AHR repressor domain of TiPARP..................................... 103
8.4 Repression of AHR by TiPARP required zinc finger function....................................... 112
8.5 Repression of AHR transactivation by TiPARP required catalytic domain and function ........................................................................................................................... 115
8.6 TiPARP interaction with AHR ....................................................................................... 120
8.7 Identification of nuclear localization signal of N-terminus of TiPARP. ........................ 125
8.8 TiPARP is a labile factor that negatively regulates AHR transactivation ...................... 128
9 Comparison of TiPARP- and AHRR-mediated negative regulation of AHR transactivation132
9.1 Kinetics of AHRR induction by TCDD.......................................................................... 133
9.2 AHRR knockdown did not affect AHR transactivation or protein levels ...................... 136
9.3 AHRR overexpression repressed AHR-regulated reporter gene activity ....................... 141
9.4 Expression patterns of the AHRR................................................................................... 144
Chapter 5: Discussion ................................................................................................................. 146
10 TiPARP is a mono-ADP-ribosyltransferase........................................................................... 146
11 TiPARP is a novel negative regulator of AHR transactivation.............................................. 148
11.1 Potential co-repression by ADP-ribosylation of AHR................................................... 150
11.2 Potential targeting of histones and other co-regulatory factors...................................... 151
11.3 TiPARP is a candidate AHR degradation promoting factor (ADPF) ............................ 152
11.3.1 TiPARP and the nuclear proteasome .................................................................. 153
11.3.2 Connecting nuclear localization with repression of AHR transactivation.......... 154
11.4 Identification of an AHR repressive domain.................................................................. 155
12 Comparing TiPARP- with AHRR-mediated transrepression ................................................ 156
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12.1 Gene expression and recruitment.................................................................................... 156
12.2 Overexpression of ARNT and co-regulators.................................................................. 157
12.3 Subcellular localization.................................................................................................. 157
12.4 Knockdown and knockout models ................................................................................. 158
12.5 Potential overlap of TiPARP and the AHRR repressive pathways................................ 159
13 TiPARP is an in vivo negative regulator of AHR transactivation.......................................... 160
13.1 Role of TiPARP in TCDD-mediated toxicity................................................................. 161
13.1.1 Purported effects of TiPARP on the NAD+ pool and SIRT activity.................... 161
14 TiPARP-mediated regulation of other transcription factors .................................................. 163
15 Limitations and future considerations.................................................................................... 163
15.1 TiPARP antibody ............................................................................................................ 163
15.2 ARTD catalytic activity assays ...................................................................................... 164
16 Overall significance of research............................................................................................. 165
References................................................................................................................................... 167
Copyright Acknowledgements.................................................................................................... 204
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List of Tables Table 1. Organization and enzymatic activities of mammalian ARTD family. ........................... 12
Table 2. Transcription factors co-regulated by ARTD1. .............................................................. 26
Table 3. Partial list of proteins and co-regulator proteins that interact with AHR/ARNT to
modulate transcription. ................................................................................................................. 35
Table 4. PCR primers used for generation of TiPARP and AHRR inserts................................... 54
Table 5. PCR primers used for site-directed mutagenesis of TiPARP. ........................................ 55
Table 6. List of qPCR primers used in RNA expression assays. .................................................. 60
Table 7. List of qPCR primers used for ChIP assays.................................................................... 62
Table 8. Frequency of co-localization of GFP-TiPARP and AHR in COS7 cells ..................... 120
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List of Figures Figure 1. Schematic representation of the mono-ADP-ribosylation reaction................................. 5
Figure 2. Metabolism of poly(ADP-ribose).................................................................................... 6
Figure 3. Domain architecture of the 18 human ARTD family members. ................................... 13
Figure 4. ARTD1 domain architecture. ........................................................................................ 15
Figure 5. Schematic of the modular domains of three bHLH-PAS family members. .................. 30
Figure 6. Selected exogenous AHR ligands. ................................................................................ 32
Figure 7. General mechanism of AHR transactivation. ................................................................ 36
Figure 8. Three proposed mechanisms of negative regulation of AHR transactivation. .............. 37
Figure 9. Functional domain structure of TCDD-inducible poly(ADP-ribose) polymerase. ....... 46
Figure 10. TiPARP is a mono-ADP-ribosyltransferase with auto-ribosylation activity. ............. 72
Figure 11. Putative auto-ribosylation domain of TiPARP is between residues 275 and 328. ...... 74
Figure 12. TiPARP is a mono-ADP-ribosyltransferase with heteroribosylation activity............. 78
Figure 13. Temporal induction of TiPARP mRNA in T47D and Hepa1c1c7 cell lines treated with
TCDD............................................................................................................................................ 80
Figure 14. TiPARP knockdown in NCI-N87, HuH7 and T47D cells increased TCDD-induced
AHR transactivation...................................................................................................................... 83
Figure 15. TiPARP knockdown increased TCDD-induced AHR transactivation. ....................... 85
Figure 16. TiPARP knockdown increased AHR/ARNT recruitment to CYP1A1 and CYP1B1
regulatory regions and reduced TCDD-induced degradation of AHR. ........................................ 87
Figure 17. TiPARP overexpression reduced AHR-regulated reporter gene activity.................... 90
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Figure 18. Overexpression of TiPARP repressed TCDD-induced AHR transactivation. ............ 92
Figure 19. Tiparp knockout fibroblasts exhibited increased TCDD-induced AHR transactivation.
....................................................................................................................................................... 95
Figure 20. Female Tiparp-/- mice demonstrated increased TCDD-induced AHR transactivation.97
Figure 21. Male Tiparp-/- mice demonstrated increased TCDD-induced AHR transactivation. .. 98
Figure 22. CYP1A1-luc reporter gene activity was preferentially inhibited by TiPARP. ......... 100
Figure 23. Overexpression of AHR but not ARNT prevented TiPARP-mediated inhibition of
AHR transactivation.................................................................................................................... 102
Figure 24. In vitro expression of 35S-labelled N-terminal TiPARP truncations. ........................ 104
Figure 25. Overexpression of N-terminal TiPARP truncations on AHR-regulated reporter gene
activity......................................................................................................................................... 106
Figure 26. Overexpression of N-terminal GFP-tagged TiPARP truncations on AHR-regulated
reporter gene activity. ................................................................................................................. 108
Figure 27. Overexpression of GFP-tagged TiPARP N-terminal truncations. ............................ 109
Figure 28. Effects of TiPARP double alanine point mutants on AHR-regulated reporter gene
activity......................................................................................................................................... 111
Figure 29. Repression of AHR transactivation required zinc finger function. ........................... 114
Figure 30. Repression of AHR transactivation required C-terminal catalytic domain............... 117
Figure 31. Repression of AHR transactivation required catalytic function................................ 119
Figure 32. TiPARP interacts with AHR in the nucleus. ............................................................. 122
Figure 33. TiPARP truncation and point mutant interaction with AHR..................................... 124
Figure 34. Identification of putative nuclear localization signal (NLS) within TiPARP N-
terminus....................................................................................................................................... 127
xiii
Figure 35. TiPARP is a labile negative regulator of AHR. ........................................................ 131
Figure 36. AHRR is induced by TCDD and recruited to AHR-regulated genes. ....................... 135
Figure 37. Effects of AHRR knockdown and TiPARP knockdown on AHR transactivation.... 139
Figure 38. AHRR, AHR and ARNT protein levels following AHRR (A) and TiPARP (B)
knockdown.................................................................................................................................. 140
Figure 39. Overexpression of AHRRΔ8 repressed AHR-regulated reporter gene activity. ....... 143
Figure 40. Expression of N-terminal and C-terminal GFP-tagged AHRR in MCF7, HuH7 and
COS7 cells. ................................................................................................................................. 145
Figure 41. Proposed model of repression of AHR transactivation by TiPARP.......................... 149
Figure 42. Updated schematic of the structure of TiPARP. ....................................................... 155
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List of Abbreviations 3MC 3-methylcholanthrene ADP adenosine diphosphate ADPF AHR degradation promoting factor ARTD ADP-ribosyltransferase diphtheria toxin-like AHR aryl hydrocarbon receptor AHRE aryl hydrocarbon receptor response element AHRR aryl hydrocarbon receptor repressor ARNT aryl hydrocarbon receptor nuclear translocator ARH ADP-ribosylhydrolase B[a]P benzo[a]pyrene BAL B-aggressive lymphoma bHLH-PAS basic Helix-Loop-Helix-Period, Aryl hydrocarbon receptor nuclear
translocator Single minded BRCT BRCA1-carboxy terminus-like Brg-1 brahma-related gene 1 CA-AHR constitutively active aryl hydrocarbon receptor ChIP chromatin immunoprecipitation CYP1A1 cytochrome P450 1A1-human CYP1B1 cytochrome P450 1B1-human DBD DNA-binding domain e-MART ecto-mono-ADP-ribosyltransferase ERα estrogen receptor α ERβ estrogen receptor β FICZ 6-formalindolo-[3,2-b]carbazole G6Pase glucose-6-phosphatase GFP green fluorescent protein GST glutathione S-transferase HAH halogenated aromatic hydrocarbon HDAC histone deacetylase HES1 hairy and enhancer of split 1 Hsp90 90 kDa heat shock protein IC3 indole-3-carbinol ICZ indolo[3,2-b]carbazole LBD ligand binding domain MART mono-ADP-ribosyltransferase MEF mouse embryonic fibroblast NAD+ nicotinamide adenine dinucleotide NASH non-alcoholic steatohepatitis NCoA nuclear co-activator NCoR nuclear receptor co-repressor NES nuclear export signal NF-κB nuclear factor kappa-light-chain-enhancer of activated B-cells NLS nuclear localization signal NQO1 NAD(P)H: quinone oxidoreductase 1 NFE2L2 nuclear factor (erythroid-derived 2)-like 2
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PAR poly(ADP-ribose) PARG poly(ADP-ribose) glycohydrolase PARP poly(ADP-ribose) polymerase PAH polycyclic aromatic hydrocarbon PCB polychlorinated biphenyl PCG1α peroxisome proliferator-activated receptor gamma co-activator 1 alpha PEPCK phosphoenolypyruvate carboxykinase RIP140 140 kDa receptor interacting protein siRNA small inhibitory RNA SHP short heterodimer partner SIRT sirtuin SMRT silencing mediator of retinoid and thyroid hormone receptor Sp1 stimulating protein 1 TAD transcriptional activation domain TCDD 2,3,7,8-tetrachlorodibenzo-p-dioxin TIF2 translation initiation factor 2 TiPARP TCDD-inducible poly(ADP-ribose) polymerase qPCR quantitative real-time PCR XAP2 hepatitis B virus X-associated protein 2 WWE tryptophan-tryptophan-glutamate ZAP zinc finger antiviral protein Zn zinc finger
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Chapter 1: Introduction 1 Statement of Research Problem 2,3,7,8-tetracholorodibenzo-p-dioxin (TCDD)-inducible poly(ADP-ribose) polymerase
(TiPARP) is a member of the diphtheria-like ADP-ribosyltransferase (ARTD) family of
nicotinamide adenine dinucleotide (NAD+)-catabolizing enzymes, formerly known as the
poly(ADP-ribose) polymerase (PARP) family. ARTD family members are best known for their
ability to catalyze ADP-ribosylation reactions, a process that uses NAD+ as a substrate to add
ADP-ribose onto target proteins. ADP-ribosylation has been implicated in numerous cellular
processes including DNA damage responses, inflammation, apoptosis and transcriptional
regulation. The function of ARTD1 (also known as PARP1), the founding ARTD, is well
established; however, the potential function(s) of the majority of other ARTD members are just
starting to be understood.
Currently, little is known about TiPARP/ARTD14 (will be referred to as TiPARP within this
thesis). As a class 2 ARTD, TiPARP is predicted to be a mono-ADP-ribosyltransferase (MART);
however, this has not been experimentally determined. As its name suggests TiPARP gene
expression is induced by TCDD. The ligand-activated transcription factor, the aryl hydrocarbon
receptor (AHR), is activated by TCDD to regulate its target genes. TiPARP expression is
regulated by numerous AHR ligands in both in vitro and in vivo models, demonstrating it to be
an important AHR regulated gene. Despite being an important AHR target gene, the potential
function of TiPARP in AHR-mediated transcriptional regulation has not been investigated.
While the activation of AHR transcription has been extensively studied, its negative regulation
has not received equal attention and is poorly understood. Proposed mechanisms of the negative
regulation of AHR include the targeted proteasomal degradation of activated-AHR and induction
of repressive factors such as the AHR repressor (AHRR) to down-regulate AHR-mediated
transactivation; however, these models are controversial and not fully understood.
In the present study we set out to characterize TiPARP and determine its role in AHR
transactivation.
2
2 ADP-ribosylation Reactions
2.1 Historical Overview The presence of poly(ADP-ribose) was first described fifty years ago by Chambon et al. (1963)
who discovered that the addition of nicotinamide adenine dinucleotide (NAD+) to hen liver
extracts stimulated the synthesis of polyadenylic acid which was later identified as poly(ADP-
ribose) (Chambon, et al., 1963). This discovery launched the research of poly(ADP-ribose)
metabolism. The structure of poly(ADP-ribose) (PAR) was later determined by three
independent laboratories (Doly and Mandel, 1967; Reeder, et al., 1967; Sugimura, et al., 1967).
The enzyme responsible for synthesizing PAR was termed poly(ADP-ribose) polymerase or
PARP. The gene encoding PARP (also referred to as poly-ADP-synthase or poly-ADP-
ribosyltransferase) was isolated in the late 1980s (Alkhatib, et al., 1987; Kurosaki, et al., 1987;
Uchida, et al., 1987). Today, this gene is known as PARP1 and for many years PARP1 was
thought to be the only enzyme with ADP-ribosylation activity in mammalian cells. Fifteen years
later, five different genes encoding PARP enzymes were identified indicating PARP1 belongs to
a family of enzymes (Ame, et al., 1999; Johansson, 1999; Kaminker, et al., 2001; Kickhoefer, et
al., 1999; Smith, et al., 1998). To date, eighteen human PARP genes have been cloned or
described based on exhaustive searches of non-redundant protein databases using the PARP1
catalytic domain. However, the PAR synthesizing activity of only a few of these putative PARPs
has been verified (Aguiar, et al., 2005; Aguiar, et al., 2000; Ame, et al., 2004; Ma, et al., 2001;
Otto, et al., 2005; Yu, et al., 2004).
Not long after the structure of PAR was determined, mono-ADP-ribosylation reactions were
discovered during studies of bacterial toxins such as diphtheria, pertussis and cholera toxins
(Corda and Di Girolamo, 2003; Di Girolamo, et al., 2005; Holbourn, et al., 2006). These
enzymes were found to be mono-ADP-ribosyltransferases (MARTs) (Gill, et al., 1969; Honjo, et
al., 1968). Subsequently, extracellular MARTs or e-MARTs (or ecto-MARTs) were identified in
mammalian cells and their enzymatic activities characterized (Haag and Koch-Nolte, 1998;
Okazaki and Moss, 1996; Okazaki and Moss, 1999; Seman, et al., 2004). Several reports have
predicted distinct families of mono-ADP-ribosylating enzymes with no obvious sequence
similarity to e-MARTs (Corda and Di Girolamo, 2002; Corda and Di Girolamo, 2003; Seman, et
al., 2004). Members of the sirtuin family (SIRTs) of NAD+-dependent histone deacetylases were
3
reported to contain mono-ADP-ribosyltransferase activities and represent a potential family of
intracellular mammalian MARTs (Frye, 1999; Garcia-Salcedo, et al., 2003; Haigis, et al., 2006;
Liszt, et al., 2005; Tanny, et al., 1999). However, more recently twelve PARP family members
have been predicted to be mono-ADP-ribosylating rather than poly-ADP-ribosylating enzymes,
based on subtle differences within their catalytic domains (Hottiger, et al., 2010; Kleine, et al.,
2008). These twelve PARP-like MARTs represent potential candidates of a family of
intracellular MART and only recently are their enzymatic activities being verified. A new
nomenclature for the PARP family has been proposed that reflects their true transferase rather
than polymerase activity, renaming it to ADP-ribosyltransferase diphtheria toxin-like family or
ARTD that takes into account their resemblance to bacterial mono-ADP-ribosylating enzymes
(Hottiger, et al., 2010).
2.2 NAD+ Metabolism In eukaryotes, NAD+ is best known for its essential role as a coenzyme/transmitter molecule in
bioenergetic processes such as cellular metabolism and respiration (Magni, et al., 2004; Ziegler,
2000). The synthesis of adenosine triphosphate (ATP) and balance of redox potential are directly
dependent on cellular NAD+ levels. NAD+ serves as an electron acceptor and its reduced form
NADH as an electron donor in reactions catalyzed by enzymes of the mitochondrial electron
transport chain leading to the generation of ATP through oxidative phosphorylation (Pollak, et
al., 2007; Rich, 2003).
In addition to its vital role in energy metabolism, NAD+ is utilized by ADP-ribosyltransferases to
transfer ADP-ribose to a substrate protein or to itself (Hassa, et al., 2006). In mono-ADP-
ribosylation reactions, NAD+ is hydrolyzed by mono-ADP-ribosyltransferases and the ADP-
ribose moiety is transferred to an acceptor protein (the enzyme itself or a target protein) and the
vitamin nicotinamide is released (Figure 1). Poly-ADP-ribosylation is similar to mono-ADP-
ribosylation but also includes an additional elongation reaction where the ADP-ribose units are
linked by glycosidic ribose (1’-2’) ribose bonds creating long, irregular, branched polymers of
ADP-ribose (Desmarais, et al., 1991; Kawaichi, et al., 1981; Mendoza-Alvarez and Alvarez-
Gonzalez, 2004; Miwa, et al., 1981) (Figure 2). Polymers can reach 200-400 ADP-ribose units
in vitro and in vivo and long polymers are branched with a frequency of approximately one
branch point per linear section of 20-50 ADP-ribose units (Alvarez-Gonzalez and Jacobson,
4
1987; Juarez-Salinas, et al., 1982; Juarez-Salinas, et al., 1983; Kanai, et al., 1982; Miwa, et al.,
1981). Poly-ADP-ribosylation is proposed to be the most important factor affecting the
maintenance of the NAD+ pool in cells (Elliott and Rechsteiner, 1975; Rechsteiner, et al., 1976;
Williams, et al., 1985). The catabolism of NAD+ in mammalian cells occurs primarily through
poly-ADP-ribosylation reactions (Berger, et al., 2004). Activation of poly-ADP-ribosylation
reactions, coinciding with increased poly-ADP-ribose polymer generation depletes intracellular
NAD+ levels to 10-20% of their normal levels within 5-15 min upon initiation of stimulus
(Goodwin, et al., 1978; Skidmore, et al., 1979). Cellular NAD+ depletion consequently results in
ATP depletion, since NAD+ is an essential coenzyme/transmitter for the generation of ATP
through oxidative phosphorylation (Berger, 1985). Because NAD+ connects bioenergetics to
ADP-ribosylation reactions and cellular NAD+ levels have important physiological consequences
in the regulation of multiple cellular processes, the control of NAD+-dependent enzymes must be
tightly regulated.
5
Figure 1. Schematic representation of the mono-ADP-ribosylation reaction.
The chemical structure of the mono-ADP-ribosylation reaction to an acceptor protein is shown.
Mono-ADP-ribosyltransferase (MART) catalyzes the addition of ADP-ribose to an acceptor
protein using nicotinamide adenine dinucleotide (NAD+) as a precursor resulting in the release of
nicotinamide. ADP-ribosylhydrolases (ARH) remove the ADP-ribose moiety from an acceptor
protein.
6
Figure 2. Metabolism of poly(ADP-ribose).
PARPs hydrolyse NAD+ and catalyze the successive addition of ADP-ribose units to acceptor
proteins. Poly-ADP-ribose size and complexity is indicated by the x and y labels that represent
values from 0 to >200. Poly-ADP-ribose-glycohydrolase (PARG) and ADP-ribosylhydrolase 3
(ARH3) can both hydrolyze poly-ADP-ribose at the indicated positions. ADE; adenine, NAM;
nicotinamide, Rib; ribose. Modified from (Hakme, et al., 2008).
7
2.3 Mono-ADP-ribosylation Mono-ADP-ribosylation of proteins is a phylogentically ancient and reversible posttranslational
modification. This modification was originally identified as an important aspect of bacterial
physiopathology, catalyzed by several toxins including diphtheria, pertussis and cholera toxins
(Corda and Di Girolamo, 2003; Di Girolamo, et al., 2005; Holbourn, et al., 2006). So far, six
subclasses of bacterial MARTs have been identified based on amino acid sequence identity
(Hassa, et al., 2006). The amino acid residues of host cell acceptor proteins modified by bacterial
MARTs include: arginine, asparagine, glutamate, aspartate, cysteine and modified histidine
(diphthamide) (Corda and Di Girolamo, 2002; Corda and Di Girolamo, 2003; Okazaki and Moss,
1996; Okazaki and Moss, 1999). Diphtheria toxin can be used as an example of the specificity of
mono-ADP-ribosylation and the important physiological consequences of a single residue
modification by a MART. Diphtheria toxin, is an exotoxin secreted by the bacterium
Corynebacterium diphtheria, which mono-ADP-ribosylates host cell elongation factor 2 (eEF2)
at diphthamide residue 699 blocking the interaction of eEF2 with other factors involved in
translation and effectively preventing host cell protein synthesis (Collier, 2001).
Mono-ADP-ribosylation reactions in multicellular eukaryotes were later discovered. Endogenous
MART activities that modify arginine, glutamate, cysteine, phosphoserine and potentially
aspartate and asparagine on acceptor proteins have been detected in mammalian cells (Corda and
Di Girolamo, 2003; Okazaki and Moss, 1996; Ord and Stocken, 1977; Seman, et al., 2004; Smith
and Stocken, 1975). Intracellular mono-ADP-ribosylation in mammalian cells has been
suggested to play important roles in processes such as: regulation of intracellular signalling
cascades, transcriptional regulation, unfolded protein response, DNA repair, insulin secretion,
immunity and cellular differentiation and proliferation (Corda and Di Girolamo, 2003; Di
Girolamo, et al., 2005; Feijs, et al., 2013; Koch-Nolte, et al., 2008).
2.3.1 Mono-ADP-ribosyltransferase families
2.3.1.1 ecto-MARTs (e-MARTs)
Human and mouse e-MARTs that have been identified are glycosylphosphatidylinositol-
anchored surface proteins or secretory proteins (Glowacki, et al., 2002; Koch-Nolte, et al., 2008;
Okazaki and Moss, 1999). E-MARTs transfer ADP-ribose to target proteins within the
8
extracellular compartment and are involved in cell communication (Koch-Nolte, et al., 2008).
Five e-MARTs (e-MART-1 to 5) have been identified in mammalian cells. Humans only express
four e-MARTs due to a defective e-MART-2 gene while mice express six e-MARTs as a result of
a duplication of the e-MART-2 gene (Seman, et al., 2004). e-MART-1, -2a, -2b and -5 transfer
ADP-ribose to arginine residues of extracellular target proteins on cell surfaces, or circulating in
body fluids (Corda and Di Girolamo, 2003; Glowacki, et al., 2002; Seman, et al., 2004). E-
MART-3 is expressed on the surface of unstimulated human monocytes whereas e-MART-4 is
only expressed on the cell surface once monocytes are stimulated by lipopolysaccharide
(Grahnert, et al., 2002). Cell surface mono-ADP-ribosylated proteins on human monocytes are
covalently modified at cysteine residues, suggesting that e-MART-3 and -4 may be cysteine-
specific e-MARTs (Grahnert, et al., 2002).
2.3.1.2 Sirtuins
Silent information regulator SIR2-like proteins (sirtuins or SIRTs) are a conserved family of
NAD+-dependent protein deacetylases (SIRT1 to 7) present in eukaryotes, prokaryotes and
archaea (Blander and Guarente, 2004; Denu, 2005; Grubisha, et al., 2005). The SIRT family
regulates a broad range of cellular processes including development, metabolism, DNA repair,
DNA recombination, cellular differentiation, transcriptional regulation, apoptosis and lifespan
(Blander and Guarente, 2004; Denu, 2005; Grubisha, et al., 2005). Most mammalian SIRTs are
reported to exhibit in vivo and in vitro histone deacetylation activity (Blander and Guarente,
2004; Denu, 2005; Michishita, et al., 2005). Of the seven mammalian SIRTs, SIRTs 1, 2, 3 and 5
have been shown to have NAD+-dependent deacetylase activities in vitro (Blander and Guarente,
2004; Denu, 2005; Michishita, et al., 2005). SIRTs also have non-histone substrates and not all
mammalian SIRTs localize to the nucleus.
The NAD+-dependent deacetylation reaction is an unusual mechanism performed by SIRTs
where NAD+ is hydrolyzed into nicotinamide and ADP-ribose (Abdellatif, 2012). The ADP-
ribose moiety is essential for the deacetylation reaction and serves as the acceptor of the acetyl
group removed from the target protein forming the intermediates 2’-O-AADP-ribose and 3’-O-
AADP-ribose (Smith and Denu, 2006; Zhao, et al., 2003; Zhao, et al., 2004). Several reports
have suggested mammalian SIRT family members to contain mono-ADP-ribosyltransferase
activities. SIRTs 1, 2 and 6 were reported to transfer mono-ADP-ribose to bovine serum albumin
9
and histones in vitro (Frye, 1999; Mostoslavsky, et al., 2006; Tanny, et al., 1999). SIRT6 has
auto-mono-ADP-ribosylation (or self ribosylation) but does not possess deacetylase activity
indicating enzymatic mechanisms of each SIRT may differ (Liszt, et al., 2005). Currently, the
potential mono-ADP-ribosyltransferase activities of the different SIRT family members has not
been fully examined and further investigation is required to determine if indeed the SIRTs are a
class of bona fide intracellular mono-ADP-ribosyltransferases (Denu, 2005).
2.3.2 ADP-ribosylhydrolases
ADP-ribosylhydrolases (ARH) reverse mono-ADP-ribosylation reactions by catalyzing the
removal of ADP-ribose from modified proteins (Moss, et al., 1992). The best characterized
ADP-ribosylhydrolase is ADP-ribosylhydrolase 1 (ARH1), which specifically hydrolyzes ADP-
ribose-arginine bonds, leading to the release of free mono-ADP-ribose and regeneration of the
guanidino group of arginine (Moss, et al., 1986; Takada, et al., 1993). In mammalian cells, most
ARH1 activities are cytosolic, although some are also located on the cell surface (Hassa, et al.,
2006). Besides ARH1, additional unidentified proteins exhibiting ADP-ribosyl lyase activities
have been reported, which are thought to function on glutamate, lysine or cysteine residues,
releasing a deoxy form of ADP-ribose (Oka, et al., 1984; Okayama, et al., 1978; Tanuma and
Endo, 1990). In silico screening revealed two ARH-like genes, ARH2 and ARH3 homologous to
the human ARH1 gene (Glowacki, et al., 2002). ARH2 does not appear to possess hydrolase
activity, but ARH3 was shown to hydrolyze poly(ADP-ribose) but not mono-ADP-ribose (see
section 2.4.2.2)(Oka, et al., 2006).
Recently, macrodomain-containing proteins were discovered to catalyze the hydrolysis of mono-
ADP-ribose from modified proteins (Jankevicius, et al., 2013; Rosenthal, et al., 2013; Sharifi, et
al., 2013). Macrodomains are a family of evolutionarily conserved proteins that bind mono- or
poly-ADP-ribose, polyadenylation or O-acetyl-ADP-ribose (Han, et al., 2011; Karras, et al.,
2005; Kim, et al., 2012; Neuvonen and Ahola, 2009). Human proteins MacroD1, MacroD2 and
C6orf130 hydrolyzed mono-ADP-ribose from modified proteins at acidic residues (Jankevicius,
et al., 2013; Rosenthal, et al., 2013). MacroD-like hydrolases inactivated MART target proteins
by removing mono-ADP-ribose and rendering them inactive and thus form the functional
antagonists of intracellular MARTs (Rosenthal, et al., 2013). These findings define
10
macrodomain-containing proteins as a novel group of specific mono-ADP-ribosylhydrolases and
establish mono-ADP-ribosylation as a fully reversible posttranslational modification.
2.4 Poly-ADP-ribosylation Poly-ADP-ribosylation was discovered in multicellular eukaryotes and appears to be less widely
used compared to mono-ADP-ribosylation (Hassa, et al., 2006). Poly-ADP-ribosylation is a
rapid and reversible posttranslational modification in which linear and branched sequences of
ADP-ribose units are covalently attached onto acceptor proteins (Otto, et al., 2005). DNA strand
breakage was long considered to be the main trigger of poly-ADP-ribosylation of proteins;
however, more recent studies suggest poly-ADP-ribosylation is important in a much broader
spectrum of cellular and molecular functions including regulation of chromatin structure,
transcriptional regulation, stress response, inflammation, cell proliferation and differentiation,
apoptosis and maintenance of genome stability (Hakme, et al., 2008; Kraus, 2008; Krishnakumar
and Kraus, 2010b; Luo and Kraus, 2012). Poly-ADP-ribosylation reactions are mediated by
some members of the ARTD family that catalyze the transfer of ADP-ribose units from NAD+ to
form a polymer of ADP-ribose units onto glutamate, aspartate and lysine residues of acceptor
proteins (Hakme, et al., 2008). ARTD1 (formerly known as PARP1), the founding and best-
studied ARTD family member, was first thought to be the only enzyme in mammalian cells to
generate poly(ADP-ribose) polymers. However in recent years, in silico studies have identified
seventeen other proteins that share a catalytic PARP domain with PARP1 and that may contain
poly-ADP-ribosylation activity (Ame, et al., 2004; Otto, et al., 2005).
2.4.1 ADP-ribosyltransferase diphtheria toxin-like (ARTD) family
2.4.1.1 Structural features and nomenclature
ARTDs are a family of ADP-ribosyltransferases that consists of eighteen members all encoded
by different genes. All ARTD family members contain a conserved catalytic domain
homologous to the ARTD1 catalytic domain (Ame, et al., 2004). Besides the catalytic domain, a
wide spectrum of modular domains is present in the different ARTD family members (Figure 3),
suggesting that these members are involved diverse physiological responses (Kleine, et al.,
2008). The catalytic domain is located at the C-terminus of the protein (except in ARTD4) other
modular domains are involved in DNA or RNA binding, protein-protein interactions or cell
11
signalling (Schreiber, et al., 2006). The catalytic domain binds NAD+ via a protein fold termed
the ‘PARP signature’ that shares homology with bacterial mono-ADP-ribosylating exotoxins,
such as diphtheria toxin (Gibson and Kraus, 2012). Based on both structural homology of the
diphtheria toxin in complex with NAD+ and in silico characterization of ARTDs, histidine and
tyrosine (H862 and Y896 in ARTD1) are critical for the proper positioning the ribose moiety of
NAD+ for ADP-ribosylation (Bell and Eisenberg, 1996; Otto, et al., 2005). In poly-ADP-
ribosylating ARTDs, a catalytic glutamate (E988 in ARTD1) is essential for the catalysis of
additional ADP-ribose units or poly-ADP-ribose chain elongation onto acceptor residues
(Marsischky, et al., 1995; Rolli, et al., 1997; Ruf, et al., 1998). These three residues, histidine-
tyrosine-glutamate, make up the catalytic ‘HYE’ triad that appears in all bona fide poly-ADP-
ribosylating ARTDs (Gibson and Kraus, 2012). The absence of the catalytic glutamate of the
ARTD catalytic domain restricts activity to mono-ADP-ribosylation (Kleine, et al., 2008;
Marsischky, et al., 1995; Rolli, et al., 1997; Ruf, et al., 1998). Other structural features such as
the loop length between β-4 and β-5 sheets of the catalytic core have been correlated with lack of
poly-ADP-ribosylation activity (Han and Tainer, 2002; Kleine, et al., 2008; Ruf, et al., 1998;
Sun, et al., 2004). Because the HYE triad of ARTDs is most similar to that of diphtheria toxin,
PARPs were renamed diphtheria-like ADP-ribosyltransferases, abbreviated ARTDs, which better
reflects their transferase rather than polymerase activity (Hottiger, et al., 2010). Of the eighteen
ARTD family members only six contain the catalytic glutamate in the HYE triad, while in the
remaining twelve the catalytic glutamate is replaced by isoleucine, leucine, threonine, valine or
tyrosine residues, suggesting that they will exhibit mono- rather than poly-ADP-
ribosyltransferase activity (Otto, et al., 2005). ARTD9 and ARTD13 lack both the catalytic
glutamate as well as the histidine of the HYE triad and are predicted to be catalytically inactive
(Otto, et al., 2005). Thus ARTDs can be divided into three classes based on their catalytic
activities (Table 1). Class 1 ARTDs are poly-ADP-ribosyltransferases, class 2 are mono-ADP-
ribosyltransferases and class 3 are catalytically inactive.
12
Table 1. Organization and enzymatic activities of mammalian ARTD family.
Class Transferase Name
PARP Name Alternative Name
Triad Motif Enzymatic Activity*
1 ARTD1 PARP1 HYE P & B 1 ARTD2 PARP2 HYE P & B 1 ARTD3 PARP3 HYE M (P predicted) 1 ARTD4 PARP4 VPARP HYE P 1 ARTD5 PARP5a Tankyase 1 HYE P & O 1 ARTD6 PARP5b Tankyase 2 HYE P & O 2 ARTD7 PARP15 BAL3 HYL M 2 ARTD8 PARP14 BAL2 HYL M 2 ARTD10 PARP10 HYI M 2 ARTD11 PARP11 HYI M (predicted) 2 ARTD12 PARP12 ZC3HDC1 HYI M 2 ARTD14 PARP7 TiPARP HYI M (predicted) 2 ARTD15 PARP16 HYI M 2 ARTD16 PARP8 HYI M (predicted) 2 ARTD17 PARP6 HYY M (predicted) 2 ARTD18 TPT1 HHV M (predicted) 3 ARTD9 PARP9 BAL1 QYT I 3 ARTD13 PARP13 ZAP YYV I
*Known or predicted enzymatic activity: mono- (M), oligo- (O) or poly-ADP-ribosylation (P) or
branching (B) or inactive (I). (Adapted from (Gibson and Kraus, 2012))
13
Figure 3. Domain architecture of the 18 human ARTD family members.
Schematic comparison of the domain architecture of the human ARTD (PARP) family (adapted
from (Hottiger, et al., 2010)). The following domains are indicated: The ART domain is the
catalytic core required for ART activity. Within each ART domain, the region that is
homologous to the PARP signature (residues 859–908 of ARTD1) as well as the equivalent of
the ARTD1 catalytic E988 is shaded. The PARP regulatory domain (PRD) might be involved in
regulation of the ADP-ribose branching activity. The WGR domain named after a conserved
central motif (W-G-R) is also found in a variety of polyA polymerases and in proteins of
unknown function. The BRCT domain (BRCA1 carboxy-terminal domain) is found within many
DNA damage repair and cell cycle checkpoint proteins. The sterile alpha motif (SAM), a domain
found within many proteins, can mediate homo- or hetero-dimerization. The ankyrin repeat
domains (ARD) mediate protein–protein interactions. The vault protein interalphatrypsin (VIT)
14
and von Willebrand type A (vWA) domains are conserved domains found in all inter-alpha-
trypsin inhibitor family members. Both of these domains are presumed to mediate protein–
protein interactions. The WWE domain is named after three conserved residues (W-W-E), and is
predicted to mediate specific protein–protein interactions. The Macro domains can serve as
ADPr or O-acetyl-ADP-ribose binding module. ZFD: zinc finger domains. SAP:
SAF/Acinus/PIAS-DNA-binding domain, MVP-ID: Major-vault particle interaction domain,
NLS: nuclear localization signal. CLS: centriole-localization signal. HPS: histidine-proline-
serine region. RRM is an RNA-binding/recognition motif. UIM: ubiquitin interaction motif.
MVP-ID: M-vault particle interaction domain. TPH: Ti-PARP homologous domain. GRD:
glycine-rich domain. TMD: transmembrane domain.
2.4.1.1.1 Class 1 ARTDs
ARTD1/PARP1 is involved in a number of cellular activities including DNA repair, apoptosis,
chromatin remodeling and transcriptional regulation (D'Amours, et al., 1999; Kim, et al., 2005;
Kraus and Lis, 2003; Schreiber, et al., 2006; Tulin and Spradling, 2003). ARTD1 accounts for
approximately 85-90% of mammalian cell poly-ADP-ribosylation activity (Shieh, et al., 1998).
ARTD1 is a nuclear enzyme that is characterized mainly by three modular domains: an N-
terminal DNA binding domain consisting of three zinc finger domains, an automodification
domain and a C-terminal catalytic ADP-ribosyltransferase domain containing the conserved
HYE motif (Figure 4)(D'Amours, et al., 1999). The DNA binding domain plays a critical role in
the recognition of DNA strand aberrations and concurrent activation of ARTD1 (Benjamin and
Gill, 1980; D'Amours, et al., 1999; Ikejima, et al., 1990). The automodification domain is
comprised of a BRCA1-carboxy terminus-like module that mediates several protein-ARTD1 and
DNA-ARTD1 interactions and is also auto-ADP-ribosylation domain (de Murcia and Menissier
de Murcia, 1994; de Murcia, et al., 1994; Mazen, et al., 1989; Schreiber, et al., 1995; Smith,
2001). ARTD1 is best known for its function in the base excision repair pathway during DNA
damage (D'Amours, et al., 1999; de Murcia and Menissier de Murcia, 1994). ARTD1 is a DNA
nick-sensor that is activated by single- and double-strand DNA breaks, DNA hairpins, cruciform
DNA and stable DNA loops (Durkacz, et al., 1980; Lonskaya, et al., 2005; Purnell, et al., 1980).
DNA binding by ARTD1 leads to a conformational change of the enzyme, followed by extensive
auto-ADP-ribosylation and hetero-ADP-ribosylation of histones H1 and H2B (Langelier, et al.,
15
2012; Poirier, et al., 1982). ARTD1-dependent ADP-ribosylation has two effects. One, the
poly(ADP-ribose) chains displace histones leading to decondensation of chromatin (Poirier, et
al., 1982; Zahradka and Ebisuzaki, 1982) and two, the poly(ADP-ribose) chains function as a
binding platform for DNA repair enzymes (Dantzer, et al., 2000; El-Khamisy, et al., 2003;
Leppard, et al., 2003). The accumulation of negative charges added to ARTD1 through auto-
ADP-ribosylation leads to an electrostatic repulsion from the DNA and subsequent loss of
activity (D'Amours, et al., 1999; Schreiber, et al., 2006). Poly(ADP-ribose) chains are then
degraded through hydrolysis by poly(ADP-ribose) glycohydrolase (PARG) (D'Amours, et al.,
1999; Zahradka and Ebisuzaki, 1982). Apart from its role in DNA repair ARTD1 is also
involved in apoptosis, inflammation and transcriptional regulation (Altmeyer, et al., 2010; Kraus,
2008; Kraus and Lis, 2003; Liaudet, et al., 2002).
Figure 4. ARTD1 domain architecture.
ARTD1 uses NAD+ to form polymers of ADP-ribose onto acceptor proteins or to itself
(automodification). Major domains include: DNA-binding domain, a nuclear localization signal
(NLS), an automodification domain containing a BRCT (BRCA1-carboxy terminus-like) motif
and a catalytic ADP-ribosyltransferase domain containing the conserved ‘PARP signature’
NAD+ fold. Adapted from (David, et al., 2009).
ARTD2/PARP2 was discovered as a result of the residual DNA-dependent PARP activity in
embryonic fibroblast derived from Artd1-deficient mice (Ame, et al., 1999; Shieh, et al., 1998).
16
The ARTD2 catalytic domain displays the highest sequence homology (69% similarity) to that of
ARTD1 (Ame, et al., 2004). Similar to ARTD1, ARTD2 is a nuclear protein that is activated by
DNA nicks leading to its auto-ADP-ribosylation and synthesis of long branched chains of
poly(ADP-ribose) (Ame, et al., 1999; Schreiber, et al., 2002). Both Artd1- and Artd2-deficient
mice display sensitivity to DNA damaging agents associated with defects in DNA damage
processing and cell cycle progression but to different extents. Artd1-deficient mice and cell lines
are more sensitive to high-dose ionizing radiation and alkylating agents, whereas Artd2-deficient
mice display hyper-radiosensitivity to low-dose irradiation, demonstrating that ARTD1 and
ARTD2 functions are complementary but do not fully overlap (Chalmers, et al., 2004). Double
mutant Artd1- and Artd2-deficient mice are embryonically lethal highlighting a critical role in
early embryogenesis and underlying partial functional redundancy in maintaining genome
integrity (Menissier de Murcia, et al., 2003; Schreiber, et al., 2002). In addition to its role in
maintenance of genomic stability and DNA repair, ARTD2 has been implicated in various
differentiation processes, including spermatogenesis, adipogenesis and immune cell development
(Bai, et al., 2007; Dantzer, et al., 2006; Robert, et al., 2009; Yelamos, et al., 2006).
ARTD3/PARP3 was initially identified as a core component of the centrosome preferentially
located at the daughter centriole throughout the cell cycle (Augustin, et al., 2003). ARTD3
overexpression interfered with the G1/S phase cell cycle progression and it was described to
interact with ARTD1 at the centrosome (Augustin, et al., 2003). A later study disputed the
centrosomal localization of ARTD3 and suggested that ARTD3 localized to the nucleus and
associated with polycomb group proteins involved in gene silencing and DNA repair networks
including DNA protein kinases, DNA ligase III and IV, Ku70 and Ku80 and ARTD1 (Boehler, et
al., 2011; Rouleau, et al., 2007). ARTD3-depleted cells and Artd1-/- and Artd3-/- double knockout
mice showed increased hypersensitivity to ionizing radiation suggesting a functional synergy of
ARTD3 and ARTD1 in the cellular response to DNA damage (Boehler, et al., 2011). These
studies suggested that ARTD3 is an important partner in the maintenance of genomic stability.
Auto-ADP-ribosylation and hetero-ADP-ribosylation activities of ARTD3 were initially
described as mono-ADP-ribosyltransferase activity (Loseva, et al., 2010); however, a later report
has described ARTD3 to possess poly-ADP-ribosyltransferase activity (Boehler, et al., 2011).
ARTD3 was described to poly-ADP-ribosylate the mitotic factor NuMA directly and indirectly
through ARTD5, suggesting that ARTD3 was required for mitotic spindle integrity during
17
mitosis (Boehler, et al., 2011). Collectively, these reports implicate ARTD3 in the maintenance
of genomic integrity, mitotic spindle integrity and transcriptional repression.
ARTD4/PARP4/VPARP is the largest member of the ARTD family and was originally
identified as a component of mammalian cytoplasmic ribonucleoprotein complexes called vault
particles that have been proposed to be involved in multidrug resistance of human tumours and
to function in intracellular transport (Kickhoefer, et al., 1999). ARTD4 associates with two
essential proteins of the vault particle, major vault protein (MVP) and telomerase-associated
protein (TEP1) (Kickhoefer, et al., 1999). ARTD4 is also present in the nucleus and at the
mitotic spindle suggesting that it may play multiple roles not yet identified (Kickhoefer, et al.,
1999). The structure of ARTD4 is unusual in that it is the only ARTD member to have its
catalytic domain located at the N-terminal portion of the protein. Despite this unique feature,
ARTD4 is catalytically active and poly-ADP-ribosylates MVP and as well as itself (Kickhoefer,
et al., 1999).
ARTD5/PARP5a/Tankyrase 1 and ARTD6/PARP5b/Tankyrase 2 are two closely related
ARTD family members that share 83% sequence identity (Kaminker, et al., 2001; Kuimov, et
al., 2001; Smith, et al., 1998). ARTD5 and ARTD6 have been implicated in a diverse range of
functions including telomere maintenance, WNT signalling, mitosis and mediation of insulin
stimulated glucose uptake (Chi and Lodish, 2000; Cook, et al., 2002; Dynek and Smith, 2004;
Guo, et al., 2012; Hsiao and Smith, 2008; Huang, et al., 2009; Smith and de Lange, 2000; Smith,
et al., 1998). ARTD5 was first discovered as a factor that regulated telomere length by binding
the negative regulator of telomere length telomeric repeat binding factor 1 (TRF1) (Smith, et al.,
1998). ARTD5 was originally named tankyrase 1 due to its interaction with TRF1, the presence
of 24 consecutive ankyrin repeats which make up the major portion of the enzyme, and the
presence of ARTD catalytic domain (Smith, et al., 1998). ARTD5 catalyzed auto-poly-ADP-
ribosylation and poly-ADP-ribosylation of TRF1, implicating ARTD5 in telomere elongation
(Smith, et al., 1998). A careful analysis of ARTD5 auto-ADP-ribosylation revealed that ARTD5
synthesized ADP-ribose polymers with an average length of 20 ADP-ribose units but polymers
lacked branching (Rippmann, et al., 2002). ARTD6 was also reported to associate with and poly-
ADP-ribosylate TRF1, indicating a potential redundant role of ARTD5 and ARTD6 in telomere
regulation (Cook, et al., 2002; Kaminker, et al., 2001). ARTD6 also associates with ARTD5 and
both enzymes share most of their protein partners including insulin-responsive aminopeptidase
18
(IRAP), NuMA and 182 kDa tankyrase-binding protein (TAB182) (Kaminker, et al., 2001;
Sbodio and Chi, 2002; Sbodio, et al., 2002; Seimiya and Smith, 2002). Artd5- and Artd6-
deficient mice do not display observed defects in development; however, the Artd5-/- and Artd6-/-
double knockout was embryonical lethal (Chiang, et al., 2008).
2.4.1.1.2 Class 2 ARTDs
ARTD7/PARP15/BAL3 and ARTD8/PARP14/BAL2 were originally identified as two genes
closely related to ARTD9/PARP9/BAL1 (B-aggressive lymphoma 1, see section 2.4.1.1.3)
(Aguiar, et al., 2005; Aguiar, et al., 2000). The encoded proteins both contain N-terminal
macrodomains and a C-terminal PARP catalytic domain (Aguiar, et al., 2005). Macrodomains
are protein domains known to bind mono- and poly-ADP-ribose (Forst, et al., 2013; Han and
Tainer, 2002; Kleine and Luscher, 2009; Welsby, et al., 2012). Recently, macrodomains 2 and 3
of ARTD8 were reported to recognize and read mono-ADP-ribosylated ARTD10 and substrates
of ARTD10 (Forst, et al., 2013). Both ARTD7 and ARTD8 demonstrated auto-mono-ADP-
ribosylation activity (Aguiar, et al., 2005). ARTD7 was reported to have a transcriptionally
repressive function through its N-terminal macrodomains and its auto-ADP-ribosylation activity
was suggested to counteract the repressive effect of the macrodomains (Aguiar, et al., 2005).
Whereas ARTD7 remains poorly characterized, ARTD8 is better understood and has been
implicated in STAT6 (Signal Transducer and Activator of Transcription 6)-dependent
transcriptional control and cytokine-regulated control of cellular metabolism (Cho, et al., 2011;
Goenka, et al., 2007). ARTD8 potentiated interleukin 4 (IL4)-induced STAT6 transactivation via
its macrodomains and catalytic activity (Goenka and Boothby, 2006; Goenka, et al., 2007).
ARTD8 was also recently identified as a downstream effector of the Jun N-terminal kinase 2
(JNK2)-dependent pro-survival signal by binding to and inhibiting JNK1 pro-apoptotic activity
promoting the survival of myeloma cells (Barbarulo, et al., 2012).
ARTD10/PARP10 is the founding member of the class 2 ARTDs and was discovered through in
silico screening ARTD family members (Ame, et al., 2004). The domain structure of ARTD10,
with the exception of the catalytic domain is unique from the other ARTDs. The N-terminus
contains a putative RNA recognition motif (RRM) followed by a glycine-rich region, which is
present in other RNA-binding proteins (Burd and Dreyfuss, 1994). The C-terminus contains a
functional nuclear export signal (NES) within its glutamate-rich region and two functional
19
ubiquitin-interaction motifs (UIMs) (Yu, et al., 2005). ARTD10 is a MART that auto-ADP-
ribosylates itself as well as each of the four core histones (Kleine, et al., 2008; Yu, et al., 2005).
ARTD10 was also reported to be an interacting partner of the oncoprotein MYC (Yu, et al.,
2005). MYC/HA-RAS- and E1A/HA-RAS-dependent transformation of rat embryo fibroblasts
was inhibited by ARTD10 (Yu, et al., 2005). Inhibition was independent of ARTD10 catalytic
activity and neither MYC nor its heterodimerization partner MAX were ADP-ribosylated by
ARTD10 (Yu, et al., 2005). Recently, ARTD10 has also been reported to be a regulator of the
NF-κB pathway by mono-ADP-ribosylating NF-κB essential modulator, reducing its poly-
ubiquitination and activation of NF-κB (Verheugd, et al., 2013). Unlike the regulation of MYC,
the regulation of NF-κB was dependent on ARTD10 catalytic activity and the UIMs (Verheugd,
et al., 2013).
ARTD12/PARP12/ZC3HDC1 has been implicated in inflammatory processes in relation to
various pathogens. During infection by the alphavirus, Venezuelan equine encephalitis virus
(VEEV), the long isoform (containing the catalytic domain) of ARTD12 was up-regulated
(Atasheva, et al., 2012). ARTD12 exhibited inhibitor effects on replication of VEEV as well as
other alphaviruses and RNA viruses (Atasheva, et al., 2012). Interferon stimulation up-regulated
ARTD12 gene expression to counteract infections which was suggested as being a cellular
defence against invading viral pathogens, although this was not mechanistically determined
(Schoggins, et al., 2011). In addition to its role in viral pathogen infection inhibition, ARTD12
has been implicated in post-transcriptional gene expression (Leung, et al., 2011). In that study,
overexpressed ARTD12 assembled in cytoplasmic mRNA-protein complexes called stress
granules and alleviated microRNA (miRNA)-mediated repression (Leung, et al., 2011).
ARTD12 exhibited auto-mono-ADP-ribosylation activity and associated with both long and
short isoforms of ARTD13 (short isoform is missing catalytic domain), ARTD5 and ARTD7
within stress granules (Leung, et al., 2011).
ARTD14/PARP7/TiPARP (will be referred to as TiPARP) was first identified as a 2,3,7,8-
tetrachlorodibenzo-p-dioxin (TCDD or dioxin)-induced gene under the control of TCDD-
activated aryl hydrocarbon receptor (AHR) (Ma, et al., 2001). TiPARP is broadly expressed in
murine tissues, suggesting that TiPARP may be involved in diverse physiological functions (Ma,
et al., 2001). In its initial study, TiPARP exhibited enzymatic ADP-ribosylation capacity but it
20
was not determined whether it undergoes auto- or hetero-ADP-ribosylation and whether this was
mono-ADP-ribosylation activity (Ma, et al., 2001). A more recent report has identified TiPARP
as a mediator the TCDD-induced suppression of hepatic glucose production (Diani-Moore, et al.,
2010). This study proposed the TCDD-induced expression of TiPARP contributed to NAD+
depletion by TiPARP catalytic activity, which decreased SIRT1 activity leading to decreased
gluconeogenic gene expression (Diani-Moore, et al., 2010). TCDD-induced TiPARP gene
expression and NAD+ depletion has also been implicated in the deactivation of SIRT3 which
leads to decreased superoxide dismutase 2 activity resulting in heightened sensitivity to non-
alcoholic steatohepatitis (He, et al., 2013). Both proposed mechanisms of TiPARP-mediated
NAD+ depletion were derived from observation that TCDD induces TiPARP gene expression
and decreases NAD+ levels. The enzymatic activity of TiPARP was not properly investigated in
these studies and neither study confirmed the mono-ADP-ribosyltransferase activity of TiPARP
and/or if this was required for the observed outcomes.
ARTD15/PARP16 was the first ARTD member found to associate with the endoplasmic
reticulum (ER) (Di Paola, et al., 2012). ARTD15 was the only ARTD member to contain a
predicted C-terminal transmembrane domain and was found to be an ER-tailed anchored enzyme
that associated with the nuclear transport factor karyopherin-β1/importin-β1 (Kapβ1) at the ER
and the nuclear envelope (Di Paola, et al., 2012). Kapβ1 was selectively mono-ADP-ribosylated
by ARTD15 and this modification was hypothesized to control nuclear transport (Di Paola, et al.,
2012). ARTD15 also demonstrated auto-mono-ADP-ribosylation that could be inhibited with
canonical PARP inhibitors (Di Paola, et al., 2012; Karlberg, et al., 2012). Mono-ADP-
ribosylation was reported to not occur at arginine, glutamate or cysteine residues and could not
be reverted by ARH1 or ARH3 (Di Paola, et al., 2012). A recent report has described that
ARTD15 was required for activation of the ER stress response kinases PERK and IREα during
the unfolded protein response (UPR), an ER stress signal that ultimately leads to apoptosis (Hetz,
2012; Jwa and Chang, 2012). During ER stress, ARTD15 was found to auto-ADP-ribosylate
itself and hetero-ADP-ribosylate PERK and IREα (Jwa and Chang, 2012). ADP-ribosylation of
PERK and IREα increased their kinase activities and the endoribonuclease activity of IREα,
which appeared necessary for proper execution of the UPR (Jwa and Chang, 2012).
21
ARTD17/PARP6 was recently described as a negative regulator of cell proliferation and
proposed tumour suppressor involved in colorectal cancer development (Tuncel, et al., 2012).
ARTD17 overexpression was reported to arrest cells in S-phase and was dependent on the
presence of the catalytic domain (Tuncel, et al., 2012). The catalytic activity of ARTD17 has not
been evaluated and its potential activity in other cellular functions has not been fully determined.
ARTD11/PARP11, ARTD16/PARP8 and ARTD18/TPT1 (the single member of the NAD+-
dependent tRNA 2’-phosphotransferase family) have no known domains outside of their
catalytic domains (with the exception of the single WWE domain of ARTD11) and their
functions have not been determined.
2.4.1.1.3 Class 3 ARTDs
ARTD9/PARP9/BAL1 is a nucleocytoplasmic shuttling protein that has been identified as a
risk-related gene product in aggressive diffuse large B-cell lymphoma (DLBCL) (Aguiar, et al.,
2000; Juszczynski, et al., 2006). ARTD9 contains two prototypical macrodomains within the N-
terminus, which can bind mono- and poly-ADP-ribose (Karras, et al., 2005). ARTD9 was
described to possess transcriptional repressive activity that was dependent on interaction through
its macrodomains but independent of its catalytic activity (Aguiar, et al., 2005). Overexpression
of ARTD9 promoted lymphocyte migration, indicating a tumour-promoting role in high-risk
DLBCL (Aguiar, et al., 2000). Promotion of lymphocyte migration by ARTD9 has been
suggested through modulation of interferon gamma (IFNγ) signalling-related gene expression
(Juszczynski, et al., 2006). ARTD9 was identified as a novel co-repressor of transcription of
interferon response factor 1 (IRF1), a tumour suppressor (Camicia, et al., 2013). ARTD9 directly
interacted with STAT1β (signal transducer and activator of transcription 1 isoform β) to inhibit
IRF1 expression, repressing the anti-proliferative and pro-apoptotic INFγ-STAT1-IRF1-p53
complex (Camicia, et al., 2013). ARTD9 has also been linked to the DNA damage response
pathway (Yan, et al., 2013). In response to DNA strand breaks, ARTD9 and its partner BBAP
(B-lymphoma and BAL-associated protein), an E3 ligase were recruited to DNA damage sites
and co-localized with ARTD1 and its product poly(ADP-ribose) (Yan, et al., 2013). ARTD9 and
BBAP at DNA damage sites mediate the specific recruitment of the adaptor protein RAP80 and
checkpoint mediators 53BP1 and BRCA1 through BBAP-mediated ubiquitylation, which limits
early and delayed DNA damage and enhances cellular viability (Yan, et al., 2013).
22
ARTD13/PARP13/ZAP/ZC3HAV1 is a type 1 interferon-inducible host factor that regulates
viral RNA transcripts and was initially identified as zinc finger antiviral protein (ZAP) in a
screen for host factors that confer resistance to the retrovirus murine leukemia virus (MLV)
infection (Bick, et al., 2003; Gao, et al., 2002; Muller, et al., 2007). ARTD13 antiviral activities
were later confirmed in other retroviruses including HIV-1 and other RNA virus families,
including alphaviruses and filoviruses (Bick, et al., 2003; Muller, et al., 2007; Zhu, et al., 2011).
ARTD13 binds directly to specific viral mRNAs through its N-terminal zinc finger domains and
recruits cellular mRNA degradation factors to promote degradation of the target viral mRNA and
the inability of the virus to replicate efficiently (Guo, et al., 2004; Guo, et al., 2007; Zhu, et al.,
2011; Zhu and Gao, 2008). These antiviral properties are not due to ADP-ribosylation since full-
length ARTD13 is not catalytically active nor is its short isoform lacking the PARP catalytic
domain (Kleine, et al., 2008; Leung, et al., 2011). Both isoforms of ARTD13 were recently
reported to localize to cytoplasmic stress granules along with ARTD5, ARTD7 and ARTD12
(Leung, et al., 2011). Interestingly, when overexpressed, both isoforms decreased miRNA-
mediated silencing (Leung, et al., 2011).
2.4.2 Poly(ADP-ribose) Catabolism
2.4.2.1 Poly(ADP-ribose) glycohydrolase (PARG)
The removal of poly(ADP-ribose) chains from modified proteins is catalyzed by poly-ADP-
ribose glycohydrolase (PARG), which is the primary enzyme responsible for poly(ADP-ribose)
degradation in vivo (Alvarez-Gonzalez and Althaus, 1989; Davidovic, et al., 2001; Jonsson, et
al., 1988). PARG has both exo- and endoglycosidase activities that hydrolyze the glycosidic
bond between ADP-ribose units that generate large amounts of free ADP-ribose (Desnoyers, et
al., 1995; Heeres and Hergenrother, 2007; Oka, et al., 1984). The structure of a bacterial PARG
revealed that the PARG catalytic domain is a distant member of the macrodomain family
suggesting a potential role in the regulation by poly-ADP-ribosylation (Slade, et al., 2011). The
single mammalian PARG gene encodes several PARG proteins that localize to various cellular
compartments that also contain ARTDs (Meyer-Ficca, et al., 2004). Four PARG isoforms have
been identified so far, but the two predominant isoforms include PARG-110/111 (110 kDa), the
full-length and very active nuclear isoform and PARG-59/60 (65 kDa), a shorter isoform
(Brochu, et al., 1994; Meyer-Ficca, et al., 2004). PARG-102 (102 kDa), a splice variant that
lacks exon 1 and PARG-99 (99 kDa), a splice variant that lacks exons 1 and 2 are cytoplasmic
23
but their roles are not fully understood (Meyer-Ficca, et al., 2004). Because of its low abundance
in eukaryotic cells and its extreme sensitivity to proteases, PARG has been difficult to study
(Bonicalzi, et al., 2005). Furthermore, a detailed analysis of the biological role of PARG has
been precluded by the lack of specific and selective inhibitors (Schreiber, et al., 2006).
Much of the current understanding of the biological role of PARG is largely based on gene
disruption studies and data obtained from Parg-deficient mice (Burkle and Virag, 2013).
Targeted deletion of exon 4 resulted in complete deletion of all PARG isoforms in Parg-deficient
mice (Koh, et al., 2004). Mice lacking all PARG isoforms arrest during development at
embryonic day 3.5 as a result of apoptosis that was induced by poly(ADP-ribose) accumulation
(Koh, et al., 2004). In vitro gene knockdown studies have revealed that PARG, like ARTD1, is
required for efficient DNA repair of single- and double-strand breaks and mitotic spindle
checkpoints since knockdown of either PARG or ARTD1 sensitized cells to apoptotic cell death
following DNA damage (Ame, et al., 2009; Erdelyi, et al., 2009; Fisher, et al., 2007; Keil, et al.,
2006). Mice that are deficient of full-length PARG (PARG110/111, Parg110-deficient) are
viable with no obvious phenotype and normal poly(ADP-ribose) metabolism (Cortes, et al.,
2004). However, Parg110-deficient mice are hypersensitive to ionizing radiation and alkylating
agents, similar to Artd1-deficient mice (Cortes, et al., 2004). Data obtained from Parg110-
deficient embryonic fibroblasts are similar to those reported for ARTD1 inhibition/knockout
studies, including defects in DNA repair, genomic instability and chromosomal aberrations (Min,
et al., 2010). These observations support a model in which a balance between poly(ADP-ribose)
synthesis and degradation is necessary for efficient repair of DNA strand breaks and cell
survival.
2.4.2.2 ADP-ribosylhydrolase 3 (ARH3)
ADP-ribosylhydrolase 3 (ARH3) is a member of the family of dinitrogenase reductase-activating
glycohydrolase-related proteins and shares very little similarity with the PARG catalytic domain
sequence (Mueller-Dieckmann, et al., 2006; Oka, et al., 2006). ARH3 localizes to the
mitochondria but is also present in the nucleus and cytosol (Niere, et al., 2008; Oka, et al., 2006).
ARH3 catalyzes the hydrolysis of both poly(ADP-ribose) and O-acetyl-ADP-ribose (OAADPR),
a metabolite that results from the activity of sirtuins (Imai and Guarente, 2010; Oka, et al., 2006;
Ono, et al., 2006). ARH3 was shown to degrade poly(ADP-ribose) in vitro and in cell but its
24
specific activity was significantly less than that of PARG (Niere, et al., 2008; Oka, et al., 2006).
Embryonic fibroblasts from Arh3-deficient mice lacked most of the mitochondrial poly(ADP-
ribose) degrading activity and ARH3 appears to be the only identified enzyme to degrade PAR in
the mitochondrial matrix (Niere, et al., 2012). Because of its localization to the cytosol and
nucleus as well as its poly(ADP-ribose) degrading activity, ARH3 has also been proposed to
assist PARG in degrading poly(ADP-ribose) in the nucleus and cytosol (Niere, et al., 2012).
2.5 Regulation of chromatin structure and transcription by ARTDs
The disruption of chromatin structure by ADP-ribosylation of histones and destabilization of
nucleosomes were some of the earliest characterized effects of ARTD1 (Huletsky, et al., 1989;
Mathis and Althaus, 1987; Poirier, et al., 1982). The ability of ARTD1 to modulate chromatin
structure and function underlies its contribution to transcriptional regulation (Krishnakumar and
Kraus, 2010b). Early studies reported that ARTD1 activity is preferentially associated with
regions of chromatin that are actively transcribed (De Lucia, et al., 1996; Levy-Wilson, 1981;
Mullins, et al., 1977). ARTD1 was identified as a factor that could increase specificity of RNA
polymerase transcriptional initiation, supporting ARTD activity to directly regulate transcription
(Matsui, et al., 1980; Slattery, et al., 1983). Transcriptional regulation by ARTD activity has
been proposed to occur by at least two mechanisms that are not mutually exclusive: (1)
modification of histones to alter chromatin structure and (2) functioning as part of the regulatory
region (enhancer/promoter) binding complexes in conjunction with other DNA-binding factors
and co-activators (Kraus and Lis, 2003).
2.5.1 Modulation of chromatin structure by ARTD1 and poly(ADP-ribose)
Poly-ADP-ribosylation of chromatin protein can cause profound effects on nucleosome
architecture and on the stability of individual nucleosomes (D'Amours, et al., 1999). Histones H1
and H2B show the most poly-ADP-ribosylation in vivo and are the preferred targets of ARTD1
in vitro, although all histones are modified to some extent (Adamietz and Rudolph, 1984;
D'Amours, et al., 1999; Huletsky, et al., 1989; Poirier, et al., 1982). As seen directly by electron
microscopy, native polynucleosomes when poly-ADP-ribosylated by purified ARTD1 led to
complete decondensation and mimicked the effects of linker histone H1 depletion from higher-
order chromatin (Poirier, et al., 1982). Further studies suggested polyanionic poly(ADP-ribose),
25
attached to target proteins or as a free polymer acts as an attractive core histone binding matrix
that can act as a histone acceptor to further destabilize nucleosomes (Mathis and Althaus, 1987;
Realini and Althaus, 1992). Moreover, ARTD1-dependent accumulation of poly(ADP-ribose) at
decondensed, transcriptionally active loci in native chromatin has been reported (Tulin and
Spradling, 2003). Collectively, these data support a model of transcriptional regulation whereby
ARTD1 promotes the decondensation of chromatin by causing the disassembly of nucleosomes
through poly-ADP-ribosylation of H1 and core histones, causing destabilization of nucleosomes
(D'Amours, et al., 1999; Kraus and Lis, 2003; Rouleau, et al., 2004; Tulin and Spradling, 2003).
More recent biochemical studies have demonstrated that in the absence of NAD+ or significant
auto-ADP-ribosylation, ARTD1 binds to nucleosomes and promotes the condensation of
chromatin by bringing together neighbouring nucleosomes (Kim, et al., 2004; Wacker, et al.,
2007). Addition of NAD+, which led to considerable auto-ribosylation of ARTD1, promoted the
release of ARTD1 from chromatin, leading to nucleosome decondensation and the restoration of
transcription (Kim, et al., 2004; Wacker, et al., 2007). These results suggest a new NAD+-
dependent model of modulation of chromatin structure and transcription whereby ARTD1 can
regulate chromatin structure without modifying histones or promoting disassembly of
nucleosomes (Kim, et al., 2004; Wacker, et al., 2007).
2.5.2 Enhancer-binding of actions of ARTD1
In addition to modulating transcription through alterations in chromatin structure, ARTD1 can
regulate transcription by functioning as a direct enhancer-binding factor. Many of the initial
studies describing the effects of ARTD on transcriptional regulation of target genes focused on
the binding of ARTD1 to specific DNA sequences or structures (Akiyama, et al., 2001; Butler
and Ordahl, 1999; Huang, et al., 2004; Nirodi, et al., 2001; Zhang, et al., 2002). Although no
consensus DNA binding sequence for ARTD1 has been established, the ability of ARTD1 to
bind DNA in a sequence-specific manner has been demonstrated where ARTD1 preferentially
bound to DNA oligonucleotides containing the 5’-TGTTG-3’ nucleotide sequence motif (Huang,
et al., 2004; Simbulan-Rosenthal, et al., 2003). Two independent studies demonstrated ARTD1
binds to specific sequences immediately upstream of chemokine (CXC motif) ligand 1 (CXCL1)
and in the first intron of B-cell lymphoma 6 (BCL6) to repress transcription (Ambrose, et al.,
2007; Amiri, et al., 2006).
26
2.5.3 Promoter-binding and transcriptional co-regulation by ARTD1
Genomic studies have revealed that ARTD1 localizes to as many as 90% of expressed Pol II-
transcribed promoters in vitro which suggested that it plays a role in promoting the formation of
chromatin structures that are receptive to transcription (Krishnakumar, et al., 2008). The
enrichment of ARTD1 at these promoters correlated with the depletion of linker histone H1, and
a high ARTD1/H1 ratio specified genes that are actively transcribed (Krishnakumar, et al.,
2008). These findings indicated that ARTD1 localized to sites of ongoing transcription, where it
may exert stimulatory or inhibitory effects (Krishnakumar, et al., 2008). Roles of ARTD1 as a
promoter-specific co-regulator (either co-activator or co-repressor) of different sequence specific
DNA-binding transcriptional factors have been reported (Table 2). In some cases, the catalytic
activity of ARTD1 is required, while in other cases it is not (Ju, et al., 2004; Olabisi, et al., 2008;
Zaniolo, et al., 2007; Zhang, et al., 2013). These studies highlight the diverse mechanisms of
ARTD1 co-regulation, which are likely to vary in a gene- or promoter-specific manner.
Table 2. Transcription factors co-regulated by ARTD1.
Transcription factor
Co-regulatory effect
Catalytic function required?
Reference
AP2 stimulatory n/d (Kannan, et al., 1999) b-MYC stimulatory no (Cervellera and Sala, 2000) Elk-1 stimulatory yes (Cohen-Armon, et al., 2007) ERα stimulatory yes (Zhang, et al., 2013) HES1 stimulatory yes (Ju, et al., 2004)
HTLV Tax1 stimulatory no (Anderson, et al., 2000) NFAT stimulatory yes (Olabisi, et al., 2008) NF-κB stimulatory no (Hassa and Hottiger, 2002) Oct-1 stimulatory n/d (Nie, et al., 1998) p53 inhibitory yes (Mendoza-Alvarez and Alvarez-
Gonzalez, 2001) RAR stimulatory no (Pavri, et al., 2005) Sp1 inhibitory yes (Zaniolo, et al., 2007)
TEF-1 inhibitory no (Butler and Ordahl, 1999) TR/RXR inhibitory yes (Miyamoto, et al., 1999)
YY1 inhibitory n/d (Oei, et al., 1997)
AP2; activating protein 2, Elk-1; ETS domain-containing protein, ERα; estrogen receptor α,
HES1; hairy and enhancer of split 1, HTLV; human T-cell leukemia virus type 1, NFAT; nuclear
factor of activated T-cells, NF-κB; nuclear factor kappa-light-chain-enhancer of activated B-
cells, Oct-1; octamer transcription factor 1, RAR; retinoic acid receptor, Sp1; specificity protein
27
1, TEF-1; transcriptional enhancer factor 1, TR/RXR; thyroid hormone receptor/retinoid X
receptor heterodimer, YY1; yin ying 1. n/d; not determined.
2.5.4 Transcriptional regulation by other ARTDs
Studies have emerged examining the regulation of transcription by other ARTD members,
including, ARTD2, ARTD8, ARTD9 and ARTD10 (Aguiar, et al., 2005; Camicia, et al., 2013;
Szanto, et al., 2012; Yu, et al., 2005).
Similar to ARTD1, ATRD2 has been demonstrated to co-regulate DNA-binding transcription
factors (Bai, et al., 2011; Bai, et al., 2007; Maeda, et al., 2006). The activity of the peroxisome
proliferator-activated receptor γ (PPARγ) was modulated by ARTD2 (Bai, et al., 2007). ARTD2
demonstrated direct interaction and co-occupancy with PPARγ/RXR heterodimers at promoter
regions of PPARγ target genes and acts as a co-activator of transcription leading to triglyceride
accumulation (Bai, et al., 2007). Estrogen receptor α (ERα) but not ERβ-dependent transcription
was decreased by ARTD2 knockdown suggesting a co-activator role of ARTD2 in ERα-
dependent transcription (Bai, et al., 2007; Szanto, et al., 2012). ARTD2 along with ARTD1 have
also been reported to serve as co-activators of thyroid transcription factor 1 (TTF1) and regulate
expression of surfactant protein B in lung (Maeda, et al., 2006). ARTD2 has been reported to be
a potent transcriptional repressor of the SIRT1 promoter and the reduction or knockout of
ARTD2 expression induced SIRT1 mRNA expression, protein and activity (Bai, et al., 2011).
The mono-ADP-ribosyltransferase ARTD8 was reported to activate a STAT6-regulated reporter
gene, which was dependent on its macro- and catalytic domains (Goenka and Boothby, 2006;
Goenka, et al., 2007). ARTD8 acted as a transcriptional switch, where under non-stimulating
conditions ARTD8 was bound to STAT6-responsive promoters and recruited histone deacetylase
2 (HDAC2) and HDAC3 to keep genes silent (Mehrotra, et al., 2011). Upon IL4 stimulation
ARTD8 catalyzed the ADP-ribosylation of itself, HDAC2 and HDAC3 leading to their
dissociation from the promoter thereby allowing STAT6 dimers binding (Mehrotra, et al., 2011).
ARTD10 may possibly play a role in MYC-induced transcription. ARTD10 was identified as an
interacting partner of MYC and this interaction was demonstrated to occur in the nucleus
(Kleine, et al., 2012; Yu, et al., 2005). ARTD10 has also recently been reported to negatively
regulate NF-κB signalling in response to IL-1β and TNFα stimulation by inhibiting NF-κB
28
target gene and protein expression (Verheugd, et al., 2013). ARTD10-mediated inhibition of NF-
κB signalling was dependent on its catalytic activity and mono-ADP-ribosylation of NF-κB
essential modulator by ARTD10 reduced its poly-ubiquitination, leading to reduced expression
of NF-κB target genes (Verheugd, et al., 2013). ARTD10 also prevented the nuclear
translocation of the NF-κB transcription factor p65 (Verheugd, et al., 2013).
Recently, ARTD9 has been identified as a novel co-repressor of transcription of IRF1, a tumour
suppressor (Camicia, et al., 2013). ARTD9 directly interacted with STAT1β through its
macrodomains mediated by trans-mono-ADP-ribosylation to inhibit IRF1 expression (Camicia,
et al., 2013). This was shown to repress the anti-proliferative and pro-apoptotic INFγ-STAT1-
IRF1-p53 complex and mediated proliferation, survival and chemo-resistance in DLBCL
(Camicia, et al., 2013).
The transcriptional regulatory roles of the remaining ARTD members, including TiPARP have
not been evaluated. The expression of TiPARP is regulated by aryl hydrocarbon receptor (AHR),
a ligand activated transcription factor, but its potential function in AHR transactivation has not
been evaluated.
3 Aryl hydrocarbon receptor The AHR is a ligand-activated transcription factor and a member of the basic-helix-loop-helix
Per (Period)-ARNT (aryl hydrocarbon receptor nuclear translocator)-SIM (single-minded)
(bHLH/PAS) family of heterodimeric transcriptional regulators (Burbach, et al., 1992). Other
members of the bHLH/PAS family of transcription factors include ARNT, hypoxia factor 1α
(HIF1α), SIM, Clock, Per, and the p160 family of co-activators (Hankinson, 1995). bHLH/PAS
family of transcription factors are involved in the control of a variety of physiological processes
which include: circadian rhythms, organ development, neurogenesis, metabolism and the stress
response to hypoxia (Crews, 1998; Fernandez-Salguero, et al., 1997; Gonzalez and Fernandez-
Salguero, 1998; Whitlock, 1999). Although AHR plays an important role in physiology it is most
well-known for mediating the toxic effects of environmental contaminants (Abbott, et al., 1995;
Hahn, 2002; Lahvis, et al., 2000; Lin, et al., 2001; Mimura, et al., 1997; Schmidt, et al., 1996;
Van den Berg, et al., 1998).
29
3.1 Structure The human AHR is composed of several modular domains with distinct functions (Figure 5A).
AHR shares similar domain architecture to other bHLH/PAS proteins, with a highly conserved
N-terminal bHLH motif, adjacent PAS region, and loosely conserved C-terminal transactivation
or transrepression regions (Figure 5B and C) (Kewley, et al., 2004). The basic region of the
bHLH motif mediates DNA binding and the HLH domain is involved in dimerization (Kikuchi,
et al., 2003). Within the N-terminus of AHR is a bipartite nuclear localization signal (NLS),
which overlaps with the basic region of the bHLH (Ikuta, et al., 1998). Moreover, AHR also
contains a leucine-rich NES with helix 2 of the bHLH motif (Ikuta, et al., 1998). The PAS region
of AHR contains two tandem PAS domains designated PAS A and PAS B and both PAS
domains along with the bHLH motif contribute to dimerization with AHR obligatory interaction
partner ARNT and interaction with chaperone protein HSP90 (heat shock protein 90 kDa)
(Lindebro, et al., 1995; Pongratz, et al., 1992). The PAS A domain controls dimerization
specificity and stability, and strengthens DNA binding (Chapman-Smith, et al., 2004; Lindebro,
et al., 1995; Pongratz, et al., 1998). The PAS B domain contains the ligand binding domain and
deletion of this domain results in a constitutively active receptor (McGuire, et al., 2001).
The C-terminal half of AHR contains a modular transactivation domain (TAD) that is composed
of three distinct subdomains: acidic, glutamine-rich (Q-rich) and proline-serine-threonine-rich
(P/S/T) (Jain, et al., 1994; Rowlands, et al., 1996; Sogawa, et al., 1995). Individually, each of the
three subdomains exhibits low levels of transactivation; however, any combination of two
subdomains can synergistically activate transcription (Rowlands, et al., 1996). The Q-rich sub-
domain is critical for transcriptional activation and its deletion completely inactivates the
receptor (Kumar, et al., 2001). Additionally the Q-rich subdomain also appears to be required for
interaction with co-regulatory proteins nuclear co-activator 1 (NcoA1) and receptor interacting
protein 140 (RIP140) (Kumar and Perdew, 1999; Kumar, et al., 2001; Kumar, et al., 1999).
30
Figure 5. Schematic of the modular domains of three bHLH-PAS family members.
(A) Aryl hydrocarbon receptor (AHR), (B) Aryl hydrocarbon nuclear translocator (ARNT), and
the (C) Aryl hydrocarbon receptor repressor (AHRR). bHLH (basic helix-loop-helix), PAS
(Per/ARNT/Sim), P/S/T (Proline/Serine/Threonine).
3.2 AHR ligands AHR is a promiscuous receptor that can bind and be activated by a wide variety of structurally
diverse exogenous and natural compounds that produce a spectrum of the biological and toxic
effects (Denison and Heath-Pagliuso, 1998; Denison and Nagy, 2003; Denison, et al., 2011;
Nguyen, et al., 1999; Poland and Knutson, 1982; Safe, 1990). AHR ligands can be divided into
two major categories: exogenous and naturally occurring.
31
3.2.1 Exogenous AHR ligands
The majority of the high affinity AHR ligands that have been identified and characterized are
exogenous, which include planar hydrophobic halogenated aromatic hydrocarbons (HAHs) and
polycyclic aromatic hydrocarbons (PAHs) and related compounds (Figure 6) (Denison and
Heath-Pagliuso, 1998; Gillner, et al., 1993; Poland and Knutson, 1982; Safe, 1990). Many HAHs
and PAHs are environmental contaminants that are by-products of chlorine chemistry, waste
incineration, metal production and fossil fuel or wood burning (Denison and Nagy, 2003;
DeVito, et al., 1994; Hankinson, 2005; Kulkarni, et al., 2008; Safe, 1986; Van den Berg, et al.,
1998). The HAHs represent the most potent class of AHR ligands and include chlorinated and
brominated dibenzo-p-dioxins, dibenzofurans and biphenyls. HAHs are metabolically stable and
have binding affinities in the picomolar to nanomolar range (Denison and Nagy, 2003). PAHs
include: 3MC, B[a]P, benzoanthracenes and benzoflavones (Denison and Nagy, 2003; Poland
and Knutson, 1982). PAHs are metabolically less stable than HAHs and tend to have lower
binding affinities usually within the nanomolar to micromolar range. Structure-activity
relationship analysis of many HAHs and PAHs congeners has demonstrated toxicity is positively
correlated to the affinity of the congener for AHR (Kafafi, et al., 1993; Mhin, et al., 2002;
Poland and Knutson, 1982; Safe, 1986; Tuppurainen and Ruuskanen, 2000; Waller and
McKinney, 1995). The HAH, 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD, dioxin) has the
highest affinity for AHR and is the most toxic of the known AHR ligands and remains the
prototype AHR ligand for which all subsequent ligands are compared (Nebert, et al., 2004).
32
Figure 6. Selected exogenous AHR ligands.
3.2.2 Naturally occurring AHR ligands
The greatest source of human exposure to AHR ligands (exogenous and naturally occurring)
comes from the diet (Denison and Nagy, 2003; Jeuken, et al., 2003). A variety of naturally
occurring dietary constituents are AHR ligands (Bjeldanes, et al., 1991; MacDonald, et al., 2001;
Wattenberg and Loub, 1978). Indole-3-carbinol (IC3) is found in cruciferous vegetables and is
hydrolyzed under the acidic conditions of the gastrointestinal tract to many products including
3,3’-diindolylmethane (DIM) and indolo[3,2-b]carbazole (ICZ) (Bjeldanes, et al., 1991; Grose
and Bjeldanes, 1992). DIM is a partial agonist for AHR while ICZ is a potent activator of AHR
activity (Broadbent and Broadbent, 1998; Chen, et al., 1996; Kleman, et al., 1994). ICZ has high
33
affinity for AHR but it does not cause toxicity (Pohjanvirta, et al., 2002). Naturally occurring
AHR ligands can also be antagonists and numerous have been previously described as such.
Resveratrol (3,5,4’-trihydroxystilbene), a polyphenol present in grapes, peanuts and berries and
kaempferol (3,4’,5,7,-tetrahydroxyflavone), a flavonol found in teas, broccoli, grapefruit and
other plant sources are two potent AHR antagonists (Casper, et al., 1999; Ciolino, et al., 1999;
MacPherson and Matthews, 2010).
To date no definitive endogenous AHR ligand has been identified; however, numerous
endogenous chemicals have been identified that bind to AHR and/or activate AHR-dependent
gene expression (Denison and Nagy, 2003). Bilirubin, tryptophan metabolites, arachidonic acid
metabolite lipoxin A4, several prostaglandins, retinoids, indigo and indirubin are some examples
of endogenous compounds known to activate AHR (Adachi, et al., 2001; Bittinger, et al., 2003;
Gambone, et al., 2002; McMillan and Bradfield, 2007; Phelan, et al., 1998; Schaldach, et al.,
1999; Seidel, et al., 2001; Sugihara, et al., 2004). FICZ (6-formylindolo[3,2-b]carbazole), a
tryptophan photoproduct has been reported to be the most potent and highest affinity candidate
endogenous AHR ligand known to date, capable of competitively displacing TCDD (Fritsche, et
al., 2007; Helferich and Denison, 1991; Rannug, et al., 1987; Wincent, et al., 2009). Femtomolar
levels of FICZ were reported to sufficiently induce AHR target mRNA and enzyme activity
(Wincent, et al., 2012). Unlike TCDD, however, FICZ is a substrate for Phase I enzymes
cytochrome P450 1A1, CYP1A2 and CYP1B1 and is readily metabolized (Wincent, et al.,
2009).
3.3 AHR-mediated toxicity Most of what is known about the toxicity of TCDD and related compounds come from studies in
experimental animals, but effects are also observed in humans (Birnbaum, 1994; Birnbaum and
Tuomisto, 2000; Pohjanvirta and Tuomisto, 1994; Sewall and Lucier, 1995; Tuomisto, 2005;
White and Birnbaum, 2009). One of the most dramatic effects from a single TCDD exposure in
experimental animals is the lethal wasting syndrome that is characterized by progressive weight
loss, hypophagia and gluconeogenic suppression (Pohjanvirta and Tuomisto, 1994; Seefeld, et
al., 1984; Seefeld and Peterson, 1984; Stahl, et al., 1993). While wasting accompanies death,
wasting per se is not likely the sole cause of death since maintenance of body weight by
parenteral nutrition does not prevent mortality (Gasiewicz, et al., 1980). Lethality of exposed
34
animals is delayed and does not ensue until 1-8 weeks after exposure with the time course
depending on the sensitivity of the animal (Birnbaum and Tuomisto, 2000; Poland and Knutson,
1982). A wide range of other short-term toxic effects following TCDD exposure are known,
including thymic atrophy, hepatic necrosis or hypertrophy, lipid accumulation within
hepatocytes, a variety of endocrine imbalances and immunosuppression (Birnbaum and
Tuomisto, 2000; Pohjanvirta and Tuomisto, 1994). Ahr-null mice challenged with TCDD are
resistant to TCDD-induced toxicities as well as to TCDD-induced transcriptional alterations
(Boutros, et al., 2009; Fernandez-Salguero, et al., 1995; Mimura, et al., 1997; Schmidt, et al.,
1996; Tijet, et al., 2006). Mice expressing a mutated form of AHR that is unable to translocate to
the nucleus or a DNA binding mutant are also essentially refractory to TCDD-mediated toxicity
(Bunger, et al., 2008; Bunger, et al., 2003). Furthermore, mice hypomorphic for ARNT are
phenotypically non-responsive to dioxins (Walisser, et al., 2004). Collectively, these genetic
manipulations indicate that TCDD toxicities are dependent on the transcriptional regulatory
function and DNA-binding capacity of AHR.
3.4 AHR transactivation
In its latent state, AHR is found in the cytosol where it is stably associated with a multi-protein
complex consisting of two molecules of chaperone protein HSP90, hepatitis B virus X-associated
protein 2 (XAP2) and a 23 kDa co-chaperone protein referred to as p23 (Figure 7)(Carver, et al.,
1994; Kazlauskas, et al., 1999; Ma and Whitlock, 1997; Meyer, et al., 1998; Perdew, 1988;
Pollenz, et al., 1994). HSP90 interacts with AHR via both the bHLH motif and the PAS B
domain and is an essential component of the unliganded AHR complex as its presence is
necessary for the high affinity ligand binding conformation of AHR and retention of AHR in the
cytosol by masking its NLS sequence (Antonsson, et al., 1995a; Antonsson, et al., 1995b;
Dolwick, et al., 1993; Pongratz, et al., 1992; Whitelaw, et al., 1993; Whitelaw, et al., 1995).
XAP2 has a multifunctional role which includes maintaining proper cytosolic folding of AHR,
improving chaperone complex stability, subcellular localization and ligand binding ability of
AHR and repressing AHR transcriptional activity (Kazlauskas, et al., 2000; LaPres, et al., 2000;
Meyer and Perdew, 1999; Meyer, et al., 1998; Petrulis, et al., 2000). Co-chaperone p23 stabilizes
the chaperone complex and is important for nuclear import of AHR (Kazlauskas, et al., 1999).
35
Most AHR ligands are highly lipophilic and therefore can enter the cell through simple diffusion
(Delescluse, et al., 2000; Goryo, et al., 2007). Ligand binding induces a conformational change
in AHR that exposes its NLS (Henry and Gasiewicz, 2003; Ikuta, et al., 2004; Whitlock, 1999).
Liganded AHR translocates into the nucleus where it dissociates from its chaperone protein
complex and then heterodimerizes with ARNT and is converted to its high affinity DNA
conformation (Hankinson, 2005; Okey, et al., 1980; Tomita, et al., 2000). The AHR/ARNT
heterodimer recognizes a DNA element designated aryl hydrocarbon receptor response element
(AHRE) located within the regulatory regions of target genes such as cytochrome P450 1A1
(CYP1A1) (Mimura and Fujii-Kuriyama, 2003; Whitlock, 1999). The AHRE is composed of two
half-sites, TnGC and GTG recognized by AHR and ARNT, respectively (Denison, et al., 1988;
Swanson, et al., 1995). Once bound to an AHRE the AHR/ARNT complex interacts with
multiple co-regulator proteins to regulate transcription (Hankinson, 2005). A list of co-regulator
proteins known to interact and/or modulate AHR/ARNT activity is provided in Table 3.
Table 3. Partial list of proteins and co-regulator proteins that interact with AHR/ARNT to
modulate transcription.
Factors Response Co-activators NCoA1/NCoA2/NCoA3/p300/CBP, interact with AHR and/or ARNT to facilitate gene NCoA6 activation (Hankinson, 2005; Harper, et al., 2006) RIP140/NRIP enhances AHRE-driven transcription (Hankinson,
2005; Harper, et al., 2006) Brg-1/Brm1 enhances transcription mediated by AHR/ARNT
(Hankinson, 2005) Co-repressors SMRT, SHP inhibits transcriptional activity of AHR/ARNT
complex (Jepsen and Rosenfeld, 2002; Nguyen, et al., 1999)
General Transcription Factors Pol II, TBP, TFIIB, TFIIF key components of the general transcription
machinery that are directly involved in gene activation by AHR/ARNT (Hankinson, 2005; Harper, et al., 2006)
For a complete list of AHR co-regulators see reference (Hankinson, 2005).
36
Figure 7. General mechanism of AHR transactivation.
Ligands bind (TCDD) to AHR in the cytosol and triggers translocation into the nucleus. AHR is
released from its chaperone complex and then binds with its dimerization partner ARNT. The
AHR/ARNT complex binds AHREs located within the regulatory region of target genes. Once
bound to the AHRE, AHR/ARNT associates with co-activators to activate transcription.
3.5 Negative regulation of AHR transactivation There are several mechanisms by which AHR transactivation can be attenuated (Figure 8). One
mechanism involves ligand-dependent degradation of AHR protein through a proteasomal
pathway (Pollenz, 2002; Wentworth, et al., 2004). The decrease in AHR protein leads to reduced
AHR-dependent transcription (Pollenz, et al., 1998). Other proposed mechanisms involve an
autoregulatory feedback loop created by induction of AHR-dependent transcriptional repressor.
The aryl hydrocarbon receptor repressor (AHRR) is induced by activated-AHR and is a proposed
transcriptional repressor of AHR (Mimura, et al., 1999). Another and more hypothetical
mechanism combines ideas from the two previous mechanisms where a yet to be identified
AHR-dependent labile factor promotes the proteolytic degradation of AHR protein and the
attenuation of transcription (Ma, 2002; Ma and Baldwin, 2002).
37
Figure 8. Three proposed mechanisms of negative regulation of AHR transactivation.
(A) Targeted proteolytic degradation of AHR protein through the ubiquitin 26S proteasome
pathway. (B) Induction of the transcriptional repressor AHRR. (C) Induction of a labile factor
that promotes the proteolytic degradation of AHR.
3.5.1 Proteolytic Degradation of AHR
Numerous studies have established AHR ligand binding leads to rapid and sustained reduction in
the level of endogenous AHR protein in both in vitro and in vivo models (Pollenz, 2002). The
ligand-induced degradation of AHR has been observed in many cell lines of different tissue
lineages, although the time-course and range of maximal degradation varied amongst cell lines
(Pollenz and Buggy, 2006). AHR degradation has also been observed in vivo. For example, AHR
protein concentration was reduced in liver, spleen, thymus and lung from Sprague-Dawley rats
given an acute dose of TCDD (Pollenz, et al., 1998). The ubiquitin-proteasome pathway has
been implicated in the ligand-induced degradation of AHR due the ability of the proteasome
inhibitors, MG-132 and lactacystin, two specific inhibitors of the 26S proteasome, to completely
block degradation (Davarinos and Pollenz, 1999; Ma and Baldwin, 2000). Inhibitors of calpains,
38
serine/cysteine, lysosomal and serine proteases failed to block AHR degradation (Davarinos and
Pollenz, 1999; Ma and Baldwin, 2000; Roberts and Whitelaw, 1999; Wormke, et al., 2000). In
addition, AHR degradation is inhibited in cells that contain a temperature-sensitive mutation in
the E1 ubiquitin conjugating enzyme (Ma and Baldwin, 2000). The inhibition of AHR
degradation resulted in sustained levels of AHR in the nucleus over time and significantly
increased in the concentration of DNA-bound AHR/ARNT heterodimers (Ma and Baldwin,
2000; Roberts and Whitelaw, 1999). These studies also demonstrated that AHR degradation
required the C-terminal transactivation domain and DNA-binding activity, as AHR lacking the
C-terminus and a DNA-binding mutant were more resistant to degradation than wildtype AHR
(Ma and Baldwin, 2000). TCDD induced the accumulation of high molecular weight AHR that
contained ubiquitin; however, specific ubiquitination sites are yet to be identified (Ma and
Baldwin, 2000). Taken together, these studies provide compelling evidence that AHR is
degraded via the 26S proteasome.
The ligand-induced degradation of AHR was connected to down-regulation of AHR target gene
expression by analyzing the duration and magnitude of gene expression following proteasomal
inhibition. AHR-regulated reporter gene experiments demonstrated pretreatment of cells with
MG-132 resulted in significantly greater reporter gene activity following TCDD treatment
compared to cells treated with TCDD alone (Davarinos and Pollenz, 1999). Inhibition of AHR
degradation also resulted in higher endogenous AHR target mRNA induction compared to non-
inhibited cells when exposed to TCDD (Ma and Baldwin, 2000). Reporter genes were also used
to assess whether down-regulation of endogenous AHR rendered cells non-responsive. Cell lines
propagated in the presence of TCDD for 12 days and then re-challenged with TCDD
demonstrated reductions in AHR protein and the absolute level of reporter gene activity (Pollenz,
et al., 1998). Collectively, these results correlate ligand-induced gene expression increases with
AHR degradation inhibition.
3.5.2 Aryl hydrocarbon receptor repressor (AHRR)
The aryl hydrocarbon receptor repressor (AHRR) is a bHLH-PAS protein that was discovered
because of its similarity to AHR (Mimura, et al., 1999). The AHRR shares high amino acid
identity with the AHR in the N-terminal region containing the bHLH (81%) and PAS A domain
(60%) but diverges significantly towards the C-terminus and does not contain a ligand binding
39
(PAS B) and transactivation domain (Figure 5C) (Mimura, et al., 1999). Due to the lack of a
ligand binding domain, the AHRR does not bind AHR ligands and appears to function in a
ligand-independent manner (Karchner, et al., 2009). Like AHR, the AHRR can heterodimerize
with ARNT and bind AHREs through its HLH-PAS A domains and basic region respectively
(Mimura, et al., 1999). Consistent with this, the initial mechanism of AHRR-mediated repression
was hypothesized to involve competition between AHR and the AHRR for ARNT
heterodimerization and competition between AHR/ARNT and AHRR/ARNT heterodimers for
AHRE binding (Mimura, et al., 1999). AHRR/ARNT binding to AHREs do not transcriptionally
activate due to the AHRR lacking a transactivation domain (Mimura, et al., 1999). Four putative
AHREs have been identified in the 5’ proximal to the promoter of the AHRR gene, and not
surprisingly AHR ligands have been reported to induce AHRR gene expression (Baba, et al.,
2001; Evans, et al., 2005; Karchner, et al., 2002; Mimura, et al., 1999; Tsuchiya, et al., 2003;
Yamamoto, et al., 2004). Induction of the AHRR by AHR agonists and AHRR-mediated
repression of AHR transactivation implicates that the AHRR forms a negative feedback loop
with AHR.
AHRR mRNA is constitutively expressed in a variety of tissues with the testis, lung, spleen,
heart and kidneys expressing the highest levels (Bernshausen, et al., 2006; Korkalainen, et al.,
2004; Nishihashi, et al., 2006; Tsuchiya, et al., 2003; Yamamoto, et al., 2004). In some tissues
that express high AHR levels such as the liver, constitutive AHRR mRNA is low and only
becomes measurable after induction with potent AHR ligands such as TCDD, B[a]P or 3MC
(Bernshausen, et al., 2006; Korkalainen, et al., 2005; Korkalainen, et al., 2004; Mimura, et al.,
1999). Conversely, constitutively high AHRR mRNA levels have been attributed to the lack of
AHR ligand inducibility of targets in primary cells and some cell lines (Gradin, et al., 1999;
Tsuchiya, et al., 2003). Reduction of AHRR expression by siRNA can in some cases restore
inducibility of AHR targets, but low constitutive expression of AHRR does not always correlate
with high AHR target inducibility (Haarmann-Stemmann, et al., 2007; Tsuchiya, et al., 2003;
Yamamoto, et al., 2004). For instance, constitutive AHRR mRNA is decreased in the heart and
brain tissue of Ahr-/- mice (Bernshausen, et al., 2006). Additionally, B[a]P induces significant
CYP1A1 mRNA in murine kidney tissue but does not induce detectable levels of AHRR mRNA
(Bernshausen, et al., 2006). The ability of AHR to transactivate the AHRR gene and the ability
of AHRR to repress AHR activity is likely to be tissue-, cell- or context-specific.
40
The AHRR has been demonstrated to be an effective transcriptional repressor of AHR activity;
however, its repressive activity on AHR transactivation has been largely based on in vitro
overexpression reporter gene experiments (Evans, et al., 2008; Evans, et al., 2005; Karchner, et
al., 2002; Karchner, et al., 2009; Mimura, et al., 1999; Oshima, et al., 2007). Overexpression
experiments using GFP-tagged AHRR were used to determine GFP-AHRR and presumably
endogenous AHRR localize primarily to the nucleus in MCF7, Hepa1c1c7 and COS7 cells
(Evans, et al., 2008; Kanno, et al., 2007; Mimura, et al., 1999). Using a series of GFP-tagged
AHRR truncations the AHRR was found to contain a functional NLS and NES and was shown to
undergo nucleocytoplasmic shuttling that favoured nuclear localization (Kanno, et al., 2007).
Overexpression experiments determined that the C-terminus of the AHRR was required for
repression, suggesting the presence of a ‘transrepression domain’ (Oshima, et al., 2007). These
observations support the hypothesis that AHRR is a nuclear transcriptional repressor. The exact
molecular mechanisms by which the AHRR represses AHR transactivation is currently subject to
debate. The initial mechanism proposed for AHRR-mediated repression involved competition
between the AHRR and AHR for heterodimerization with ARNT; however, subsequent studies
have reported that overexpression of ARNT had no effect on AHRR-mediated repression,
demonstrating that competition for ARNT is not the primary mechanism of repression (Evans, et
al., 2008; Karchner, et al., 2009). Moreover, a DNA-binding point mutant of the AHRR
effectively repressed AHR activity indicating that AHRE binding was not required for repression
(Evans, et al., 2008). Furthermore, interaction experiments demonstrated the AHRR interacted
with ARNT and AHR in a ligand-independent manner (Karchner, et al., 2009). These studies
illustrate the complexity of AHRR-mediated repression and the mechanism by which the AHRR
represses requires further clarification.
3.5.3 Induction of a labile AHR degradation promoting factor (ADPF)
A third proposed mechanism by which AHR transactivation can be terminated is through a
hypothetical labile factor that promotes the proteasomal degradation of AHR, termed the AHR
degradation promoting factor (ADPF) (Ma, 2007; Ma and Baldwin, 2002; Ma, et al., 2000). The
existence of a potential labile factor was first proposed based on the observation that protein
synthesis inhibition of liver cells increased the AHR ligand-induced target gene expression well
above levels of AHR ligand treatment alone (Whitlock and Gelboin, 1973). Cycloheximide, a
potent protein synthesis inhibitor, enhanced the TCDD-induced CYP1A1 mRNA to levels
41
greater than the sum of each compound alone, a phenomenon termed superinduction (Ma and
Baldwin, 2002; Ma, et al., 2000). Superinduction results from increased transcriptional rate and
subsequent generation of AHR target mRNA and protein but not from increased mRNA or
protein stability (Israel, et al., 1985; Israel and Whitlock, 1983). Superinduction requires
substantial (>90%) protein synthesis inhibition and functional AHR and ARNT suggesting that
the synthesis of a repressor protein, which is labile and induced during AHR target induction is
inhibited (Lusska, et al., 1992). CHX co-treatment was later found to fully inhibit the
proteasomal degradation of AHR that correlated with superinduction in a time- and dose-
dependent manner (Ma, et al., 2000). Similarly, proteasomal inhibition with MG-132 also
superinduced AHR targets following AHR agonist treatment further supporting the notion that
increasing the stability of AHR by decreasing its degradation superinduced target gene
expression (Ma and Baldwin, 2000; Ma, et al., 2000). Together these observations establish the
hypothesis that a repressor induced by agonist-activated AHR and sensitive to CHX mediates the
proteasomal degradation of AHR, this unknown factor is referred to as the ADPF (Ma, et al.,
2000). Two possible mechanisms, which are not mutually exclusive, have been proposed for the
ADPF based on results from inhibitor studies. One is an autoregulatory mechanism in which the
ADPF is an AHR target protein that is induced by AHR agonists (Ma and Baldwin, 2002).
Blockage of the induction of the ADPF by protein synthesis disrupts the autoregulation (Ma and
Baldwin, 2002). The second is the ADPF is a labile factor that requires constant protein synthesis
for function and therefore sensitive to cycloheximide (Ma and Baldwin, 2002). Despite the
evidence supporting its existence, an ADPF has not yet been identified.
3.5.4 In vivo models used to study negative regulation of AHR
3.5.4.1 Ahrr-deficient mice
Ahrr-deficient mice were generated to study the function of the AHRR in AHR transactivation in
vivo (Hosoya, et al., 2008). The AHR-dependent induction of CYP1A1 was expected to be
greater in Ahrr-deficient mice than wildtype mice following 3MC treatment; however, this
pattern of induction was highly tissue-specific and limited to only the heart, spleen and skin
(Hosoya, et al., 2008). Tissues with high AHR expression including the liver, lung, kidney and
thymus did not display significant increases in 3MC-treated Ahrr-deficient mice compared with
treated wildtype mice nor did the brain or stomach (Hosoya, et al., 2008). Constitutive
expression of CYP1A1 in Ahrr-deficient mice was not significantly different than wildtype mice
42
for all the tissues examined (Hosoya, et al., 2008). Ahrr-deficient mice treated with subcutaneous
B[a]P showed a significant delay in skin tumour formation than wildtype (Hosoya, et al., 2008).
It was suggested that the increased CYP1A1 expression in the skin shifted the balance of
metabolic activity of CYP1A1 in favour of detoxification and this attributed to the delay in skin
carcinogenesis (Hosoya, et al., 2008). B[a]P-treated Ahrr-deficient mice demonstrated a slightly
lower mortality rate trend compared with treated-wildtype mice (Hosoya, et al., 2008).
3.5.4.2 Constitutively active AHR mice
Deletion of the entire PAS B domain of AHR results in a protein that does not interact with
HSP90, shows constitutive nuclear localization, binds ARNT and exhibits potent transactivation
in the absence of ligand i.e. TCDD, a constitutively active AHR (CA-AHR) (McGuire, et al.,
2001). Transgenic mice expressing CA-AHR were generated and used to study the in vivo
consequences of lack of AHR transactivation termination (Andersson, et al., 2002; Andersson, et
al., 2003; Brunnberg, et al., 2006; Brunnberg, et al., 2011; McGuire, et al., 2001; Moennikes, et
al., 2004; Wejheden, et al., 2010). Transgenic CA-AHR animals expressed CA-AHR in
lymphatic organs including thymus, spleen, lymph nodes, enriched T and B cells and bone
marrow as well as in lung, liver, muscle, skin, brain, heart, kidney and throughout the
gastrointestinal tract (Andersson, et al., 2002). All tissues that showed CA-AHR expression also
demonstrated induced CYP1A1 mRNA expression at varying levels (Andersson, et al., 2002).
Additionally, homozygous CA-AHR mice demonstrated significantly greater CYP1A1, CYP1A2
and AHRR mRNA expression in the liver compared with wildtype animals (Andersson, et al.,
2002). CYP1A1 mRNA induction in homozygous CA-AHR mice was similar to that of low to
mid-dose TCDD-treated wildtype mice in the liver, thymus and stomach (Andersson, et al.,
2002). CA-AHR mice also showed several symptoms that are similar to TCDD toxicity such as
thymic atrophy and liver enlargement (Andersson, et al., 2002). CA-AHR mice showed a
significantly reduced lifespan where mice began dying as early as 6 months and often without
symptoms (other than body weight loss a day before death), only a few animals survived after 12
months (Andersson, et al., 2002). Another distinguishing feature of CA-AHR mice was the
development of invasive tumours of the glandular stomach at an early age that correlated with
increased mortality (Andersson, et al., 2002). Other findings in these CA-AHR mice included:
decreased population of B lymphocytes in the peritoneal cavity, enlarged heart and kidneys and
43
increased hepatocarcinogenesis (Andersson, et al., 2003; Brunnberg, et al., 2006; Brunnberg, et
al., 2011; Moennikes, et al., 2004).
3.6 AHR target genes The up-regulation of a battery of six enzymes involved in biotransformation of xenobiotics and
endogenous compounds in response to TCDD and related AHR ligands has been well established
and studied in several species (Nebert, 1989). This battery includes cytochrome P450 1A1
(CYP1A1), CYP1A2, glutathione S-transferase Ya (GSTYa), aldehyde-3-dehydrogenase
(ALDH-3) and NAD(P)H: quinone oxidoreductase (NQO1) (Nebert, et al., 1990; Nebert, et al.,
2000; Schrenk, 1998). A third cytochrome P450 enzyme, CYP1B1 was later identified to be up-
regulated by AHR and added to the gene battery (Savas, et al., 1994; Shen, et al., 1993; Sutter, et
al., 1994). CYP1A1, CYP1A2 and CYP1B1 are NADPH-dependent monooxygenases that
mediate phase I metabolism rendering end-products more reactive for subsequent reactions. The
other four enzymes mediate phase II metabolism by conjugating charged groups onto xenobiotics
to facilitate their elimination. Many AHR ligands are metabolized by these enzymes and the idea
of induction of these enzymes as an adaptive response to AHR ligand exposure has been
proposed (Gu, et al., 2000; Nebert, et al., 2004; Okey, 1990). The induction of these enzymes,
however, cannot explain the plethora of toxic effects that are associated with TCDD exposure.
TCDD is poorly metabolized and thus causes chronic and sustained induction of cytochrome
P450 enzymes, leading to oxidative stress, formation of reactive oxygen species, lipid
peroxidation and DNA damage (Hassoun, et al., 2002; Hassoun, et al., 2001; Nebert, et al.,
2000; Shertzer, et al., 1998; Slezak, et al., 2000).
Several different methodological approaches have been used to discover new AHR target genes,
such as microarrays, differential display, representational difference analysis and serial analysis
of gene expression (Boverhof, et al., 2006; Dong, et al., 1997; Fletcher, et al., 2005; Frericks, et
al., 2006; Frueh, et al., 2001; Hayes, et al., 2007; Kurachi, et al., 2002; Ohbayashi, et al., 2001;
Oikawa, et al., 2002; Puga, et al., 2000; Svensson and Lundberg, 2001; Tijet, et al., 2006;
Zeytun, et al., 2002). These studies revealed that AHR ligands induce the expression of a number
of genes that do not encode metabolizing enzymes. Some of these genes have been shown to be
‘true’ AHR target genes, being transcriptionally regulated via a genomic AHR pathway
involving AHRE binding within the gene regulatory region while others are affected by a non-
44
AHRE binding or non-genomic pathway (Dere, et al., 2011; Lo, et al., 2011; Lo and Matthews,
2012). TiPARP is an example of a novel AHR target gene that encodes a non-xenobiotic
metabolizing enzyme and is regulated via the genomic AHR pathway (Hao, et al., 2012; Ma, et
al., 2001).
3.6.1 TCDD-inducible poly(ADP-ribose) polymerase (TiPARP): a novel AHR target
TCDD-inducible poly(ADP-ribose) polymerase (TiPARP) was first identified in mouse
hepatoma cells by differential display as a TCDD-induced mRNA (Ma, et al., 2001). The human
orthologue of the mouse Tiparp gene was later identified and contains six exons and human
TiPARP protein displayed 91.8% total amino acid identity with mouse TiPARP (Katoh and
Katoh, 2003). Induction of TiPARP gene expression by TCDD is mediated via the AHR (Ma,
2002; Ma, et al., 2001). TiPARP induction by TCDD demonstrated characteristics similar to
other AHR target genes such as superinduction in the presence of proteasomal and protein
synthesis inhibitors (Ma, 2002). Subsequent microarray studies have shown that TiPARP mRNA
is consistently up-regulated by TCDD both in vitro and in vivo (Dere, et al., 2006; Forgacs, et
al., 2013; Frericks, et al., 2006; Henry, et al., 2010; Lo, et al., 2011; Tijet, et al., 2006).
Additionally, TiPARP gene regulation by other AHR ligands both exogenous and naturally
occurring as well as AHR antagonists has been previously observed (de Waard, et al., 2008;
Hao, et al., 2012; Ito, et al., 2006; Pansoy, et al., 2010; Wahl, et al., 2008). Together these
studies support TiPARP as an AHR target gene.
3.6.1.1 AHR regulation of TiPARP expression
TiPARP induction by TCDD has been reported to be rapid reaching maximal levels after 1.5 h to
2 h of treatment leading to TiPARP sometimes being referred to as an immediate early gene
(Diani-Moore, et al., 2010; Hao, et al., 2012; Ma, 2002; Ma, et al., 2001). Currently, the
regulation of TiPARP by activated-AHR is not fully understood. Previous studies have reported
regions upstream of the TiPARP transcriptional start that are highly bound by ligand activated-
AHR suggesting the presence of an AHRE-containing regulatory region (Ahmed, et al., 2009;
Hao, et al., 2012; Lo, et al., 2011; Lo and Matthews, 2012; Pansoy, et al., 2010). There are a
total of 24 AHRE core elements (GCGTG) located 10 kb upstream of the mouse Tiparp
transcriptional start site (Hao, et al., 2012). Six of these AHREs are clustered together within a
45
45 bp fragment including four tandem repeats of AHRE core elements arranged in a concatemer
configuration (Hao, et al., 2012). This AHRE concatemer was bound by activated-AHR and
when expressed as a reporter gene displayed activity, although the level of ligand-induced
activity was not substantial due to high constitutive activity (Hao, et al., 2012).
3.6.1.2 TiPARP Structure
TiPARP encodes a protein of 657 amino acid residues that contains three predicted modular
domains: a C-terminal PARP catalytic domain, a central tryptophan-tryptophan-glutamate
(WWE) domain and a single CX8CX5CX3-type zinc-finger (Figure 9)(Schreiber, et al., 2006).
The PARP catalytic domain shares 43% sequence homology with the catalytic domains of
ARTDs (Ma, et al., 2001). The catalytic triad of the catalytic domain is H-Y-I and TiPARP is
predicted to possess mono-ADP-ribosyltransferase (MART) activity (Otto, et al., 2005). During
its initial characterization, TiPARP was reported to be a catalytically active enzyme; however, its
potential mono-ADP-ribosyltransferase activities were not evaluated and remain to be
determined (Ma, et al., 2001).
The WWE domain is composed of two conserved tryptophan residues and one glutamic acid
residue and is the putative protein-protein interaction motif of TiPARP that also occurs in classes
of proteins associated with ubiquitination (Aravind, 2001). Other ARTD family members
including ARTD8, ARTD11, ARTD12 and ARTD13 contain at least one WWE domain and it
was recently reported that the WWE domain of ARTD11 binds ADP-ribose (He, et al., 2012).
The role of the TiPARP WWE domain is currently unknown.
The CCCH-type zinc finger is a putative RNA binding motif characterized by three cysteines and
one histidine, which together coordinate a zinc ion to form a local peptide structure (Blackshear,
2002; Liang, et al., 2008). ARTD13 contains four CCCH-type zinc fingers in tandem and has
demonstrated to bind specific viral mRNA sequences (Gao, et al., 2002). CCCH-type containing
proteins are best known for their RNA binding ability but they also bind single-stranded DNA
(Bai and Tolias, 1996; Collart, et al., 2005). The functionality and nucleic acid binding potential
of CCCH-type zinc finger of TiPARP have not been determined.
46
Figure 9. Functional domain structure of TCDD-inducible poly(ADP-ribose) polymerase.
TiPARP/ARTD14/PARP7 contains a single CCCH-type zinc-finger, which is a putative RNA-
binding module, a centrally located WWE (tryptophan-tryptophan-glutamate) domain and a
carboxy-terminal PARP catalytic domain. The catalytic domain contains a ‘HYI’ catalytic triad,
which is found in mono-ADP-ribosyltransferases and not poly-ADP-ribose polymerases.
3.6.1.3 Potential role of TiPARP in TCDD-mediated toxicity
TiPARP has been implicated in the suppression of hepatic gluconeogenesis, a prominent feature
of the lethal wasting syndrome seen in experimental animals exposed to TCDD (Diani-Moore, et
al., 2010). Phosphoenolpyruvate carboxykinase (PEPCK) and glucose-6-phosphatase (G6Pase)
are two key gluconeogenic genes known to be down-regulated by TCDD (Fan and Rozman,
1994; Hsia and Kreamer, 1985; Stahl, et al., 1993; Weber, et al., 1991). Because TCDD
treatment of chick embryo livers reduced NAD+ levels, increased total cellular PARP activity
and TiPARP expression is induced by TCDD, it was hypothesized that TiPARP was responsible
for NAD+ and NADH depletion (Diani-Moore, et al., 2010). The depletion of NAD+ was
suggested to decrease the activity of SIRT1, which is dependent on NAD+. PEPCK and G6Pase
are transcriptionally regulated by peroxisome proliferator-activated receptor γ co-activator 1α
(PGC1α), which is activated by deacetylation by SIRT1 (Diani-Moore, et al., 2010). The
decreased SIRT1 activity would lead to decreased PGC1α activity and decrease PEPCK and
G6Pase expression levels. TiPARP overexpression resulted in decreased glucose output, NAD+
levels, PEPCK mRNA, PGC1α protein levels and increased acetylated PGC1α (Diani-Moore, et
al., 2010). Knockdown of TiPARP resulted in increased glucose output and NAD+ levels (Diani-
47
Moore, et al., 2010). It appeared that TiPARP mediated the TCDD-induced suppression of
gluconeogenesis and its overexpression mimicked the effects of TCDD in a chick embryo model.
Similarly, TiPARP has been recently suggested to be involved in heightened sensitivity to non-
alcoholic steatohepatitis (NASH) through depletion of NAD+ (He, et al., 2013). Activation of
AHR has been reported to lead to spontaneous fatty liver disease, lipid peroxidation and
oxidative stress, all of which are risk factors for NASH (Dalton, et al., 2002; Lee, et al., 2010;
Lu, et al., 2011). The increase in reactive oxygen species (ROS) production in mouse liver that is
associated with activated-AHR was proposed to be a result of lowered superoxide dismutase 2
(SOD2) activity and compromised clearance of ROS (He, et al., 2013). In another elaborated
model, TiPARP gene induction by TCDD was connected to a decrease in NAD+ levels by
catalytic consumption, this in turn deactivated mitochondrial SIRT3, which reduced SOD2
deacetylation decreasing its activity (He, et al., 2013). The decreased activity of SOD2 led to
increased ROS production and increased sensitivity to NASH (He, et al., 2013).
These two studies based their proposed models of TiPARP-mediated hepatic depletion of NAD+
to TiPARP gene induction by TCDD. Both studies failed to measure increased TiPARP protein
levels following TCDD treatment or increased catalytic activity of TiPARP, making models
highly presumptuous. Additionally, ARTD1 when activated accounts for 90% cellular
poly(ADP-ribose) formation leading one to question whether TiPARP, a predicted MART could
be solely responsible for NAD+ depletion via catalytic consumption (Schreiber, et al., 2002).
48
Chapter 2: Aims of the present study The ARTD superfamily consists of eighteen members and their potential activities in diverse
cellular processes are just now being explored. TiPARP, the seventh ARTD family member to be
identified, has received little attention since its discovery over ten years ago. TiPARP is better
known as an AHR target gene that is induced by a vast array of AHR ligands in both in vivo and
in vitro models. Despite being an apparently important AHR target gene, the potential role of
TiPARP in AHR-mediated transcriptional regulation is not known. Moreover, the predicted
functional domains of TiPARP also give little insight into a potential role in AHR transcriptional
regulation. The TiPARP catalytic triad motif predicts that TiPARP is a class 2 ARTD with
mono-ADP-ribosyltransferase activity rather than a class 1 poly-ADP-ribosylating ARTD;
however, this has not been experimentally confirmed. Because TiPARP is an important AHR
target and a potential mono-ADP-ribosyltransferase, we were interested in whether TiPARP and
mono-ADP-ribosylation were involved in the AHR-mediated transcription.
The repression of AHR-mediated transcription is an aspect that is less understood than
transactivation. Three mechanisms of negative regulation have been proposed: the targeted
proteasomal degradation of activated-AHR via the 26S proteasome, the AHRR and an
autoregulatory feedback loop involving an unidentified labile factor that promotes the proteolytic
degradation of AHR. Because TiPARP expression is induced by activated-AHR we examined its
potential as negative regulator of AHR transactivation.
The overall objective of this thesis research was to understand TiPARP and its potential role in
AHR transactivation. Specific aims were:
• Demonstrate TiPARP is a mono-ADP-ribosyltransferase
• Determine role of TiPARP in AHR transactivation and understand its mechanism
• Compare TiPARP- and aryl hydrocarbon receptor repressor (AHRR)-mediated regulation
of AHR transactivation
49
Chapter 3: Materials and Methods 4 Materials
4.1 Chemicals and biological reagents 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) was purchased from Wellington Laboratories
(Guelph, ON, Canada). Dimethyl sulfoxide (DMSO) was purchased from Sigma-Aldrich (St.
Louis, MO). Cell culture reagents: Dulbecco’s Modified Eagle’s medium (DMEM)
supplemented with 1 g/L, high glucose (4.5 g/L glucose) DMEM, 1:1 mixture of DMEM and
Ham’s F-12 nutrient mixture (DMEM/F-12), RPMI-1640 medium, fetal bovine serum (FBS),
penicillin/streptomycin antibiotic mixture, 0.25% Trypsin-EDTA, Phosphate Buffered Saline,
were all purchased from Sigma-Aldrich (St. Louis, MO). Protein A-Agarose Fast Flow 50%
(v/v) was purchased from Sigma-Aldrich. Lipofectamine LTX with Plus reagent was purchased
from Life Technologies (Burlington, ON) and FuGENE HD tranfection reagent was purchased
from Roche (Laval, QC).
Antibodies used for chromatin immunoprecipitation (ChIP), co-immunoprecipitation (IP),
indirect immunofluorescence (IF) or Western blot (WB) experiments were: anti-AHR (H-211,
ChIP, IF, WB), anti-AHR (N-19, WB), anti-ARNT (H-172, ChIP), anti-ARNT (C-19, WB), anti-
rabbit immunoglobin (IgG, sc-2027, ChIP, IP) (all from Santa Cruz Biotechnology, Santa Cruz,
CA), anti-PARP7 (Abcam, 84664, Cambridge, MA, IF), anti-AHRR (HPA019614, ChIP, WB)
and anti-β-actin (A2228, WB) both from Sigma-Aldrich, anti-AHR (SA-210, Biomol
International Inc., Plymouth Meeting, PA), anti-GFP (632460, ChIP) and anti-GFP (JL-8, WB)
both from Clontech (Mountain View, CA). Conjugated secondary antibodies used were:
Horseradish Peroxidase (HRP)-conjugated anti-rabbit IgG whole antibody (NA934, GE
Healthcare; Buckinghamshire, UK), HRP-conjugated normal anti-mouse IgG (sc2954) and HRP-
conjugated normal anti-goat IgG (sc2741) all from Santa Cruz, and HRP-conjugated Clean Blot
IP Reagent (ThermoScientific) and Alexa 568 Fluor-conjugated anti-rabbit IgG (Life
Technologies, Burlington, ON, Canada). ECL Advance Western blotting detection system and
ECL Plus both from GE Healthcare and SuperSignal West Dura (ThermoScientific) were used to
visualize proteins for both Western blots and Co-IP experiments. For purification of PCR
products Buffer PB (Qiagen, Toronto, ON) and EZ-10 Spin Column PCR Purification Kit (Bio
50
Basic Inc. Markham, ON) were used. For RNA extraction and cDNA synthesis, Aurum™ Total
RNA Mini kit (Bio-Rad, Hercules, CA) and Superscript III reverse transcriptase (Life
Technologies) were used. Pfu Ultra DNA polymerase (Agilent, Mississauga, ON) and Pfx50
DNA polymerase (Life Technologies) were used for site-directed mutagenesis. High Fidelity
PCR enzyme mix (ThermoScientific) was used for amplification of PCR products. QIAquick Gel
Extraction Kit and HiSpeed Plasmid Maxi Kit were purchased from Qiagen. SsoFast EvaGreen®
Supermix (Bio-Rad) was used for all quantitative real-time PCR reactions. ONE-glo™
Luciferase Assay System, TNT® Coupled Reticulocyte Lysate System and pGEM®-T Easy
Vector System were purchased from Promega (Madison, WI). 35S-radiolabelled L-methionine
and 32P-radiolabelled β-NAD+ were purchased from Perkin-Elmer (Woodbridge, ON). All
primers used for real-time PCR and for the generation of expression constructs were synthesized
by Integrated DNA Technologies (IDT, Coralville, IA).
4.2 Plasticware Plasticware used for cell culture was purchased from Sarstedt (Newton, CT) which include: T-
25, T-75, and T-175 tissue culture flasks, 10 cm tissue culture dishes, 6-well tissue culture plates,
and 2 ml cryovials. The 12-well and 96-well clear flat bottom plates were purchased from BD
Biosciences-Falcon (San Jose, CA). The 96-well black flat bottom plates used for luciferase
assays were purchased from Corning (Lowell, MA). The 1.5 ml (Axygen) and 1.7 ml low-
binding (Progene) microcentrifuge tubes were purchased from Ultident Scientific (St. Laurent,
QC). 96-well non-skirted PCR plates were purchased from D-Mark Biosciences (Toronto, ON).
4.3 Instruments For cell culture, HERAcell® 150 incubators (Kendro, Langenselbold, Germany) were used. The
water bath (model # 1228, Sheldon manufacturing, Cornelius, OR) was purchased from VWR
International (Plainfield, NJ). Cells were centrifuged with Centrifuge 5702 (Eppendorf,
Hamburg, Germany). All centrifuge steps in PCR Purification and RNA extraction processes
used Centrifuge 5415D (Eppendorf). Locator JR Cryo Biological Storage System (Thermolyne)
was used to freeze, and store human immortalized cells was purchased from VWR International
(Plainfeld, NJ). To count cells, a Bright-Line Hemocytometer (Hausser Scientific, Horsham, PA)
and Vistavision Light Microscope (VWR) were used. The purity and concentration of RNA and
protein were measured with an Ultrospec 2100 pro Spectrophotometer (Biochrom, Cambridge,
51
England). Branson Digital Sonifier 450 (Danbury, CT) was used to sonicate cells. The rotator
used for the ChIP assay and Western blotting was from Mini LabRoller (Labnet Inc., Edison,
NJ). Real-time PCR amplification of ChIP DNA and cDNA was performed on Chromo4 Real-
Time PCR detector (Bio-Rad). A DNAEngine Peltier Thermal Cycler (Bio-Rad) was used for
site-directed mutagenesis and cDNA synthesis of extracted RNA. Protein and β-galactosidase
levels were determined using a ThermoScientific 96 well Multiskan EX Photometer (Waltham,
MA). Luciferase levels from reporter gene constructs were measured using GLO-max 96-well
microplate luminometer (Promega). Indirect immunofluorescence images were viewed with
Imager.Z1 epifluorescence microscope and Axiovision software (Zeiss, Toronto, ON) or an
Olympus Fluoview 1000 confocal microscope (Center Valley, PA). Mouse livers were
homogenized on a VWR PowerMax Advanced Homogenizing System 200 (VWR).
5 Methods
5.1 Maintenance of HuH7, COS7, TREx-FLP-IN 293, T47D, NCI-N87 and MCF7 cell lines
HuH7 human hepatoma, T47D human breast carcinoma, MCF7 human breast carcinoma and
NCI-N87 human gastric carcinoma cell lines were purchased from the American Type Culture
Collection (ATCC, Manassas, VA). TREx-FLP-IN 293 human embryonic kidney cells were
purchased from Life Technologies. COS7 African Green Monkey kidney cells were a generous
gift from Dr. Stephane Angers (Faculty of Pharmacy, University of Toronto). All cell lines were
cryopreserved and stored in liquid nitrogen. All cells were ~80% confluent before freezing, cells
were trypsinized and neutralized with fresh medium and centrifuged for 2 min at 4000 rpm. The
cell pellet was re-suspended in a 90% FBS containing 10% DMSO (v/v) solution and aliquoted
into labelled cryovials. Cryovials were immediately frozen at -80ºC and then transferred to liquid
nitrogen for long-term storage. Before use, HuH7, COS7 and TREx-FLN cells were rapidly
thawed at 37°C and transferred to a T-75 tissue culture flask containing high glucose DMEM
supplemented with 10% (v/v) FBS and 1% (v/v) penicillin/streptomycin (PEST). Once cell lines
were at least 80% confluent they were sub-cultured 1:6 to 1:12 twice a week. T47D cells were
thawed and transferred to a T-25 tissue culture flask containing DMEM/F12 medium
supplemented with 10% FBS and 1% PEST. Once cells were at least 80% confluent, cells were
sub-cultured 1:2 to 1:3 twice a week. NCI-N87 cells were thawed and transferred to T-25 tissue
52
culture flask containing RPMI-1640 supplemented with 10% FBS and 1% PEST. MCF7 cells
were thawed and transferred to T-25 tissue culture flask containing low glucose DMEM
containing 10% FBS and 1% PEST. Once cells were at least 80% confluent they were sub-
cultured 1:2 to 1:3 twice a week. All cell lines were sub-cultured as follows: medium was
aspirated from culture flask and cells were washed once with 5-10 ml of sterile PBS. PBS was
aspirated and 2-4 ml of pre-warmed trypsin-EDTA was added to the flask. The flask was placed
back into incubator for 1-5 min until cells detached completely. An equal volume of pre-warmed
completed medium was added to cell suspension in flask to neutralize the activity of tryspin. Cell
suspension was vigorously pipetted up and down until homogenous then 1-4 ml of the cell
solution was transferred to a T-175 flask containing 40 ml of completed media. All cell lines
were maintained at 37°C and 5% CO2.
5.2 Generation of Expression Constructs All full-length human TiPARP expression constructs, TiPARP N-terminal truncation expression
constructs and the C-terminal truncation 1-234 were created by PCR amplification of TiPARP
cDNA from pCMV6-XL4-hTiPARP (Origene, Rockwood, MD). Two-hundred micrograms of
pCMV6-XL4-hTiPARP was added to a reaction mixture containing 0.2 μM of desired forward
primer containing Kozak sequence and/or EcoRI restriction site and reverse primer containing
SalI restriction cut site (Table 4), 0.2 mM dNTPs and High Fidelity PCR enzyme mix
(ThermoScientific) and amplified using the DNAEngine Peltier Thermal Cycler (Bio-Rad). PCR
products were loaded on a 1% agarose gel and separated by gel electrophoresis. PCR products
were excised from gel and purified using QIAquick Gel Extraction Kit following manufacturer’s
instructions. One hundred nanograms of purified PCR product was ligated into 50 ng pGEM-T
Easy vector with T4 DNA ligase (New England Biolabs, Whitby, ON) in a 10 μl reaction
overnight at room temperature. Ligation reactions were transformed into 100 μl chemically
competent DH5α strain E. coli. Bacteria were plated on LB-ampicillin agar plates containing 40
μg/ml X-gal and incubated overnight at 37°C. Individual white colonies were selected and grown
in 3 ml LB-ampicillin overnight at 37°C with constant shaking. The following morning, plasmid
DNA from selected colonies was purified using GenElute Plasmid Miniprep kit (Sigma)
according to the manufacturer’s instructions. Purified plasmids were digested with EcoRI and
SalI restriction enzymes for 1 h and digestions were loaded on a 1% agarose gel and separated by
53
gel electrophoresis. Inserts were excised from gel and purified using QIAquick Gel Extraction
Kit (Qiagen). Inserts were ligated into pcDNA3.1 (Life Technologies) using EcoRI and XhoI
restriction sites, pEGFP-C2 (Clontech) using EcoRI and SalI restriction sites and pGEX-4T1
(GE Healthcare) using EcoRI and XhoI restriction sites overnight at room temperature. The
following day ligations were transformed into chemically competent DH5α strain E. coli and
grown on LB-ampicillin plates for pcDNA3.1 and pGEX-4T1 ligations or LB-kanamycin plates
for pEGFP-C2 ligations overnight at 37°C. The following day individual colonies were selected
and grown in LB-ampicillin or LB-kanamycin overnight at 37°C with constant shaking. Plasmids
were purified using GenElute Plasmid Miniprep kit (Sigma) according to the manufacturer’s
instructions. pcDNA3.1 and pGEX-4T1 constructs were screened by restriction digests using
EcoRI and XhoI restriction enzymes and pEGFP-C2 constructs were digested using EcoRI and
SalI. Digests were loaded onto a 1% agarose gel and separated by gel electrophoresis. All
plasmids that screened positive were sent to The Centre for Applied Genomics DNA Sequencing
and Synthesis Facility (Medical and Related Sciences (MaRS) building, Toronto, ON, Canada)
to verify sequence and framing. Mouse Tiparp expression constructs were generated by PCR
amplification of mTiPARP cDNA from pCMV6-kan/neo-mTiPARP (Origene) using forward
primer containing EcoRI restriction site and reverse primer containing XhoI restriction site
(Table 4). Chicken TiPARP expression vectors were generated by PCR amplification of chicken
TiPARP cDNA from total Gallus gallus liver RNA (Zyagen, San Diego, CA) using forward
primer containing EcoRI restriction site and reverse primer containing SalI site (Table 4). Both
chicken and mouse TiPARP inserts were cloned into EcoRI and XhoI sites of pcDNA3.1 and
pGEX-4TI and EcoRI and SalI sites of pEGFP-C2.
pEGFP-C2-AHRR-FL, pEGFP-C2-AHRRΔ8 (delta exon 8), pAcGFP-N1-AHRR-FL and
pAcGFP-N1- AHRRΔ8 were generated by sub-cloning from pcDNA3.1-AHRR-FL and pcDNA-
AHRRΔ8, (both gracious gifts from Dr. Mark Hahn, Woods Hole Oceanographic Institute,
Woods Hole, MA). PCR products were amplified using forward primer containing EcoRI
restriction site and reverse primer containing stop codon and XhoI restriction site or without stop
codon and BamHI restriction site (Table 4). Both AHRR inserts were cloned into EcoRI and SalI
sites of pEGFP-C2 or EcoRI and BamHI sites of pAcGFP-N1 (Clontech).
54
Table 4. PCR primers used for generation of TiPARP and AHRR inserts.
TiPARP EcoRI 5’ CAAA GAATTC ATGGAAATGGAAACCACCGAACC
TiPARP SalI 3’ CAAA GTCGAC TCAAATGGAAACAGTGTTACTGAC EcoRI TiPARP 33 5’ CAAA GAATTC ATCACTCCATTGAAGACTTGTTTTAAG EcoRI Kozak TiPARP 33 3’ CAAA GAATTC ACCATG
ATCACTCCATTGAAGACTTGTTTTAAG EcoRI TiPARP 53 5’ CAAA GAATTC CTGAGGTCTTTGAGGCCAATATT EcoRI Kozak TiPARP 53 5’ CAAA GAATTC ACCATG CTGAGGTCTTTGAGGCCAATATT EcoRI TiPARP 103 5’ CAAA GAATTC GCAGAAAATAATATGTCTGTTCTGAT EcoRI Kozak TiPARP 103 5’ CAAA GAATTC ACCATG
GCAGAAAATAATATGTCTGTTCTGAT EcoRI TiPARP 200 5’ CAAA GAATTC CTGGACACCAGTGATAAGCTGAGT EcoRI Kozak TiPARP 200 5’ CAAA GAATTC ACCATG
CTGGACACCAGTGATAAGCTGAGT EcoRI TiPARP 218 5’ CAAA GAATTC GCTTCCCTTGACCTCGTGTTTGA EcoRI Kozak TiPARP 218 5’ CAAA GAATTC ACCATG GCTTCCCTTGACCTCGTGTTTGA EcoRI TiPARP 225 5’ CAAA GAATTC GAGCTGGTGAACCAGTTGCAGTAC EcoRI Kozak TiPARP 225 5’ CAAA GAATTC ACCATG
GAGCTGGTGAACCAGTTGCAGTAC EcoRI TiPARP 245 5’ CAAA GAATTC GACTTTCTGCAAGGCACTTGTATTTA EcoRI Kozak TiPARP 245 5’ CAAA GAATTC ACCATG
GACTTTCTGCAAGGCACTTGTATTTA EcoRI TiPARP 275 5’ CAAA GAATTC ACTCAAAAGTGGCAGAGTGTATTC EcoRI Kozak TiPARP 275 5’ CAAA GAATTC ACCATG
ACTCAAAAGTGGCAGAGTGTATTC EcoRI TiPARP 328 5’ CAAA GAATTC CTACGAAGGCTGTCCACACCAC EcoRI Kozak TiPARP 328 5’ CAAA GAATTC ACCATG CTACGAAGGCTGTCCACACCAC EcoRI TiPARP 445 5’ CAAA GAATTC TCAGCAAACTTTTACCCTGAAACTT EcoRI Kozak TiPARP 445 5’ CAAA GAATTC ACCATG
TCAGCAAACTTTTACCCTGAAACTT SalI TiPARP 234 3` CAAA GTCGAC TCAAGTGTGGTACTGCAACTGGTTCAC EcoRI mouse TiPARP 5’ CAAA GAATTC ATGGAAGTGGAAACCACTGAACC XhoI mouse TiPARP 3’ CAAA CTCGAG TCAAATGGAAACAGTGTTACTGACT EcoRI chicken TiPARP 5’ CAAA GAATTC ATGGACACCGAAGCAAAGCCA SalI chicken TiPARP 3’ CAAA GTCGAC TCAAATGGAGACAGTGTTGCTA EcoRI hAHRR 5’ CAAA GAATTC ATGCCGAGGACGATGATCCCGCC XhoI hAHRR stop 3’ CAAA CTCGAG CTATGGCAGGAATGTGCACCCC BamHI AHRR no stop 3’ CAAA GGATCC CG TGGCAGGAATGTGCACCCCGAA
5.3 Site-directed mutagenesis Catalytic, zinc finger, nuclear localization point mutants, double alanine point mutants and C-
terminal 1-448 truncation were generated by site-directed mutagenesis. Fifty nanograms of
pcDNA-TiPARP, pEGFP-C2-TiPARP and pGEX-4T1-TiPARP was added to a reaction mixture
containing 0.2 mM dNTPs, 0.2 μM forward and reverse primers containing desired mutation
55
(Table 5), Pfu Ultra DNA polymerase (Agilent) or Pfx50 DNA polyermase (Life Technologies)
with 0.5 mM MgCl2 and amplified using the DNAEngine Peltier Thermal Cycler (Bio-Rad).
PCR products were digested by 1 μl DpnI restriction enzyme (New England BioLabs) for 1 h at
37°C and 10 μl of digested PCR product was transformed into chemically competent DH5α E.
coli. Bacteria were plated on LB-ampicillin or LB-kanamycin agar plates and incubated
overnight at 37°C. The following day colonies were selected and grown overnight at 37°C under
constant agitation. The following morning plasmid DNA from selected colonies was purified
using GenElute Plasmid Miniprep kit (Sigma) according to the manufacturer’s instructions.
Purified plasmid DNA was sent to The Centre for Applied Genomics DNA Sequencing and
Synthesis Facility (Medical and Related Sciences (MaRS) building, Toronto, ON, Canada) to
verify mutations.
Table 5. PCR primers used for site-directed mutagenesis of TiPARP.
TiPARP K41A 5’ ATTGAAGACTTGTTTTGCGAAAAAGGATCAGAAAAGA TiPARP K41A 3’ TCTTTTCTGATCCTTTTTCGCAAAACAAGTCTTCAAT TiPARP K41A; K42A; K43A 5’
CCATTGAAGACTTGTTTTGCGGCAGCGGATCAGAAAAGATTGGG
TiPARP K41A; K42A; K43A 3’
CCCAATCTTTTCTGATCCGCTGCCGCAAAACAAGTCTTCAATGG
TiPARP C243A 5’ AACGGAATTGAAATTGCCATGGACTTTCTGCAAGG TiPARP C243A 3’ CCTTGCAGAAAGTCCATGGCAATTTCAATTCCGTT TiPARP C251A 5’ CTTTCTGCAAGGCACTGCTATTTATGGCAGGGATT TiPARP C251A 3’ AATCCCTGCCATAAATAGCAGTGCCTTGCAGAAAG TiPARP C257A 5’ TATTTATGGCAGGGATGCTTTGAAGCACCACACTGT TiPARP C257A 3’ ACAGTGTGGTGCTTCAAAGCATCCCTGCCATAAATA TiPARP H532A 5’ AGACATTTATTTGCTGGAACATCCCAGG TiPARP H532A 3’ CCTGGGATGTTCCAGCAAATAAATGTCT TiPARP Y564A 5’ TTGGACAAGGCAGTGCTTTTGCAAAGAAGG TiPARP Y564A 3’ CCTTCTTTGCAAAAGCACTGCCTTGTCCAA TiPARP I631A 5’ TTTCTTTGAGCCTCAGGCTTTTGTCATTTTTAATG TiPARP I631A 3’ CATTAAAAATGACAAAAGCCTGAGGCTCAAAGAAA TiPARP I631E 5’ TTTCTTTGAGCCTCAGGAATTTGTCATTTTTAATGATG TiPARP I631E 3’ CATCATTAAAAATGACAAATTCCTGAGGCTCAAAGAAA TiPARP 449 stop 5’ TTCAGCAAACTTTTAGCCTGAAACTTGG TiPARP 449 stop 3’ CCAAGTTTCAGGCTAAAAGTTTGCTGAA A218; S219A 5’ AGGACAAAAGTGAAGAGGCTGCCCTTGACCTCGTGTTTGAGCTG A218; S219A 3’ CAGCTCAAACACGAGGTCAAGGGCAGCCTCTTCACTTTTGTCCT L220A; D221A 5’ ACAAAAGTGAAGAGGCTTCCGCTGCCCTCGTGTTTGAGCTGGTGAA L220A; D221A 3’ TTCACCAGCTCAAACACGAGGGCAGCGGAAGCCTCTTCACTTTTGT L222A;V223A 5’ GTGAAGAGGCTTCCCTTGACGCCGCGTTTGAGCTGGTGAACCAGTT
56
L222A; V223A 3’ AACTGGTTCACCAGCTCAAACGCGGCGTCAAGGGAAGCCTCTTCAC F224A; E225A 5’ AGGCTTCCCTTGACCTCGTGGCTGCGCTGGTGAACCAGTTGCAGTA F224A; E225A 3’ TACTGCAACTGGTTCACCAGCGCAGCCACGAGGTCAAGGGAAGCCT L226A; V227A 5’ CCCTTGACCTCGTGTTTGAGGCGGCGAACCAGTTGCAGTACCACAC L226A; V227A 3’ GTGTGGTACTGCAACTGGTTCGCCGCCTCAAACACGAGGTCAAGGG N228A; Q229A 5’ ACCTCGTGTTTGAGCTGGTGGCCGCGTTGCAGTACCACACTCACCA N228A; Q229A 3’ TGGTGAGTGTGGTACTGCAACGCGGCCACCAGCTCAAACACGAGGT L230A; Q231A 5’ TGTTTGAGCTGGTGAACCAGGCGGCGTACCACACTCACCAAGAGAA L230A; Q231A 3’ TTCTCTTGGTGAGTGTGGTACGCCGCCTGGTTCACCAGCTCAAACA Y232A; H233A 5’ AGCTGGTGAACCAGTTGCAGGCCGCCACTCACCAAGAGAACGGAAT Y232A; H233A 3’ ATTCCGTTCTCTTGGTGAGTGGCGGCCTGCAACTGGTTCACCAGCT T234A; H235A 5’ TGAACCAGTTGCAGTACCACGCTGCCCAAGAGAACGGAATTGAAAT T234A; H235A 3’ ATTTCAATTCCGTTCTCTTGGGCAGCGTGGTACTGCAACTGGTTCA Q236A; E237A 5’ AGTTGCAGTACCACACTCACGCAGCGAACGGAATTGAAATTTGCAT Q236A; E237A 3’ ATGCAAATTTCAATTCCGTTCGCTGCGTGAGTGTGGTACTGCAACT N238A; G239 5’ AGTACCACACTCACCAAGAGGCCGGAATTGAAATTTGCATGGAC N238A; G239 3’ GTCCATGCAAATTTCAATTCCGGCCTCTTGGTGAGTGTGGTACT I240A;E241A 5’ ACACTCACCAAGAGAACGGAGCTGCAATTTGCATGGACTTTCTGCA I240A;E241A 3’ TGCAGAAAGTCCATGCAAATTGCAGCTCCGTTCTCTTGGTGAGTGT
5.4 Transient transfection of HuH7 cells for RNA expression, Western blot and ChIP assays
HuH7 cells were seeded in six-well plates (35 mm diameter) at a density of 2.0 x 105 cells/well.
Following 24 h cells were transfected with 3 μg pcDNA or pcDNA-TiPARP or pEGFP or
pEGFP-C2-TiPARP using 6 μl Lipofectamine LTX and 6 μl Plus reagent. Medium was changed
the following day and cells were treated with 10 nM TCDD or DMSO (0.1%) for 6 h and RNA
isolation was performed (see below). For ChIP assays transfected cells were treated with 10 nM
TCDD or DMSO for 45 min and cells were harvested for ChIP assays (see below). For Western
blots, HuH7 were transfected with 2 μg pEGFP-TiPARP or pEGFP-TiPARP point mutant or
truncation construct with 4 μl Lipofectamine LTX + 4 μl Plus reagent. Following 24 h medium
was changed and proteins were overexpressed for another 24 h and cells were prepared for
protein extraction.
5.5 Transient transfection of T47D cells for Western blot T47D cells were plated in six-well plates at a density of 3.0 x 105 cells/well. Following 24 h cells
were transfected with 3 μg pcDNA or pcDNA-TiPARP using 6 μl Lipofectamine LTX and 6 μl
57
Plus reagent. Medium was changed the following day and cells were treated for 0-24 h with 10
nM TCDD or DMSO and cells were harvested for protein extraction.
5.6 Transient transfection of HuH7 cells and reporter gene assays
HuH7 cells were plated in 12-well plates (22 mm diameter) at a density of 1.2 x 105 cells/well.
The following day cells were transfected with 50-1500 ng pcDNA-TiPARP and/or 250-750 ng
pRc-CMV-hAHR (a generous gift from Dr. Patricia Harper) and/or 250-750 ng pcDNA4-
hARNT (a generous gift from Dr. Kevin Gardner) or 200 ng pcDNA-mTiparp, pcDNA-
chTiPARP, 200-1500 ng pCMV-XL-ARTD1 or pCMV-XL-ARTD12 (Origene) or 200 ng
pcDNA-TiPARP truncation or point mutant plasmids or 5-100 ng pcDNA-AHRRΔ8 or 500-100
ng pcDNA-AHRR-FL and 200 ng pCYP1A1-luc or pGL3-CYP1B1-luc plasmid using 2-4 μl
Lipofectamine LTX and 2-4 μl Plus reagent. All reactions included 50 ng of pCH110-β-Gal (GE
Healthcare) to normalize for transfection efficiency. pcDNA3.1 was used as carrier DNA for all
transfection experiments. After 24 h cells were treated with TCDD or DMSO for 24 h and cells
were washed once with 1 ml PBS and lysed with 250 μl 1X Passive Lysis Buffer (Promega) for
15 min with gentle rocking. Twenty-five microliters of lysate were loaded in duplicate into a 96-
well black flat bottom plate and 25 μl ONE-Glo luciferase substrate (Promega) were added to
each well and luciferase activity was determined using a GLO-max luminometer (Promega). For
β-galactosidase activity 10 μl of lysate were loaded into a 96-well clear flat bottom plate in
duplicate and 100 μl β-galactosidase buffer (0.6 M Na2HPO4, 0.04 M NaH2PO4, 0.01 M KCl, 1
mM MgSO4, 100 mM β-mercaptoethanol) and 2.5 mM ONPG (ortho-Nitrophenyl-β-galactoside)
were added to lysate and plate was incubated at 37ºC for 30 min and reaction was stopped
addition of 160 mM Na2HCO3 to reaction and 420 nm absorbance was measured using
Multiskan EX Photometer (ThermoScientific). Luciferase activity was normalized to
corresponding β-galactosidase activity.
5.7 Transient transfection and RNAi T47D and NCI-N87 cells were plated in 6-well plates at a density of 3.0 x 105 cells/well. HuH7
cells were plated in in 6-well plates at a density of 2.0 x 105 cells/well. MCF7 cells were plated
in 6-well plates at a density of 3.5 x 105 cells/well. Twenty-four hours after plating T47D, HuH7
and NCI-N87 cells were transfected with 100 nM SMARTpool: ON-TARGETplus TiPARP
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siRNAs (siTiPARP; L-013948-00) or ON-TARGETplus Non-targeting Pool (NTP; D-0011810-
10) siRNAs with 4 μl DharmaFECT1 (Dharmacon, Lafayette, CO). T47D cells were transfected
with 100 nM ON-TARGETplus TiPARP (siTiPARP-12; J-013948-12, siTiPARP-13; J-013948-
13) siRNAs or ON-TARGETplus non-targeting siRNA #2 (NT; D-0018100-02) (Dharmacon)
with 4 μl DharmaFECT1. MCF7 cells were transfected with 50 nM MISSION AHRR siRNAs
(siAHRR1; SASI_Hs02_00353810, siAHRR2; SASI_Hs01_00213384, Sigma-Aldrich) or NT2
(Dharmacon) with 2 μl DharmaFECT1. Medium of all cell lines was changed 24 h following
transfection. TiPARP was knocked down for 48 h post-transfection and dosed with 10 nM
TCDD or DMSO for 1.5, 3 h and 24 h and cells were harvested for Western blot, ChIP and RNA
expression assays. AHRR was knocked down for 24 h post-transfection and dosed with 10 nM
TCDD or DMSO for 24 h and cells were harvested for protein and RNA expression assays.
5.8 mRNA time-course T47D cells were seeded in 6-well plates at a density of 3.0 x 105 cells/well. Following 24 h cells
were treated with 1 μg/ml actinomycin D (Sigma) for 1 h to 24 h or 10 nM TCDD for 0.25 h to
24 h. MCF7 cells were seeded in 6-well plates at a density of 3.5 x 105 cells/well. Following 24 h
cells were treated with 10 nM TCDD or DMSO for 0.25 h to 24 h.
5.9 Tiparp-null mice and generation of mouse embryonic fibroblasts (MEF)
Tiparp-/- mice (strain B6;129S4-TiparpGt(ROSA)79Sor) were purchased from Jackson Laboratory
(Bar Harbor, ME) and housed and bred at the Division of Comparative Medicine (University of
Toronto) under the management of our lab technician, Debbie Brenneman. Six to eight week old
male and female wildtype (Tiparp+/+), knockout (Tiparp-/-) and heterozygous (Tiparp+/-) mice
were treated with a single dose of 30 μg/kg b.w. TCDD or corn oil (control) by intraperitoneal
injection. Mice were sacrificed after 6 h of treatment and livers excised and snap frozen in liquid
nitrogen and transferred to a -80ºC freezer. Four mice of each group were tested.
Wildtype, Tiparp-deficient (Tiparp-/-) and Tiparp+/- fibroblasts were prepared from E14.5
embryos derived from matings of mice heterozygous for disruption of the Tiparp allele. Primary
fibroblasts from wildtype, Tiparp-/- and Tiparp+/- siblings were immortalized at passage 2 after
transfection with Simian virus large T antigen (SV40gp6) in pSG5 (Stratagene) and a puromycin
59
resistance plasmid, and selected in puromycin-containing medium. The genotypes of the MEFs
were verified by PCR. Once immortalized, all MEF cell lines were plated into 10 cm dishes at a
density of 2.0 million cells/dish. Twenty-four hours after plating cells were treated with 10 nM
TCDD or DMSO for 45 min for ChIP assays and 24 h for Western blots. For RNA expression
experiments MEFs were seeded in 6-well plates at a density of 1.5 x 105 cells/well and treated 24
h following plating with 10 nM TCDD or DMSO for 6 h or 24 h or pre-treated with 10 μg/ml
cycloheximide (CHX, Sigma) for 1 h followed by 10 nM TCDD for 6 h or pre-treated with 10
μM MG132 (Sigma) for 30 min followed by 10 nM TCDD for 4 h. Following all treatments cells
were harvested for RNA isolation.
5.10 RNA isolation and cDNA synthesis
5.10.1 Mammalian cells For RNA isolation of all transfected and/or treated cells, medium was aspirated and cells were
washed once with PBS. Three hundred and fifty microliters of Total RNA Lysis Solution (Bio-
Rad) containing 1% β-mercaptoethanol was added to each well to lyse cells. Following 10 min
cell lysates were scraped and transferred to microcentrifuge tubes. An equal volume of 70%
ethanol was added to each tube and was mixed by gentle pipetting to bind RNA. RNA was
purified using Aurum™ Total RNA Mini Kit (Bio-Rad) following the manufacturer’s protocol.
Briefly, RNA was bound to spin columns by centrifugation at 13,000 rpm for 30 s. The column
membranes were washed once with Total RNA Wash Low Stringency solution and 80 μl DNAse
I solution (DNAse I stock solution diluted 1:40 in DNAse I Dilution Solution) was added
directly to column membranes and allowed to digest for 15 min at room temperature. DNAse I
was removed by addition of Total RNA Wash High Stringency solution followed by
centrifugation at 13,000 rpm for 30 s. Column membranes were then washed with Total RNA
Wash Low Stringency solution and centrifuged at 13 000 rpm for 1 min. Columns were dried by
centrifugation at 13,000 rpm for 2 min prior to elution. To elute, columns were transferred to
new microcentrifuge tubes and 40 μl of Total RNA Elution Solution was placed directly onto
column membrane and allowed to incubate for 1 min. Tubes were centrifuged at 13,000 rpm for
2 min. RNA concentration was measured using the spectrophotometer (Biochrom) and all
samples were set to 50 ng/μl concentrations by addition of DNAase/RNAse-free distilled water.
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5.10.2 Mouse liver
Approximately 40 mg of snap frozen liver was thawed on ice and re-suspended in 700 μl Total
RNA Lysis Solution (Bio-Rad) containing 1% β-mercaptoethanol and homogenized for 20 s
using a VWR PowerMax AHS homogenizer. Homogenates were centrifuged at 13,000 rpm for 3
min and supernatant was transferred to a new microcentrifuge tube. An equal volume of 70%
ethanol was added to supernatant and mixed by gentle pipetting until homogenous. RNA was
purified using Aurum™ Total RNA Mini Kit (Bio-Rad) following the manufacturer’s protocol.
5.10.3 cDNA synthesis
For cDNA synthesis, 500 ng of extracted RNA was reversed transcribed using 2500 U
SuperScriptIII (Life Technologies) in a total volume of 20 μl containing 2.5 μM random
hexamers and 1 μM dNTPs. cDNA was synthesized for 1 h at 50°C then heated to 70°C for 15
min to inactivate enzyme. After synthesis cDNA samples were diluted with 60 μl
DNAse/RNAse-free distilled water. Real-time PCR (qPCR) was performed on 1 μl of cDNA
synthesis sample using SsoFast EvaGreen® SYBR Supermix (Bio-Rad) and primers to amplify
CYP1A1, CYP1B1, TiPARP, AHRR, NFE2L2, NQO1 and β-actin mRNAs (Table 6) using
Chromo4 Real-Time PCR detector (Bio-Rad) and analyzed using Opticon Monitor 3 (Bio-Rad).
All target gene transcripts were normalized to β-actin mRNA and run in triplicate. Fold change
was calculated using the difference between cycle threshold (Ct) value of treatment and control
using the formula comparative method. ΔCt sample - ΔCt control = ΔΔCt. The expression level
of each gene was then calculated using 2-ΔΔCt and compared to DMSO or 0 h treatment (=1). PCR
was performed using specific primers (Table 6). Cycling conditions consisted of an initial
denaturation step of 95°C for 3 min followed by 45 cycles of 95 °C for 15 s, and the 60°C
annealing temperature for 30 s.
Table 6. List of qPCR primers used in RNA expression assays.
hCYP1A1 mRNA 5' TGGTCTCCCTTCTCTACACTCTTGT hCYP1A1 mRNA 3' ATTTTCCCTATTACATTAAATCAATGGTTCT hCYP1B1 mRNA 5’ CCTATGTCCTGGCCTTCCTTT hCYP1B1 mRNA 3’ TGAGGAATAGTGACAGGCACAAA TiPARP mRNA 5' AGGGACCACTTTGGATGGAGAGAGT TiPARP mRNA 3' TCAGACCCCGAGAGTTGGCTT hAHRR mRNA 5’ CCCCGCCCTTGGAGACAGGA
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hAHRR mRNA 3’ AGTACTCGGTGGGCGTGCCT
β-actin mRNA 5’ CAGAGCAAGAGAGGCATCCT
β-actin mRNA 3’ GGTCTCAAACATGATCTGGGTC mCYP1A1 mRNA 5’ CGTTATGACCATGATGACCAAGA mCYP1A1 mRNA 3’ TCCCCAAACTCATTGCTCAGAT mCYP1B1 mRNA 5’ CCAGATCCCGCTGCTCTACA mCYP1B1 mRNA 3’ TGGACTGTCTGCACTAAGGCTG mTiPARP mRNA 5’ CCTTCAACCAAGAACAAAGCATCT mTiPARP mRNA 3’ GGATGTCTGTATTTTGTGGCTGAAT mAHRR mRNA 5’ CTGCCCAGGTACTCTGAACC mAHRR mRNA 3’ ACTGTCCACAAAGCCTGACC mNFE2L2 mRNA 5` GTGACGGTGGCAGCATGCCT mNFE2L2 mRNA 3` TGCCTTCAGCGTGCTTCGGG mNQO1 mRNA 5` GCAGGATTTGCCTACACATATGC mNQO1 mRNA 3` AGTGGTGATAGAAAGCAAGGTCTTC
5.11 Chromatin Immunoprecipitation (ChIP) For ChIP assays, all three MEF cell lines were plated in 10 cm dishes at a density of 2.0 million
cells/dish. MCF7 cells were plated in 10 cm dishes at a density of 3.5 million cells/dish. Cells
were grown for 24 h and treated with 10 nM TCDD or DMSO for 45 min. Treated HuH7 cells
overexpressing EGFP-TiPARP or EGFP and treated T47D cells with TiPARP knockdown (from
sections 5.4 and 5.7 respectively) were used for ChIP assays. Following treatment protein-DNA
complexes were cross-linked with 1% formaldehyde (Sigma) for 10 min and then quenched with
125 mM glycine for 5 min. The medium was aspirated and cells were washed twice with 2 ml
PBS. Five hundred microliters of PBS + 0.1% (v/v) Tween 20 (PBS/T) was added to each well
and cells were scraped and centrifuged for 5 min at 7 500 rpm at 4°C. Supernatant was aspirated
off and cells were re-suspended in ice cold 1 ml Buffer A (10 mM HEPES [pH 7.9], 1.5 mM
MgCl2, 10 mM KCl, 0.05% NP-40) containing 1X protease inhibitor cocktail (PIC, Sigma) and
rotated at 4ºC for 15 min to remove cytosol. Nuclei were pelleted by centrifugation at 7,500 rpm
for 5 min and supernatant containing cytosolic fraction was aspirated. Nuclei were re-suspended
in 400 μL TSE I buffer (50 mM Tris-Base [pH 8.0], 150 mM NaCl, 1 mM EDTA, 1% Triton X-
100) containing 1X PIC and incubated on ice for 10 min. Nuclei were sonicated on ice for 10
times for 10 sec each time, resulting in the shearing of chromatin to an average size of ~500 bp.
Soluble chromatin was collected by centrifugation at 13 000 rpm for 10 min at 4°C and
supernatant transferred to a new microcentrifuge tube. A volume of 30 μl of Protein A-Agarose
(50% slurry) was added to chromatin and incubated at 4°C under gentle rotation for 2 h. An
62
aliquot of chromatin was stored at -20°C for the 5% total input sample. Agarose beads were
pelleted by centrifugation and chromatin was aliquoted into fresh microcentrifuge tubes
containing 1 μg of antibody, 0.5 μg/μl BSA and 0.05 μg/μl salmon sperm DNA in TSE I buffer.
Chromatin was immunoprecipitated overnight at 4°C under gentle rotation. The following
morning Protein A-Agarose (25 μl of 50% slurry) was added and chromatin was incubated for
another 1.5 hours at 4°C under gentle rotation. Protein A beads were pelleted by centrifugation
and washed successively for 10 min in 3 X 1 ml of TSE I, 1 ml of TSE II (20 mM Tris-Base [pH
8.0], 500 mM NaCl, 2 mM EDTA, 1% Triton X-100, 0.1% SDS), 1 ml of LiCl buffer (20 mM
Tris-Base [pH 8.0], 250 mM LiCl, 1 mM EDTA, 1% NP-40, 1% Na-deoxycholate), and 2 X 1 ml
of TE (10 mM Tris-Base [pH 8.0], 1 mM EDTA). Protein-DNA complexes were eluted in 110 µl
of elution buffer (TE + 1% SDS) for 30 min under constant rotation, and the cross-links were
reversed by overnight incubation at 65°C. The following morning, eluted DNA was bound using
5X volume of Buffer PB (Qiagen) and purified using EZ-10 Spin Column PCR Purification Kit
(Bio Basic) according to manufacturer’s instructions with slight modification. Briefly, bound
DNA was added to column and centrifuged at 11 000 rpm for 1 min. Columns were washed
twice with wash solution (containing 95% ethanol) and dried by centrifugation. DNA was eluted
from column using 100 μl of elution buffer. ChIP DNA was quantified by qPCR using SsoFast
EvaGreen® SYBR Supermix (Bio-Rad) and primers (Table 7) to amplify human CYP1A1 and
CYP1B1 enhancer regions and mouse Cyp1a1 and Cyp1b1 enhancer regions and downstream
negative binding regions of each gene. ChIP DNA samples were run in triplicate on Chromo4
Real-Time PCR detector (Bio-Rad) and analyzed using Opticon Monitor 3 (Bio-Rad). Results
were normalized to 5% total input and reported as percent recruitment relative to 100% total
input.
Table 7. List of qPCR primers used for ChIP assays.
hCYP1A1 enhancer 5’ AGGCGTGGACCGAAAATG hCYP1A1 enhancer 3’ CTAGGTCTGCGTGTGGCTTCT hCYP1B1 enhancer 5’ ATATGACTGGAGCCGACTTTCC hCYP1B1 enhancer 3’ GGCGAACTTTATCGGGTTGA hCYP1A1 5 782 bp downstream 5’ TGGTCTCCCTTCTCTACACTCTTGT hCYP1A1 5 782 bp downstream 3’ ATTTTCCCTATTACATTAAATCAATGGTTCT hCYP1B1 8 437 bp downstream 5’ AGTACATTCTGGGGAAACAACA hCYP1B1 8 437 bp downstream 3’ TTTTGGTAATGGTGTCCCAGT mCyp1a1 enhancer 5’ GAGGATGGAGCAGGCTTACG mCyp1a1 enhancer 3’ GGGCTACAAAGGGTGATGCTT
63
mCyp1b1 enh XRE 5’ rt GGAGCCGACTTCCCAGAAG mCyp1b1 enh XRE 3’ rt AACAAACGGTTGGGTTGAAAAG mCyp1a1 12 791 bp downstream 5’ CGTTATGACCATGATGACCAAGA mCyp1a1 12 791 bp downstream 3’ TCCCCAAACTCATTGCTCAGAT mCyp1b1 7 576 bp downstream 5’ GTGCGGCAAAAGCATGTCTC mCyp1b1 7 576 bp downstream 3’ GGGGAAAAGCAACGTTCTGAC
5.12 Western blot
All transfected and/or treated cells were washed once with PBS and scraped in 350-750 μl
PBS/T and cells were pelleted by centrifugation at 7 500 rpm for 5 min at 4ºC and supernatants
were aspirated and pellets were frozen at -80ºC. For detection of AHR, cell pellets were thawed
on ice and re-suspended in 200 μl cold Cell Lysis Buffer (50 mM HEPES [pH 7.5], 10%
glycerol, 100 mM KCl, 2 mM EDTA, 0.1% NP-40) containing 1X PIC and 2 mM DTT. Cell
suspensions were incubated on ice for 10 min then sonicated for 15 s at 30% amplitude to lyse
cells. Cell lysates were placed back on ice and incubated for 10 min and soluble proteins were
quantified by Bradford assays. Fifty micrograms of cell lysates were resolved by SDS-8%-PAGE
and transferred to a nitrocellulose blotting membrane (Pall Life Sciences) in 25 mM Tris base
(pH 8.3) containing 19.2 mM glycine and 20% (v/v) methanol. The membranes were blocked in
2% fat-free milk PBS/T for 2 h at room temperature with constant rocking and then
immunoblotted with a 1:5000 dilution of anti-AHR (H-211, Santa Cruz) or 1:10 000 AHR (SA-
210, Biomol International Inc.) antibody in 2% fat-free milk PBS/T overnight at 4ºC with
constant rocking. The membrane was then washed with PBS/T for 30 min and then incubated
with ECL Anti-rabbit IgG, Horseradish Peroxidase linked antibody (NA934, GE Healthcare) for
1 h at room temperature with constant rocking. Proteins were visualized using ECL Select
Western Blotting Detection System (GE Healthcare) according to the manufacturer’s
instructions. The membranes were exposed to an autoradiography film for 1 to 5 min. β-actin
was used as a loading control. AHR protein levels were normalized to β-actin levels and
quantified using ImageJ analysis software (NIH).
For detection of AHRR, ARNT or GFP-TiPARP fusions, frozen cell pellets were thawed on ice
and then re-suspended in 100 μl cold TENG buffer (50 mM Tris-Base [pH 8.0], 2 mM EDTA,
150 mM NaCl, 5% glycerol, 1% NP-40) containing 2.5X PIC and 2 mM DTT. Cell suspensions
were incubated on ice for 10 min and then frozen at -80ºC for 10 min to lyse cells. Lysates were
thawed on ice for 10 min and then centrifuged at 13 000 rpm for 10 min at 4ºC to pellet cell
64
debris. Supernatants were collected and transferred to new microcentrifuge tubes and solubilized
proteins were quantified by Bradford assays. For AHRR, ARNT and GFP blots 200 μg whole
cell lysates were resolved on by SDS-8%-PAGE transferred to a nitrocellulose blotting
membrane as described above. For AHRR immunoblots, membranes were blocked for 2 h at
room temperature in 2% fat-free milk TNT buffer (100 mM Tris-Base [pH 7.5], 150 mM NaCl,
0.1% (v/v) Tween 20) and then immunobloted with 1:2000 anti-AHRR antibody (HPA019614,
Sigma) in 2% milk TNT overnight at 4ºC with constant rocking. For ARNT immunoblots
membranes were blocked with 5% milk TNT for 2 h at room temperature then immunoblotted
with 1:200 anti-ARNT antibody (C-19, Santa Cruz) in 5% milk TNT overnight at 4ºC with
constant rocking. The membranes were washed with TNT for 30 min and then membranes were
incubated with ECL Anti-rabbit IgG, Horseradish Peroxidase linked antibody (GE Healthcare)
for AHRR immunoblots or HRP-conjugated normal anti-goat IgG (sc2741, Santa Cruz) for
ARNT immunoblots for 1 h at room temperature with constant rocking. Proteins were visualized
using SuperSignal West Dura (Thermo Scientific) and ECL Plus Western Blotting Detection
System (GE Healthcare) for AHRR and ARNT immunoblots respectively, according to the
manufacturers’ instructions. The membranes were exposed to an autoradiography film for 1 to 5
min. For GFP immunoblots membranes were blocked in 5% fat-free milk PBS/T for 2 h and
immunoblotted for GFP-fusion proteins using 1:2000 anti-EGFP antibody (JL-8, Clontech) in
2.5% milk PBS/T overnight at 4ºC with constant rocking. The following morning membranes
were washed with PBS/T for 30 min and membranes were incubated with HRP-conjugated
normal anti-mouse IgG (sc2954) in 2.5% milk PBS/T for 1 h at room temperature with constant
rocking. Proteins were visualized using SuperSignal West Dura (Thermo Scientific) according to
manufacturer’s instructions. Membranes were exposed to autoradiography film for 1 to 5 min.
5.13 Co-immunoprecipitation TREx-FLP-IN 293 cells were seeded in 6-well plates at a density of 4.0 x 105 cells/well were
transfected with 650 ng pRc-CMV-AHR, pRc-CMV-AHR-1-425, pCMV-AHR-TAD (464-848)
(all generous gifts from Dr. Patricia Harper), 200 ng pcDNA4-hARNT and 650 ng pEGFP-
TiPARP-FL or 650 ng pEGFP-TiPARP truncation with 3 μl FuGENE HD (Roche) according to
the manufacturer’s instructions. Twenty-four hours following transfection cells were treated with
10 nM TCDD or DMSO for 1.5 h and cells were scraped in PBS/T and centrifuged at 7 500 rpm
for 5 min at 4ºC. Whole cell extracts were prepared by two freeze-thaw cycles using 200 μl ice
65
cold TENG buffer containing 1X PIC and 2 mM DTT as described above. Two micrograms of
anti-AHR antibody (H-211) or anti-GFP (632460, Clontech) 1:200 dilution were incubated with
whole cell extracts for 1 h in 4ºC with constant rotation followed by incubation with protein-A
beads for 1 h. Beads were washed five times with NP-40 buffer (20 mM Tris Base [pH 8.0], 150
mM NaCl, 10% glycerol, 1% NP-40, 2 mM EDTA) and eluted in 1X sample buffer, separated by
SDS-PAGE and transferred to nitrocellulose membrane. Membranes were exposed to anti-GFP
(JL-8, Clontech), anti-AHR (H-211) or anti-AHR (N-19) antibodies. All membranes were then
washed and incubated with the appropriate secondary antibodies for 1 h at room temperature.
After washing, bands were visualized using SuperSignal West Dura substrate or ECL Select or
ECL Plus.
5.14 In vitro translation and 35S PARP assays All pcDNA-TiPARP constructs were in vitro translated and labelled with 35S-methionine using
the TNT® Coupled Reticulocyte Lysate System according to manufacturer’s instructions to
verify expression. Briefly, one microgram of pcDNA-TiPARP construct was added to rabbit
reticulocyte lysate containing 1X TNT buffer, 1 mM amino acid mixture minus methionine, 40 U
RNAseOUT (Invitrogen), T7 RNA polymerase and 35S-methionine (Perkin-Elmer) and reactions
were incubated at 30ºC for 90 min. Five percent of each reaction was diluted in TENG buffer
and 1X loading dye was added to each tube. Proteins were resolved by SDS-8%-12% PAGE and
gels were incubated in KODAK™ ENLIGHTNING™ Rapid Autoradiography Enhancer
(Perkin-Elmer) for 30 min with gentle rocking. Gels were then incubated in gel drying solution
(10% (v/v) ethanol and 5% acetic acid) for 20 min with gentle rocking and gels were dried at
80ºC for 2 h. Proteins were visualized by autoradiography.
35S PARP assays were carried out in a 30 μl reaction volume at 30ºC in 1X PARP buffer (5 mM
Tris Base [pH 8.0], 0.2 mM DTT, 4 mM MgCl2, 1% BSA) or 500 μM β-NAD+ (Sigma) or 1 μg
activated DNA (Sigma) with 5% in vitro translated proteins. Reactions were stopped with 1X
sample buffer and resolved by SDS-8%-12% PAGE.
5.15 Indirect immunofluorescence Glass cover slips (VWR No. 1, 22 mm x 22 mm) were sterilized in 70% ethanol for 20 min in the
cell culture hood. Cover slips were transferred to 6-well cell culture plates and allowed to dry
66
against well wall for 15 min. HuH7 and COS7 cells were seeded onto sterile cover slips at a
density of 2.0 x 105 cells/well and MCF7 cells were seeded onto cover slips at a density of 2.0 x
105 cells/well and cells were allowed to adhere overnight. Cells were transfected with 700 ng
pEGFP-TiPARP constructs and/or 500 ng pRc-CMV-AHR and 300 ng pcDNA4-ARNT with 3
μl Lipofectamine LTX or 100 ng pEGFP-AHRR constructs or 500 ng pAcGFP-AHRR
constructs and all proteins were allowed to overexpress for 24 h. Cells co-transfected with
pEGFP-TiPARP, pRc-CMV-AHR and pcDNA4-ARNT were treated with 10 nM TCDD for 1.5
h. Cover slips were washed once with PBS and fixed in 4% paraformaldehyde for 10 min. Cells
were PBS washed three times for 5 min each and then incubated with PBS + 0.4% Triton-X in
PBS for 20 min to permeabilize nuclei. Cover slips were washed three times with PBS for 5 min
and mounted with Vectashield containing 4,6-diamidino-2-phenylindole (DAPI) (Vector
Laboratories).
For cells co-overexpressed with GFP-TiPARP constructs, AHR and ARNT coverslips were
blocked in 1% goat serum/0.2% Triton-X in PBS for 1 h and incubated with 1:100 anti-AHR
antibody (H-211, Santa Cruz) in 1% goat serum/0.2% Triton-X in PBS overnight at 4ºC. The
following morning cover slips were then incubated with Alexa 568-conjugated secondary
antibody (Life Technologies) for 45 min and mounted with Vectashield containing DAPI.
For detection of endogenous TiPARP, HuH7 cells were fixed with methanol/acetone for 20
min/1 min at -20ºC. Cells were permeabilized with 0.4% Triton-X in PBS for 20 min and then
blocked with 1% goat serum/0.2% Triton-X in PBS for 1 h. Cover slips were incubated
overnight with anti-TiPARP (84664, Abcam) diluted 1:100 in 1% goat serum/0.2% Triton-X in
PBS overnight at 4ºC. The following morning cover slips were then incubated with Alexa 568-
conjugated secondary antibody for 45 min and mounted with Vectashield containing DAPI.
All images were acquired using an Imager.Z1 epifluorescence microscope and Axiovision
software (Zeiss) or an Olympus Fluoview 1000 confocal microscope.
5.16 GST purification pGEX-4T1-TiPARP constructs were transformed into BL21 stain E. coli. The following day
single colonies were selected and cultured overnight in 10 ml LB. Overnight cultures were
transferred to 500 ml LB-ampicillin and cultured at 30ºC under constant shaking. Once cultures
67
reached OD600= 0.8-1.0 GST fusion protein expression was induced with 0.1 mM isopropyl-β-D-
thio-galactoside (IPTG, BioShop) for 3-4 h at 37ºC with shaking. Bacteria were pelleted at 8 000
rpm for 10 min at 4ºC and frozen overnight at -20ºC. Pellets were thawed on ice and re-
suspended in 10 ml GST lysis buffer (20 mM Tris Base [pH 7.6], 10 mM MgCl2, 1 mM MnCl2,
1 mM EDTA, 150 mM NaCl, 5% Glycerol) containing 1X PIC and then incubated with 1 mg/ml
lysozyme (BioShop), 1 μg/ml DNAse I (Life Technologies), 10 mg/ml RNAse A (BioShop) for
15 min at room temperature. Extracts were sonicated for 2 min at 45% amplitude and rotated at
4ºC for 30 min. Extracts were centrifuged at 14 000 rpm for 15 min and soluble protein were
transferred to pre-cooled 50 ml conical tubes. One milliliter of equilibrated Glutathione
Sepharose 4 Fast Flow beads (50% slurry in GST lysis buffer) were added to extracts and rotated
overnight at 4ºC. The following morning beads were loading into 20 ml Econo-Column
Chromatography Columns (Bio-Rad). Beads were consecutively washed at 4ºC with cell lysis
buffer (20X bed volume) 3X and NETN buffer (50 mM Tris Base [pH 7.6], 100 mM NaCl, 1
mM MgCl2, 10% glycerol, 0.5% NP-40) 3X. GST fusion proteins were eluted from beads in 2 ml
10 mM glutathione for 30 min and concentrated and purified using Amicon Ultra-0.5 ml
Centrifugal Filters 50 kDa cut off (Millipore).
5.17 32P-NAD+ auto-ADP-ribosylation and hetero-ADP-ribosylation assays
Purified ARTD1 was purchased from Trevigen (Gaithersburg, MD). Auto-ribosylation assays
were carried out in a 30 μl reaction volume at 30ºC in 1X PARP buffer (5 mM Tris Base [pH
8.0], 0.2 mM DTT, 4 mM MgCl2, 0.1 μg/ml BSA), 2 μCi 32P-NAD+ (Perkin-Elmer,
Woodbridge, ON) and/or 500 μM β-NAD+ (Sigma), 1 μg activated DNA (Sigma) or RNA.
Approximately 5 μg of GST fusion partially purified protein or 1 μg ARTD1 were used. For
hetero-ribosylation experiments 8 μg of core histones (H2A, H2B, H3, H4) (New England
Biolabs) were added to the reaction. Reactions were incubated for 30 min and stopped by
addition of 1X sample buffer, separated by SDS-PAGE and visualized by autoradiography.
5.18 Statistical Analysis All real-time PCR results are expressed as means ± standard error of the means (SEM).
Statistical analysis was calculated using GraphPad Prism statistical software (San Diego, CA).
68
One-way Analysis of Variance (ANOVA) followed by Tukey’s multiple comparison tests were
used to assess statistical significance (P<0.05).
69
Chapter 4: Results 6 TiPARP is a mono-ADP-ribosyltransferase
6.1 Auto-ADP-ribosylation The catalytic activities of ARTDs are dependent on a well-conserved glutamate residue within
the catalytic active site of the HYE triad sequence (Holbourn, et al., 2006; Marsischky, et al.,
1995). Sequence alignment of the ARTD catalytic core sequences of the 18 ARTD family
members showed that H532, Y564 and I631 of TiPARP are equivalent to the ARTD catalytic
HYE triad. Since TiPARP has an isoleucine in place of the catalytic glutamate and a shortened
connecting loop, it is predicted to possess MART rather than poly-ADP-ribosyltransferase
activity (Kleine, et al., 2008); however, this has not been experimentally determined. To examine
the catalytic activity of TiPARP, we compared the enzymatic properties of TiPARP to that of
ARTD1 using an auto-ADP-ribosylation assay. ARTD1 demonstrated auto-ADP-ribosylation
with the incorporation of 32P-NAD+ and a marked shift in mobility and smearing pattern which
was indicative of poly(ADP-ribose) polymer formation (Figure 10A). Partially purified GST-
TiPARP also showed evidence of auto-modification but no apparent change in mobility,
suggesting that TiPARP exhibits mono-ADP-ribosyltransferase rather than poly(ADP-ribose)
polymer synthase activity (Figure 10B). This activity was competed away with unlabelled β-
NAD+. We cannot exclude the possibility that TiPARP generates short oligo-ADP-ribose units
since they would not alter the migration of the protein. Because ARTD1/PARP1 activity is
stimulated by its interaction with DNA (D'Amours, et al., 1999) and TiPARP contained a zinc-
finger domain with putative RNA binding capacity (Kelly, et al., 2007), we tested whether the
presence of DNA or RNA modified TiPARP auto-ribosylation activity. Addition of activated
DNA or DNAse-treated total RNA did not alter the migration pattern of GST-TiPARP (Figure
10B). We next tested mutants of the TiPARP catalytic triad. GST-TiPARP-H532A and GST-
TiPARP-Y564A did not show any evidence of auto-ribosylation or shift in mobility (Figure
10B). GST-TiPARP-I631A demonstrated similar auto-ribosylation activity to that of wildtype,
suggesting that this residue was not required for enzymatic activity (Figure 10B).
Next we evaluated the catalytic activity of mouse and chicken (Gallus gallus) TiPARP as GST
fusion proteins. GST-mouse TiPARP (GST-mTiPARP) demonstrated auto-ribosylation that
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could be mono-ADP-ribosyltransferase activity due to the lack of shift in mobility (Figure 10B).
GST-chicken TiPARP (GST-chTiPARP) demonstrated weak auto-ribosylation activity compared
to human, despite the loading of similar amounts of protein (Figure 10B). Addition of activated
DNA or DNAse-treated RNA did not alter the migration pattern or auto-ribosylation ability of
GST-mTiPARP or GST-chTiPARP (Figure 10C).
To determine if mutation of isoleucine 631 to a glutamate would convert TiPARP MART
activity to poly-ADP-ribosytransferase activity, we analyzed the auto-ribosylation activity of
GST-TiPARP-I631E mutant. The auto-ribosylation activity of GST-TiPARP-I631E was similar
to that of wildtype (Figure 10D), exhibiting MART but not polymer synthase activity.
To further examine TiPARP catalytic activity we used an in vitro 35S-labelled protein shift assay,
which allowed us to directly compare mobility shift of modified and unmodified ARTD proteins.
In vitro translated 35S-labelled ARTD1 (35S-ARTD1) incubated with β-NAD+ (and activated
DNA) resulted in a marked shift in its mobility compared to unmodified 35S-ARTD1 in the
absence of β-NAD+ (Figure 11A). Conversely, only a small change in mobility was observed for 35S-TiPARP when incubated with β-NAD+, which may represent mono-ADP-ribosylation of
multiple residues or short oligo-ADP-ribose units. None of the 35S-labelled TiPARP catalytic
point mutants demonstrated altered mobility compared to wildtype TiPARP (Figure 11B). The
absence of a modest mobility shift of 35S-labelled TiPARP-I631A in the presence of β-NAD+ is
in contrast to results obtained from 32P-NAD+ PARP assays with GST-TiPARP I631A (Figure
11B). This discrepancy could be due to differences in sensitivity between the two assays. The
I631A mutant may have reduced auto-ADP-ribosylation activity that could be below the
threshold of sensitivity of the 35S protein shift assay. A similar mono-ribosylation pattern was
observed for mouse 35S-TiPARP but not chicken 35S-TiPARP (Figure 11C). 32P-NAD+ catalytic
assays performed on GST-chTiPARP demonstrated very weak auto-ribosylation that may not
have been detected by the 35S protein shift assay. Taken together, these data revealed that
TiPARP was a mono-ADP-ribosyltransferase.
We next used the 35S-labelled in vitro protein shift assay to assess the catalytic activity of the N-
terminal truncations of TiPARP. Deletion mutants from amino acid residue 33 to residue 275
showed a slight migration or shift comparable to full-length TiPARP when incubated with β-
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NAD+ (Figure 11D). However, this shift was not apparent in N-terminal deletion mutants 328-
657 and 445-657, suggesting that the auto-modification domain of TiPARP is between residues
275 to 328.
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Figure 10. TiPARP is a mono-ADP-ribosyltransferase with auto-ribosylation activity.
(A) Auto-ADP-ribosylation of ARTD1, purified ARTD1 was incubated with 32P-NAD+ and 1 μg
activated DNA then analyzed the SDS-PAGE and autoradiography (B) Auto-ADP-ribosylation
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of GST-TiPARP, GST-TiPARP catalytic mutants (GST-TiPARP-H532A, –Y564A and –I631A),
GST-mouse TiPARP (GST-mTiPARP) and GST-chicken TiPARP (GST-chTiPARP). Auto-
ribosylation reactions were carried out with semi-purified GST-TiPARP fusion proteins and 32P-
NAD+ or co-incubated 500 μM β-NAD+ or 1 μg activated DNA or RNA. Automodified GST-
TiPARP fusion proteins were analyzed by SDS-PAGE and autoradiography. (C) Auto-ADP-
ribosylation of mouse TiPARP (mTiPARP) and chicken TiPARP (chTiPARP) incubated with
DNA or RNA. Mouse GST-tagged TiPARP (GST-mTiPARP) and chicken GST-tagged TiPARP
(GST-chTiPARP) were incubated with 32P-NAD+ and activated DNA or DNAse-treated RNA
and analyzed by SDS-PAGE and autoradiography. (D) Comparison of auto-ADP-ribosylation of
GST-TiPARP-I631E (GST-I631E) with wildtype GST-TiPARP (GST-wt). All figures represent
data from three independent experiments. Closed arrow denotes mobility shifted ARTD1.
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Figure 11. Putative auto-ribosylation domain of TiPARP is between residues 275 and 328.
(A) Mobility shift comparisons of modified and unmodified 35S-labelled ARTD1 and TiPARP.
In vitro translated 35S-labelled ARTD1 and TiPARP were incubated with or without 500 μM β-
NAD+, activated DNA or RNA and analyzed by SDS-PAGE and autoradiography. (B) TiPARP
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catalytic point mutants are inactive. (C) Auto-ribosylation of mouse TiPARP and chicken
TiPARP. In vitro translated 35S-labelled TiPARP, catalytic point mutants, mouse and chicken
TiPARP were incubated with or without 500 μM β-NAD+ and analyzed by SDS-PAGE and
autoradiography. (D) TiPARP automodification domain is between residues 275 and 328. In
vitro translated 35S-labelled TiPARP and N-terminal truncations were incubated with or without
500 μM β-NAD+ and analyzed by SDS-PAGE and autoradiography. These figures represent data
from three independent experiments. Closed arrow denotes shifted ARTD1.
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6.2 Hetero-ADP-ribosylation
We then compared the ability of GST-TiPARP and ARTD1 to hetero-ribosylate core histones. In
agreement with a previous report, ARTD1 hetero-ADP-ribosylated each of the core histones
(Figure 12A)(Messner, et al.). GST-TiPARP also mono-ADP-ribosylated each of the core
histones as evidenced by the lack of a marked mobility shift in molecular weight of the histones
(Figure 12B). No ADP-ribosylation of the core histones was observed in the presence of the
catalytic mutant GST-TiPARP-H532A (Figure 12C). Similar to wildtype, GST-TiPARP-I631A
also ribosylated core histones in the presence of 32P-NAD+ (Figure 12D). These results
demonstrated TiPARP hetero-ribosylation activity was mono-ADP-ribosyltransferase.
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Figure 12. TiPARP is a mono-ADP-ribosyltransferase with heteroribosylation activity.
(A) Hetero-ribosylation of core histones by ARTD1. Purified ARTD1 was incubated with 2 μCi 32P-NAD+, individual or mixed core histones (core) and hetero-ribosylation was analyzed by
SDS-PAGE followed by autoradiography. (B) Hetero-ribosylation of core histones by GST-
TiPARP. GST-TiPARP was incubated with 32P-NAD+ and histones H2A, H2B, H3 or H4 and
analyzed by SDS-PAGE and autoradiography. (C) Catalytic mutant GST-TiPARP-H532A did
not hetero-ribosylation core histones. GST-TiPARP-H532A (H532A) or wildtype GST-TiPARP
(wt) were incubated with mixed core histones and hetero-ribosylation was determined by SDS-
PAGE followed by autoradiography. (D) Hetero-ribosylation of core histones by TiPARP
catalytic mutant I631A. GST-TiPARP-I631A was incubated with 32P-NAD+ and histones H2A,
H2B, H3 or H4 and analyzed by SDS-PAGE and autoradiography.
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7 Identification of TiPARP as a novel negative regulator of AHR
7.1 Temporal induction of TiPARP mRNA by TCDD TiPARP was first reported to be a TCDD-responsive gene that was regulated by AHR in mouse
hepatoma Hepa1c1c7 cell line (Ma, et al., 2001); however, little is known about the regulation of
TiPARP in human cell lines. With this in mind, we determined the temporal changes in TiPARP
expression after treatment of T47D human breast carcinoma cells with TCDD. The results were
compared to those obtained for CYP1A1. TCDD-dependent increases in TiPARP mRNA levels
peaked at 1.5 h, whereas CYP1A1 mRNA levels increased over the 24 h time course (Figure
13A). Similarly, TiPARP mRNA induction peaked at 1.5 h TCDD treatment in Hepa1c1c7 cells
(Figure 13B) (Ma, 2002). However, the TiPARP mRNA declined less rapidly compared to
T47D cells, suggesting possible cell type differences in TiPARP regulation or mRNA stability.
Similar kinetic differences between TCDD-induced TiPARP and CYP1A expression levels were
observed in chick embryo hepatocytes (Diani-Moore, et al., 2010), demonstrating that these
regulatory differences between TiPARP and CYP1A genes are conserved across species.
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Figure 13. Temporal induction of TiPARP mRNA in T47D and Hepa1c1c7 cell lines
treated with TCDD.
(A) Temporal analysis of TCDD-induced TiPARP and CYP1A1 mRNA expression levels. T47D
cells were treated with 10 nM TCDD at the times indicated. (B) Temporal analysis of TCDD-
induced TiPARP mRNA expression levels in Hepa1c1c7 cells. Hepa1c1c7 cells were treated
with 10 nM TCDD for the times indicated. Data are presented as means ± S.E.M. of three
independent replicates.
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7.2 TiPARP knockdown increased TCDD-induced AHR transactivation
To test whether TiPARP modulates AHR activity, we used RNAi-mediated knockdown of
TiPARP using pooled siTiPARP siRNA (containing four different siTiPARP sequences) in
human gastric carcinoma NCI-N87 cells, human hepatoma HuH7 cells and human breast
carcinoma T47D cells and examined TCDD-dependent changes in CYP1A1 and CYP1B1
mRNA levels. TiPARP mRNA levels were reduced to approximately 25-38% compared to
control cells from the cell lines with a non-targeting pool (NTP) of siRNA (Figure 14A).
Endogenous TiPARP protein levels could not be assessed by immunoblot due to the lack of a
suitable antibody. RNAi-mediated knockdown of TiPARP resulted in significantly greater
TCDD-dependent induction of CYP1A1 and CYP1B1 mRNA expression levels compared to
NTP control cells after 24 h exposure (Figure 14B and C). Next we verified these results by
using single siTiPARP sequences (siTiPARP12 and siTiPARP13) to knockdown TiPARP in
T47D cells. TiPARP mRNA levels were reduced to approximately 23% and 34% compared to
non-targeting (NT) control cells using siTiPARP12 and siTiPARP13 sequences, respectively
(Figure 15A). Due to the lack of suitable antibody to detect endogenous TiPARP protein, we
instead co-transfected GFP-tagged TiPARP with siTiPARP sequences to demonstrate
knockdown of overexpressed GFP-TiPARP protein with anti-GFP antibody (Figure 15B).
Single-strand RNAi-mediated knockdown of TiPARP resulted in significantly greater TCDD-
dependent induction of CYP1A1 and CYP1B1 mRNA expression levels compared to NT control
cells after 24 h exposure which is in agreement with that we observed with pooled siTiPARP
knockdown (Figure 15C). We next tested how soon after treatment TCDD-induced gene expression
was increased in T47D cells following 48 h knockdown of TiPARP. T47D cells were treated with TCDD
for 1.5 h and 3 h. Increased TCDD-dependent induction of CYP1A1 and CYP1B1 mRNA levels
following TiPARP knockdown was observed after 3 h but not 1.5 h after treatment (Figure 15D
and E).
To examine the TiPARP-dependent mechanism of AHR repression we performed ChIP assays in
T47D cells after RNAi-mediated TiPARP knockdown of TiPARP and treatment with TCDD for
1 h. In agreement with increased CYP1A1 and CYP1B1 mRNA levels, knockdown of TiPARP
resulted in significantly greater recruitment of AHR and ARNT to the CYP1A1 and CYP1B1
enhancer regions compared to the NT control (Figure 16A and C). No significant differences in
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AHR or ARNT recruitment to downstream genomic control regions for both genes were
observed (Figure 16B and D).
We next determined the effect of TiPARP knockdown on ligand-induced AHR protein
degradation. Cells with TiPARP knocked down trended toward increased constitutive AHR
protein levels (Figure 16E). TiPARP knockdown cells treated with TCDD for 24 h demonstrated
reduced AHR degradation compared with TCDD-treated non-targeting cells, which is consistent
with elevated AHR-regulated gene expression and AHR recruitment results (Figure 16E). These
data suggested TiPARP mechanism of AHR repression to potentially involve reduced proteolytic
degradation of AHR.
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Figure 14. TiPARP knockdown in NCI-N87, HuH7 and T47D cells increased TCDD-
induced AHR transactivation.
Cell lines were transfected with siRNA pool (containing four different siRNAs) targeting against
TiPARP for 48 h. Cells were then treated with 10 nM TCDD and DMSO (control) for 24 h and
RNA was isolated and reverse transcribed as described in the Materials and Methods. Changes in
(A) TiPARP; (B) CYP1A1; and (C) CYP1B1 mRNA expression was then determined using
qPCR. Data were normalized to non-targeting pool (NTP) DMSO (control) and to β-actin levels
of each cell line. Results shown are means ± S.E.M for at least four independent experiments.
mRNA expression levels for each cell line significantly (P < 0.05) different than NTP transfected
cells are denoted with an asterisk.
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85
Figure 15. TiPARP knockdown increased TCDD-induced AHR transactivation.
(A) TiPARP mRNA expression levels in T47D cells following 48 h knockdown with single
siRNAs against TiPARP (siTP12 and siTP13) and non-targeting control (NT). Expression levels
significantly (P < 0.05) lower than NT were denoted with asterisks. (B) Western blot analysis of
GFP-TiPARP protein overexpression following RNAi-mediated TiPARP knockdown in TREx-
FLP-IN 293 cells. (C) TiPARP knockdown increased TCDD-induced CYP1A1 and CYP1B1
mRNA expression levels in T47D cells following 24 h treatment. Gene expression levels
significantly (P < 0.05) greater than NT were denoted with asterisks. (D) TiPARP knockdown
did not affect CYP1A1 and CYP1B1 mRNA levels following 1.5 h TCDD treatment. Asterisks
denote TiPARP mRNA levels significantly lower than NT. (E) TiPARP increased TCDD-
induced CYP1A1 and CYP1B1 mRNA levels following 3 h treatment. Gene expression levels
significantly (P < 0.05) different than NT were denoted with asterisks.
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Figure 16. TiPARP knockdown increased AHR/ARNT recruitment to CYP1A1 and
CYP1B1 regulatory regions and reduced TCDD-induced degradation of AHR.
TiPARP knockdown increased TCDD-induced AHR and ARNT recruitment to CYP1A1 (A-B)
and CYP1B1 (C-D). AHR and ARNT to CYP1A1 regulatory region (A), CYP1A1 distal
downstream region (B), CYP1B1 regulatory region (C) and CYP1B1 distal downstream region
(D). T47D cells with TiPARP knocked down were treated with 10 nM TCDD for 1 h and
harvested for ChIP assays. Percent recruitment significantly greater (P < 0.05) than NT was
denoted with an asterisk. (E) TiPARP knockdown reduced TCDD-induced AHR degradation in
T47D cells. T47D cells following TiPARP knockdown were treated with TCDD (T) or DMSO
(D) for 24 h and AHR protein expression was determined by Western blot. Left panel;
representative Western blot from three independent experiments, right panel; Quantification of
AHR protein levels, densitometry was performed using ImageJ analysis software. AHR protein
levels were normalized to pcDNA-transfected and DMSO treated cells. Asterisks denoted AHR
protein levels significantly (P < 0.05) greater than treatment-matched non-targeting-transfected
cells. All data were presented as means ± S.E.M for three independent experiments. Statistical
analysis was determined by two-tailed Student’s t-test.
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7.3 Overexpression of TiPARP repressed AHR-mediated transactivation
Because the RNAi experiments suggested that TiPARP might act as a negative regulator of AHR
transactivation, we investigated the ability of TiPARP overexpression to repress AHR-dependent
gene expression in HuH7 cells transiently transfected with a CYP1A1-regulated luciferase
reporter plasmid (CYP1A1-luc). Transient transfection of increasing amounts of TiPARP
resulted in a dose-dependent reduction of TCDD-induced reporter gene activity (Figure 17A).
Similar results were observed using a CYP1B1-regulated luciferase reporter plasmid (CYP1B1-
luc) (Figure 17B). To examine whether TiPARP-mediated repression was limited to CYP1
regulatory regions we used an artificial 3XAHRE-luc reporter (3X repeat of Hairy and Enhancer
of Split-1 (HES1) AHRE sequence) and determined TCDD-induced reporter gene activity.
Overexpression of increasing amounts of TiPARP dose-dependently repressed 3XAHRE-luc
reporter gene activity, which was similar to the CYP1-luc reporter constructs (Figure 17C).
Transfection of up to one microgram of the catalytic inactive mutant TiPARP-H532A plasmid
did not repress CYP1A1-luc reporter gene activity but rather demonstrated dominant negative
activity, indicating catalytically active TiPARP is required for repression (Figure 17D). We next
tested mouse TiPARP, which shares 92.8% sequence homology with human TiPARP and
demonstrated it to repress induction of CYP1A1-luc activity similar to human TiPARP (Figure
17E). Chicken TiPARP, which only exhibits 68.5% sequence homology with human TiPARP
did not repress reporter gene activity. The lack of repression of chicken TiPARP may have been
due the use of mammalian cells and/or reporter gene in our model. Similar findings were
observed after transfection with GFP-tagged TiPARP fusion proteins (Figure 17F). Transfection
of equal amounts of GFP-TiPARP plasmid DNA resulted in similar protein expression levels of
human and mouse GFP-tagged TiPARP, whereas higher protein levels of chicken TiPARP were
observed (Figure 17G). Collectively these data showed that TiPARP was a negative regulator of
AHR transactivation, but this effect may exhibit species-specificity.
To examine if TiPARP overexpression inhibited endogenous CYP1A1 and CYP1B1 mRNA
expression, we transiently transfected HuH7 cells with TiPARP, treated them with TCDD for 24
h prior to isolating RNA and performing qPCR. We observed a significant repression of TCDD-
dependent increases in CYP1A1 and CYP1B1 mRNA levels in cells overexpressing TiPARP,
supporting our reporter gene assay results (Figure 18A and B).
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We then determined if TiPARP was recruited to AHR target gene enhancer regions using ChIP
assays. We transiently transfected HuH7 cells with GFP-tagged TiPARP and ChIP assays were
performed with an anti-GFP antibody. HuH7 cells transfected with GFP-TiPARP revealed a
small but significant TCDD-dependent increase in GFP-TiPARP recruitment to CYP1A1 after 45
min of TCDD treatment (Figure 18C), but not to a downstream chromatin region that was not
bound by AHR (Figure 18D). Interestingly, reduced AHR occupancy at CYP1A1 was also
observed in cells overexpressing GFP-TiPARP, which supported the increased AHR and ARNT
recruitment we observed with TiPARP knockdown (Figure 18C). We next examined if TiPARP
overexpression affected AHR protein levels. Time course studies showed a reduction in AHR
protein levels in cells overexpressing TiPARP in the presence or absence of TCDD (Figure
18E), which was in support of the increased AHR protein levels observed after RNAi-mediated
knockdown of TiPARP (Figure 16E).
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Figure 17. TiPARP overexpression reduced AHR-regulated reporter gene activity.
(A-C) Concentration-dependent reduction of TCDD-induced CYP1A1-luc, CYP1B1-luc and
3xAHRE/HES1-luc reporter gene activity with TiPARP overexpression. (D) Dose-response of
TiPARP catalytic mutant H532A in HuH7 cells. (E-F) Mammalian TiPARP inhibited TCDD-
induced CYP1A1-luc reporter gene activity. Cells were co-overexpressed with reporter gene and
increasing amounts of pcDNA-TiPARP (50 ng to 200 ng), pcDNA-TiPARP-H532A (500 ng to
1000 ng), 200 ng pcDNA-chicken TiPARP (chTiPARP), 200 ng pcDNA-mouse TiPARP
(mTiPARP), pEGFP-hTiPARP, pEGFP-chTiPARP or pEGFP-mTiPARP. All transfected cells
were treated with TCDD or DMSO for 24 h and reporter gene activity was determined. Values
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were normalized to 100% TCDD treatment. Data are represented as means ± S.E.M. of three
independent experiments. Statistical significance was analyzed by one-way ANOVA and
Tukey’s multiple comparisons test. Reporter gene activity significantly different than (P < 0.05)
TCDD-treated empty vector was denoted with an asterisk. (G) Western blot of overexpressed
GFP-hTiPARP, GFP-chTiPARP and GFP-mTiPARP in HuH7 cells.
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Figure 18. Overexpression of TiPARP repressed TCDD-induced AHR transactivation.
(A-B) TiPARP overexpression repressed TCDD induction of CYP1A1 (A) and CYP1B1 (B)
mRNA in HuH7 cells. Asterisks denote mRNA expression significantly lower (P < 0.05, two-
tailed Student’s t-test) than treatment-matched empty vector transfected cells. (C) Overexpressed
GFP-TiPARP recruitment to CYP1A1 regulatory region was induced by TCDD and repressed
AHR recruitment. (D) Recruitment of GFP-TiPARP and AHR to CYP1A1 distal downstream
region. HuH7 cells were transfected with pEGFP empty vector or pEGFP-TiPARP and dosed
with TCDD for 45 min and cell were harvested for ChIP assays. Percent recruitment of GFP-
TiPARP of significantly greater (P < 0.05, two-tailed Student’s t-test) than transfection-matched
DMSO-treated cells was denoted with an asterisk. Percent recruitment of AHR significantly
lower (P < 0.05, two-tailed Student’s t-test) than treatment-matched GFP-transfected cells is
denoted with a pound sign. Data shown are representative of three independent experiments. (E)
TiPARP overexpression increased TCDD-induced AHR proteasomal degradation. T47D cells
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were transfected with vector or pcDNA-TiPARP and dosed with TCDD for the indicated times
and AHR protein levels were determined by Western blot. Left panel; representative Western
blots from three independent experiments. Right panel; quantification of AHR protein levels,
densitometry was performed using ImageJ analysis software (NIH). Asterisks denote AHR
protein levels significantly (P < 0.05, two-tailed Student’s t-test) lower than treatment-matched
vector-transfected cells.
7.4 Tiparp knockout increased TCDD-dependent AHR target transactivation and AHR protein levels
To investigate the impact of TiPARP loss on AHR transactivation and protein expression we
created immortalized mouse embryonic fibroblasts (MEFs) from wildtype, Tiparp-/- and
Tiparp+/- siblings (strain B6;129S4). As expected, TiPARP mRNA levels were induced in
wildtype and Tiparp+/- cells after 24 h treatment with TCDD, but no TiPARP was detected in
Tiparp-/- cells (Figure 19A). TCDD treatment of Tiparp-/- MEFs resulted in greater increases in
CYP1A1, CYP1B1 and AHRR mRNA levels compared to wildtype and Tiparp+/- cells, while
GAPDH mRNA levels were unaffected by TCDD treatment or by genotype (Figure 19A and
B). Ectopic expression of TiPARP into Tiparp-/- MEFs reduced TCDD-dependent AHR
transactivation compared to vector control (Figure 19C). ChIP assays revealed significantly
greater TCDD-induced AHR recruitment to Cyp1a1 and Cyp1b1 in Tiparp-/- compared to
wildtype or Tiparp+/- cells (Figure 19D and E). Because we observed increased AHR
transactivation in Tiparp-/- compared to wildtype or Tiparp+/- MEFs, we examined the relative
AHR protein levels and ligand-induced AHR degradation in the different MEF lines. Tiparp-/-
cells had higher constitutive AHR protein levels compared to wildtype and Tiparp+/- cells
(Figure 19F). TCDD-induced degradation of AHR was reduced in Tiparp-/- compared to
wildtype and Tiparp+/- cells (Figure 19F). TCDD treatment for 24 h caused a 72% and 81%
reduction in AHR protein levels in wildtype and Tiparp+/- cells, respectively, but only a 53%
reduction in Tiparp-/- MEFs (Figure 19F). In contrast to increased AHR protein levels, small but
significant TCDD-dependent decreases in AHR mRNA levels were detected in Tiparp-/- cells
(Figure 19G).
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Figure 19. Tiparp knockout fibroblasts exhibited increased TCDD-induced AHR
transactivation.
(A) Wildtype (Tiparp+/+), Tiparp-/- and Tiparp+/- immortalized mouse embryonic fibroblast
(MEF) lines were treated with 10 nM TCDD for 24 h and gene expression determined. Gene
expression was normalized relative to DMSO treated wildtype cells. Gene expression levels
significantly (P < 0.05) different than treatment-matched wildtype cells are denoted with an
asterisk, and gene expression levels significantly (P < 0.05) different than treatment-matched
Tiparp+/- (+/-) cells are denoted with a pound sign. (B) Expression of GAPDH mRNA in
immortalized mouse embryonic fibroblast lines. (C) Genetic complementation of Tiparp-/- MEFs
with mouse TiPARP overexpression. Tiparp-/- MEFs were transfected with 2 μg pcDNA-
mTiPARP or pcDNA (vector). Following 24 h transfected cells were treated with TCDD for 24 h
and CYP1A1, CYP1B1 and AHRR mRNA levels were determined. Data presented were
representative of means ± S.E.M. of three independent experiments. mRNA expression
significantly (P<0.05) different than vector-transfected cells was denoted with an asterisk. (D-E)
Tiparp-/- cells exhibit increased TCDD-induced AHR recruitment to Cyp1a1 and Cyp1b1. (D)
AHR recruitment to Cyp1a1 regulatory region and distal downstream control region. (E) AHR
recruitment to Cyp1b1 regulatory region and distal downstream control region. MEF lines were
treated with 10 nM TCDD for 45 min and harvested for ChIP assays. Percent recruitment
significantly greater (P < 0.05) than treatment-matched wildtype cells was denoted with an
asterisk, and percent recruitment significantly greater (P < 0.05) than treatment-matched
Tiparp+/- (+/-) cells was denoted with a pound sign. (F) Tiparp-/- fibroblasts have reduced
TCDD-induced AHR degradation. MEF lines were treated with TCDD (T) or DMSO (D) for 24
h and AHR protein expression was determined by Western blot. Left panel; representative
Western blot from three independent experiments. Right panel; quantification of AHR protein
levels, densitometry was performed using ImageJ analysis software (NIH). Asterisks denoted
AHR protein levels significantly (P < 0.05, two-tailed Student’s t-test) greater than treatment-
matched wildtype cells. Pound sign denotes AHR protein levels significantly (P < 0.05, two-
tailed Student’s t-test) less than genotype-matched DMSO-treated cells. (G) AHR mRNA
expression levels of MEF cell lines treated with DMSO or TCDD for 24 h. Asterisks denoted
AHR mRNA levels significantly (P < 0.05, two-tailed Student’s t-test) greater than treatment-
matched wildtype cells. Pound sign denoted AHR mRNA levels significantly (P < 0.05, two-
tailed Student’s t-test) less than genotype-matched DMSO-treated cells.
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7.5 Tiparp knockout increased AHR-mediated gene expression in vivo
To determine if TiPARP was a negative regulator of AHR transactivation in vivo we treated male
and female 6-8 week old Tiparp+/+, Tiparp+/- and Tiparp-/- mice (strain B6;129S4-
TiparpGt(ROSA)79Sor) with 30 μg/kg b.w. TCDD or corn oil (vehicle control) for 6 h. Mice were
sacrificed, livers excised and AHR-regulated gene expression was determined. As expected
TiPARP expression was detected in hepatic tissues from wildtype and Tiparp+/- mice but not in
Tiparp-/- mice (Figures 20A and 21A). Following 6 h TCDD treatment both female and male
Tiparp-/- mice demonstrated significantly greater AHR target gene CYP1A1, CYP1B1, AHRR,
nuclear factor (erythroid-derived 2)-like 2 (NFE2L2) and NAD(P)H dehydrogenase (quinone) 1
(NQO1) mRNA expression compared to wildtype mice (Figures 20B-F and Figure 21B-F).
These data supported our TiPARP knockdown and Tiparp-/- MEF data and indicated that
TiPARP is a negative regulator of AHR transactivation in vivo.
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Figure 20. Female Tiparp-/- mice demonstrated increased TCDD-induced AHR
transactivation.
Six week old female Tiparp+/+, Tiparp+/- and Tiparp-/- mice were treated with corn oil (CO) or 30
μg/kg b.w. TCDD (TCDD) for 6 h and livers were excised, RNA was isolated and mRNA
reversed transcribed. Gene expression was normalized relative to CO-treated wildtype mice.
Each bar represented means ± S.E.M. from four animals. Data were analyzed by One-way
ANOVA and Tukey’s multiple comparisons test. Gene expression levels significantly (P < 0.05)
different than treatment-matched wildtype mice were denoted with an asterisk and gene
expression levels significantly (P < 0.05) different than treatment-matched Tiparp+/- (+/-) mice
were denoted with a pound sign.
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Figure 21. Male Tiparp-/- mice demonstrated increased TCDD-induced AHR
transactivation.
Six week old male Tiparp+/+, Tiparp+/- and Tiparp-/- mice were treated with corn oil (CO) or 30
μg/kg b.w. TCDD (TCDD) for 6 h and livers were excised, RNA was isolated and mRNA
reversed transcribed. Gene expression was normalized relative CO-treated wildtype mice. Each
bar represented the means ± S.E.M. from four animals. Data were analyzed by One-way
ANOVA and Tukey’s multiple comparisons test. Gene expression levels significantly (P < 0.05)
different than treatment matched wildtype mice were denoted with an asterisk and gene
expression levels significantly (P < 0.05) different than treatment- matched Tiparp+/- (+/-) mice
were denoted with a pound sign.
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8 Understanding the mechanism of TiPARP-mediated repression of AHR transactivation
8.1 TiPARP selective repression of AHR activity Because ARTD1 and other ARTD family members can function as transcriptional repressors or
activators (Kraus, 2008), we examined the ability of ARTD1 and ARTD12 to modulate AHR
transactivation. ARTD1 was selected since it is the most studied ARTD, whereas ARTD12 was
chosen because it shared the greatest sequence homology (46.8%) with TiPARP compared to
other members of the ARTD family (Schreiber, et al., 2006). We co-transfected 200 ng and 1500
ng of ARTD1 and ARTD12 with CYP1A1-luc into HuH7 cells and determined reporter gene
activity after 24 h TCDD treatment. Transfection of HuH7 cells with 200 ng of TiPARP caused a
~50% reduction in TCDD-dependent induction of CYP1A1-luc reporter gene activity, whereas
transfection with the same amount of ARTD1 or ARTD12 had no effect (Figure 22). Increasing
the amount of transfected plasmid DNA to 1500 ng caused a greater than 85% reduction in
CYP1A1-luc reporter gene activity by TiPARP, but only a ~40% reduction by ARTD1 or
ARTD12. These results suggested that TiPARP exhibited selective repression of AHR
transactivation compared to other ARTD family members.
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Figure 22. CYP1A1-luc reporter gene activity was preferentially inhibited by TiPARP.
HuH7 cells were co-overexpressed with 200 ng or 1500 ng of pcDNA-TiPARP, pCMV-ARTD1
or pCMV-ARTD12 and pCYP1A1-luc and treated with TCDD. Reporter gene activity
significantly lower (P < 0.05) than empty vector transfected cells was denoted with an asterisk.
Reporter gene activity significantly (P < 0.05) greater than TiPARP (1500 ng) transfected cells
was denoted with a pound sign. All luciferase data were presented as means ± S.E.M from three
independent experiments and statistical significance analyzed by One-way ANOVA and Tukey’s
multiple comparisons test. Western blots confirming the presence of increased ARTD1 and
ARTD12 protein expression following transient transfection are presented to the right of bar
graph.
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8.2 Ectopic AHR but not ARNT prevented TiPARP-mediated inhibition of AHR transactivation
The AHRR negatively regulates AHR transactivation by competing with AHR for ARNT
dimerization and binding to the AHRE (Mimura, et al., 1999), but also through ARNT-
independent mechanisms (Evans, et al., 2008; Karchner, et al., 2009). To evaluate the role of
ARNT in TiPARP-mediated repression of AHR transactivation we assessed the ability of
TiPARP to repress AHR transactivation in the presence of increasing amounts of AHR or
ARNT. Transient co-transfection of AHR but not ARNT rescued the TiPARP-dependent
repression of CYP1A1-regulated reporter gene induction (Figure 23A and B).
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Figure 23. Overexpression of AHR but not ARNT prevented TiPARP-mediated inhibition
of AHR transactivation.
HuH7 cells were co-transfected with pCYP1A1-luc, TiPARP (200 ng) and (A) AHR (250-750
ng) or (B) ARNT (250-750 ng). All transfected cells were treated with 10 nM TCDD for 24 h
and reporter gene activity was determined. Results were shown as means ± S.E.M from three
independent experiments and significance analyzed by One-way ANOVA and Tukey’s multiple
comparisons test. Reporter gene activity of cells co-overexpressing TiPARP significantly lower
than (P < 0.05) transfection- and treatment-matched control cells were denoted with an asterisk.
Western blots confirming the presence of increased AHR and ARNT protein expression
following transient transfection were presented to the right of respective bar graphs.
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8.3 Identification of a putative AHR repressor domain of TiPARP To determine the mechanism of TiPARP-dependent repression of AHR, we created a series of
TiPARP truncations and site-directed point mutants, and determined their ability to repress
AHR-dependent reporter gene activity in HuH7 cells. The expression of each truncated product
was confirmed by in vitro translation (Figure 24). N-terminal truncation constructs of TiPARP
expressing amino acids 33-657 to 218-657 repressed CYP1A1-regulated reporter gene activity
similar to full length TiPARP, with the exception of 200-657, which exhibited greater repressor
activity (Figure 25A). The 225-657 truncation construct exhibited a significantly reduced ability
to repress reporter gene activity, whereas overexpression of 245-657 to 445-657 truncations did
not repress AHR-regulated reporter gene activity. A similar pattern of repression was observed
with the CYP1B1-luc reporter (Figure 25B). We created GFP-tagged versions of all N-terminal
truncations to verify expression in transfected cells by Western blot with anti-GFP antibody. We
first tested the ability of the GFP-tagged N-terminal truncations to repress CYP1A1- and
CYP1B1-luc. The GFP-tagged N-terminal truncations displayed a similar pattern of repression of
both reporter genes to their untagged counterparts (Figure 26A and B). Western blot analysis of
the GFP-TiPARP N-terminal truncations showed that the lack of repression was not due to
reduced protein expression of the different GFP-TiPARP truncations since the protein expression
of all N-terminal truncations was greater than full-length GFP-TiPARP despite transfecting and
loading equal amounts (Figure 27).
Because loss of AHR repressive ability was lost between residues 218 and 245, we were then
interested in whether this region of TiPARP was a putative AHR repressor domain that contained
specific residues pertinent to its ability to repress AHR transactivation. We performed an alanine
scan study whereby we mutated every two consecutive residues to alanines between 218 and 241
and determined the ability of each double alanine point mutant to repress TCDD induction of
CYP1A1- and CYP1B1-luc reporter gene activity. Endogenous glycines were not mutated. All
double point mutants significantly repressed CYP1A1- and CYP1B1-luc activity to varying
degrees except for the Y232A; H233A mutant which failed to repress induction of both reporter
genes (Figure 28A and B). These findings indicated that Y232 and H233 were important for the
ability of TiPARP to repress AHR transactivation. Point mutations between residues 220 and 235
significantly reduced the AHR repressive activity of TiPARP indicating these residues may
contribute to repression of AHR as well.
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Figure 24. In vitro expression of 35S-labelled N-terminal TiPARP truncations.
N-terminal pcDNA-TiPARP constructs were in vitro translated with 35S-methionine and detected
by SDS-PAGE and autoradiography.
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106
Figure 25. Overexpression of N-terminal TiPARP truncations on AHR-regulated reporter
gene activity.
Reporter gene activity of CYP1A1-luc (A) and CYP1B1-luc (B). HuH7 cells were transiently co-
transfected with pcDNA-TiPARP N-terminal truncation construct and pCYP1A1-luc or
pCYP1B1-luc reporter genes. Cells were treated with DMSO or TCDD for 16 h and reporter
gene activity was determined. Values were normalized to 100% TCDD treatment. Data are
represented as means ± S.E.M. of three independent experiments. Statistical significance was
analyzed by One-way ANOVA and Tukey’s multiple comparisons test. Reporter gene activity
significantly different than (P < 0.05) TCDD-treated empty vector was denoted with an asterisk.
Reporter gene activity significantly different than (P < 0.05) TCDD-treated full-length TiPARP
(1-657) was denoted with a pound sign.
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Figure 26. Expression of N-terminal GFP-tagged TiPARP truncations on AHR-regulated
reporter gene activity.
Reporter gene activity of CYP1A1-luc (A) and CYP1B1-luc (B). HuH7 cells were transiently co-
transfected with pEGFP-TiPARP N-terminal truncation construct and pCYP1A1-luc or
pCYP1B1-luc reporter genes. Cells were treated with DMSO or TCDD for 16 h and reporter
gene activity was determined. Values were normalized to 100% TCDD treatment. Data are
represented as means ± S.E.M. of three independent experiments. Statistical significance was
analyzed by One-way ANOVA and Tukey’s multiple comparisons test. Reporter gene activity
significantly different than (P < 0.05) TCDD-treated empty vector was denoted with an asterisk.
Reporter gene activity significantly different than (P < 0.05) TCDD-treated full-length TiPARP
(1-657) was denoted with a pound sign.
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Figure 27. Overexpression of GFP-tagged TiPARP N-terminal truncations.
HuH7 cells were overexpressed with pEGFP-TiPARP N-terminal truncation constructs for 48 h
and proteins were detected by Western blot. GFP-TiPARP fusions were detected with anti-GFP
antibody. β-actin was used as a loading control.
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Figure 28. Effects of TiPARP double alanine point mutants on AHR-regulated reporter
gene activity.
Reporter gene activity of CYP1A1-luc (A) and CYP1B1-luc (B). HuH7 cells were transiently co-
transfected with pcDNA-TiPARP double alanine point mutant and pCYP1A1-luc or pCYP1B1-
luc. Cells were treated with DMSO or TCDD for 16 h and reporter gene activity was determined.
Values were normalized to 100% TCDD treatment. Data were represented as means ± S.E.M. of
three independent experiments. Statistical significance was analyzed by One-way ANOVA and
Tukey’s multiple comparisons test. Reporter gene activity significantly different than (P < 0.05)
TCDD-treated empty vector was denoted with an asterisk. Reporter gene activity significantly
different than (P < 0.05) TCDD-treated wildtype TiPARP was denoted with a pound sign.
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8.4 Repression of AHR by TiPARP required zinc finger function Because the zinc-finger domain (residues 237-264) overlapped with the putative repressor
domain (residues 218-245), we tested the importance of the zinc finger domain in TiPARP-
mediated repression by creating single point mutants of the cysteine residues of CCCH-type
zinc-finger motif. We verified the expression of all zinc finger mutants by in vitro translation
with 35S-methionine (Figure 29A). All three TiPARP cysteine point mutants failed to inhibit
TCDD-induced CYP1A1-luc and CYP1B1-luc reporter gene activity (Figure 29B and C). We
created GFP-tagged versions of all three zinc finger point mutants to test expression in
transiently transfected HuH7 cells. We compared the ability of the GFP-tagged zinc finger point
mutants to untagged. All GFP-tagged point mutants failed to repress induction of CYP1A1- and
CYP1B1-luc, which was similar to the untagged mutants (Figure 29D and E). Expression of
GFP-tagged zinc finger point mutants was greater than wildtype GFP-TiPARP despite equal
loading and transfection of the same amount. These observations demonstrated that the lack of
repression of the zinc finger point mutants was not due to lack of expression (Figure 29F). Zinc
finger function was required for TiPARP-mediated repression of AHR transactivation.
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Figure 29. Repression of AHR transactivation required zinc finger function.
(A) In vitro translated 35S-labelled TiPARP zinc finger point mutants. pcDNA-TiPARP zinc
finger point mutant constructs were in vitro translated with 35S-methionine and proteins were
detected by SDS-PAGE and autoradiography. (B-C) CYP1A1-luc and CYP1B1-luc reporter
gene activity was not repressed by TiPARP zinc finger point mutants. HuH7 cells were
transiently co-transfected with pcDNA-TiPARP zinc finger point mutant constructs and
pCYP1A1-luc or pCYP1B1-luc. Cells were treated with DMSO or TCDD for 16 h and reporter
gene activity was determined. (D-E) CYP1A1-luc and CYP1B1-luc reporter gene activity was
not repressed by GFP-tagged TiPARP zinc finger point mutants. HuH7 cells were transiently co-
transfected with pEGFP-TiPARP zinc finger point mutant constructs and pCYP1A1-luc or
pCYP1B1-luc. Cells were treated with DMSO or TCDD for 16 h and reporter gene activity was
determined. All luciferase data were normalized to TCDD treatment, which was set to 100%.
Data were represented as means ± S.E.M. of three independent experiments. Statistical
significance was analyzed by One-way ANOVA and Tukey’s multiple comparisons test.
Reporter gene activity significantly different than (P < 0.05) TCDD-treated empty vector was
denoted with an asterisk. Reporter gene activity significantly different than (P < 0.05) TCDD-
treated wildtype TiPARP was denoted with a pound sign. (F) Western blot of GFP-tagged
TiPARP zinc finger mutants. pEGFP-TiPARP zinc finger point mutant constructs were
overexpressed in HuH7 cells for 48 h. Proteins were detected by Western blot using anti-GFP
antibody. Representative Western blot of three independent replicates. β-actin was used as a
loading control.
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8.5 Repression of AHR transactivation by TiPARP required catalytic domain and function
To evaluate the importance of TiPARP catalytic domain in mediating the repression of AHR
transactivation, we created C-terminal truncation and catalytic point mutants, and evaluated them
in our transient transfection reporter gene assay. We in vitro translated both C-terminal
truncations of catalytic mutants constructs with 35S-methionine to confirm expression (Figures
30A and 31A). Deletion of the TiPARP C-terminus containing the catalytic domain (residues
449-657) abolished the ability of TiPARP to repress AHR-dependent reporter gene activity
(Figure 30B). The ability of TiPARP to repress CYP1A1- and CYP1B1-mediated reporter gene
activity was prevented with the H532A and Y564A point mutants, but not I631A (Figure 31B),
which retained catalytic activity (Figure 10B). Similar results were observed with GFP-tagged
TiPARP mutants and immunoblotting confirmed that the lack of repression was not due to lack
of protein expression (Figures 30C-D and 31C-D). Together these data demonstrated TiPARP-
mediated repression of AHR transactivation required catalytic activities.
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Figure 30. Repression of AHR transactivation required C-terminal catalytic domain.
(A) In vitro translated 35S-labelled TiPARP C-terminal truncations. pcDNA-TiPARP C-terminal
truncation constructs were in vitro translated with 35S-methionine and proteins were detected by
SDS-PAGE and autoradiography. (B-C) CYP1A1-luc and CYP1B1-luc reporter gene activity
was not repressed by TiPARP C-terminal truncations. HuH7 cells were transiently co-transfected
with pcDNA-TiPARP C-terminal truncation constructs and pCYP1A1-luc or pCYP1B1-luc.
Cells were treated with DMSO or TCDD for 16 h and reporter gene activity was determined. (D-
E) CYP1A1-luc and CYP1B1-luc reporter gene activity was not repressed by GFP-tagged
TiPARP C-terminal truncations. HuH7 cells were transiently co-transfected with pEGFP-
TiPARP C-terminal truncation constructs and pCYP1A1-luc or pCYP1B1-luc. Cells were treated
with DMSO or TCDD for 16 h and reporter gene activity was determined. All luciferase data
were normalized to 100% TCDD treatment. Data are represented as means ± S.E.M. of three
independent experiments. Statistical significance was analyzed by One-way ANOVA and
Tukey’s multiple comparisons test. Reporter gene activity significantly different than (P < 0.05)
TCDD-treated empty vector was denoted with an asterisk. (F) Western blot of GFP-tagged
TiPARP C-terminal truncations. pEGFP-TiPARP C-terminal truncation constructs were
overexpressed in HuH7 cells for 48 h. Proteins were detected by Western blot using anti-GFP
antibody. Representative Western blot of three independent replicates. β-actin was used as a
loading control.
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Figure 31. Repression of AHR transactivation required catalytic function.
(A) In vitro translated 35S-labelled TiPARP catalytic point mutants. pcDNA-TiPARP catalytic
point mutant constructs were in vitro translated with 35S-methionine and proteins were detected
by SDS-PAGE and autoradiography. (B-C) CYP1A1-luc and CYP1B1-luc reporter gene activity
was not repressed by TiPARP catalytic point mutants. HuH7 cells were transiently co-transfected
with pcDNA-TiPARP catalytic mutant constructs and pCYP1A1-luc or pCYP1B1-luc. Cells
were treated with DMSO or TCDD for 16 h and reporter gene activity was determined. (D-E)
CYP1A1-luc and CYP1B1-luc reporter gene activity was not repressed by GFP-tagged TiPARP
catalytic point mutants. HuH7 cells were transiently co-transfected with pEGFP-TiPARP
catalytic point mutant constructs and pCYP1A1-luc or pCYP1B1-luc. Cells were treated with
DMSO or TCDD for 16 h and reporter gene activity was determined. All luciferase data were
normalized to 100% TCDD treatment. Data are represented as means ± S.E.M. of three
independent experiments. Statistical significance was analyzed by One-way ANOVA and
Tukey’s multiple comparisons test. Reporter gene activity significantly different than (P < 0.05)
TCDD-treated empty vector was denoted with an asterisk. (F) Western blot of GFP-tagged
TiPARP catalytic point mutants. pEGFP-TiPARP catalytic point mutant constructs were
overexpressed in HuH7 cells for 48 h. Proteins were detected by Western blot using anti-GFP
antibody. Representative Western blot of three independent replicates. β-actin was used as a
loading control.
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8.6 TiPARP interaction with AHR To examine direct interactions between AHR and TiPARP we performed co-localization and co-
immunoprecipitation experiments. We first determined the subcellular localization of TiPARP.
Endogenous TiPARP localized as small nuclear foci in HuH7 cells (Figure 32A, panel i).
Overexpressed GFP-TiPARP exhibited focal nuclear expression patterns similar to endogenous
TiPARP (Figure 32A, panel ii). GFP-tagged zinc finger mutant C243A displayed cytosol foci,
suggesting the zinc finger domain is important for nuclear localization (Figure 32A, panel iii).
The GFP-tagged catalytic mutant H532A displayed a diffuse nuclear expression pattern,
suggesting catalytic function is important for foci organization (Figure 32A, panel iv). GFP-
tagged mouse TiPARP expressed a similar nuclear focal pattern to human (Figure 32A, panel
v), while GFP-tagged chicken TiPARP diffusely expressed in the cytosol (Figure 32A, panel
vi).
We then tested whether AHR co-localized with GFP-TiPARP in transfected HuH7 cells treated
with TCDD 1.5 h. In co-transfected cells, a fraction of the nuclear AHR was enriched in nuclear
foci containing GFP-TiPARP (Figure 32B). To verify interaction, experiments were repeated in
COS7 cells to score co-localization (Figure 32C). COS7 were selected for scoring because they
contain low levels of endogenous AHR (Xu, et al., 2007) and cells overexpressing AHR could be
easily distinguished from untransfected cells. GFP-TiPARP and AHR co-localization was scored
based on co-transfection and AHR focal staining. COS7 cells transfected with only AHR and
ARNT and stained for AHR were scored for focal staining. We observed AHR focal staining in
75.6% of GFP-TiPARP and AHR and ARNT co-transfected cells, whereas only 5.5% of cells
expressing AHR and ARNT demonstrated focal staining (Table 8).
Table 8. Frequency of co-localization of GFP-TiPARP and AHR in COS7 cells
Transfection Total number of cells AHR focal staining No AHR focal staining
GFP-TiPARP + AHR/ARNT
113
90 (75.6%)
23 (24.4%)
AHR/ARNT
238
13 (5.5%)
225 (94.5%)
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Co-immunoprecipitation experiments were then performed using TREx FLP-IN 293 cells
transfected with GFP-TiPARP, AHR and ARNT and treated with DMSO or TCDD for 1.5 h to
support interaction. Similar to co-localization experiments GFP-TiPARP and AHR co-
immunoprecipitated (Figure 33A). Deletion analysis revealed that AHR interacted with residues
275-448 of TiPARP (Figure 33B). Reciprocal co-immunoprecipitation experiments
demonstrated that both the N-terminus (residue 1-425) and C-terminal transactivation domain
(residue 464-848) of AHR co-immunoprecipitated with GFP-TiPARP (Figure 33C and D).
We then tested the ability of the catalytically inactive and non-repressive point mutant GFP-
TiPARP H532A to interact with AHR. COS7 cells co-overexpressed with GFP-TiPARP H532A
and AHR and ARNT demonstrated co-localization within the nucleus (Figure 33E). Co-
immunoprecipitation experiments demonstrated interaction between GFP-TiPARP H532A and
AHR in the presence and absence of TCDD treatment (Figure 33F). Similarly, GFP-TiPARP
C243A, a mutant that did not repress AHR-dependent reporter gene activity, co-
immunoprecipitated with AHR in the presence and absence of TCDD (Figure 33F).
Interestingly, the repressive N-terminal truncation, GFP-TiPARP 200-657, which co-
immunoprecipitated with AHR (Figure 33B) localized within the cytosol and co-localized with
cytosolic AHR (Figure 33G). These observations suggested that TiPARP interaction with AHR
is not dependent on subcellular localization or the repressive function of TiPARP.
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Figure 32. TiPARP interacts with AHR in the nucleus.
(A) Nuclear localization of endogenous TiPARP using anti-PARP7 (84664) (panel i),
overexpressed GFP-TiPARP (panel ii), zinc finger mutant GFP-TiPARP-C243A (panel iii),
catalytic mutant GFP-TiPARP-H532A (panel iv), GFP-mouse TiPARP (GFP-mTiPARP, panel
v) and GFP-chicken TiPARP (GFP-chTiPARP, panel vi) in HuH7 cells. Images were acquired
using an Olympus Fluoview 1000 confocal microscope. (B-C) Co-localization of overexpressed
GFP-TiPARP and AHR in HuH7 (B) and COS7 cells (C). For co-localization with AHR cells
were treated with 10 nM TCDD for 1.5 h and fixed, immunostained for AHR, counterstained
with DAPI and mounted using Vectashield. Images were acquired using an Imager.Z1
epifluorescence microscope and Axiovision software (Zeiss) following deconvolution.
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Figure 33. TiPARP truncation and point mutant interaction with AHR.
(A) Reciprocal co-immunoprecipitation experiments of overexpressed GFP-TiPARP and AHR.
TREx-FLP-IN 293 cells were transfected and treated with TCDD (T) or DMSO (D) for 1.5 h.
(B) Co-immunoprecipitation experiments of AHR and GFP-TiPARP truncations in TREx-FLP-
IN 293 cells (upper panel). Co-immunoprecipitation experiments with AHR and GFP in TREx-
FLP-IN 293 cells (lower panel). AHR (H-211) antibody was used for immunoprecipitation and
GFP (JL-8) was used to probe. (C-D) Co-immunoprecipitation experiments of GFP-TiPARP and
AHR truncations in TREx-FLP-IN 293 cells. GFP (632460) antibody was used for
immunoprecipitation and AHR (N-19 or H-211) antibodies were used to probe. (E) Co-
localization of overexpressed GFP-tagged TiPARP catalytic mutant H532A (GFP-H532A) and
AHR in COS7 cells. (F) Co-immunoprecipitation experiments of GFP-TiPARP catalytic
(H532A) and zinc finger (C243A) point mutants with AHR in TREx-FLP-IN 293 cells. (G) Co-
localization of overexpressed GFP-tagged TiPARP N-terminal truncation 200-657 (GFP-200-
657) with AHR. For all co-localization experiments transfected cells were treated with 10 nM
TCDD for 1.5 h and fixed, immunostained for AHR, counterstained with DAPI and mounted
using Vectashield. Images were acquired using an Imager.Z1 epifluorescence microscope and
Axiovision software (Zeiss).
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8.7 Identification of nuclear localization signal of N-terminus of TiPARP.
Our co-localization observations suggested the presence of a nuclear localization signal (NLS)
within the N-terminus of TiPARP between residues 1 and 200. We then examined the expression
patterns of the N-terminal and C-terminal GFP-TiPARP truncations. We transiently transfected
GFP-tagged TiPARP truncations into HuH7 cells and observed the subcellular localization. We
first overexpressed GFP-tagged N-terminal truncation 33-657 in HuH7 cells and observed
expression patterns similar to full-length GFP-TiPARP (Figure 34A panels i and ii). GFP-
TiPARP 53-657 expressed in the cytosol as foci within the perinuclear region (Figure 34A panel
iii), whereas GFP-TiPARP 103-657 expressed within the perinuclear region with less defined
foci (Figure 34A panel iv). Both GFP-TiPARP 235-657 and GFP-TiPARP 445-657 truncations
expressed in less defined pattern in the perinuclear region (Figure 34A panels v and vi). We
then overexpressed the C-terminal truncations in HuH7 cells and both GFP-TiPARP 1-448 and
1-234 expressed diffusely and exclusively in the nucleus (Figure 34A panels vii and viii). These
data suggested the presence of a nuclear localization signal between residues 33 and 53 of
TiPARP. Between residues 33 and 53 we identified a short sequence consisting of basic amino
acids (KKKDQKR), which is consistent with a putative NLS sequence (Marfori, et al., 2011).
We created a triple mutant of the three tandem lysines (K41A; K42A; K43A) as well as a single
mutant of the first lysine (K41A) and examined their cellular localization in HuH7 cells. Both the
triple and single NLS mutants expressed in the cytosol suggesting that these residues are part of a
functional NLS sequence (Figure 34A panels ix and x).
We next tested the ability of TiPARP NLS mutants to repress AHR transactivation. Using
reporter gene assays we determined both the triple and single NLS mutants retained the ability to
repress AHR-regulated reporter gene activity similar to wildtype (Figure 34B and C). TiPARP
53-657 also retained the ability to repress reporter gene activity similar to wildtype, which was
expected based on our previous findings that TiPARP truncations 33-657 through 225-657
repressed reporter gene activity. Despite the expression of the NLS mutants and N-terminal
truncations, 53-657 through 225-657, in the cytosol, AHR transactivation was still repressed.
These findings suggested that nuclear localization is not required for TiPARP-mediated
repression of AHR transactivation.
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Figure 34. Identification of putative nuclear localization signal (NLS) within TiPARP N-
terminus.
(A) Localization of EGFP-TiPARP N-terminal and C-terminal truncations and nuclear
localization single point mutants in HuH7 cells. Panel i: Full-length GFP-TiPARP (1-657), N-
terminal truncations- panel ii: GFP-TiPARP 33-657, panel iii: GFP-TiPARP 53-657, panel iv:
GFP-TiPARP 103-657, panel v: GFP-TiPARP 235-657, panel vi: GFP-TiPARP 445-657, C-
terminal truncations- panel vii: GFP-TiPARP 1-448, panel viii: GFP-TiPARP 1-234, NLS point
mutants- panel ix: GFP-TiPARP K41A; K42A; K43A, panel x: GFP-TiPARP K41A. All GFP
fusions were overexpressed for 24 h and then fixed in 4% paraformaldehyde. Images were
acquired using an Olympus Fluoview 1000 confocal microscope. (B-C) TiPARP 53-657
truncation and NLS mutants repressed CYP1A1-luc reporter gene activity. HuH7 cells were
transiently co-transfected with pcDNA-TiPARP truncation or point mutant expression constructs
(B) or pEGFP-TiPARP truncation or point mutant expression constructs (C). Cells were treated
with DMSO or TCDD for 16 h and reporter gene activity was determined. All luciferase data
were normalized to 100% TCDD treatment. Data are represented as means ± S.E.M. of three
independent experiments. Statistical significance was analyzed by one-way ANOVA and
Tukey’s multiple comparisons test. Reporter gene activity significantly different than (P < 0.05)
TCDD-treated empty vector was denoted with an asterisk.
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8.8 TiPARP is a labile factor that negatively regulates AHR transactivation
TCDD-induced proteolytic degradation of AHR has been proposed to be under the control of a
labile factor, referred to as an AHR degradation promoting factor (ADPF) that when inhibited
increases the expression of AHR target genes, such as CYP1A1 (Ma, 2007; Ma and Baldwin,
2000). In support of these findings, pre-treatment with cycloheximide (CHX) resulted in
enhanced TCDD-dependent CYP1A1 mRNA levels (Ma, 2007; Ma and Baldwin, 2000).
Because we observed that TiPARP was a negative regulator of AHR, we were interested to
determine if TiPARP was a labile factor regulating AHR protein levels. To this end, we tested
the stability of TiPARP mRNA by treating T47D cells with actinomycin D and isolating RNA at
the time indicated (Figure 35A). We observed a 50% loss in TiPARP mRNA after 1 h
actinomycin D treatment (Figure 35A), which was in agreement with a previous report (Ma and
Baldwin, 2002). Due to the lack of a suitable antibody to detect endogenous TiPARP protein
levels, we examined the stability of TiPARP by overexpressing GFP-tagged TiPARP in HuH7
and treating with the 26S proteasome inhibitor, MG-132. After 6 h MG-132 treatment we
observed increased GFP-TiPARP expression compared to transiently transfected but untreated
cells (Figure 35B).
These data suggested that TiPARP was a candidate for the labile regulator of AHR protein levels
and responsible for the increased expression of AHR target genes after pre-treatment with CHX
(Ma, 2007; Ma and Baldwin, 2000). To test this notion, we pre-treated wildtype, Tiparp-/- and
Tiparp+/- MEFs for 1 h with CHX prior to treatment with 6 h TCDD. Because we were interested
in determining the effect of CHX on TCDD-dependent target gene expression, we also
determined the fold increase of TCDD+CHX over TCDD alone for each genotype (Figure 35C
and D). CYP1A1 mRNA levels increased approximately 480- and 300-fold following
TCDD+CHX compared to TCDD alone in wildtype and Tiparp+/- MEFs, respectively. CYP1B1
and AHRR mRNA levels were also significantly increased following TCDD+CHX compared to
TCDD alone, but the fold increases were an order of magnitude lower than those observed for
CYP1A1 (Figure 35C). For Tiparp-/- cells, treatment with CHX+TCDD resulted in significant
but modest increases of approximately 2-fold for each of AHR target gene examined compared
to TCDD alone (Figure 35C). A similar pattern of CYP1A1 and CYP1B1 mRNA
129
superinduction after TCDD treatment was observed after MG-132 pre-treatment, although the
magnitude of induction was lower than that observed with CHX (Figure 35D). AHRR mRNA
was not significantly induced when co-treated with MG-132 for all MEF lines (Figure 35D).
These data suggested TiPARP was an important factor regulating the CHX- and MG-132-
dependent increases in ligand-activated AHR transactivation.
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Figure 35. TiPARP is a labile negative regulator of AHR.
(A) TiPARP mRNA is rapidly degraded after actinomycin D treatment. T47D cells were treated
with 1 μg/ml actinomycin D for the times indicated and TiPARP mRNA expression levels
determined by qPCR. (B) GFP-TiPARP overexpression is increased after proteasome inhibition.
HuH7 cells were transfected with pEGFP-TiPARP, pcDNA (vector) and pEGFP for 24 h and
treated with 25 μM MG-132 for 6 h and GFP-TiPARP protein levels were determined by
Western blot using anti-GFP antibody. The data presented were from a representative Western
blot from three independent experiments. (C) AHR target gene induction in MEF lines pre-
treated with 10 μg/ml cycloheximide (CHX). MEF cell lines were pre-treated with 10 μg/ml
CHX for 1 h then treated with TCDD for 6 h and gene expression was determined. Data were
normalized to wildtype DMSO. Fold changes between TCDD alone and CHX+TCDD (CHX+T)
were provided for each gene. Gene expression results were shown as means ± S.E.M for three
independent experiments and significance analyzed by One-way ANOVA and Tukey’s multiple
comparisons test. Gene expression levels significantly different (P < 0.05) than TCDD alone
within each genotype were denoted with an asterisk. (D) MEFs lines were pre-treated with 25
μM MG-132 (MG) for 30 min then treated with TCDD for 4 h and gene expression was
determined. Data were normalized to wildtype DMSO. Fold changes between TCDD alone and
MG-132+TCDD (MG+T) were provided for each gene. Gene expression results were shown as
means ± S.E.M for three independent experiments and significance analyzed by One-way
ANOVA and Tukey’s multiple comparisons test. Gene expression levels significantly different
(P < 0.05) than TCDD alone within each genotype were denoted with an asterisk.
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9 Comparison of TiPARP- and AHRR-mediated negative regulation of AHR transactivation
Much of our understanding of the negative regulation of AHR comes from studies of the aryl
hydrocarbon receptor repressor (AHRR), which is a part of a negative feedback loop regulating
AHR transactivation (Evans, et al., 2008; Karchner, et al., 2009; Mimura, et al., 1999). Similar
to TiPARP, AHRR expression is induced by ligand-activated AHR (Mimura, et al., 1999). The
AHRR has been proposed to negatively regulate AHR transactivation by competing with AHR
for ARNT heterodimerization and AHRE binding (Mimura, et al., 1999). However, we have
observed in both Tiparp-/- MEFs and Tiparp-/- mouse liver tissue the TCDD-mediated induction
of AHRR mRNA was increased. This observation contradicts the proposed mechanism of
AHRR-mediated repression of AHR transactivation since greater expression of the AHRR in
Tiparp-/- mice should result in greater repression of AHR target transcription rather than greater
induction. To better understand the mechanism(s) of AHR repression we compared TiPARP- and
AHRR-mediated repression of AHR transactivation.
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9.1 Kinetics of AHRR induction by TCDD Because both TiPARP and AHRR gene expression are induced by TCDD we first compared the
temporal changes in TiPARP and AHRR mRNA following TCDD treatment in MCF7 breast
carcinoma cells. TCDD-dependent increases in TiPARP mRNA levels were rapid and peaked
after 1.5 h similar to T47D cells, whereas AHRR mRNA an increased over 24 h (Figure 36A).
We detected AHRR protein after 24 h TCDD treatment (Figure 36B); due to lack of suitable
antibody for TiPARP we were unable to detect TiPARP protein expression following TCDD
treatment. We next examined AHRR recruitment to CYP1A1 and CYP1B1 enhancer regions.
Because we detected AHRR protein after 24 h TCDD treatment we treated cells up to 24 h and
then performed ChIP assays. We observed recruitment of AHRR to the CYP1A1 and CYP1B1
regulatory regions only after 24 h TCDD treatment (Figure 36C). AHR/ARNT recruitment
peaked after 45 min TCDD and declined following 6 h, similar to our previous reports (Figure
36D) (Ahmed, et al., 2009; Pansoy, et al. 2010). We previously observed recruitment of
overexpressed GFP-TiPARP to the CYP1A1 enhancer region following TCDD treatment (Figure
18C), indicating that both TiPARP and AHRR are present at AHR regulatory regions in response
to TCDD.
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Figure 36. AHRR is induced by TCDD and recruited to AHR-regulated genes.
(A) Temporal induction of AHRR mRNA and TiPARP mRNA in MCF7 cells. MCF7 cells were
treated with TCDD for the times indicated. (B) AHRR protein expression following 24 h TCDD
or DMSO treatment. (C) Recruitment of AHRR to the CYP1A1 and CYP1B1 regulatory regions.
MCF7 cells were treated with TCDD for the times indicated and AHRR recruitment was
determined by ChIP. (D) Recruitment of AHR and ARNT to the CYP1A1 and CYP1B1
regulatory regions. MCF7 cells were treated with TCDD for the times indicated and AHR and
ARNT recruitment was determined.
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9.2 AHRR knockdown did not affect AHR transactivation or protein levels
We have demonstrated that RNAi-mediated knockdown of TiPARP resulted in increased TCDD-
induced CYP1A1 and CYP1B1 mRNA expression and we were then interested in the effects of
AHRR knockdown on CYP1A1 and CYP1B1 mRNA levels. We transfected MCF7 cells with
three siAHRR strands for 24 h and treated cells with DMSO or TCDD for 24 h and determined
AHRR mRNA knockdown. All three siAHRR strands reduced AHRR mRNA levels by 38%-
48% following DMSO treatment (Figure 37A). AHRR mRNA levels were also reduced
following TCDD treatment (Figure 37B). Because AHRR protein was not detected in non-
induced (DMSO) MCF7 cells (Figure 36B), we induced AHRR expression by treating
transfected cells with TCDD 24 h post-transfection to determine AHRR protein knockdown. We
observed substantial knockdown of TCDD-induced AHRR protein by all three siAHRR
sequences tested (Figure 37B). We chose to use siAHRR2 and siAHRR3 strands for all
subsequent experiments because they showed greatest AHRR knockdown. We then examined
the effect of AHRR knockdown on AHR target gene expression. AHRR knockdown did not
significantly affect the TCDD-mediated induction of CYP1A1, CYP1B1 or TiPARP mRNA
levels (Figure 37C). We repeated TiPARP knockdown experiments in MCF7 cells to compare
the effects of TiPARP knockdown to AHRR knockdown. Similar to our previous data (Figure
15C) TiPARP knockdown increased TCDD-induced CYP1A1, CYP1B1 and AHRR mRNA
levels (Figure 37D), indicating the mechanism of AHRR-mediated repression of AHR
transactivation may differ from that of TiPARP.
We were then interested in whether AHRR knockdown affected AHR protein levels.
Knockdown of AHRR did not increase constitutive AHR protein levels (DMSO) nor was
TCDD-induced AHR proteasomal degradation reduced, which is in contrast to the effects of
TiPARP knockdown on AHR protein levels (Figure 38A). These results were, however, in
agreement with previous reports demonstrating AHRR overexpression did not alter AHR protein
levels (Evans, et al., 2008). Since ARNT has been previously proposed to play an important role
in AHRR-mediated repression of AHR (Mimura, et al., 1999) we tested ARNT protein levels
following AHRR knockdown. Basal ARNT protein levels following TCDD treatment were not
affected by AHRR knockdown (Figure 38A). TiPARP knockdown increased constitutive AHR
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and ARNT levels and decreased TCDD-induced degradation of both proteins, similar to that
observed in T47D cells (Figure 38B). Both basal and TCDD-induced AHRR protein levels were
not affected by TiPARP knockdown despite elevated TCDD-induced AHRR mRNA levels
(Figure 38B). A potential explanation for the discrepancy between AHRR mRNA and protein
levels could be that the increase in AHRR mRNA levels may not be sufficient to observe
increases in AHRR protein.
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Figure 37. Effects of AHRR knockdown and TiPARP knockdown on AHR transactivation.
(A) AHRR mRNA and (B) protein knockdown in MCF7 cells. MCF7 cells were transfected with
non-targeting (NT) or three siAHRR strands, following 24 h knockdown cells were treated with
DMSO (D) or TCDD (T) for 24 h and Western blots were performed. (C) Effects of AHRR
knockdown on AHR target gene induction. AHRR was knocked down in MCF7 cells, treated
with DMSO or TCDD for 24 h and AHR target mRNA expression determined. (D) TiPARP
mRNA knockdown in MCF7 cells (E) Effect of TiPARP knockdown on AHR target gene
induction. TiPARP was knocked down in MCF7 cells for 48 h and cells were treated with
DMSO or TCDD for 24 h. Gene expression data were presented as means ± S.E.M. of three
independent replicates. Statistical significance was determined by One-way ANOVA and
Tukey’s multiple comparisons test. mRNA significantly (P < 0.05) different than treatment-
matched non-targeting (NT) was denoted with an asterisk.
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Figure 38. AHRR, AHR and ARNT protein levels following AHRR (A) and TiPARP (B)
knockdown.
AHRR and TiPARP were knockdown in MCF7 cells. Cells were transfected with non-targeting
(NT), siAHRR2 and siAHRR3 or siTiPARP12 and siTiPARP13. Following knockdown cells
were treated with DMSO (D) or TCDD (T) for 24 h and protein levels determined by Western
blot.
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9.3 AHRR overexpression repressed AHR-regulated reporter gene activity
We next complemented the AHRR knockdown studies with overexpression studies. Based on a
previous report identifying two isoforms of the AHRR we tested both full-length AHRR
(AHRR719) and the predominantly expressed active form of the AHRR, which lacks exon 8
(AHRRΔ8) (Karchner, et al., 2009). We overexpressed increasing amounts of both forms of
AHRR into HuH7 cells and measured the ability of each form to repress TCDD-induced
CYP1A1-luc activity. Similar to a previous report, overexpression of increasing amounts of
AHRR719 did not affect CYP1A1-luc reporter gene activity, whereas AHRRΔ8 potently inhibited
activity (Figure 39A and B) (Karchner, et al., 2009). We next tested the ability of AHR and
ARNT overexpression to rescue AHRRΔ8 repression. Increasing amounts of AHR but not
ARNT prevented the AHRRΔ8-mediated repression of CYP1A1-luc reporter gene activity as
described previously (Karchner, et al., 2009). These results were also similar to rescue
experiments with TiPARP where AHR but not ARNT rescued TCDD-dependent induction of
CYP1A1-luc reporter gene activity (Figure 39C and D).
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Figure 39. Overexpression of AHRRΔ8 repressed AHR-regulated reporter gene activity.
HuH7 cells were co-transfected with pCYP1A1-luc and pcDNA-AHRR719 (A) or pcDNA-
AHRRΔ8 (B) at the amounts indicated. Cells were treated with TCDD for 16 h and reporter gene
activity was determined. Transfection efficiency was normalized to β-glactosidase activity. Data
are presented as means ± S.E.M. of three independent replicates. Co-overexpression of AHR (C)
but not ARNT (D) rescued CYP1A1-luc activity. HuH7 cells were co-transfected with 5 ng
pcDNA-AHRRΔ8 construct and 100-500 ng pRc-CMV-AHR or pcDNA4-ARNT. Transfected
cells were treated with DMSO or TCDD for 16 h and CYP1A1-luc reporter gene activity was
determined. Transfection efficiency was normalized to β-glactosidase activity. Data are
presented as means ± S.E.M. of three independent replicates. Reporter gene activity significantly
different than AHRRΔ8 alone was denoted with an asterisk.
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9.4 Expression patterns of the AHRR
To examine the cellular localization patterns of the AHRR we created both N- and C-terminal
GFP-tagged expression constructs of AHRRΔ8 and AHRR719. We overexpressed all constructs
in HuH7, COS7 and MCF7 cells. Unexpectedly, both GFP-tagged AHRRΔ8 and AHRR719
expressed in the cytosol in HuH7 and COS7 cells (Figure 40). The location (N- or C-terminally
linked) of the GFP tag on either AHRRΔ8 or AHRR719 did not affect their expression or
localization pattern. We then repeated overexpression of GFP-tagged AHRR constructs in MCF7
cells, GFP-tagged AHRRΔ8 localized to the nucleus, whereas the GFP-tagged AHRR719
localized diffusely to both the nucleus and cytosol (Figure 40). GFP-tagged TiPARP, when
overexpressed in MCF7 cells, appeared as nuclear foci, similar to that observed in HuH7 and
COS7 cells (Figure 32A). Despite the differences in expression patterns of the AHRR in the
three cell lines tested the AHRR significantly repressed AHR-regulated reporter gene activity in
all three cell lines (Karchner, et al., 2009).
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Figure 40. Expression of N-terminal and C-terminal GFP-tagged AHRR in MCF7, HuH7
and COS7 cells.
MCF7, HuH7 and COS7 cells were transfected with pEGFP-C2-AHRRΔ8, pEGFP-C2-
AHRR719, pAcGFP-N1-AHRRΔ8, pAcGFP-N1-AHRR719 or pEGFP-C2-TiPARP. All GFP
fusions were overexpressed for 24 h then fixed with 4% paraformaldehyde. Images were
acquired using an Olympus Fluoview 1000 confocal microscope.
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Chapter 5: Discussion 10 TiPARP is a mono-ADP-ribosyltransferase Class 2 ARTDs are predicted to be mono-ADP-ribosyltransferases based on the sequence and
structural features of their catalytic domains (Otto, et al., 2005). The presence of a glutamate
within the catalytic HYE triad motif, which is essential for poly(ADP-ribose) chain elongation is
absent from class 2 ARTD members and restricts their enzymatic activities to mono-ADP-
ribosylation (Marsischky, et al., 1995; Rolli, et al., 1997; Ruf, et al., 1998). In agreement with
this notion, the class 2 ARTDs, ARTD7, 8, 10, 12 and 15 have been shown to exhibit mono-
ADP-ribosyltransferase activity (Aguiar, et al., 2005; Di Paola, et al., 2012; Kleine, et al., 2008;
Leung, et al., 2011). TiPARP (ARTD14) has previously demonstrated to be a catalytically active
enzyme; however, whether this catalytic activity was mono-ADP-ribosylation was not
determined (Ma, et al., 2001). We demonstrated that TiPARP is a mono-ADP-ribosyltransferase
that ADP-ribosylates itself and core histones (section 6). Our results indicate the catalytic
activities of TiPARP are dependent on the catalytic histidine and tyrosine of its HYI motif,
because mutation of these residues abolished auto- and hetero-ADP-ribosylation activities. Auto-
ADP-ribosylation of TiPARP is not affected by the presence of activated DNA or RNA, which is
similar to previous reports for ARTD7 and ARTD10 (Aguiar, et al., 2005; Kleine, et al., 2008).
These results suggest TiPARP is a nucleic acid-independent mono-ADP-ribosyltransferase.
We used two assays to examine the auto-ribosylation activity of TiPARP, the 32P-NAD+ assay
and the 35S protein shift assay. The 32P-NAD+ assay uses purified TiPARP incubated with 32P-
labelled NAD+ is an established methodology to detect catalytic activity of ARTDs by indicating
substrate incorporation and shift in mobility by SDS-PAGE (Shah, et al., 2011). The 35S protein
shift assay, uses in vitro translated 35S-labelled TiPARP to detect changes in mobility of 35S-
TiPARP following incubation with or without NAD+ by SDS-PAGE. This small shift in mobility
(or smearing) that we observed for 35S-TiPARP when incubated with β-NAD+ could be
indicative of short oligo-ADP-ribose polymers, dimers or trimers, or mono-ADP-ribose on
multiple residues of TiPARP (Lindahl, et al., 1995; Satoh, et al., 1994). The actual polymer sizes
of auto-ADP-ribosylated TiPARP can be analyzed by sequencing gel (Alvarez-Gonzalez and
Jacobson, 1987; Tanaka, et al., 1978).
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For all variants and species of TiPARP as well as conditions tested (with nucleic acids) results
from both 32P-NAD+ and 35S protein shift assays were in agreement except for the catalytic
isoleucine mutant (I631A) and chicken TiPARP. Results from the 32P-NAD+ assay demonstrated
I631A mutant to be catalytically active while the 35S protein shift assay did not show a mobility
shift when incubated with NAD+. This discrepancy may be due to sensitivity differences between
the two assays with the 32P-NAD+ assay being more sensitive than the 35S protein shift assay. A
mutation of the catalytic isoleucine of ARTD10 to a glutamate greatly reduced auto-ADP-
ribosylation activity but did not abolish it (Kleine, et al., 2008). The 35S protein shift assay may
not be sensitive enough to detect a reduction in the level of TiPARP catalytic activity. This
hypothesis may also explain the discrepancy between the two assays with respect to chicken
TiPARP. We observed weaker auto-ADP-ribosylation activity of chicken TiPARP than
mammalian TiPARPs by the 32P-NAD+ assay but no apparent activity in the 35S protein shift
assay, suggesting that sensitivity may be limited in this assay.
We used the 35S protein shift assay to determine whether TiPARP contains an auto-ribosylation
domain. We observed auto-ADP-ribosylation activity was greatly reduced between residues 275
and 328 of TiPARP suggesting the presence of an auto-ribosylation domain. The BRCT (BRCA1
C-terminus-like) domain of ARTD1 is generally referred to as the auto-ribosylation (or auto-
modification) domain (Loeffler, et al., 2011). Careful dissection of the ARTD1 auto-ribosylation
domain revealed three specific residues that are modified (one aspartate and two glutamates)
(Tao, et al., 2009). Other reports have described lysines within the auto-ribosylation and DNA
binding domain to be target residues for modification (Altmeyer, et al., 2009). A recent study
identified 12 auto-ribosylation sites of ARTD1, namely glutamates, aspartates and lysines, 8 of
these auto-ribosylation sites were found outside of the so-called auto-ribosylation domain
(Chapman, et al., 2013). Based on these findings auto-ribosylation of TiPARP may not be
restricted to a specific domain but is rather dispersed throughout the enzyme on multiple
residues. It has been suggested that there are preferred sites of auto-ribosylation of ARTD1 and
when these preferred or primary sites are unavailable secondary sites become prevalent (Tao, et
al., 2009). Results from our 35S mobility shift assay may suggest TiPARP may contain multiple
primary and secondary sites of auto-ribosylation throughout the protein. Currently it is not clear
which residues of ARTD family members serve as modification sites. Glutamate and aspartate
residues were reported to be auto-ribosylation sites on ARTD10 (Kleine, et al., 2008; Rosenthal,
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et al., 2013). Other residues that have been reported to be modified by eukaryotic MARTs
include arginine, cysteine, phosphoserine and asparagine and together with glutamate and
aspartate represent potential auto-ribosylated sites on TiPARP (Corda and Di Girolamo, 2003;
Okazaki and Moss, 1996; Ord and Stocken, 1977; Seman, et al., 2004; Smith and Stocken,
1975). Our laboratory is currently determining specific sites of auto-ADP-ribosylation of
TiPARP by mass spectrometry. We have identified 6 distinct peptides that are mono-ADP-
ribosylated. These findings confirm the mono-ADP-ribosyltransferase activity of TiPARP, but
we have not mapped or identified the modified amino acid residues. We have initiated electron
dissociation transfer mass spectrometry to specifically identify the ribosylated residues.
We have demonstrated that TiPARP possesses hetero-ADP-ribosylation activity and ADP-
ribosylates core histones. ARTD1, 2, 3 and 10 have also been described so far to ADP-ribosylate
histones (Kleine, et al., 2008; Messner, et al., 2010; Rulten, et al., 2011). Histone ribosylation
links ADP-ribosylation to other modifications such as acetylation, methylation and
phosphorylation, which constitute an epigenetic code for histones (Messner and Hottiger, 2011).
The contribution of histone ribosylation to the histone code and epigenetic regulation is not well
known. Moreover, ribosylated histones only constitute a small fraction (< 1%) of total histones
(Boulikas, 1989). Recently, macrodomain-containing proteins such as ARTD8, ARTD9,
MacroD1, MacroD2 and C6orf130 have been reported to interact with ADP-ribosylated target
proteins and represent potential ‘readers’ of ADP-ribosylation (Jankevicius, et al., 2013;
Rosenthal, et al., 2013; Sharifi, et al., 2013). MacroD1, MacroD2 and C6orf130 have been
identified as mono-ADP-ribosylhydrolases that bind mono-ADP-ribose and mediate its removal
from target proteins (Rosenthal, et al., 2013). Potential ‘readers’ and mono-ADP-
ribosylhydrolases of ADP-ribosylated histones by TiPARP have not been determined, but would
be an interesting future direction as it will help expand our understanding of the function of
histone ribosylation.
11 TiPARP is a novel negative regulator of AHR transactivation
We determined that TiPARP is a novel negative regulator of AHR transactivation. Our studies
using human cell lines and MEFs isolated from wildtype and Tiparp-deficient mice reveal that
TiPARP modulates ligand-induced proteolytic degradation of AHR and functions as a repressor
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of AHR transactivation. In this model, TCDD-activated AHR induces TiPARP expression,
TiPARP is recruited to AHR target genes either through direct DNA binding or tethering through
AHR, increased levels of TiPARP enhance the proteolytic degradation of AHR, by either direct
ribosylation or through the ribosylation of unidentified intermediary factor(s) to repress AHR
target gene transcriptional activation (Figure 41).
Figure 41. Proposed model of repression of AHR transactivation by TiPARP.
The AHR mediates the TCDD-induced expression of TiPARP (1). TiPARP is recruited to the
AHR/ARNT-bound AHRE of targets to repress transcription (CYP1A1 in this example) (2A).
TiPARP represses its own transcription and the regulation of other AHR target genes by binding
to AHR/ARNT-bound AHRE or potentially through direct DNA binding, creating a negative
feedback loop (2B) in which AHR dissociates from AHRE, translocates from the nucleus and is
proteolytically degraded by the 26S proteasome (26S) (3). BTF; basal transcription factors, Co-
act; co-activators.
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11.1 Potential co-repression by ADP-ribosylation of AHR The ability of TiPARP to repress AHR transactivation requires its catalytic domain and the direct
mono-ADP-ribosylation of AHR by TiPARP is an obvious mechanism to consider. Previous
studies have reported that ARTDs act as co-regulators (co-activators or co-repressors) for a
number of different transcriptional regulators (see section 2.5.4 for further information). In some
cases, enzymatic activity is required while in others it is not. ADP-ribosylation of Oct-1 and Sp1
electrostatically repels them from DNA resulting in altered transcript expression profiles (Gagne,
et al., 2008; Nie, et al., 1998; Smith, 2001). ADP-ribosylation of p53 and NF-κB prevents their
association with other nuclear factors, which also affected gene expression (Kanai, et al., 2007;
Zerfaoui, et al., 2010). It is possible that mono-ADP-ribosylation of AHR by TiPARP prevents
AHR binding to AHREs or its association with co-regulatory proteins. AHR that is not
complexed with ARNT is susceptible to degradation which could explain the changes in AHR
protein levels we observe when TiPARP expression altered (Davarinos and Pollenz, 1999; Heid,
et al., 2000; Kazlauskas, et al., 2000; Shetty, et al., 2003). The impact of mono-ADP-
ribosylation on transcription factor activity is not well understood and it is unclear whether
mono-ADP-ribosylation can alter transcription factor function by the same mechanism or extent
as poly(ADP-ribose). However it should be noted that mono-ADP-ribosylation of a single amino
acid residue by bacterial toxins, such as diphtheria and cholera toxins, can drastically alter target
protein function leading to severe physiological consequences (Collier, 2001).
The potential that ADP-ribosylation represents a novel posttranslational modification of AHR is
extremely exciting. AHR is subject to posttranslational modifications including SUMOylation,
ubiquitination and phosphorylation (Chen and Tukey, 1996; Pongratz, et al., 1991; Xing, et al.,
2012). Phosphorylation of specific serine and tyrosine residues of AHR is important for DNA-
binding and maximal transactivation, demonstrating modification of AHR affects its function
(Chen and Tukey, 1996; Park, et al., 2000; Pongratz, et al., 1991). Therefore ADP-ribosylation
of AHR could alter many processes involved in the AHR transactivation such as ARNT
dimerization, DNA-binding, association with co-regulatory or basal transcription factors, nuclear
import/export and transactivation. Because the structure and function of the modular domains of
AHR are well understood, mapping of potential mono-ADP-ribosylation sites by TiPARP will be
very informative in determining the effects ADP-ribosylation has on its function. We are
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currently optimizing purification of AHR protein to determine if it is hetero-ribosylated by
TiPARP and if so identify ADP-ribosylation using in vitro assays and mass spectrometry.
11.2 Potential targeting of histones and other co-regulatory factors
Core histones, linker histone H1 and chromatin-associated proteins are all targets of ADP-
ribosylation by ARTD1 (Krishnakumar and Kraus, 2010b). ADP-ribosylation (mono- or poly-)
of H1, H2A and H2B by ARTD1 may play a role in the regulation of chromatin structure,
although the general extent of histone modification to transcriptional regulation remains to be
clarified (D'Amours, et al., 1999). ARTD1 can displace the H1 by ADP-ribosylation and block
binding of H1 to promoter chromatin creating a structure permissive to transcription (Ju, et al.,
2004; Kim, et al., 2004; Krishnakumar, et al., 2008). The hetero-ADP-ribosylation of core
histones by TiPARP is a potential mechanism by which TiPARP modulates chromatin structure
at active AHR regulatory regions to repress transcription. TiPARP could also target co-
regulatory proteins with histone-modifying activities that are known to interact with AHR/ARNT
(see Table 3) altering their intrinsic histone-modifying or chromatin-remodelling activities by
ADP-ribosylation. Many of the known AHR co-activators have histone acetyltransferase (HAT)
activity such as NCoA1, 2, 3 and p300 (Hankinson, 2005; Harper, et al., 2006). Histone ADP-
ribosylation by TiPARP could reduce co-activator activity by competing with or blocking
histone acetylation and thus reducing transactivation. Poly-ADP-ribosylation of histone lysine
demethylase KDM5B has been shown to block binding of KDM5B to chromatin and inhibit its
demethylase activity supporting the hypothesis that hetero-ADP-ribosylation by TiPARP may
reduce AHR co-activator function (Krishnakumar and Kraus, 2010a). TiPARP could also
modulate other chromatin-associated proteins to alter chromatin to architecture more favourable
for repression. ARTD1 also hetero-ribosylates DEK, a chromatin-associated protein and
promotes its release from chromatin suggesting TiPARP may potentially modulate other
chromatin-associated proteins besides co-activators to alter chromatin to architecture in favour of
repression (Gamble and Fisher, 2007; Kappes, et al., 2008). Proteomics will be very useful to
identify interacting proteins with TiPARP and provide insights into the mechanism of TiPARP-
mediated repression.
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The auto-ribosylation activity of TiPARP may also be important for its repressive function.
Auto-ribosylation accounts for approximately 90% of ARTD1 total catalytic activity indicating
the enzyme itself is the primary target (D'Amours, et al., 1999). Auto-ribosylated ARTD1
provides a binding platform for DNA repair enzymes and scaffolding proteins (Gagne, et al.,
2008; Hegde, et al., 2008). Therefore it is possible that auto-ribosylated TiPARP recruits co-
repressor proteins including known AHR co-repressors SMRT, SHP or AHRR to promote
transcriptionally repressive complex assembly at the AHREs of target genes (Harper, et al.,
2006; Mimura, et al., 1999; Nguyen, et al., 1999). Alternatively, it is equally possible that auto-
ribosylated TiPARP at AHREs could also prevent the association of AHR/ARNT or basal
transcription factors with co-activators. Identification of potential auto-ribosylation sites on
TiPARP will help clarify their importance in mediating the repressive function of TiPARP.
11.3 TiPARP is a candidate AHR degradation promoting factor (ADPF)
Our proposed model of TiPARP-mediated repression (Figure 41) is reminiscent of the ADPF
hypothesis which proposes the induction of a labile factor that is sensitive to proteasomal and
protein synthesis inhibition and promotes the proteasomal degradation of AHR (Ma and
Baldwin, 2000; Ma and Baldwin, 2002; Ma, et al., 2000). We show by use of transcriptional and
proteasomal inhibitors that TiPARP mRNA and protein is rapidly degraded indicating TiPARP is
labile. Protein synthesis inhibition of Tiparp-/- MEFs did not result in superinduction of AHR
target genes when cells were treated with TCDD which suggested newly generated TiPARP
represses AHR transactivation. We also observed TiPARP knockdown or knockout increases
AHR protein levels and transactivation mimicking the effects of protein synthesis and
proteasomal inhibition (Ma and Baldwin, 2000). Overall, our data support TiPARP as a
candidate labile ADPF.
The ubiquitin-proteasome pathway plays an important role in TCDD-mediated degradation of
AHR (Ma and Baldwin, 2000; Pollenz, 2007). ARTD1 plays a role in the reactive oxygen-
induced activation of the proteasome (Ullrich, et al., 1999). During oxidative stress auto-
ribosylated ARTD1 interacts with and activates nuclear proteasome pathways leading to the
rapid up-regulation of 20S proteasome activity to remove damaged histones more efficiently
(Ullrich, et al., 1999). Inhibition of ARTD5 and ARTD6 activity prevents the degradation of
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axin through the ubiquitin-proteasome pathway and this increased half-life of axin promotes the
β-catenin degradation (Mayer-Kuckuk, et al., 1999). These findings demonstrate there is
interplay between the nuclear proteasome proteins and poly-ADP-ribosylation. It is therefore
tempting to hypothesize TiPARP activates components of the proteasome to enhance the TCDD-
mediated AHR degradation. It is also equally possible that TiPARP directly ADP-ribosylates
AHR, targeting it for degradation. Iduna is a poly(ADP-ribose)-dependent E3 ubiquitin ligase
that binds and ubiquitinates both poly-ADP-ribosylated and poly(ADP-ribose)-binding proteins
labelling these proteins for ubiquitin proteasomal degradation and provides an example of how
ADP-ribosylation can be used to target proteins for degradation (Kang, et al., 2011).
11.3.1 TiPARP and the nuclear proteasome
TiPARP localizes to the nucleus and exhibits a focal expression pattern, which can be described
as nuclear bodies (von Mikecz, 2006). It has been established that proteasomes occur in the
nucleus and are associated with nuclear bodies including PML bodies, nucleoplasmic speckles
and focal clusters throughout the nucleoplasm representing proteolytic centres in the nucleus
(Chen, et al., 2002; Fabunmi, et al., 2001; Rockel, et al., 2005). TiPARP expression is stabilized
by proteasomal inhibition resulting in its accumulation in the nucleus (data not shown) and
suggests that TiPARP is subject to proteolytic degradation via the 26S proteasome. All TiPARP
catalytic mutants and cytosolic N-terminal truncations and point mutants (both zinc finger and
NLS mutants) are more highly expressed compared with wildtype indicating that components
within the nucleus and the catalytic activity of TiPARP are responsible for its instability.
Because TiPARP affects AHR protein levels and AHR interacts with TiPARP within the nuclear
foci it is reasonable to hypothesize that TiPARP associates with nuclear bodies containing
proteasomes to target AHR for proteasomal degradation. The catalytic mutant TiPARP-H532A
focal expression pattern was greatly altered but still interacted with AHR. This suggests that
catalytic activity of TiPARP is required for association with proteolytic centres in the nucleus to
degrade AHR, repressing its transactivation. The effects of overexpression of TiPARP catalytic
mutants on AHR protein levels will help clarify whether catalytic activity is required for the
proteolytic degradation of AHR. Together, our findings indicate TiPARP is associated with the
nuclear proteasome and this association is dependent upon its catalytic activity. Currently, we are
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determining which nuclear body types are associated with TiPARP to provide evidence that
TiPARP is associated with the nuclear proteasome.
11.3.2 Connecting nuclear localization with repression of AHR transactivation
We identified a functional NLS of TiPARP within its N-terminus between residues 41-47
(Figure 42). The zinc finger domain is also important for nuclear localization of TiPARP. Zinc
finger motifs have been identified as the NLS of several zinc finger proteins (Kuwahara, et al.,
2000; Matheny, et al., 1994; Racca, et al., 2011; Spittau, et al., 2007). Mutation or removal of
either the NLS or zinc finger results in the cytosolic localization of TiPARP. The purpose of
having two different NLSs is currently not known. One significant difference between the NLS
and zinc finger is the NLS mutants still repress AHR transactivation whereas zinc finger mutants
do not. These findings suggest repression is not dependent on nuclear localization but rather zinc
finger function. Zinc finger mutants still interact with AHR presumably in the cytosol indicating
the zinc finger is important for repression and potential proteasomal degradation of AHR. Zinc
finger domains of proteins are known to play a role in protein-protein interactions and thus the
zinc finger domain of TiPARP could play a role in the association with the cytosolic proteasome
rather than with AHR (Kang, et al., 2004; Zhang, et al., 2011). Determining interacting partners
by proteomics and AHR protein levels following overexpression of zinc finger TiPARP mutants
can be used to test this hypothesis.
All C-terminal truncations of TiPARP are nuclear but do not repress AHR transactivation. In
constrast, N-terminal truncations 53-657 to 225-657 are cytosolic and still repess AHR activity.
TiPARP truncation 200-657 which is cytosolic and functional and interacts with AHR in the
cytosol. Because AHR exists in the nucleus and cytosol this observation is not completely
unexpected. It is possible the TiPARP 200-657 truncation interacts with the cytosolic proteasome
to degrade AHR and repress transactivation, representing an alternative pathway to nuclear AHR
degradation. Determining AHR protein levels in cells overexpressed with TiPARP 200-657 will
clarify this alternative pathway. The association with AHR and repression of transactivation by
TiPARP 200-657 may also be independent of proteasomal degradation and instead TiPARP 200-
657 prevents the nuclear translocation of AHR. As far as we know all the truncations and
mutants we created do not exist endogenously in cells and their activity, or lack thereof, is
observed experimentally using overexpression but may not occur biologically. During apoptosis
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ARTD1 is cleaved into 24 and 89 kDa fragments and each fragment is functional (Chaitanya, et
al., 2010). The 24 kDa fragment containing two of the zinc fingers localizes to the nucleus and
binds DNA irreversible and the 89 kDa contains the auto-modification and catalytic domains and
retains its catalytic activity but localizes to the cytosol (D'Amours, et al., 2001; Soldani, et al.,
2001). Therefore it is also not completely unexpected that truncated cytosolic TiPARP variants
repress AHR and that potential endogenous TiPARP truncations may exist and exhibit a similar
ability to repress AHR.
Figure 42. Updated schematic of the structure of TiPARP.
NLS; nuclear localization signal, Zn; CCCH-type zinc finger domain, WWE; tryptophan-
tryptophan-glutamate.
11.4 Identification of an AHR repressive domain The use of N-terminal truncations identified a putative AHR repressive domain between residues
218 and 245 (Figure 42). Alanine scan results narrowed this region to between residues 220 and
235. Alanine scanning also identified tyrosine 232 (Y232) and histidine 233 (H233) to be
important for repression of AHR transactivation. The exact function and contribution of Y232
and H233 to repression is not clear and we can only speculate their roles. Both tyrosine and
histidine residues are phosphorylated which can alter protein function and conformation (Miller,
2012; Steeg, et al., 2003). For example, AHR phosphorylation at tyrosine residues is important
for its AHRE-binding activity (Minsavage, et al., 2003; Park, et al., 2000). Therefore it is
possible that phosphorylation of Y232 and/or H233 is important for TiPARP-mediated
repression of AHR transactivation. Tyrosines are commonly sulfated which strengthens protein-
protein interactions, Y232 may also be important for stabilizing interactions with intermediary
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proteins involved in the repression of AHR (Seibert and Sakmar, 2008). It is unlikely that Y232
or H233 are important for interaction with AHR since the TiPARP 275-657 truncation interacted
with AHR. Given the proximity of H233 to the zinc finger domain (four residues away) and the
fact that histidine is commonly involved in coordinating the zinc ion of zinc fingers, H233 could
be involved with zinc finger function and its loss resembles a zinc finger mutation (Alberts, et
al., 1998).
12 Comparing TiPARP- with AHRR-mediated transrepression
Much of our understanding of AHR repression comes from studies of AHRR, which is also part
of negative feedback loop regulating AHR function (Evans, et al., 2008; Karchner, et al., 2009;
Mimura, et al., 1999). Despite its discovery more than ten years ago and several reports from
independent laboratories, the precise mechanism of AHRR-mediated transrepression of AHR is
not well understood. Conflicting reports between independent laboratories has been a major
contributing factor for our lack of understanding of the AHRR. We compared the AHR
repressive functions of TiPARP to the AHRR to better understand the mechanisms of negative
regulation of AHR transactivation.
12.1 Gene expression and recruitment As expected, both TiPARP and the AHRR gene expression are induced by TCDD treatment. The
AHRR expression gradually increased over 24 h, whereas TiPARP induction peaked after 1.5 h
and declined thereafter. In line with gene expression kinetics, we detected endogenous AHRR
protein and its recruitment to endogenous AHRE-containing regulatory regions after 24 h TCDD
treatment. This recruitment can be supported by previous reports that used in vitro methods to
determine AHRR interaction with AHRE-containing sequences (Evans, et al., 2008; Mimura, et
al., 1999). Recruitment of the AHRR to AHR target regulatory regions coincides with reduced
AHR/ARNT recruitment, suggesting that AHRR displaces AHR/ARNT binding which is similar
to a previous study that describes the AHRR recruitment to an AHRE-containing regulatory
region of AHRR correlated with AHR/ARNT recruitment loss (Haarmann-Stemmann, et al.,
2007). Similarly, we observed TCDD-dependent recruitment of GFP-TiPARP and AHRR to the
CYP1A1 regulatory region suggesting that both TiPARP and the AHRR can influence the
AHR/ARNT complexes at AHREs. The mechanism of how the AHRR and TiPARP alter
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AHR/ARNT complexes at AHREs may differ. TiPARP alters AHR protein levels and the
enhanced AHR proteolytic degradation which could lead to reduced AHR recruitment to its
target genes. The AHRR does not affect the proteolytic degradation of AHR therefore the
reduced AHR/ARNT recruitment we observed after 6 h and 24 h is not likely due AHR
degradation (Evans, et al., 2008).
The direct AHRE-binding of the AHRR is not involved in the mechanism of repression of the
AHRR. This is supported by studies using an AHRR DNA-binding mutant that localizes to the
nucleus and still substantially represses AHR transactivation (Evans, et al., 2008). Direct AHRE-
binding is also not essential for TiPARP-mediated repression as TiPARP NLS mutants and N-
terminal truncations from residue 53 to residue 225 still repress AHR-regulated reporter gene
activity despite localizing to the cytosol. Both TiPARP and the AHRR can directly interact with
AHR; therefore, it is possible that TiPARP and the AHRR directly interact with AHRE-bound as
well as unbound AHR to repress transactivation.
12.2 Overexpression of ARNT and co-regulators Overexpression of AHR but not ARNT significantly reduces repression by AHRR or TiPARP,
suggesting that AHR rather than ARNT is the primary target of both repressors (Evans, et al.,
2008; Karchner, et al., 2009). The lack of involvement of ARNT does not support the original
model AHRR-mediated repression, which proposed the AHRR competes with AHR for ARNT
heterodimerization and AHRE binding (Mimura, et al., 1999). Additionally, overexpression of
many known AHR co-activators did not rescue the inducibility of AHR-mediated reporter gene
activity indicating competition with co-activators is not involved in the AHRR mechanism of
repression. Histone deacetylases 4 and 5 (HDAC4 and HDAC5) are co-repressors that are
associated with the AHRR perhaps recruitment of co-repressors is a mechanism by which the
AHRR can negatively repress AHR transactivation (Oshima, et al., 2009; Oshima, et al., 2007).
12.3 Subcellular localization Previous reports have described that the AHRR is a nuclear protein with an active NLS located
within its N-terminus (Evans, et al., 2008; Kanno, et al., 2007). However we observe GFP-
tagged AHRR constructs to localize to the nucleus in a cell context-dependent manner. Only in
MCF7 cells did we observe the nuclear localization of the AHRR, AHRR localized to the cytosol
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in HuH7 and COS7 cells. TiPARP localized to the nucleus in all three cell lines. The reason for
the different localization patterns of the AHRR among the cell lines is currently not known.
ARNT expression has been proposed to influence the localization of the AHRR (Kanno, et al.,
2007). The expression of an AHRR NLS mutant in COS7 cells was shifted from the cytosol to
the nucleus when ARNT was co-overexpressed suggesting ARNT expression can influence
AHRR localization (Kanno, et al., 2007). We observed that AHRR localizes to the cytosol in
both COS7 cells which have low levels of ARNT and in HuH7 cells which express ARNT, so it
does not appear that ARNT expression influences the localization of the AHRR. However, we
did not co-overexpress ARNT with the AHRR in COS7 and HuH7 cells therefore it is possible
that overexpressed ARNT influences subscellular localization of the AHRR. Despite the
difference in localization of the AHRR, the AHRR is a potent repressor of AHR transactivation
in all three cell lines (Evans, et al., 2008; Karchner, et al., 2009). This observation is similar to
that observed with the TiPARP NLS mutants and N-terminal truncations from residue 53 up to
residue 225 that repress AHR transactivation but localize to the cytosol. The AHRR has
previously been reported to not alter nuclear translocation of AHR by comparing the nuclear
translocation of AHR when AHRR was co-overexpressed although these reports did not examine
AHR and AHRR co-localization. Although the subcellular localization of endogenous AHRR
has not yet been determined, our recruitment findings suggest that a proportion of endogenous
AHRR is nuclear.
12.4 Knockdown and knockout models Our knowledge on the AHRR function is largely based on overexpression models, and when
overexpressed the AHRR is an effective repressor of AHR-regulated reporter gene activity.
However, findings from AHRR overexpression studies are not fully supported by knockout
models (Hosoya, et al., 2008; Tigges, et al., 2013). Previous studies that have reported that cell
lines that have low AHR ligand-dependent inducibility of CYP1A1 have high basal levels of
AHRR and knockdown of the AHRR increases basal CYP1A1 expression and AHR ligand
inducibility (Haarmann-Stemmann, et al., 2007; Oshima, et al., 2007). We observed the
constitutive levels of CYP1A1 mRNA in MCF7 cells were not significantly affected by AHRR
knockdown. Our results are supported by a recent report that describes a non-causal relationship
between AHRR expression and CYP1A1 induction and activity human skin fibroblasts (Tigges,
et al., 2013). Reports using fibroblasts isolated from Ahrr-deficient mice demonstrated
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impressive CYP1A1 induction by AHR ligands which suggest that the knockdown levels we
achieved in MCF7 cells were not sufficient to significantly increase inducibility of AHR target
genes (Hosoya, et al., 2008; Tigges, et al., 2013). Alternatively, given that the increased
CYP1A1 expression levels by AHR ligand treatment was limited to the heart, spleen and skin of
Ahrr-deficient mice our choice of cell type (MCF7 breast carcinoma) may have limited results.
TiPARP knockdown in all cell types that we examined which includes two human breast
carcinoma, hepatoma and gastric carcinoma lines and the Tiparp-deficient MEFs displayed
significantly greater AHR target inducibility which suggests TiPARP to be a general negative
regulator of AHR transactivation and the AHRR to be a tissue/cell type-specific repressor.
Interestingly, Ahrr-deficient MEFs demonstrated significantly greater constitutive AHR mRNA
levels compared to wildtype MEFs (Tigges, et al., 2013). In contrast, Tiparp-deficient MEFs do
not display significantly different basal AHR mRNA levels and altered transcriptional expression
of the AHR represents a potential difference between the modes of repression of TiPARP and
AHRR.
12.5 Potential overlap of TiPARP and the AHRR repressive pathways
A comparison of what is known about the AHR repressive functions of TiPARP and the AHRR
reveals similarities but also some notable differences. Similarities between TiPARP- and AHRR-
mediated inhibition of AHR transactivation include: (1) both genes are induced by TCDD and
regulated by AHR; (2) both interact with AHR; and (3) increased ARNT expression fails to
prevent AHRR- or TiPARP-dependent repression of AHR transactivation (Evans, et al., 2008;
Karchner, et al., 2009; Lo, et al., 2011; Mimura, et al., 1999). In contrast, AHRR repression of
AHR action does not involve increased AHR degradation, direct binding to AHREs, or inhibition
of the AHR nuclear translocation (Evans, et al., 2008; Hahn, et al., 2009). These similarities and
differences suggest there could be pathway overlap and interplay between TiPARP and the
AHRR.
The kinetic profiles of TCDD-regulated expression of TiPARP and AHRR suggest that TiPARP
induction could be an immediate mechanism to repress AHR transactivation, while the AHRR is
a more long-term mechanism of repression. We observe the expression and recruitment of
TiPARP after 45 min of TCDD treatment, whereas the expression and recruitment of the AHRR
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occurs after 24 h TCDD treatment. The rapid induction and recruitment of TiPARP can be a
primary response to repress AHR and the more gradual induction of AHRR expression and
recruitment to AHR target genes may represent a secondary mechanism of repression. TiPARP
knockdown and knockout increased TCDD-induced expression of AHRR mRNA and other AHR
target genes. Initially this might seem counterintuitive since greater expression of AHRR should
decrease AHR target expression rather than enhance it; however, this could indicate that AHRR
requires TiPARP for optimal repression. Because in the absence of TiPARP the increase of
AHRR expression alone is not sufficient to repress AHR target induction this suggests TiPARP
and the AHRR function in tandem or co-operatively. Both TiPARP and AHRR can directly
interact with AHR and overexpression of AHR can restore induction of AHR targets when co-
overexpressed with either TiPARP or AHRR suggesting that AHR is a common target of both
repressors (Karchner, et al., 2009). Our finding that TCDD induction of AHR targets in Tiparp-
deficient MEFs with protein synthesis inhibition was approximately 2-fold greater than TCDD
treatment alone indicates that TiPARP induction is not solely responsible for the observed
repression and that inhibition of AHRR synthesis might also contribute to the increases. Double
knockout of TiPARP and AHRR will be very helpful in determining potential co-operatively
between TiPARP and the AHRR to repress AHR transactivation.
13 TiPARP is an in vivo negative regulator of AHR transactivation
Hepatic tissue from Tiparp-deficient mice show significantly greater TCDD-induced AHR target
gene expression compared with tissue isolated from wildtype mice, supporting that TiPARP is an
in vivo negative regulator of AHR transactivation. This AHR target induction profile was not
observed in hepatic tissue from 3MC-treated Ahrr-deficient mice (Hosoya, et al., 2008).
Increased CYP1A1 induction of Ahrr-deficient mice was only observed in spleen, heart and skin
tissues, which indicates that AHRR is a tissue-specific repressor (Hosoya, et al., 2008).
Preliminary results indicate that kidney and spleen tissue from 6 h TCDD-treated Tiparp-
deficient mice show a similar AHR target induction profile to hepatic tissue suggesting TiPARP
to potentially be a more wide-spread repressor than the AHRR (data not shown). The tissue
distribution and increased induction CYP1A1 of Tiparp-deficient mice is similar to CA-AHR
transgenic mice which display heightened CYP1A1 mRNA in many tissues including liver,
kidney and spleen (Andersson, et al., 2002; Brunnberg, et al., 2006; Moennikes, et al., 2004).
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However it should be noted that beyond CYP1A1, the gene induction profile of hepatic tissue
from CA-AHR mice is very different from hepatic tissue isolated from TCDD-treated wildtype
mice and most of the TCDD-responsive genes, including TiPARP, were not detected in CA-
AHR mouse livers (Dere, et al., 2006; Kurachi, et al., 2002; Moennikes, et al., 2004; Zeytun, et
al., 2002). TCDD-treated Tiparp-deficient mice demonstrate significantly greater induction of
many TCDD-responsive genes indicating that uninhibited AHR transactivation by the lack of
repression of AHR is different from continuous AHR activation as observed in the CA-AHR
mouse model.
13.1 Role of TiPARP in TCDD-mediated toxicity
13.1.1 Purported effects of TiPARP on the NAD+ pool and SIRT activity
Our knowledge of a role of TiPARP in TCDD-mediated toxicity has been limited to studies that
use chicken (Gallus gallus) TiPARP to examine the effects of TiPARP on the suppression of
gluconeogenesis that is associated with wasting syndrome (Diani-Moore, et al., 2010; Diani-
Moore, et al., 2013). Chicken TiPARP was identified as a mediator of the TCDD-induced
suppression of gluconeogenesis that consumes hepatic NAD+, lowering SIRT activity, repressing
gluconeogenic genes G6Pase and PEPCK, and reducing glucose output (Diani-Moore, et al.,
2010). TiPARP induction was also purported to be involved in AHR-mediated sensitivity to
NASH by depleting NAD+ levels which lowers SIRT3 activity leading decreased SOD2 activity
and less superoxide scavenging (He, et al., 2013). Conclusions from both of these studies were
largely based observations that TCDD treatment increases TiPARP gene expression and
decreases NAD+ levels suggesting that NAD+ levels control gene expression (Diani-Moore, et
al., 2010; He, et al., 2013). We considered whether lower NAD+ levels rather than TiPARP
directly could be responsible for the repression of AHR transactivation; however, addition of
nicotinamide, a NAD+ precursor also represses induction of AHR targets indicating that NAD+
levels and TiPARP-mediated repression of AHR targets are independent (Diani-Moore, et al.,
2010).
TiPARP was found to contribute to TCDD-mediated suppression of gluconeogenesis because
TCDD treatment induces TiPARP mRNA and decreases gluconeogenic gene expression and
subsequently reduces glucose output (Diani-Moore, et al., 2010). TiPARP overexpression
reduces PEPCK mRNA and glucose output, mimicking the effects of TCDD, leading the authors
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to conclude that TiPARP mediates TCDD-induced suppression of gluconeogenesis (Diani-
Moore, et al., 2010). If TiPARP is a mediator of TCDD toxicity, Tiparp-deficient mice should be
more resistant to TCDD-induced gluconeogenesis suppression and wasting; however,
preliminary studies suggest that Tiparp-deficient mice are more sensitive to TCDD suggesting
TiPARP plays a protective rather than a mediator role in TCDD-mediated toxicity (data not
shown). Toxicity studies with Tiparp-deficient mice are currently ongoing.
Chicken TiPARP is reported to be a poly-ADP-ribosylating ARTD that is able to deplete the
cellular NAD+ pool (Diani-Moore, et al., 2010; Diani-Moore, et al., 2013). We have
demonstrated using catalytic assays that chicken TiPARP is a mono-ADP-ribosyltransferase,
which agrees with the structural prediction of its catalytic activity (Otto, et al., 2005). The choice
of assay to demonstrate catalytic activity is most likely the reason for this discrepancy. We use
purified GST-tagged chicken TiPARP incubated with 32P-NAD+, the generally accepted method
to examine catalytic activity of a specific ARTD whereas Diani-Moore et al. used an indirect
method by immunoprecipitating poly(ADP-ribose) from whole cell extracts transfected with
TiPARP or treated with TCDD as an indirect means to study TiPARP catalytic activity (Diani-
Moore, et al., 2010; Diani-Moore, et al., 2013). The anti-poly(ADP-ribose) antibodies are able to
detect six or more ADP-ribose units and cannot detect mono-ADP-ribose accurately, therefore it
is unclear why an anti-poly(ADP-ribose) antibody is able to detect catalytic activity directly from
TiPARP (Kleine, et al., 2008). It is still unclear how TiPARP, which is a MART, is capable of
significantly depleting the NAD+ pool. ARTD1 accounts for 85-90% of cellular ADP-ribose
synthesis suggesting that TiPARP is not the primary consumer of cellular NAD+ (Shieh, et al.,
1998). The reported depletion of cellular NAD+ could be due to stress caused by transfection of
TiPARP which leads to the activation of ARTD1 and increase in poly(ADP-ribose) activity and
thus decreases cellular NAD+ levels. Additionally, we observe chicken TiPARP is less
catalytically active than mammalian TiPARPs making TiPARP less likely of a candidate in
chicken cells to substantially deplete the NAD+ pool.
Another difference between chicken TiPARP and mammalian TiPARPs is when transfected into
mammalian cells chicken TiPARP did not repress AHR transactivation or form nuclear foci. The
lack of activity could be due to cell context, since we transfected chicken TiPARP into
immortalized human cell lines. These findings suggest that TiPARP exhibits species- or model-
specific differences. Chicken TiPARP is also less catalytically active than mammalian TiPARPs
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illustrating another species difference between mammalian and chicken TiPARP. Since chickens
are more sensitive to TCDD than mice (LD50 <25 μg/kg bw for chickens versus 182 μg/kg bw
for B6 strain mice), it is possible that the lower TiPARP activity could be a potential contributing
factor to the higher TCDD sensitivity reported in chickens (Chapman and Schiller, 1985; Greig,
et al., 1973).
14 TiPARP-mediated regulation of other transcription factors
The ability of TiPARP to regulate transcription is most likely not limited to AHR. ARTD1 and
ARTD2 regulate many transcriptional factors acting as co-activators and co-repressors in
different pathways (section 2.5.4) (Krishnakumar and Kraus, 2010b; Szanto, et al., 2012).
Despite its name, the AHRR has also shown activity toward transcription factors other than AHR
including hypoxia inducible factor 1 alpha (HIF-1α) (Karchner, et al., 2009). TiPARP is
regulated by other transcription factors including estrogen receptor alpha (ERα) and
glucocorticoid receptor supporting the possibility that TiPARP could be involved in these
pathways (Kininis, et al., 2007; Reddy, et al., 2009). Ongoing studies in our laboratory reveal
that TiPARP interacts with ERα and negatively regulates ERα-mediated transactivation,
whereas TiPARP co-activates liver X receptor (LXR)-mediated transactivation (data not shown).
All together our results indicate that TiPARP is a multifunctional transcription factor co-
regulator.
15 Limitations and future considerations
15.1 TiPARP antibody Our studies of TiPARP function were highly dependent on transient transfection to overexpress
or knockdown TiPARP. We were unable to verify TiPARP protein overexpression, knockdown
of endogenous TiPARP or TCDD-induced endogenous TiPARP due to the lack of suitable
commercially available antibody able to detect TiPARP. To verify knockdown we resorted to
overexpressing GFP-tagged TiPARP with siTiPARP sequences and using an anti-GFP antibody
to detect GFP-TiPARP protein levels. Similarly, we overexpressed GFP-TiPARP to perform
ChIP assays and test GFP-TiPARP protein stability. Plasmid-derived overexpressed protein is
not identical to endogenous protein, which is why a high quality anti-TiPARP antibody is critical
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for studying the endogenous TiPARP protein behaviour. An anti-TiPARP antibody would also
be extremely useful to detect TCDD-induced TiPARP expression, recruitment to AHR target
regulatory regions and co-immunoprecipitation with endogenous AHR. Due to the lack of a
suitable antibody we were also unable to confirm complete knockout in the MEFs isolated from
Tiparp-deficient mice at the protein level, leaving open the possibility Tiparp-deficient mice to
express a truncated version of TiPARP. To date the only evidence we have of endogenous
TiPARP protein are confocal images of putative TiPARP staining in the nuclei of HuH7 cells by
a commercially available anti-TiPARP antibody that in our hands only works for
immunofluorescence. The specificity of the antibody was tested with overexpressing GFP-
TiPARP and staining for TiPARP (data not shown). We are currently purifying TiPARP protein
for generation of our own antibody. Once generated, we intend to use the antibody to detect and
confirm TiPARP protein expression in all our models. We also plan to use the antibody to
perform ChIP-sequencing to determine TiPARP recruitment and potential overlapping
recruitment with AHR and ARNT. A proper anti-TiPARP antibody will certainly be an
important tool for our laboratory.
15.2 ARTD catalytic activity assays The in vitro ADP-ribosylation assays, which involve incubation of purified ARTD protein and 32P-NAD+ are relatively simple and is a well-accepted method to demonstrate ARTD activity.
Although catalytic activity assays provide insights into auto- and hetero-ADP-ribosylation of
ARTDs, they do not identify specific sites/residues of ADP-ribosylation. Furthermore, although
ARTD catalytic assays do illustrate extensive poly-ADP-ribosylation with the reduced mobility
of modified protein by SDS-PAGE they do not provide information about polymer length.
Because the molecular weight of a single ADP-ribose moiety is quite small (540 Da) mono-
ADP-ribosylated and short oligo-ADP-ribosylated proteins are visually indistinguishable (Laing,
et al., 2011). Sequencing gels of modified proteins can be used to illustrate the lengths of ADP-
ribose chains. We used catalytic activity assays to demonstrate TiPARP has auto- and hetero-
ADP-ribosylation activities and TiPARP is most likely a mono-ADP-ribosyltransferase.
However, we did not quantify the amount of ADP-ribose units TiPARP is able to synthesize
leaving open the possibility that TiPARP can synthesis short oligo-ADP-ribose polymers that
display no visual slowing of migration by SDS-PAGE. Because catalytic activity assays are non-
quantitative, small differences in the level of activity may not be visually observed. For instance,
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the TiPARP-I631A and –I631E mutants are catalytically active; however, it is not clear if the
level of catalytic activity of either mutant differs from that of wildtype. Our GST-purified
TiPARP and mutant protein preparations contain breakdown products making quantification of
full-length enzyme difficult. To compare the activities of wildtype TiPARP and these isoleucine
mutants requires highly pure protein to ensure equal quantities of wildtype and mutants are used
for each assay. We are currently optimizing our protein purification protocol for TiPARP to
reduce breakdown products and increase purity.
Our catalytic assays with TiPARP also did not identify specific sites or residues on TiPARP that
were auto-ADP-ribosylated or sites of core histones that were hetero-ADP-ribosylated by
TiPARP. Determining the sites or residues of modification will demonstrate whether TiPARP
has a preference for a particular amino acid. We are currently using mass spectrometry to
determine sites of auto-ADP-ribosylation on TiPARP, by using mass spectrometry we are able to
identify specific residues of TiPARP that are auto-ADP-ribosylated and confirm that these sites
are in fact mono-ADP-ribosylated. Once we purify large enough quantities of AHR protein we
will determine whether it is hetero-ADP-ribosylated by TiPARP and use mass spectrometry to
identify specific sites of AHR that are mono-ADP-ribosylated.
In vitro catalytic assays may be artificial and may not accurately reflect modification in a cellular
context or in vivo. Direct evidence of intracellular protein mono-ADP-ribosylation is quite
limited due to the lack of antibody to detect mono-ADP-ribosylated proteins (Feijs, et al., 2013).
The challenge of providing direct evidence for intracellular mono-ADP-ribosylation is
compounded by the lack of specific mono-ARTD inhibitors (Feijs, et al., 2013). Although
TiPARP displays auto- and hetero-ADP-ribosylation in vitro our understanding of whether this
occurs in cell or in vivo is limited.
16 Overall significance of research The objective of this research was to understand TiPARP and its function in AHR
transactivation. We provided evidence that TiPARP is a nuclear MART that mono-ADP-
ribosylates itself and core histones. We identified TiPARP as a novel AHR ligand-induced
negative regulator of AHR transactivation in vitro and in vivo. We also identified its NLS
sequence providing further evidence for its nuclear localization. TiPARP specifically targets
AHR and promotes its proteolytic degradation and represents a candidate for the elusive ADPF.
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The catalytic and zinc finger functions are essential for TiPARP to repress AHR transactivation.
Comparison with the AHRR reveals some notable similarities and differences suggesting
TiPARP and the AHRR use partially overlapping but also distinct mechanisms to repress AHR
activity. Overall we have established a new mechanism of negative feedback regulation of AHR
transactivation and identified a nuclear mono-ADP-ribosyltransferase.
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