Effects of Arsenic on Dna Repair and Cell Cell Checkpoints: Involvement in Arsenic Co-Mutagenesis and Co-Carcinogenesis

UNIVERSITY OF CINCINNATI

Date:______

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

It is entitled:

This work and its defense approved by:

Chair: ______

EFFECTS OF ARSENIC ON DNA REPAIR AND CELL CELL CHECKPOINTS: INVOLVEMENT IN ARSENIC CO-MUTAGENESIS AND CO-CARCINOGENESIS

A dissertation submitted to the

Dvision of Research and Advanced Studies of the University of Cincinnati

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

in the Department of Environmental Health of the College of Medicine

2005

SHENGQIN LIU

M.S., Sun Yat-Sen University of Medical Sciences, 1998

Committee Chair: Kathleen Dixon, Ph.D.

ABSTRACT ABSTRACT

Arsenic is a well-documented human carcinogen associated with a variety of cancers, though the mechanism of action is unknown. Arsenic is also currently being used to treat some cancers. The paradoxical action of arsenic as both a carcinogen and a chemotherapeutic agent drug may be due to its ability to synergize with other genotoxic agents. Such synergy has been hypothesized to involve effects on DNA repair and/or cell cycle checkpoints.

Although arsenic is a carcinogen, attempts to prove arsenic carcinogenesis in animals models have been unsuccessful, leading to the hypothesis that arsenic might act as a co- carcinogen instead of a carcinogen itself. Consistent with a co-carcinogenic model, arsenic by itself does not induce mutations in most bacterial or mammalian systems tested, instead it potentiates the mutagenicity of other agents such as ultraviolet (UV), Benzo(a)Pyrene and

N-methyl-N-nitrosourea (NMU). While the mechanism of this co-mutagenicity remains unknown, a possible explanation is that arsenic inhibits DNA repair, specifically nucleotide excision repair (NER). To test this hypothesis, we have completed multiple experiments using low concentrations of arsenite in cultured human cells. Our results showed that at a concentration of 2.5 µM, arsenic induced insertion/deletion mutations in a mutagenesis model using a shuttle vector pZ189, and synergistically enhanced UV mutagenicity in the same model, while it did not alter the UV mutation spectra. In cell culture, arsenic did not increase the generation of UV-induced thymine dimers, but did inhibit the removal of the dimers. In addition, arsenic at 1-5 µM enhanced and prolonged RPA-p34 hyper-

ABSTRACT phosphorylation induced by UV irradiation, suggesting the persistence of DNA damage.

Arsenic did not alter expression of several critical NER proteins or inhibit the incision of

UV-induced photoproducts in an in vitro incision assay. Together, these results suggest that arsenic inhibits NER, and perhaps only one subtype of NER, transcription-coupled repair.

Further work though is required to identify the mechanism of this inhibition, and the exact process(es) affected.

Cell cycle checkpoint pathways are surveillance mechanisms that help maintain genomic integrity. The absence of normal checkpoint functions can lead to premature progression through the cell cycle, insufficient time for DNA repair or failure to eliminate damaged cells. Any of these events will lead to an increased risk of genomic instability and its associated risk of malignant transformation. Current evidence suggested that arsenic may suppress cell cycle checkpoints, leading to genomic instability under insults of DNA- damaging agents. In order to investigate the effects of arsenic on cell cycle checkpoints, we studied the ability of arsenic treatment alone to activate/deactivate checkpoints, as well as arsenic’s ability to modulate checkpoints induced by ultraviolet light (UV) irradiation. Our study showed that sodium arsenite alone at 5 µM did not markedly alter cell cycle progression in HeLa cells, other than initiating an M-phase arrest. In contrast, following UV irradiation arsenite restored the G1 checkpoint and enhanced the S and G2 checkpoints in HeLa cells. These results suggest that arsenic does not inhibit the activation of DNA damage checkpoints after UV, and therefore its function as a co- mutagen or co-genotoxin most likely does not occur via cell cycle checkpoint suppression.

ABSTRACT Instead, the observed enhancement of cell cycle checkpoints suggests an increase in DNA damage signaling, perhaps due to an inhibition of DNA repair.

iii

ACKNOWLEDGMENTS

I am very thankful to my research advisor, Dr. Kathleen Dixon, for her excellent and patient guidance and continued encouragement throughout the course of this study.

Appreciation is also expressed to the members of the dissertation committee, Dr. G.

Talaska, Y. Xia, M. Medvedovic, and Y. Sanchez, for their valuable advises.

The friendship of G. G. Oakley is much appreciated and has led to many interesting and good-spirited discussions relating to this research. I am also grateful to my colleague J. G.

Robison for reading drafts of this dissertation and providing many valuable comments that improved this dissertation.

My thanks also go to Dr. S. Patrick and Dr. J. Turchi in Wright State University for helping me out with the in vitro incision assay.

Last, but not least, I would like to thank my family for their understanding and love

during the past few years. Their support and encouragement was in the end what made

this dissertation possible.

TABLE OF CONTENTS

ABSTRACT………………………………………………………….…………………....i

ACKNOWLEDGEMENTS…………….……………………………….………………..iv

TABLE OF CONTENTS………………………………………………………...………..v

LIST OF TABLES…………………………………………………...…………………viii

LIST OF FIGURES………………………………………………………..……………..ix

ABBREVIATIONS……………………………………………………………………..xiii

1. INTRODUCTION…………………………………………………..………………….1

1.1 Arsenic Co-mutagenicity and Co-carcinogenicity…………………………………1

1.1.1 Background and Significance…………………………………...……………1

1.1.2 Arsenic Chemistry and Metabolism…………………………………...……..2

1.1.3 Arsenic Mutagenicity and Co-mutagenicity………………………...………..2

1.1.4 Arsenic Carcinogenicity and Co-carcinogenicity…………………………….3

1.2 Arsenic and DNA Repair……………………………………………….………….3

1.3 Arsenic and Cell Cycle Checkpoints………………………………………………4

1.3.1 Cell Cycle Checkpoints………………………………………………..……..4

1.3.2 Cell Cycle Checkpoint Defect and Carcinogenesis……………………....…..6

1.3.3 Arsenic and Chemotherapy………………………………………..………….6

1.3.4 Arsenic and Cell Cycle…………………………………………...…………..7

1.4 Hyhothesis Tested………………………………………………………..………...8

2. REPORTS………………………………………………………………………………9

2.1 REPORT-1 Inhibition of DNA Repair

As a Mechanism of Arsenic Carcinogenesis ……………………….…..9

2.1.1 Abstract……………………………………………………………………...10

2.1.2 Introduction………………………………………………………………….11

2.1.3 Materials and methods………………………………………………………14

2.1.4 Results……………………………………………………………………….20

2.1.5 Discussion…………………………………………………………………...30

2.1.6 References…………………………………………………………………...37

2.1.7 Tables……….……………………………………………………………….43

2.1.8 Figures legends…………….....……………………….…………………….45

2.1.9 Figures………………………...……………………….…………………….50

2.2 REPORT-2 Arsenic Initiates a UV-Induced G1 Checkpoint, and

Enhances the S-Phase and G2/M Checkpoints in HeLa Cells…………58

2.2.1 Abstract……………………………………………………………………...59

2.2.2 Introduction………………………………………………………………….60

2.2.3 Materials and methods………………………………………………………64

2.2.4 Results……………………………………………………………………….68

2.2.5 Discussion…………………………………………………………………...76

2.2.6 References…………………………………………………………………...86

2.2.7 Figure legends…………………………………………………………….....95

2.2.8 Figures…………………………………...……….……………………….....98

3. DISCUSSION AND FUTURE DIRECTIONS…………………………….……..…113

3.1 Summary……………………………………………………………………...... 113

3.2 Cell cycle checkpoints and DNA repair……………………………..…………114

3.3 Consistency between the experiments……………………...…………………..117

3.4 A possible unified explanation for both DNA repair inhibition and cell cycle alteration by arsenic………………………...…………………………………..119

3.5 Relevance in human health effects: carcinogenesis and chemotherapy……...... 121

3.6 Future directions………………………………..…………...………..………...122

4. REFERENCES…………………………………..……………..……………………126

vii

LIST OF TABLES

Analysis of mutant frequencies of pZ189 following UV and/or arsenic treatment…………………………………………………..43

Distribution of UV-induced mutations in the supF gene in the pZ189

plasmids recovered from arsenic-treated or mock-treated GM637i……………………..44

viii

LIST OF FIGURES

Diagrammatic representation of the pZ189 mutagenesis protocol……………………....50

Mutant frequency of E. coli MLB100 transformed with pZ189 plasmids from the mutagenesis protocol…………………………………………………50

Mutation spectra of the supF gene obtained from replication of

UV-irradiated or non-irradiated pZ189 plasmids in arsenic- treated GM637i fibroblasts………………………………………………………………51

RPA-p34 is hyper-phosphorylated following UV irradiation……………………………52

5 µM arsenic enhanced and prolonged RPA-p34 hyper- phosphorylation induced by UV irradiation……………………………………………..52

1 µM arsenic enhanced and prolonged RPA-p34 hyper- phosphorylation induced by UV irradiation……………………………………………..52

Arsenic prolonged BPDE-induced RPA-p34 hyper-phosphorylation…………………..53

Arsenic treatment did not affect UV-induced RPA-p34 hyper-phosphorylation in NER-deficient XPA cells……………………………………53

FACS analysis of thymine dimers following UV irradiation……………………………54

Arsenic treatment did not increase the generation of thymine dimers in UV treated HeLa cells………………………………………………..54

Arsenic decreased the removal of thymine dimers………………………………………54

Comets were classified into one of three different categories based on appearance………………………………………………………….55

Arsenic treatment prolonged the appearance of type III comets following UV irradiation……………………………………………….55

Arsenic did not affect the expression level of several critical proteins in the NER pathway……………………………………………56

Autoradiograph of an 8% polyacrlyamide-7M urea sequencing gel from the in vitro incision assay………………………………………….57

Quantification of the intensity of the incised adducts……………………………………57

Protocol for overall assessment of the effects of arsenic, UV, and a combination of arsenic plus UV on cell cycle responses………………………….98

Overall assessment of the effects of arsenic, UV, and a combination of arsenic plus UV on cell cycle responses………………………………98

Graphic illustration of overall assessment of the effects of arsenic,

UV, and a combination of arsenic plus UV on cell cycle responses…………………….99

Arsenic arrests cells at G1 checkpoint demonstrated by BrdU labeling………………..100

FACS profiles of propidium iodide-labeled cells and time course of cell cycle progression of cells irradiated in G1…………………………101

Time course of cell cycle progression of cells irradiated at the G1/S border…………..102

FACS profiles of cell cycle progression following

UV irradiation in the presence and absence of p53…………………………………….103

Western immunoblot demonstrating the effect of arsenic on the expression of p53 in HeLa cells…………………………………………104

Treatment protocol used to evaluate arsenic’s effect on S-phase checkpoint…………………………………………………105

An image from FACS analysis with anti-BrdU FITC showing

active S-phase cells and arrested S-phase cells…………………………………………105

Effect of arsenic on S-phase checkpoint………………………………………………..106

Treatment protocol for investigating the effect of arsenic on G2 checkpoint …………107

An image from the FACS analysis with anti-phospho-Histone H3 immunostaining……………………………………………..107

Effect of arsenic on the G2 checkpoint…………………………………………………108

Arsenic alone arrests cells at M phase, not G2…………………………………………109

Cytogenetic analysis of cell arrested by nocodazole and arsenic………………………110

Effect of arsenic on nocodazole-arrested M phase cells………………………………..111

Effect of arsenic on M phase arrest in primary human keratinocytes…………………..112

xii

ABBREVIATIONS

NER: Nucleotide Excision Repair

GGR: Global Genomic Repair

TCR: Transcription-Coupled Repair

UV: Ultraviolet Light

RPA: Replication Protein A

IPTG: Isopropyl-β- D-thiogalactoside

FACS: Fluorescence Activated Cell Sorting

IR: Ionizing Irradiation

BPDE: Benzo(a)pyrene Diol-Epoxide

DSB: Double-stranded DNA break

SSB: Single-Stranded DNA Break

ROS: Reactive Oxygen Species

AT: Ataxia Telangiectasia

NBS: Nijmegen Breakage Syndrome

APL: Acute Promyelocytic Leukemias

DMEM: Dulbecco’s Modified Eagle Media

BrdU: Bromodeoxyuridine

PI: Propidium Iodide

CDK: Cyclin-Dependent Kinase

HPV: Human Papillomavirus

APC/C: Anaphase-Promoting Complex/Cyclosome

xiii

INTRODUCTION 1. INTRODUCTION

1.1 Arsenic Co-mutagenicity and Co-carcinogenicity

1.1.1 Background and Significance

Arsenic is a metalloid widely distributed in the environment. The oxidation states of arsenic vary from -3 to +5. The most commonly found oxidative states of arsenic are the trivalent As(III) and pentavalent As(V) forms, with the As(III) being more toxic and implicated in most of the health problems associated with arsenic exposure[1]. In the

environment, arsenic is generated from both natural and human sources. The principal

natural source is volcanic activity, while human sources are mainly from industries like

metal smelting, combustion of fuels, and use of pesticides [2]. Arsenic can enter the human body by skin contact, inhalation, or gastrointestinal uptake. Arsenic contamination in drinking water is a major public health concern throughout the world, especially in

Chile, Taiwan, China, India, and Bangladesh. It is also a problem in some areas of the southwestern United States [3].

According to epidemiological studies, exposure to arsenic has been associated with an

increased risk of a variety of cancers, including skin, liver, bladder, kidney, and lung cancer.

These studies have led the Environmental Protection Agency (EPA) to classify arsenic as a

“known human carcinogen.” However, attempts to prove arsenic carcinogenesis in animal

models have been unsuccessful, making arsenic the only human carcinogen not shown to

cause cancers in experimental animal models [3].

INTRODUCTION The current arsenic drinking water standard is 10 µg/L. A few years ago, when the standard was still 50 µg/L, an article published in American Journal of Epidemiology in 2001 by a

Taiwanese group drew much attention. The investigators surveyed 8,102 people who had been exposed to arsenic-contaminated drinking water. They found at exposure levels as low as 10-50 µg/L, the risk of cancer of the urinary organs was still significantly higher than in the reference group [4]. That triggered a fierce debate on the appropriateness of the previous

standard of arsenic in drinking water of 50µg/L. Although the new standard was adopted,

the debate goes on which calls for more studies on arsenic.

1.1.2 Arsenic Chemistry and Metabolism

In mammals, arsenic is metabolized primarily by methylation in the liver [5]. Arsenic

biotransformation is catalyzed by monomethylarsonic acid (MMA(V)) reductase and

arsenic methyltransferase using S-adenosyl methionie (SAM) as a methyl donor [6]. The

process follows the order of As(V)-As(III)-MMA(V)-MMA(III)-DMA(V)-DMA(III) and is

generally accepted as a detoxification pathway. Arsenic (III) has been proven to be the most

toxic, while new studies are showing that DMA could be genotoxic. Arsenic metabolites

are mainly excreted through urine with a ratio of inorganic arsenic 10-20%, MMA 10-20%,

DMA 60-80% [7]. Recently an arsenic methyltransferase has been identified in human and

cloned [8]. Current data show that the activity of this enzyme seems to vary among species, which could be the biological foundation for variation in tumor susceptibility in different species [7].

1.1.3 Arsenic Mutagenicity and Co-mutagenicity

INTRODUCTION Unlike most of carcinogens which are also mutagens, arsenic does not seem to cause mutation in single gene loci in most bacterial and mammalian mutagenesis models tested, while arsenic has been shown to potentiate the mutagenicity of other mutagens. So far this co-mutagenic effect has been tested with UV [9], Benzo(a)Pyrene [10], and N-methyl-N-

nitrosourea (NMU) [11;12] in mammalian systems.

1.1.4 Arsenic Carcinogenicity and Co-carcinogenicity

Although listed as a known human carcinogen, arsenic alone has never been proven to induce tumors in experimental animal models. It has been proposed that arsenic might actually act as a co-carcinogen. Recently Roseman’s group treated hairless Skh1 mice with ultraviolet (UV) light in the presence or absence of arsenic. They found that arsenic treatments alone did not induce tumor formation; however, mice treated with UV and arsenite developed cancer at earlier time points, and tumor number and size were increased compared to the animals treated with UV alone [13]. This study provided the first

experimental evidence supporting the long-proposed hypothesis of arsenic acting as a co-

carcinogen.

1.2 Arsenic and DNA Repair

Although evidence of arsenic co-mutagenicity is strong, the mechanism of this co-

mutagenicity is unknown; one hypothesis is that arsenic inhibits DNA repair. The mutagens

that show a co-mutagenic effect with arsenic generate damage that is repaired by the

nucleotide excision repair (NER) pathway, therefore making this pathway a possible target

of arsenic action. Supportive evidence for arsenic inhibition of NER comes from a study by

INTRODUCTION Okui et al [14] that showed arsenic potentiated UV killing in NER-proficient human

fibroblasts, but not in NER-deficient XPA cells. NER can be divided into 4 steps: (1)

recognition, (2) dual incision, (3) gap-filling (repair synthesis), and (4) ligation. Each of

these steps is carried out by different proteins/enzymes and could be a potential target for

arsenic activity. Reduced excision of thymine dimers was also detected indirectly by

measuring [3H] labeled TT dimers chromatographically. There are also reports that suggest

arsenic may act at one or more of the specific steps of NER. Hartwig et al observed a

reduced incision frequency of DNA strands after cells were treated with UV in the presence

of arsenite [15] and Li Jih-Heng et al reported that DNA ligase II was inhibited by arsenic

in Chinese hamster V79 cells [12]. Recently, a report by Hamilton’s group investigated the expression of repair genes and arsenic exposure in a subset of individuals in a population- based case-control study investigating arsenic exposure and cancer risk. They found that arsenic levels were inversely correlated with the expression of critical members of the nucleotide excision repair complex, including ERCC1, XPF, and XPB. This suggests that arsenic might affect DNA repair by altering the expression of repair proteins [16]; however,

contrasting results were also reported [17]. Another mechanism by which arsenic may

inhibit DNA repair is that arsenic directly interacts with repair proteins through thiol groups;

for example, XPA was found to release zinc in the presence of arsenic [18]. In summary,

whether arsenic inhibits DNA repair is still controversial, and a mechanistic theory has yet

to be proven. Therefore, more studies in this area are needed.

1.3 Arsenic and Cell Cycle Checkpoints

1.3.1 Cell Cycle Checkpoints

INTRODUCTION In eukaryotic cells, DNA is under constant attack from endogenous and exogenous agents and processes such as DNA replication errors, metabolic by-products, exposure to environmental DNA-damaging agents, [e.g., ionizing radiation (IR), and ultraviolet light

(UV)], and many chemotherapeutic agents. To maintain genomic integrity, cells have evolved a surveillance network termed the DNA damage response [19]. This network is composed of transduction cascades that sense DNA damage, transduce and amplify the signal, and activate appropriate effector pathways. The major effector pathways within the DNA damage response are cell cycle checkpoints, DNA repair, and cell death via apoptosis. These pathways work together in a coordinated and highly integrated fashion to help maintain genomic integrity in response to genotoxic insult. Checkpoint pathways arrest or delay the progression of the cell cycle to allow sufficient time for repair of the damage and apoptosis is initiated when the DNA damage is too severe to repair.

The major cell cycle checkpoints (G1, S, G2 and the mitotic assembly checkpoint) have been well studied and characterized [reviews see [20;21]]. The activation of a checkpoint

depends on the cell cycle position at which DNA damage occurs. Once a checkpoint has

been passed, the cell cycle will progress despite damage until it reaches the next

checkpoint. The G1 checkpoint guards entry into S phase from the G1 phase, and

prevents cells from initiating DNA replication using damaged DNA as a template. The S

phase checkpoint slows down DNA replication, reducing potential errors arising from

replication of a damaged DNA template. The G2 checkpoint functions to stop cells that

have not completed DNA replication or have unresolved damage from entering M phase,

avoiding errors during chromosome segregation. The mitotic assembly checkpoint

INTRODUCTION responds to unattached chromosomes or lack of tension of mitotic spindle microtubules, in order to prevent uneven segregation of chromosomes.

1.3.2 Cell Cycle Checkpoint Defects and Carcinogenesis

Absence of normal checkpoint function can lead to premature progression through the cell cycle, insufficient time for DNA repair, or failure to eliminate damaged cells via apoptosis. Any one of these events will compromise the integrity of the genome, leading to an increased risk of malignant transformation. Several well-studied inherited diseases demonstrate the importance of the DNA damage response network, and specifically cell cycle checkpoint pathways. Ataxia Telangiectasia (AT) cells are defective in multiple cell cycle checkpoints, which leads to increased sensitivity to ionizing irradiation (IR)[22].

Cells from both Nijmegen Breakage Syndrome (NBS) and Fanconi Anemia subtype D2

(FA-D2) are defective in the S checkpoint, and fail to repress DNA synthesis following

DNA damage [23;24]. Patients with these syndromes show genomic instability and increased susceptibility to cancers.

1.3.3 Arsenic and Chemotherapy

Arsenic by itself has recently proven to be useful in the chemotherapeutic treatment of

Acute Promyelocytic Leukemias (APL) [[25;26] and reviewed in [27]], and its use is now being expanded to cancers other than APL [28]. Regimens combining arsenic and other chemotherapeutic drugs are also being developed to treat cancers [29]. The paradoxical

action of arsenic as both a carcinogen and as a chemotherapeutic agent may be due to its

ability to synergize with other genotoxic agents.

INTRODUCTION

1.3.4 Arsenic and Cell Cycle

Inhibition of DNA repair by arsenic has long been proposed as an explanation of the synergism of arsenic with genotoxic agents. We and other investigators have observed that the repair of DNA damage induced by UV and other genotoxic agents was inhibited by arsenic [14;15;30]. At the same time, we also observed that cell cycle profiles differed

greatly in cells treated with UV alone and in cells treated with UV in the presence of

arsenic. Considering the effects of arsenic on DNA repair, the cell cycle changes may be

a secondary effect of the inhibition of DNA repair. An alternative hypothesis is that

arsenic might directly interfere with cell cycle regulation networks. Evidence exists that

several key proteins involved in checkpoint signaling pathways and cell cycle regulation,

such as cyclins, p53 and Cdc25 C, can be affected by arsenic. The expression of cyclin

D1 was found to increase, while cyclin A and cyclin B1 were observed to decrease in

response to arsenic treatment [31-33]. The tumor suppressor gene p53, which plays a major

role in G1 and G2 checkpoint activation, was reported to be either upregulated or

downregulated in response to arsenic treatment, depending on the cell type investigated

[32;34]. Cdc25C, a dual phosphatase that dephosphorylates Cdk1, activating it to drive the

G2/M transition, is degraded in the presence of arsenic through the KEN-box and

ubiquitin-proteasome pathway, resulting in G2/M arrest [35]. These studies suggest that even though arsenic itself is not mutagenic, arsenic could interfere with DNA damage responses, such as abolishment of checkpoint responses resulting in sensitization of cells to genotoxic agents.

INTRODUCTION Our observations that cell cycle profiles in cells treated with UV alone differed greatly from those in cells treated with UV in the presence of arsenic raised our interest to look at arsenic’s effect on cell cycle responses in greater detail. Previous studies have focused on the effect of arsenic itself on the cell cycle, but few studies have examined alterations of checkpoints activated by DNA damaging agents in the presence of arsenic.

1.4 Hypotheses Tested

In order to understand the mechanism of arsenic co-mutagenesis and co-carcinogenesis, we therefore tested the hypotheses that (1) arsenic potentiates mutagenicity via inhibition of DNA repair; (2) arsenic acts as a co-mutagen via inhibition of cell cycle checkpoint.

REPORT-1

Inhibition of DNA Repair

As a Mechanism of Arsenic Carcinogenesis

Shengqin Liu1, Steve Patrick2, Jacob G. Robison1, Mario Medvedovic1,

Greg G. Oakley1 and Kathleen Dixon1, 3, *

1Department of Environmental Health, University of Cincinnati

College of Medicine, Cincinnati, OH 45220

2Department of Biochemistry and Molecular Biology, Wright State

University School of Medicine, Dayton, OH 345435

3Present address: Department of Molecular and Cellular Biology, University

of Arizona, Tucson, AZ 85721

*Corresponding author. Mailing address: Department of Molecular and

Cellular Biology, University of Arizona, Tucson, AZ 85721. Phone: (520)

621-7563. Fax: (520) 621-3709. Email: [email protected]

REPORT-1 Abstract

Arsenic is a widely distributed metalloid associated with an increased risk of a variety of cancers. However, attempts to prove arsenic carcinogenesis in animals models have been unsuccessful, leading to the hypothesis that arsenic might act as a co-carcinogen instead of a carcinogen itself. Consistent with a co-carcinogenic model, arsenic by itself does not induce mutations in most bacterial or mammalian systems tested, instead it potentiates the mutagenicity of other agents such as ultraviolet (UV) light, benzo(a)pyrene and N-methyl-

N-nitrosourea (NMU). While the mechanism of this co-mutagenicity remains unknown, a possible explanation is that arsenic inhibits DNA repair, specifically nucleotide excision repair (NER). To test this hypothesis, we have completed multiple experiments using low concentrations of arsenite in cultured human cells. Our results showed that at a concentration of 2.5 µM, arsenic induced insertion/deletion mutations in a mutagenesis model using a shuttle vector pZ189, and synergistically enhanced UV mutagenicity in the same model, while it did not alter the UV mutation spectra. In cell culture, arsenic did not increase the generation of UV-induced thymine dimers, but did inhibit the removal of the dimers. In addition, arsenic at 1-5 µM enhanced and prolonged RPA-p34 hyper- phosphorylation induced by UV irradiation, suggesting the persistence of DNA damage.

Arsenic did not alter expression of several critical NER proteins or inhibit the incision of

UV-induced photoproducts in an in vitro incision assay. Together, these results suggest that arsenic inhibits NER, and more specifically, probably only one subtype of NER, transcription-coupled repair. Further work though is required to identify the mechanism of this inhibition and the exact process(es) affected.

REPORT-1 1. Introduction

Arsenic is a widely distributed metalloid found in the environment. The most commonly

found oxidative states of arsenic are the trivalent (As+3) and pentavalent (As+5) forms, with

the trivalent form being more toxic and implicated in most of the health problems

associated with arsenic exposure [1]. In the environment, arsenic is generated by both

natural and human sources. The principal natural source is volcanic activity, while human

sources are mainly from industries like metal smelting, combustion of fuels, and use of

pesticides [2]. Arsenic can enter the human body by skin contact, inhalation, or

gastrointestinal uptake. Arsenic contamination in drinking water is a major public health

concern throughout the world, especially in developing countries, such as Chile, Taiwan,

China, India, and Bangladesh. It is also a problem in some areas of the southwestern United

States[3].

According to epidemiological studies, exposure to arsenic has been associated with an

increased risk of a variety of cancers, including skin, liver, bladder, kidney, and lung cancer.

These studies have led the Environmental Protection Agency (EPA) to classify arsenic as a

“known human carcinogen.” However, attempts to prove arsenic carcinogenesis in animal

models have been unsuccessful, making arsenic the only human carcinogen not shown to

cause cancers in experimental animal models {Rossman TG, 1998 208 /id}.

In order to explain the negative results in animal carcinogenesis models, it has been proposed that arsenic might act as a co-carcinogen. Recently Roseman’s group treated

REPORT-1 hairless Skh1 mice with ultraviolet (UV) light in the presence or absence of arsenic. They found that arsenic treatments alone did not induce tumor formation; however, mice treated with UV and arsenite developed cancer at earlier time points, and tumor number and size were increased compared to the animals treated with UV alone [4]. This study provided the

first experimental evidence supporting the long-proposed hypothesis of arsenic acting as a

co-carcinogen.

In further support of the hypothesized co-carcinogenic activity of arsenic, arsenic itself does

not induce mutations in most of the bacterial and mammalian mutagenesis models tested.

Instead, it potentiates the mutagenicity of other agents. This co-mutagenic effect has been

tested with UV [5], benzo(a)pyrene [6], and N-methyl-N-nitrosourea (MNU) [7;8]. While the mechanism of this co-mutagenicity is unknown, one hypothesis is that arsenic inhibits DNA repair. The mutagens that show a co-mutagenic effect with arsenic generate damage that is repaired by the nucleotide excision repair (NER) pathway, therefore making this pathway a possible target of arsenic action. NER can be divided into 4 steps: (1) recognition, (2) dual incision, (3) gap-filling (repair synthesis), and (4) ligation. Each of these steps is carried out by different proteins/enzymes and could be a potential target for arsenic activity.

Supportive evidence for arsenic inhibiting NER comes from a study by Okui et al. [9] that

showed arsenic-potentiated UV killing in NER-proficient human fibroblasts, but not in

NER-deficient XPA cells. Reduced excision of thymine dimers was also detected indirectly

by measuring [3H] labeled TT dimers chromatographically. There are also reports that

suggest arsenic may act at one or more of the specific steps of NER. Hartwig et al.

observed a reduced incision frequency of DNA strands after cells were treated with UV in

REPORT-1 the presence of arsenite [10], and Li et al. reported that DNA ligase II was inhibited by arsenic in Chinese hamster V79 cells [8]. Recently, a report by Hamilton’s group

investigated the expression of repair genes and arsenic exposure in a subset of individuals

in a population-based case-control study investigating arsenic exposure and cancer risk.

They found that arsenic levels were inversely correlated with the expression of critical

members of the NER complex, including ERCC1, XPF, and XPB. This suggests that

arsenic might affect DNA repair by altering the expression of repair proteins [11]; however,

contrasting results were also reported [12]. In summary, whether arsenic inhibits DNA repair

is still controversial, and a mechanistic theory has yet to be proven. Therefore, more studies

in this area are needed.

While the current literature has not specifically answered the question of how arsenic acts

as a carcinogen, the proposed hypothesis that arsenic works as a co-carcinogen through

inhibition of NER has gained supportive evidence. We have used low concentrations of

arsenic and several techniques to address arsenic’s effect on DNA repair, particularly NER,

to shed light on the potential mechanism(s) of arsenic mutagenesis and/or co-mutagenesis.

REPORT-1 2. Materials and Methods:

Cell lines and treatments

The cell lines used in this study were purchased from Coriel Cell Repository (Camden,

NJ). HeLa cells were maintained in Dulbecco’s Modified Eagle Media (DMEM;

Invitrogen, Carlsbad, CA) with 10% Fetal Bovine Serum (FBS; Hyclone, Logan, UT) and

1% penicillin-streptomycin (PS; Invitrogen). GM637i SV-40 transformed fibroblasts

were grown in Minimum Essential Medium (MEM; Invitrogen) with 15% FBS and 1%

PS. XPA cells (XP 12BE) were grown in MEM supplemented with 15% FBS, 1% PS

and 2 mM L-glutamine (Invitrogen). Arsenic treatment was done by adding sodium

arsenite from a stock solution into the cell medium to the desired concentration. In those

cells that were pretreated, arsenic was added 3 h before UV irradiation. For UV

irradiation, cells were washed with PBS once and irradiated in PBS at a rate of 1 J/m2/s using a low pressure mercury lamp (Mineralight lamp; model UVG-11; UVP, Inc., San

Gabriel, CA) with a maximal output at 254 nm. After UV exposure, PBS was removed

and replaced with the original growth medium, and cells were incubated at 37°C until harvesting.

Antibodies

For western immunoblots, anti-RPA-p34, anti-XPA, anti-XPF, and anti-ERCC1 were all purchased from Neomarker (Fremont, CA). Anti-RPA-p34 was used at a dilution of

1:500; anti-XPA, anti-XPF, and anti-ERCC1 were all used at a dilution of 1:500. Anti-

XPB was obtained from Santa Cruz Biotechnology (Santa Cruz, CA) and used at a

REPORT-1 dilution of 1: 2000. Secondary antibodies were horseradish peroxidase-linked anti-mouse

(Amersham, Piscataway, NJ) and used at a 1:3000 dilution. For Fluorescence-Activated

Cell Sorting (FACS) analysis, mouse anti-thymine dimer was bought from Kamiya

Biomedical Company (Seattle, WA) and used at a dilution of 1:200. Biotin-linked anti- mouse IgG (H+L) and FITC-conjugated streptavidin were purchased from Zymed

Laboratories (San Francisco, CA) and used at a 1:50 dilution.

Plasmids

The pZ189 plasmid, provided by Michael Seidman (NIA, NIH), is a 5504 base pair double-stranded plasmid containing a 200-bp bacterial suppressor supF tRNA gene. The sequence of pZ189 includes a pBR327 origin of replication for bacteria, an SV-40 origin of replication for mammalian cells or cell extracts, an ampicillin-resistance gene for selection of transformed bacteria, and the supF mutagenesis target gene. Mutations in supF gene result in its failure to suppress the amber mutation in the lacZ gene in the

MLB100 E. coli strain, resulting in generation of white colonies on LB plates containing

IPTG (isopropyl-β-D-thiogalactoside) and X-gal (5-bromo-4-chloro-3-indolyl-β-D- galactoside). The pZ189 plasmid was grown in the MBM7070 strain of E. coli [13] and

purified using a Qiagen Maxiprep kit (Qiagen, Valencia, CA). The plasmid was irradiated

using a low pressure mercury lamp with a maximal output at 254 nm (UVP, Inc) at a

concentration of 100 ng/ml in TE buffer (10 mM Tris-HCl, 1 mM EDTA). The plasmid

was then diluted to 25 ng/ml in TE buffer for transfection.

Mutagenesis assay using the pZ189 shuttle vector

REPORT-1 The pZ189 plasmid irradiated with either 300 or 600 J/m2 UV was transfected into GM637i

fibroblasts using lipofectamine according to the manufacturer’s protocol (Invitrogen). Four

h after the transfection, the serum-free medium was replaced with complete medium and

the cells incubated for an additional 48 h. The cells were then washed twice with PBS,

covered with Hirt’s Buffer [14] (10 mM Tris, 10 mM EDTA, 0.6% SDS, pH7.5) and left at

room temperature for 30 min. Cells lysates were scraped from the plates, and transferred to

5M NaCl in TE buffer, inverted gently several times, and then kept at 4ºC overnight. The

following day, samples were vortexed briefly and then centrifuged for 30 min. The

supernatant, which contained the plasmid DNA, was transferred to a 50 ml-tube with TE

buffer containing 12.5% CsCl, and kept at room temperature overnight. Samples were then

filtered into 50 ml-tubes using a syringe driven 0.45 µm Millex HA filter (Millipore,

Billerica, MA). Supercoiled plasmid DNA passed through the filter, while most of the

denatured proteins and some of the plasmid replicative intermediates were trapped. After

using Centricon YM-30 concentrating columns (Millipore), plasmids were recovered and

the DNA concentration measured by spectrophotometery and adjusted to equal

concentrations using TE buffer. Aliquots of the recovered plasmid were digested with Dpn I

(New England Biolabs, Beverly, MA) to eliminate unreplicated input plasmids [15] and run

on agarose gels to confirm appropriate digestion. One µl of digested plasmid was used to

transform E. coli MBM7070 via electroporation. The E. coli cell suspension then was

transferred to LB broth and incubated at 37ºC for 1 h. The broth was then spread onto LB

plates supplemented with IPTG and X-gal using glass beads and incubated at 37ºC

overnight. The following morning, the resultant colonies were recorded as normal (blue) or

mutant (white), and the mutant frequencies calculated. Mutants were further verified by re-

REPORT-1 plating on separate LB plates supplemented with IPTG and X-gal. Plasmid DNA from the resulting mutants was prepared using QIAGEN Miniprep kits (QIAGEN), and sequenced at the University of Cincinnati DNA core. Sequence of the mutations led to compilation of a mutation spectrum generated by UV in the presence or absence of arsenic.

Western blotting

Whole cell lysates were solubilized in Laemmli sample loading buffer, placed at 100°C for

3 min, and then separated on 12% denaturing SDS–polyacrylamide gels. Proteins were transferred to Immobilon-P polyvinyl-divinyl fluoride (PVDF) membranes (Millipore,

Bedford, MA), using a semidry apparatus (Bio-Rad, Hercules, CA) at a maximum of 150 mA and 20 V for 1.5–2 h. The membranes were blocked for 0.5–1 h with TTBS (100 mM

Tris–HCl, pH 7.5; 0.9% NaCl; 0.3% Tween-20) containing 5% dry milk, and then probed with primary antibodies for 1–2 h. After washing four times with TTBS, the membranes were incubated with horseradish peroxidase-linked secondary antibody for 1 h. Membranes were then washed three times with TTBS, and the proteins visualized using chemiluminescence.

Fluorescence Activated Cell Sorting (FACS) with an anti- thymine dimmer antibody

HeLa cells were mock-treated or treated with 5 µM arsenic for 3 h, and then irradiated with

0-40 J/m2 UV. Following UV treatment, cells were incubated in fresh medium in the

presence or absence of 5 µM arsenic. 0-24 h later, 1-2x106 cells from each sample group

were fixed with 70% ethanol and permeablized with 0.1 N HCl and 0.7 % Triton X-100 on

ice for 10 min. After centrifugation, the cells were resuspended in sterile dH2O, vortexed

REPORT-1 gently, and then boiled for 5 min to denature the DNA. After immediately chilling in an ice- water bath for 15 min, the samples were incubated with anti-thymine dimer antibody, washed with PBT (0.5% Tween-20, 5% FBS), incubated with biotin-conjugated anti-mouse

IgG (H+L), and then incubated with FITC-conjugated streptavidin at room temperature for

30 minutes. Finally, cells were washed with PBT and stained with 20 µg/ml propidium iodide (PI; Sigma-Aldrich, St. Louis, MO) at 37ºC for 30 min. FACS analysis was carried out using a FACScallibor (Becton & Dickinson, Franklin Lakes, NJ) flow cytometer and data were analyzed using Cellquest software (Becton & Dickinson) .

Comet assay with T4 endonuclease

Following irradiation of HeLa cells with 10 J/m2 UV, cells were incubated in fresh medium either with or without 5 µM arsenic for 4-48 h. At the appropriate time points, aliquots of

300,000 cells (in PBS) were mixed with 1% low-melting point agarose (final concentration of 0.75%; Invitrogen) and spread on fully frosted glass microscope slides. The cell membranes were lysed open using lysis buffer (2.5 M NaCl, 100 mM EDTA, 10 mM Tris,

1% Triton X-100, 10% DMSO, pH 12.1) at 4ºC for 1 h. After a brief dH2O wash, the slides

were incubated in reaction buffer (50 mM Tris-Cl, pH 7.5; 5 mM EDTA) at room

temperature for 20 min. Two units of T4 endonuclease (Epicentre, Madison, WI) in 30 µl of

reaction buffer were dropped onto the slides, and the slides covered with parafilm, placed in

a humidified box, and incubated at 37ºC for 1 h. After washing briefly with dH2O, the slides were immersed in alkaline electrophoresis buffer (30 mM NaOH, pH 12.1; 10 mM

EDTA) for 30 min to denature the DNA, and then electrophoresed in the same buffer at 20

V, 50 mA for 20 min. Neutralization buffer (0.4 M Tris-HCl) was used to neutralize the

REPORT-1 slides for 10 min and the slides were stained with 40 µg/ml propidium iodide (PI) for 10 min, rinsed with dH2O, and covered with cover slips. Slides were examined using a Nikon inverted fluorescent microscope, and 100 cells per slide were classified in one of three groups according to the extent of damage. Type I cells, with the least amount of damage,

either had no tail or a tail whose length was less than twice the diameter of the head of the

comet. Type II comets had tails whose width was greater than twice the diameter of the

head, but remained continuous with the head. Type III comets, which contained the most

damage, had tails containing uniformly small pieces of DNA, resulting in the tail becoming

disconnected from the head.

In vitro incision assay

An in vitro DNA incision assay using cell-free HeLa extracts was carried out according to a

previously described method with slight modification [16]. The reaction mixtures contained

20 fmol internally labeled 120 bp DNA with a cisplatin GpXpG adduct. Reactions were

initiated with 150 µg of HeLa cell extracts in buffer containing 45 mM HEPES (pH 7.8); 70

mM MgCl2; 0.9 mM DTT; 0.4 mM EDTA; 3.4% glycerol; 5 µg BSA; 2 mM ATP; and 20

µM each of dATP, dCTP, dGTP, and dTTP. Three types of extracts were used. One extract

was made from non-treated control cells, and the other two were made from cells treated

with either 1 µM or 5 µM arsenic for 8 h. The reactions were incubated for 1 h at 30˚C, and

the reactions were stopped by the addition of 0.5 M EDTA and 10 mg/ml proteinase K. The

DNA was ethanol precipitated and resuspended in gel loading buffer. Products were

separated on 8% polyacrylamide-7 M urea sequencing gels and visualized by autoradiography.

REPORT-1 3. Results:

Mutagenesis assay with the pZ189 shuttle vector

The pZ189 plasmid contains the target gene supF, whose gene product suppresses the amber mutant LacZ gene of the host E. coli strain MBM 7070. Mutations in supF can cause loss of suppression of the mutant LacZ gene in the transformed E. coli cells, leading to formation of white colonies on agar plates supplemented with IPTG and X-gal

(Figure 1A). We used this system to determine if arsenic potentiated the mutagenicity of

UV, and if arsenic altered the mutation spectrum induced by UV. Data from 4 independent experiments are shown in Table 1. The background mutation frequency in the transformed E. coli was 1.9x10-4, similar to the frequency reported in previous studies

[17]. The mutation frequency increased with increasing UV dose in all of the experimental

groups. The mutant frequencies from cells treated with 1 or 2.5 µM arsenic alone were

1.1x10-4 and 1.0x10-4respectively. This was not significantly different from the background frequency, suggesting that arsenic at 1 or 2.5 µM is not mutagenic, consistent with previous reports [17]. Transfection of 300 J/m2 UV-irradiated plasmid into cells

treated with 2.5 µM arsenic resulted in a mutation frequency of 19.3x10-4, significantly

higher (p<0.05) than the sum of the mutation frequencies of un-irradiated plasmid

recovered from cells treated with 2.5 µM arsenic (1.0x10-4) and 300 J/m2-irradiated

plasmid recovered from cells with no arsenic treatment (7.3x10-4). This suggests that 2.5

µM arsenic synergistically increased the mutant frequency induced by UV. However,

such a synergistic action was not seen with an arsenic concentration of 1 µM. When the

UV dose was increased to 600 J/m2, 2.5 µM arsenic no longer caused an increase in the

REPORT-1 mutation frequency above the non-arsenic treated controls (Figure 1B). This was probably due to excessive damage of the plasmids at the higher UV dose, resulting in hindrance of plasmid replication. Since the unreplicated plasmids are digested by Dpn I during the protocol, only plasmids that had received less damage could be replicated and recovered for transformation.

Next, we wanted to investigate the effect of arsenic on the UV-induced mutation spectrum of the pZ189 plasmid. We hypothesized that inhibition of DNA repair by arsenic at different points in the repair pathway would result in different mutation spectra, offering an indication of the mode of action of arsenic. If the recognition/incision step was inhibited by arsenic, the DNA damage would not be removed, mutagenesis would be increased, but the mutational spectrum would remain relatively unchanged. If gap filling or ligation were inhibited, the DNA adducts would be removed, but strand interruptions or strand breaks would remain, leading to increased mutagenesis and a change in the mutational spectrum to include more deletions. A total of 74 mutants were sequenced, with the compiled data represented in Table 2. Sixty-seven of the 74 mutants were from

UV-irradiated plasmids transfected into cells treated with either 1 or 2.5 µM arsenic. The remaining 7 mutants were from non-irradiated plasmids recovered from cells treated with

2.5 µM arsenic. The UV-induced mutation spectra without arsenic treatment have been previously reported by our lab [18;19] and other investigators [17]. UV treatment alone

featured point mutations, predominantly located at several “hot spots” within the supF gene[19]. In this study, 56 mutants identified from UV and arsenic co-treated groups were

point mutations, 7 were deletion/insertions, and 4 carried both types of mutations. Of the

REPORT-1 84 different point mutations that we sequenced from the UV and arsenic co-treated samples, 69 (or 82%) contained GC to AT transitions (Table 2). The top 3 hot spots for point mutations were located at nucleotides 156, 168, and 122 (Figure 1C), which were among the hot spots seen in the mutation spectrum induced by UV treatment alone [19].

This suggests that arsenic treatment synergistically increased the mutagenicity of UV, but

did not alter its mutation spectrum. We also noticed that all of the insertions identified

were 1 or 2 nucleotide insertions, typical for UV-induced mutation spectra [18].

Interestingly, all of the identified deletions were large fragments (at least 80 bp), and all

of these deletions were from plasmids recovered from cells treated with 2.5 µM arsenic,

including both irradiated and non-irradiated plasmids. Furthermore, within the 7 sequenced mutants of non-irradiated plasmids recovered from cells treated with 2.5 µM arsenic, 3 of them (or 44%) carried large deletions (Table 2). This suggests that in this pZ189 model, arsenic alone at higher concentrations may be clastogenic, but does not appear to cause point mutations similar to UV irradiation.

Prolonged UV-induced RPA-p34 phosphorylation by arsenic

Replication protein A (RPA) is the major single-stranded DNA binding protein in eukaryotic cells and is involved in DNA replication, repair and recombination [20]. RPA

is composed of 3 subunits: p70, p34, and p14. The p34 subunit is hyper-phosphorylated

(highest migrating band in Figure 2A) in response to DNA damaging agents, including

ionizing irradiation (IR) and UV irradiation [21-23]. Substantial evidence has confirmed

the correlation between RPA-p34 hyper-phosphorylation and DNA damage, making it a

reliable marker for the presence of damage [24]. Here the effect of arsenic on UV-induced

REPORT-1 DNA damage and repair was examined in HeLa cells using RPA-p34 hyper- phosphorylation as a marker of damage.

In cells synchronized at the G1/S boundary and then released, hyper-phosphorylation of

RPA-p34 occurred 4 h after 10 J/m2 UV treatment and reached its peak at around 16 h

(Figure 2B). At 24 h, the amount of hyper-phosphorylated RPA-p34 started to diminish until it was completely absent at 32 h, demonstrating repair of the UV-induced damage

(Figure 2B). In cells that received the combined treatment of UV and arsenic, the amount of hyper-phosphorylated RPA was increased when compared to UV irradiation alone.

More importantly though, the presence of hyper-phosphorylated RPA persisted longer in the presence of arsenic when compared to UV treatment alone, suggesting that arsenic prevented repair of UV-induced DNA damage. We also tested this effect with a lower dose of UV (2 J/m2), which alone did not induce marked hyper-phosphorylation of RPA-

p34. When cells were treated with 2 J/m2 in the presence of 5 µM arsenic, the RPA-p34

hyper-phosphorylation response was increased to an extent comparable with, or even

stronger than 10 J/m2 UV alone. As predicted, the response to UV treatment at 5 J/m2

showed an intermediate effect between that of 2 J/m2 and 10 J/m2. When the arsenic

concentration was lowered to 1 µM, we observed a similar enhancement of RPA-p34

hyper-phosphorylation following treatment with 10 J/m2 UV (Figure 2C). These data

suggested that arsenic somehow inhibited the repair of the UV-induced damage.

It is known that UV-induced TT dimers are mainly removed by nucleotide excision repair

(NER). If arsenic inhibits NER, we predicted that arsenic would enhance and prolong

REPORT-1 RPA-p34 hyper-phosphorylation in response to other agents that also induce DNA damage repaired by NER. In order to test this hypothesis, we treated HeLa cells with benzo(a)pyrene diol-epoxide (BPDE), an active metabolite of benzo(a)pyrene, which can directly interact with DNA and form bulky adducts. As expected, RPA-p34 hyper- phosphorylation induced by BPDE was prolonged by arsenic in a manner similar to the

UV/arsenic data (Figure 2D). These data provided additional evidence that suggested arsenic may inhibit NER.

If the persistence of RPA-p34 hyper-phosphorylation were due to inhibition of NER, we would expect to see no difference in the duration of RPA-p34 hyper-phosphorylation between arsenic-treated and mock-treated groups following UV irradiation in a cell line deficient in NER. To test this, we treated XPA cells with UV and arsenic and looked for

RPA-p34 hyper-phosphorylation. XPA cells carry a mutated XPA gene and fail to express a functional XPA protein, causing an inability to carry out NER [25]. As XPA

cells are much more sensitive to UV irradiation (due to the inability to carry out NER),

we lowered the UV dose to 2 J/m2 and the arsenic concentration to 1 and 2 µM. We

found that the enhancement of UV-induced RPA-p34 hyper-phosphorylation was less

prominent in XPA cells compared to NER proficient cells (Figure 2E), again supporting

the hypothesis that NER is one of the mechanisms of DNA damage repair that is

inhibited by arsenic.

FACS analysis using a TT dimer-specific antibody

REPORT-1 After establishing a correlation between arsenic treatment and increased UV mutagenicity and persistence of DNA damage, we wanted to try and determine the direct effect of arsenic on DNA repair. We adopted a FACS protocol to detect and quantify UV-induced

TT dimers using an anti-TT dimer specific antibody. The intensity of fluorescence in individual cells correlated with the presence of UV-induced TT dimers (Figure 3A).

HeLa cells were incubated with 5 µM arsenite or mock-treated for 3 h before irradiation to ensure that the arsenic had entered the cells by the time of the UV treatment. Using 0-

40 J/m2 UV, a dose-response curve was obtained showing no difference in the amount of

TT dimers generated between the arsenic-treated and the mock-treated groups (Figure

3B). This suggested that arsenic treatment did not increase the generation of TT dimers by UV. Next, we looked at the time course of TT dimer removal. At each time point, the percentage of remaining TT dimers in the arsenic-treated group was significantly higher

than that in the group treated with UV alone (p<0.05) (Figure 3C), indicating that arsenic

inhibited removal of TT dimers.

Comet assay with T4 endonuclease

In order to confirm the FACS result, we used a modification of the alkaline comet assay.

The alkaline comet assay has been proven to be a sensitive method for detecting DNA

damage in individual cells, with damaged DNA migrating away from the nucleus during

electrophoresis, producing a “comet” image when DNA is stained with either ethidium

bromide or propidium iodide [26;27]. In our study we incorporated the use of T4

endonuclease to generate DNA single-stranded breaks (SSBs) at sites of cyclobutane

pyrimidine dimers (CPDs) following UV irradiation. T4 endonuclease binds to UV-

REPORT-1 induced CPDs and cleaves the N-glycosylic bond of the 5’-pyrimidine in the dimer, breaking the phosphodiester bond 3’ to the resulting abasic site[28]. The resulting SSB

can then be detected by the alkaline comet assay, enabling us to evaluate the presence

and/or removal of UV-induced CPDs over time. At a dose range of 0-10 J/m2 UV, there

is a dose response in the appearance of the comets, with higher UV doses resulting in

cells with longer and wider comet tails (Figure 4A). With T4 endonuclease applied,

comet tails were observed only in cells irradiated with UV, reflecting the high specificity

of the T4 endonuclease enzyme for CPDs under these assay conditions (Figure 4B). We

can also concluded that the comet tails we observed up to 24 h were resulted from T4

endonuclease cutting at CPD sites, and were not due to strand breaks directly generated

by UV irradiation. In order to qualitatively evaluate the comets, we scored the comets

based on the length and width of the tail; with Type I comets having the least amount of

damage and Type III comets having the most (Figure 4A). As expected, arsenic treatment

did not result in DNA damage that was recognized by the T4 endonuclease (Figure 4B).

Comets from cells irradiated with UV in the presence or absence of arsenic showed no

significant difference at 4 and 8 h. All of the cells were Type III cells, suggesting that

densely distributed dimers had resulted in cleavage of the DNA into small DNA

fragments by treatment with T4 endonuclease. At 16 h, cells irradiated with UV in the

absence of arsenic started to convert from Type III to Type II comets, suggesting repair

or removal of dimers. However, in UV-irradiated cells treated with arsenic, the comets

remained Type III comets, suggesting the persistence of TT dimers. This difference in

recovery was even more marked at 24 and 48 h. In conjunction with the FACS data using

the TT dimer specific antibody, we concluded that arsenic delayed the repair/removal of

REPORT-1 UV-induced dimers. These data suggested to us that arsenic inhibits NER at the stage of damage recognition or incision. These data are also consistent with a report by Jay et al, who used a similar assay with rodent cells [29].

Expression of several critical proteins involved in the NER pathway in response to UV and arsenic treatment

Generally DNA repair enzymes are constitutively expressed in cycling mammalian cells

[30]. However, in response to DNA damage, cells not only activate checkpoints to halt

cell cycle progression, they also transcriptionally activate specific proteins involved in

corresponding repair pathways to facilitate DNA repair [31], although generally DNA

repair enzymes are constitutively expressed in cycling mammalian cells [30]. It has been

reported that the expression of critical members of the NER complex, including ERCC1,

XPF, and XPB, was inversely correlated with toenail arsenic levels in individuals

exposed to arsenic [11]. To determine if exposure to arsenic down-regulated the

expression of proteins involved in the NER pathways, we examined the expression levels

of XPB, XPA, XPF, and ERCC1 proteins after UV irradiation in the presence or absence

of arsenic. As shown in Figure 5, the expression of XPB, XPF, and ERCC1 was

constitutive and constant, independent of arsenic treatment. The only difference we

observed was that the expression of XPA appeared to be decreased in the presence of 5

µM arsenic at the 16-32 h time points. The timing of the decrease in XPA levels

correlated with the inhibition of DNA repair by arsenic in the comet assay data, but we

do not know if this represents a causal relationship or not.

REPORT-1 In vitro incision assay

Our data, along with previous studies, suggest that arsenic inhibits NER, specifically

targeting the damage recognition/incision step(s) [9;10]. To assess whether arsenic inhibits the excision step of NER, we performed an in vitro DNA excision assay using HeLa cell- free extracts from cells either treated with arsenic or mock-treated. Additionally, we used extracts from mock-treated control cells and added arsenic to the reactions in vitro. The

120 bp DNA substrate used in the assay contained a single cisplatin adduct, with the 14th bp 5’ to the adduct internally labeled with 32P. Once excision has occurred via NER, the

20-30 bp excision product carrying the cisplatin adduct and labeled nucleotide can be

detected with auto-radiography. If arsenic inhibits excision by directly interacting with

proteins involved in the NER pathway, we would expect to see fewer excised products in

the reactions using extracts from control cells with arsenic added in vitro and extracts from arsenic-treated cells. If arsenic inhibits excision by interfering with DNA damage signaling pathways, we would expect to see decreased excision only in reactions using extracts from arsenic-treated cells. The excision assay depicted in Figure 6 measured the ability of several different cell extracts, in the absence or presence of in vivo arsenic treatment (1 or 5 µM for 8h) or addition of arsenic to the reaction mixture in vitro (0, 5, or 50 µM), to carry out excision of the labeled cisplatin adduct. Lane 1 is a negative control, containing labeled substrate with no cell extract. Lanes 2-4, 5-7 and 10-12 used extracts from control cells with the addition of arsenic in vitro. Lanes 8, 9, 13 and 14 used extracts prepared from cells pre-treated with arsenic. Pre-treatment of cells with arsenic in vivo or addition of arsenic to the reaction mixture in vitro did not affect the excision of the cisplatin-labeled adduct. The lack of an effect in the reactions to which arsenic was

REPORT-1 added in vitro suggested that arsenic did not interact directly with, and inhibit, NER protein activities, consistent with the finding by Hu et al. that arsenic does not directly inhibit DNA repair enzymes [12]. No difference in excision activity between extracts from

arsenic-treated cells and control extracts suggested that arsenic treatment may not affect

NER activity via interference with DNA damage signaling pathways.

REPORT-1 4. Discussion:

Inhibition of DNA repair by arsenic has long been proposed as one of the mechanisms of

arsenic co-mutagenesis and co-carcinogenesis. In this study using the pZ189 plasmid

shuttle vector model, we demonstrated that arsenic alone at 1 µM or 2.5 µM did not induce

any increase in mutation frequency. However, we demonstrated that arsenic synergistically

enhanced UV-induced mutations, with the maximum potentiation at 2.5 µM. These results

were similar to those reported by Wienche et al., though they observed an increased

mutation frequency at 5 µM arsenic [32]. Both studies demonstrated the synergistic enhancement of UV-induced mutation frequency by arsenic at a non-mutagenic concentration. In our study, comparison of sequenced mutations derived from cells treated with 1 or 2.5 µM arsenic demonstrated that 2.5 µM arsenic induced a higher rate of deletions and insertions, suggesting that arsenic may be clastogenic at higher concentrations.

Consistent with this notion, the deletions we identified were almost exclusively large fragment deletions, similar to what Hei et al. observed in human-hamster hybrid (AL) cells

[33]. Since the mutations derived from the combined UVand arsenic treatment were mainly

point mutations, and arsenic alone generated only deletions, the increased UV-induced

point mutation frequency in the presence of 2.5 µM arsenic is most likely due to a

synergism between arsenic and UV, and not due to arsenic’s ability to cause deletions.

Mutagens generate specific mutation spectra, and mutagens with similar mutation spectra

are likely to induce damage via similar mechanisms. We compared UV-induced mutation

spectra in the presence and absence of arsenic and found that although arsenic increased the

REPORT-1 UV-induced mutation frequency, arsenic did not alter the UV-induced mutation spectrum, suggesting arsenic works in a manner similar to UV or accentuates the action of UV. Our data from the pZ189 plasmid mutagenesis model suggest that arsenic does not induce point mutations similar to UV, but the idea of accentuating the action of UV fits well with the hypothesis that arsenic inhibits DNA repair. In order to continue investigating the possible

inhibition of DNA repair by arsenic, we moved into a human cell culture model system.

RPA-p34 is hyper-phosphorylated in response to UV irradiation. Rodrigo et al have

demonstrated that stalled replication forks at the site of UV-induced photoproducts, but not

NER intermediates (DNA strand breaks) generated during dimer removal, are the essential

signals for RPA-p34 hyper-phosphorylation [34]. Our data showed an enhanced and

prolonged hyper-phosphorylation of RPA-p34 following UV irradiation in the presence of

arsenic. Three mechanisms could explain this enhanced and prolonged hyper-

phosphorylation of RPA-p34: arsenic induces replication fork stalling, arsenic inhibits

resolution of stalled forks, or arsenic inhibits removal of UV-induced photoproducts. Since

arsenic alone even up to 10 µM does not induce detectable RPA-p34 hyper-

phorsphorylation (data not shown), we can exclude the possibility that arsenic induces

replication fork stalling. Resolution replication fork stalling occurs by mechanisms that

result in bypass instead of actual removal of the damage [35]. Inhibition of resolution of stalled replication forks could explain prolonged RPA-p34 hyper-phosphorylation, but it would not account for the observed reduced dimer removal. Thus, we went to the next step, to determine if arsenic inhibited the removal of UV-induced thymine dimers.

REPORT-1 FACS quantitation of dimers induced by UV irradiation revealed that TT dimer removal was inhibited in the presence of arsenic. Previously, Hartwig et al. reported that arsenic inhibited the removal of UV-induced TT dimers, using generation of single strand breaks during NER as the endpoint [10]. Our method directly measured the amount of TT dimers

instead of depending on the incision mechanism to be fully functional. Our data showed

that with UV irradiation alone, 50% of the thymine dimers were removed by 24 h,

consistent with a previous report using an ELISA assay with the same antibody [36]. We

observed that the presence of arsenic reduced removal of TT dimers by approximately 20% at every time point investigated. Compared with previous studies using single-strand or double-strand breaks generated during UV-induced DNA damage repair as end points, this

FACS method provides more direct evidence of inhibition of DNA repair by arsenic, As some people have argued that strand breaks may be due to the effect of reactive oxygen species (ROS) generated during arsenic metabolism, and not inhibition of thymine dimer removal.

In order to provide supporting evidence for the FACS data, we used visualization of T4 endonuclease-induced DNA strand cleavage at sites of CPDs using the alkaline comet assay as a marker of the presence of CPDs. In the presence of arsenic, UV-induced thymine dimers persisted for a longer time, as demonstrated by the persistence of Type III comets.

Interestingly, the comet shape observed in cells collected right after UV irradiation appeared similar to comets produced by apoptotic cells. However, 3 lines of evidence confirmed that the comets we saw were due to UV-induced thymine dimers and not apoptosis. First, we saw no decrease in cell viability during the course of our experiments

REPORT-1 (and as DNA fragmentation is one of the later stages of apoptosis, we would have expected to see cell death at the time points we looked at). Second, in the absence of T4 endonuclease, we did not see the fan-shaped tail associated with extreme damage or apoptosis up to 24 h after UV treatment. And third, we saw repair of the damage as evidenced by regression of the tail and the decrease in the amount of the smallest cleaved

DNA pieces over time. Additionally, arsenic alone did not induce damage in the presence or absence of T4 endonuclease up to 24 h after the treatment. This suggested that arsenic at these concentrations was not clastogenic, similar to previous reports [29]. Together with the

FACS data, these results strongly suggest that arsenic inhibited the removal of the thymine

dimers. This may be through inhibition of dimer recognition, inhibition of incision, a

combination of both, or some other mechanism altogether.

Hamilton’s group previously reported that the expression level of several critical repair

proteins was decreased in people exposed to arsenic, suggesting arsenic might play an

inhibitory role in activation of repair proteins [11]. Although DNA repair is a constitutive

function throughout the cell cycle, a number of genes are inducible in response to DNA

damage [37;38]. Volker et al reported that XPB and XPC expression patterns changed in

response to UV treatment within the first 2 h, but only in confluent cultured cells [30], suggesting that repair protein induction may be dependent upon cell status. Although we did not observe a marked downregulation of NER proteins, we can not exclude the possibility that repair protein expression may be lowered following long-term exposure to arsenic. However, this mode of action can not explain the inhibition of DNA repair by arsenic observed in this study.

REPORT-1 Besides failure to observe changes in expression levels of NER proteins, we also failed to detect any effect of arsenic in the in vitro incision assay. One of the essential components of the in vitro incision assay is the XPA protein. The XPA protein contains a zinc finger motif

(containing thiol groups), and it has been reported that arsenic causes a release of zinc from

XPA in a dose-dependent manner [39]. It has been hypothesized that arsenic’s ability to

inhibit DNA repair may be through direct interaction of arsenic with proteins containing

zinc finger domains and/or thiol groups [40] such as XPA. Since we did not see an inhibition in the in vitro incision assay with arsenic, this suggested that direct interaction and inhibition of NER proteins is most likely not the answer. Consistent with our results,

Asmuss et al. reported that arsenic did not reduce the binding of XPA to an UV-irradiated oligonucleotide [41].

Data from the RPA-p34 hyper-phosphorylation experiments, FACS measurement of thymine dimer removal experiments, and the comet assay experiments all indicated that arsenic probably worked through inhibition of either dimer recognition and/or incision.

The results of the in vitro incision assay suggested that arsenic did not inhibit incision, but that study alone does not rule out incision as the target for arsenic activity. We need to keep in mind that our results in the incision assay may be due to differences between in vitro conditions and the real in vivo situation. Inside cells, DNA repair involves

recognition of distorted/damaged DNA and accessibility of NER proteins to the damaged

DNA. Proteins such as topoisomerase I and II are required to unwind the DNA in order to

make the damaged DNA accessible to NER proteins. If arsenic inhibits these

steps/proteins, we would not see a difference in the in vitro incision assay.

REPORT-1 An alternative explanation of the negative result in the in vitro assay is that arsenic may inhibit only one subtype of NER. There are two subtypes of NER, global genomic repair

(GGR) and transcription-coupled repair (TCR). Transcription-coupled repair requires transcription and is limited to the transcribed strand of the transcribed genes. If arsenic inhibits TCR, we would not see an effect in the in vitro incision assay since there is no transcription on the 120bp DNA oligomer substrate. The comet assay data also support the hypothesis that arsenic may only inhibit TCR. TCR is known to work earlier and repair DNA damage more rapidly and efficiently than GGR. Different from a typical comet assay where the comet length is proportional to DNA damage, we observed that the earliest change in UV-irradiated cells was the increase in DNA fragments of intermediate length appearing between the head and tail of the comets. Only at a later time points, probably following more global repair, did the comet tails start to shorten.

This early appearance of intermediate-length DNA fragments between the comet head and tail may be the result of transcription-coupled repair, and it was these intermediate- length DNA fragments that were not observed in cells co-treated with UV and arsenic.

Another possible explanation for the reduced dimer removal in the presence of arsenic is arsenic’s effect on cell cycle responses. Arsenic has been shown to enhance UV-induced checkpoints [42]. With UV treatment alone, cells arrested by checkpoint activation were

able to resume normal cell cycle progression after repair, while in the presence of arsenic,

cells seemed to arrest longer (even permanently arrested in some cases) and eventually

undergo apoptosis. Thus, the removal of dimers in cells treated with UV alone may be due

to a dilution effect on the amount of dimers after cells divide, while the amount of dimers

REPORT-1 stays constant in arsenic-treated cells since most of these cells are not able to resume normal cell cycle. This notion may also be supported by the DNA fragment distribution data from the comet assay. Cells treated with UV alone may finish S phase. After S phase, only one half of the DNA contains thymine dimers since the newly synthesized DNA is dimer-free. Cells treated with UV and arsenic showed a longer or even permanent arrest in

S-phase, which means they were not able to finish DNA replication. When being treated with T4 endonuclease, both strands would be cut because both strands still contain dimers.

Increased susceptibility to cancer due to defects in DNA repair has been well recognized.

Our study supports that the inhibition of DNA repair, specifically NER or a subset of NER, may be the mechanism underlying arsenic’s carcinogenesis/co-carcinogenesis potential.

Our data support the hypothesis that the recognition or incision steps of NER are the target of arsenic activity. However, more work is needed to identify the specific process(es) that are affected by arsenic.

Acknowledgements:

This work was supported by the NIEHS Superfund Basic Research grant ES04908. We thank Dr. John Turchi and the members in his lab in Wright State University for help with the in vitro incision assay, Daniel Marmer and Sue Vergamini in Cincinnati

Children’s Hospital flow cytometry lab for help with FACS analysis. We would also like to thank Ms. Elizabeth Kopras, Ms. Rebecca Nebert, Dr. Nika van Tilburg and Dr.

Michael Thomas for editorial suggestions on the manuscript.

REPORT-1 References

[1] P.C. Chan, J. Huff. Arsenic carcinogenesis in animals and in humans:

mechanistic, experimental, and epidemiological evidence, Environ.Carcino.&

Ecotox.Revs., C15 (2), (1997) 83-122.

[2] T.G. Rossman. Molecular and genetic toxicology of arsenic, Environ Toxicol, 17,

(1998) 171-187.

[3] T.G. Rossman. Arsenic, in: Rom WN (Ed.), Environmental and Occupational

Medicine, 1998, pp. 1011-1019.

[4] T.G. Rossman, A.N. Uddin, F.J. Burns, M.C. Bosland. Arsenite is a cocarcinogen

with solar ultraviolet radiation for mouse skin: an animal model for arsenic

carcinogenesis, Toxicol.Appl.Pharmacol., 176, (2001) 64-71.

[5] A. Hartmann, G. Speit. Effect of arsenic and cadmium on the persistence of

mutagen-induced DNA lesions in human cells, Environ Mol.Mutagen., 27, (1996)

98-104.

[6] A. Maier, B.L. Schumann, X. Chang, G. Talaska, A. Puga. Arsenic co-exposure

potentiates benzo[a]pyrene genotoxicity, Mutat.Res., 517, (2002) 101-111.

[7] J.H. Li, T.G. Rossman. Mechanism of comutagenesis of sodium arsenite with N-

methyl-N-nitrosourea, Biol.Trace Elem.Res., 21, (1989) 373-381.

REPORT-1 [8] J.H. Li, T.G. Rossman. Inhibition of DNA ligase activity by arsenite: a possible

mechanism of its comutagenesis, Mol.Toxicol, 2, (1989) 1-9.

[9] T. Okui, Y. Fujiwara. Inhibition of human excision DNA repair by inorganic

arsenic and the co-mutagenic effect in V79 Chinese hamster cells, Mutat.Res.,

172, (1986) 69-76.

[10] A. Hartwig, U.D. Groblinghoff, D. Beyersmann, A.T. Natarajan, R. Filon, L.H.

Mullenders. Interaction of arsenic (III) with nucleotide excision repair in UV-

irradiated human fibroblasts, Carcinogenesis, 18, (1997) 399-405.

[11] A.S. Andrew, M.R. Karagas, J.W. Hamilton. Decreased DNA repair gene

expression among individuals exposed to arsenic in United States drinking water,

Int.J.Cancer, 104, (2003) 263-268.

[12] Y. Hu, L. Su, E.T. Snow. Arsenic toxicity is enzyme specific and its effects on

ligation are not caused by the direct inhibition of DNA repair enzymes,

Mutat.Res., 408, (1998) 203-218.

[13] M.M. Seidman, K. Dixon, A. Razzaque, R.J. Zagursky, M.L. Berman. A shuttle

vector plasmid for studying carcinogen-induced point mutations in mammalian

cells, Gene, 38, (1985) 233-237.

[14] B. Hirt. Selective extraction of polyoma DNA from infected mouse cell cultures,

J.Mol.Biol., 26, (1967) 365-369.

REPORT-1 [15] K.W. Peden, J.M. Pipas, S. Pearson-White, D. Nathans. Isolation of mutants of

an animal virus in bacteria, Science, 209, (1980) 1392-1396.

[16] I.L. Hermanson, J.J. Turchi. Overexpression and purification of human XPA

using a baculovirus expression system, Protein Expr.Purif., 19, (2000) 1-11.

[17] J.K. Wiencke, J.W. Yager, A. Varkonyi, M. Hultner, L.H. Lutze. Study of arsenic

mutagenesis using the plasmid shuttle vector pZ189 propagated in DNA repair

proficient human cells, Mutat.Res., 386, (1997) 335-344.

[18] N.M. King, G.G. Oakley, M. Medvedovic, K. Dixon. XPA protein alters the

specificity of ultraviolet light-induced mutagenesis in vitro, Environ

Mol.Mutagen., 37, (2001) 329-339.

[19] J. Hauser, M.M. Seidman, K. Sidur, K. Dixon. Sequence specificity of point

mutations induced during passage of a UV-irradiated shuttle vector plasmid in

monkey cells, Mol.Cell Biol., 6, (1986) 277-285.

[20] M.S. Wold. Replication protein A: a heterotrimeric, single-stranded DNA-

binding protein required for eukaryotic DNA metabolism, Annu.Rev.Biochem.,

66, (1997) 61-92.

[21] G.G. Oakley, L.I. Loberg, J. Yao, M.A. Risinger, R.L. Yunker, M. Zernik-Kobak,

K.K. Khanna, M.F. Lavin, M.P. Carty, K. Dixon. UV-induced

hyperphosphorylation of replication protein A depends on DNA replication and

expression of ATM protein, Mol.Biol.Cell, 12, (2001) 1199-1213.

REPORT-1 [22] M.P. Carty, M. Zernik-Kobak, S. McGrath, K. Dixon. UV light-induced DNA

synthesis arrest in HeLa cells is associated with changes in phosphorylation of

human single-stranded DNA-binding protein, EMBO J., 13, (1994) 2114-2123.

[23] V.F. Liu, D.T. Weaver. The ionizing radiation-induced replication protein A

phosphorylation response differs between ataxia telangiectasia and normal human

cells, Mol.Cell Biol., 13, (1993) 7222-7231.

[24] S.K. Binz, A.M. Sheehan, M.S. Wold. Replication protein A phosphorylation and

the cellular response to DNA damage, DNA Repair (Amst), 3, (2004) 1015-1024.

[25] J.C. States, S.P. Myrand. Splice site mutations in a xeroderma pigmentosum

group A patient with delayed onset of neurological disease, Mutat.Res., 363,

(1996) 171-177.

[26] D. Anderson, T.W. Yu, D.B. McGregor. Comet assay responses as indicators of

carcinogen exposure, Mutagenesis, 13, (1998) 539-555.

[27] D.W. Fairbairn, P.L. Olive, K.L. O'Neill. The comet assay: a comprehensive

review, Mutat.Res., 339, (1995) 37-59.

[28] R.D. Schrock, III, R.S. Lloyd. Site-directed mutagenesis of the NH2 terminus of

T4 endonuclease V. The position of the alpha NH2 moiety affects catalytic

activity, J.Biol.Chem., 268, (1993) 880-886.

[29] D.T. Bau, J.R. Gurr, K.Y. Jan. Nitric oxide is involved in arsenite inhibition of

pyrimidine dimer excision, Carcinogenesis, 22, (2001) 709-716.

REPORT-1 [30] M. Volker, M.J. Mone, P. Karmakar, A. van Hoffen, W. Schul, W. Vermeulen,

J.H. Hoeijmakers, R. van Driel, A.A. van Zeeland, L.H. Mullenders. Sequential

assembly of the nucleotide excision repair factors in vivo, Mol.Cell, 8, (2001)

213-224.

[31] B.B. Zhou, S.J. Elledge. The DNA damage response: putting checkpoints in

perspective, Nature, 408, (2000) 433-439.

[32] T.G. Rossman, D. Stone, M. Molina, W. Troll. Absence of arsenite mutagenicity

in E coli and Chinese hamster cells, Environ Mutagen., 2, (1980) 371-379.

[33] T.K. Hei, S.X. Liu, C. Waldren. Mutagenicity of arsenic in mammalian cells: role

of reactive oxygen species, Proc.Natl.Acad.Sci.U.S.A, 95, (1998) 8103-8107.

[34] G. Rodrigo, S. Roumagnac, M.S.Wold, B.Salles, P.Calsou. DNA replication but

not nucleotide excision repair is required for UVC-induced replication protein A

phosphorylation in mammalian cells, Mol.Cell Biol., 20, (2000) 2696-2705.

[35] C.H. McGowan. Running into problems: how cells cope with replicating

damaged DNA, Mutat.Res., 532, (2003) 75-84.

[36] A. Nakagawa, N. Kobayashi, T. Muramatsu, Y. Yamashina, T. Shirai, M.W.

Hashimoto, M. Ikenaga, T. Mori. Three-dimensional visualization of ultraviolet-

induced DNA damage and its repair in human cell nuclei, J.Invest Dermatol., 110,

(1998) 143-148.

REPORT-1 [37] B.J. Hwang, J.M. Ford, P.C. Hanawalt, G. Chu. Expression of the p48 xeroderma

pigmentosum gene is p53-dependent and is involved in global genomic repair,

Proc.Natl.Acad.Sci.U.S.A, 96, (1999) 424-428.

[38] S. Adimoolam, J.M. Ford. p53 and DNA damage-inducible expression of the

xeroderma pigmentosum group C gene, Proc.Natl.Acad.Sci.U.S.A, 99, (2002)

12985-12990.

[39] T. Schwerdtle, I. Walter, A. Hartwig. Arsenite and its biomethylated metabolites

interfere with the formation and repair of stable BPDE-induced DNA adducts in

human cells and impair XPAzf and Fpg, DNA Repair (Amst), 2, (2003) 1449-

1463.

[40] A. Hartwig. Zinc finger proteins as potential targets for toxic metal ions:

differential effects on structure and function, Antioxid.Redox.Signal., 3, (2001)

625-634.

[41] M. Asmuss, L.H. Mullenders, A. Eker, A. Hartwig. Differential effects of toxic

metal compounds on the activities of Fpg and XPA, two zinc finger proteins

involved in DNA repair, Carcinogenesis, 21, (2000) 2097-2104.

[42] S. Liu, J.G. Robison, G.G. Oakley, Z. Abdel-Malek, K. Dixon. Arsenic initiates a

UV-induced G1 checkpoint, and enhances the S-phase and G2/M checkpoints in

HeLa cells, Cell Cycle, submitted, (2005).

42 Table 1. Analysis of Mutant Frequencies of pZ189 following UV and/or Arsenic Treatment

Arsenic UV Mutants/Colonies No. of Mutant Frequency* Scored Experiments (X10 -4) 0 µM 0J/m 2 14/38444 4 1.9 300J/m 2 79/205920 4 7.3 600J/m 2 154/72560 4 23.3

4 1.0 µM 0J/m 2 8/104300 4 1.1 3 300J/m 2 87/130248 4 9.4 600J/m 2 237/100040 4 27.2

2.5 µM 0J/m 2 12/104628 4 1.0 300J/m 2 113/62904 4 19.3** 600J/m 2 131/62568 4 23.2 R E P O R

*Mutant frequencies were calculated from the sum of 4 independent experiments. **Significantly higher T - than the simple addition of the mutant frequencies from 300 J/m2 UV/0 µM arsenic-treated cells (line 2) 1 and 0 J/m2 UV/2.5 µM arsenic-treated cells (line 7). Table 2. Distribution of UV-induced mutations in the supF gene in the pZ189 plasmids recovered from arsenic-treated or mock- treated Gm637i

Treatment As (µM)/UV (J/m2 ) 2.5/600 2.5/300 2.5/0 1.0/600 1.0/300 Mutations analyzed 17 28 7 12 9 Deletion 0 5 3 0 0 Insertion 2 2 1 0 0 4 4 Point Mutations Transitions GC-->AT 19 28 1 13 9 AT-->GC 0 1 0 0 0 Transversions CG-->AT 0 1 4 2 3

AT-->TA 2 1 0 1 1 R E P GC-->CG 1 2 0 0 0 O R T - 1 REPORT-1

FIGURE LEGENDS

Figure 1. A, Diagrammatic representation of the pZ189 mutagenesis protocol. The UV irradiated pZ189 plasmids were transfected into GM637i fibroblasts that had been pre- treated with arsenic for 24 h. After the transfection, cells were incubated either with or without arsenic for 48 h. pZ189 plasmids were recovered from the fibroblasts and used to transform E. coli. The total number of colonies and mutant colonies were counted to obtain mutation frequencies. Mutant colonies were collected and the pZ189 plasmids sequenced to generate mutation spectra. B, Mutant frequency of pZ189 plasmids from the mutagenesis protocol. 2.5 µM arsenic increased the mutation frequency at low doses of UV, but at higher doses; there was no difference in mutation frequency between the 3 treatment groups. C, Mutation spectrum of the supF gene obtained from replication of

UV-irradiated or non-irradiated pZ189 plasmids in arsenic treated Gm637i fibroblasts.

The first mutation spectrum (500 J/m2 ) was adapted from Hauser et al [19]. Red bars represent deletions, blue flags represent insertions, and bases above the sequence denote point mutations. The mutation hot spots are labeled at the bottom with red arrows. While

2.5 µM arsenic induced higher frequencies of large deletions than 1 µM arsenic, it synergistically increased the level of point mutations with UV irradiation, and did not alter the UV irradiation mutation spectra. 1.0 µM arsenic did not induce deletions by itself, and did not alter the UV irradiation-induced mutation spectra.

45 REPORT-1

Figure 2. Arsenic enhanced and prolonged RPA-p34 hyper-phosphorylation following

UV irradiation. A, RPA-p34 is hyper-phosphorylated following UV irradiation. B, 5 µM arsenic enhanced and prolonged RPA-p34 hyper-phosphorylation induced by UV irradiation. HeLa cells synchronized and released at the G1/S phase boundary were incubated with 5 µM arsenic for 3 h then treated with 2, 5, or 10 J/m2 UV. The cells were then incubated in complete medium either with or without 5 µM arsenic and collected at the indicated time points. RPA-p34 hyper-phosphorylation was visualized by western immunoblot. C, 1 µM arsenic enhanced and prolonged RPA-p34 hyper-phosphorylation induced by UV irradiation. Cells were treated as described for Figure 2B except 1 µM arsenic was used instead of 2.5 µM. D, Arsenic prolonged BPDE-induced RPA-p34 hyper-phosphorylation. HeLa cells were incubated with 5 µM arsenic and then treated with 1 µM BPDE in PBS for 30 min. After washing twice with serum-free medium, the cells were incubated in fresh medium either with or without 5 µM arsenic for the indicated periods of time before western immunoblot. E, Arsenic treatment did not affect

UV-induced RPA-p34 hyper-phosphorylation in NER deficient XPA cells. XPA cells synchronized and released at the G1/S phase boundary were incubated with 1 or 2 µM arsenic for 3 h and then treated with 2 J/m2 UV. The cells were then incubated in complete medium either with or without 1 or 2 µM arsenic for the indicated periods of time before western immunoblot.

46 REPORT-1

Figure 3. FACS analysis of thymine dimers following UV irradiation. A, HeLa cells treated with 10 J/m2 UV and collected immediately or 24 h later were fixed with ethanol and stained with FITC-labeled anti-thymine dimer antibody and subjected to FACS analysis. At 24 h, approximately half of the thymine dimers were removed as measured by this method. B, Arsenic treatment did not increase the generation of thymine dimers in

UV treated HeLa cells. Cells were treated with 5 µM arsenic for 3 h and then irradiated with UV at the indicated doses. Cells were then harvested and analyzed by FACS. C.

Arsenic decreased the removal of thymine dimers as measured by FACS analysis. HeLa cells were treated with 5 µM arsenic and then irradiated with 10 J/m2 UV. Cells were then allowed to recover for the indicated periods of time either with or without arsenic in the medium before being collected and analyzed by FACS.

47 REPORT-1

Figure 4. Comet assay analysis of arsenic-induced inhibition of thymine dimer removal.

A, Comets were classified into 1 of 3 different categories based on appearance. Type I comets, having the least amount of damage, did not have a tail, or had a tail with a width less than two-times the diameter of the head of the comet. Type II comets had tails with a width of more than twice the diameter of the head, but the tail remained connected with the head. Type III comets, having the greatest amount of damage, had tails with a width greater than twice the diameter of the head, and the tail was not directly connected to the head. B, Arsenic treatment prolonged the appearance of Type III comets following UV irradiation. HeLa cells were irradiated with 10 J/m2 and allowed to recover for the indicated periods of time in medium with or without 5 µM arsenic before being collected and analyzed by the comet assay. Those cells that were not treated with T4 endonuclease did not show any DNA damage regardless of treatment until the latest time point examined, demonstrating the specificity of the T4 endonuclease for CPDs and that arsenic is not clastogenic. In cells treated with UV and T4 endonuclease, Type III comets were the most abundant type of comets at the early time points. At the later time points, the comets shifted from predominantly Type III to predominantly Type II, indicating removal/repair of thymine dimers. In cells co-treated with arsenic and UV, Type III comets remained the predominant type even at the latest time point investigated, indicating inhibition of removal/repair of thymine dimers by arsenic.

48 REPORT-1

Figure 5. Arsenic did not affect the expression level of several critical proteins in the

NER pathway. HeLa cells were synchronized and released at the G1/S phase boundary, treated with 5 µM arsenic for 3 h, then treated with 10 J/m2 UV. Cells were then incubated in medium either with or without 5 µM arsenic for the indicated periods of time before being collected and analyzed by western immunoblot. cXPB, cXPF, etc stand for expression of the particular protein compared to expression of the internal loading control G3PDH (arbitrary units). The anti-XPA antibody recognizes two bands of XPA, so we used total cXPA (cXPA1+cXPA2) for comparison.

Figure 6. Arsenic treatment did not affect the incision of a cisplatin adduct in an in vitro incision assay. A, Autoradiograph of an 8% polyacrylamide-7M urea sequencing gel from the in vitro incision assay. HeLa crude extracts from cells treated with 1 or 5 µM arsenic or mock treated were added to a radiolabeled 120 bp DNA oligomer containing a cisplatin 1,3(GpXpG) DNA adduct. Incision of the DNA adduct was visualized by appearance of a 20-30 bp long radiolabeled DNA fragment. Lane 1 is the labeled oligomer in the absence of cellular extract. Lanes 2-4, 5-7 and 10-12 used extracts from control cells with the addition of 0-50 µM arsenic to the reaction in vitro. Lanes 8 and 13 are extracts from cells pre-treated in vivo with 1 µM arsenic for 8 h and lanes 9 and 14 are extracts from cells pre-treated in vivo with 5 µM arsenic for 8 h. Lane 15 is a control using a DNA substrate without a cisplatin adduct. B, Quantification of the intensity of the incised adducts in the respective lanes compared to the intensity in lane 2. Arsenic, whether from in vivo or in vitro treatment, did not affect incision of the cisplatin adduct.

49 REPORT-1 Figure 1

A Mutagenesis Protocol

25 ) % (

y 20 c n e u q

e 15 r f t n a

t 10 u

M 0uM 5 0 µ1uM AMrsenic 1 µ2.5uM ArsMenic 2.5 µM Arsenic 0 0 100 200 300 400 500 600 700 UV dose (J/m2)

50 REPORT-1 Figure 1 C

500 J/m2 UV**

2.5 µM As/0 J/m2 UV

2.5 µM As/300 J/m2 UV

2.5 µM As/600 J/m2 UV

1 µM As/300 J/m2 UV

1 µM As/600 J/m2 UV

122 Point Mutation 156 168 Insertion Deletion Hot Spot

51 REPORT-1 Figure 2

A -UV +UV

Hyper-p-RPAp34

RPAp34

B 10 J/m2 UV 10 J/m2 UV + 5 µM As Time (hr) 0 4 8 16 24 32 0 4 8 16 24 32

Hyper-p-RPAp34 RPAp34

2 2 J/m2 UV 5 J/m UV + 5 µM As 2 J/m2 UV +5 µM As - As - As - As Time (hr) 4 8 24 4 8 24 4 4 8 8 24 24 Hyper-p-RPAp34 RPAp34

Control 1 µM As C Time (hr) 0 4 8 16 24 32 0 4 8 16 24 32

RPAp34

10 J/m2 UV 10 J/m2 UV + 1 µM As Time (hr) 0 4 8 16 24 32 0 4 8 16 24 32

Hyper-p-RPAp34

RPAp34

52 REPORT-1 Figure 2

D 1 µM BPDE 1 µM BPDE+ 5 µM As Time (hr) 0 4 8 16 24 32 0 4 8 16 24 32

Hyper-p-RPAp34 RPAp34

Alfa-tubulin

2 J/m2 UV 2 J/m2 UV + 2 µM As Time (hr) 0 4 8 16 24 32 0 4 8 16 24 32

Hyper-p-RPAp34 RPAp34

2 J/m2 UV 2 J/m2 UV+1 µM As Time (hr) 8 16 24 8 16 24

Hyper-p- RPAp34 RPAp34

53 REPORT-1 Figure 3

A No UV 24 h after 10J/m 2 UV

0 h after 10J/m2UV

B 350

) 300

e y t

m i

i s

d 250

n e

T d

( n

I y 200

t e

s c

n n

e e

t c 150

n s

I e

e o

n u 100 l

c F

o 50

F No arsenic 5 µM arsenic 0 0 10 20 30 40 50 UV (J/m2)

C 120%

r 100%

T 80%

l %

a r

n e

g m

i i 60%

r d

o T 40%

20%

UV UV + As 0% 0 5 10 15 20 25 30 Hours

54 REPORT-1 Figure 4

Type I Type II Type III

B 100 s

l I

l 80 e

c 60 II 4 hr f o 40 III 20 % 0 UV+T4 As+T4 UV+As+T4 UV-T4 As-T4 UV+As-T4 100 s

l I l 80 e

c 60 II f

8 hr o 40 III 20 % 0 UV+T4 As+T4 UV+As+T4 UV-T4 As-T4 UV+As-T4 100 s

l I

l 80 e

c 60 II f

16 hr o 40 III 20 % 0 UV+T4 As+T4 UV+As+T4 UV-T4 As-T4 UV+As-T4 100 s

l I

l 80 e

c 60 II f

24 hr o 40 III 20 % 0 UV+T4 As+T4 UV+As+T4 UV-T4 As-T4 UV+As-T4 100 s

l I

l 80 e

c 60 II

48 hr f o 40 III 20 % 0 UV As UV+As UV As UV+As + T4 - T4 Treatment

55 REPORT-1 Figure 5

10 J/m2 UV 10 J/m2 UV + 5 µM As Time (hr) 0 4 8 16 24 0 4 8 16 24

XPB

XPF

ERCC1

G3PDH

10 J/m2 UV 10 J/m2 UV + 5 µM As Time (hr) 0 4 8 16 24 32 0 4 8 16 24 32

XPA1 XPA2

G3PDH

56 REPORT-1 Figure 6

A Extract #1 Extract #2 Extract #3 In Vitro As (µM) - 0 5 50 0 5 50 0 0 0 5 50 0 0 - In Vivo As (µM) - 0 0 0 0 0 0 1 5 0 0 0 1 5 -

100bp

80bp

50bp

30bp

Incision products

20bp 1 2 3 4 5 6 7 8 9 101112131415

B Control Extract 140 Liu 2 Extracts 121200 120

y Liu 1 extract t i 100 v 101000 100 i t l c o

80 80 r 80

80 t l A o n r t o n

n 60 60 c 60 o 60 o i f c s o f i 40 o 40 c 4040 % n % I 2020 20 20

00 0 0 c2ont rol cont3rol + 5 uM4control + 50uM 5 6 7 8 9 10 11 12 13 14

REPORT-2

Arsenic Initiates a UV-Induced G1 Checkpoint and Enhances the S-Phase and G2/M Checkpoints in HeLa Cells

Shengqin Liu1, Jacob G. Robison1, Greg G. Oakley1, Zalfa Abdel-Malek2 and Kathleen Dixon1, 3, *

1Department of Environmental Health, University of Cincinnati College of Medicine, Cincinnati, OH 45267 2Department of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH 45267 3Present address: Department of Molecular and Cellular Biology, University of Arizona, Tucson, AZ 85721

*Corresponding author. Mailing address: Department of Molecular and Cellular Biology, University of Arizona, Tucson, AZ 85721. Phone: (520) 621-7563. Fax: (520) 621-3709. Email: [email protected]

REPORT-2 ABSTRACT

Arsenic is a well-documented human carcinogen associated with a variety of cancers, though the mechanism of action is unknown. Arsenic is also currently being used to treat some cancers. The paradoxical action of arsenic as both a carcinogen and a chemotherapeutic agent may be due to its ability to synergize with other genotoxic agents.

Such synergy has been hypothesized to involve effects on DNA repair and/or cell cycle checkpoints. Cell cycle checkpoint pathways are surveillance mechanisms that help maintain genomic integrity. The absence of normal checkpoint functions can lead to premature progression through the cell cycle, insufficient time for DNA repair or failure to eliminate damaged cells. Any of these events will lead to an increased risk of genomic instability and its associated risk of malignant transformation. In order to investigate the effects of arsenic on cell cycle checkpoints, we studied the ability of arsenic treatment alone to activate/deactivate checkpoints, as well as arsenic’s ability to modulate checkpoints induced by ultraviolet light (UV) irradiation. Our study showed that sodium arsenite alone at 5 µM did not markedly alter cell cycle progression in HeLa cells other than initiating an M-phase arrest. In contrast, following UV irradiation arsenite restored the G1 checkpoint and enhanced the S and G2 checkpoints in HeLa cells. These results suggest that arsenic does not inhibit the activation of DNA damage checkpoints after UV, and therefore its function as a co-mutagen or co-genotoxin most likely does not occur via cell cycle checkpoint suppression. Instead, the observed enhancement of cell cycle checkpoints suggests an increase in DNA damage signaling, perhaps due to an inhibition of DNA repair.

REPORT-2 INTRODUCTION

In eukaryotic cells, DNA is under constant attack from endogenous and exogenous agents

and processes, such as DNA replication errors, metabolic by-products, exposure to

environmental DNA-damaging agents, [e.g., ionizing radiation (IR), and ultraviolet light

(UV)], and many chemotherapeutic agents. To maintain genomic integrity, cells have

evolved a surveillance network termed the DNA damage response.1 This network is

composed of transduction cascades that sense DNA damage, transduce and amplify the

signal, and activate appropriate effector pathways. The major effector pathways within

the DNA damage response are cell cycle checkpoints, DNA repair, and cell death via

apoptosis. These pathways work together in a coordinated and highly integrated fashion to help maintain genomic integrity in response to genotoxic insult. Checkpoint pathways arrest or delay the progression of the cell cycle to allow sufficient time for repair of the damage and apoptosis is initiated when the DNA damage is too severe to repair.

The major cell cycle checkpoints (G1, S, G2 and the mitotic assembly checkpoint) have been well studied and characterized.2,3 The activation of a checkpoint depends on the cell

cycle position at which DNA damage occurs. Once a checkpoint has been passed, the cell

cycle will progress despite damage until it reaches the next checkpoint. The G1

checkpoint guards entry into S phase from the G1 phase, and prevents cells from

initiating DNA replication using damaged DNA as a template. The S phase checkpoint

slows down DNA replication, reducing potential errors arising from replication of a

damaged DNA template. The G2 checkpoint functions to stop cells that have not

REPORT-2 completed DNA replication or have unresolved damage from entering M phase, avoiding errors during chromosome segregation. The mitotic assembly checkpoint responds to unattached chromosomes or lack of tension of mitotic spindle microtubules in order to prevent uneven segregation of chromosomes.

Absence of normal checkpoint function can lead to premature progression through the cell cycle, insufficient time for DNA repair, or failure to eliminate damaged cells via apoptosis. Any one of these events will compromise the integrity of the genome, leading to an increased risk of malignant transformation. Several well-studied inherited diseases demonstrate the importance of the DNA damage response network, and specifically cell cycle checkpoint pathways. Ataxia telangiectasia (AT) cells are defective in multiple cell cycle checkpoints, which leads to increased sensitivity to IR.4 Cells from both Nijmegen

breakage syndrome (NBS) and Fanconi anemia subtype D2 (FA-D2) are defective in the

S checkpoint, and fail to repress DNA synthesis following DNA damage.5,6 Patients with

these syndromes show genomic instability and increased susceptibility to cancers.

Arsenic is a well-documented human carcinogen associated with a variety of cancers.

Due to lack of actions at low concentrations in most mutagenesis assay systems, arsenic

has been viewed as a non-genotoxic carcinogen. More and more evidence has

accumulated showing that non-mutagenic concentrations of arsenic have the potential to

increase the mutagenicity of other agents,7,8 suggesting that arsenic might work as a co-

carcinogen. Recently, Rossman et al. showed that arsenic increased tumor formation in

hairless mice treated with UV irradiation.9

REPORT-2

Arsenic by itself has recently proven to be useful in the chemotherapeutic treatment of

Acute promyelocytic leukemias (APL),10-12 and its use is now being expanded to cancers other than APL.13 Regimens combining arsenic and other chemotherapeutic drugs are

also being developed to treat cancers.14 The paradoxical action of arsenic as both a

carcinogen and as a chemotherapeutic agent may be due to its ability to synergize with

other genotoxic agents.

Inhibition of DNA repair by arsenic has long been proposed as an explanation for the

synergism of arsenic with genotoxic agents. We and other investigators have observed that the repair of DNA damage induced by UV and other genotoxic agents was inhibited by arsenic.15-17 At the same time, we also observed that cell cycle profiles differed greatly

in cells treated with UV alone and in cells treated with UV in the presence of arsenic.

Considering the effects of arsenic on DNA repair, the cell cycle changes may be a

secondary effect of the inhibition of DNA repair. An alternative hypothesis is that arsenic

might directly interfere with cell cycle regulation networks. Evidence exists that several

key proteins involved in checkpoint signaling pathways and cell cycle regulation, such as

cyclins, p53 and Cdc25C, can be affected by arsenic. The expression of cyclin D1 was

found to increase, while cyclin A and cyclin B1 were observed to decrease in response to

arsenic treatment.18-20 The tumor suppressor gene p53, which plays a major role in G1

and G2 checkpoint activation, was reported to be either upregulated or downregulated in

response to arsenic treatment, depending on the cell type investigated.19,21 Cdc25C, a dual

phosphatase that dephosphorylates Cdk1 activating it to drive the G2/M transition, is

REPORT-2 degraded in the presence of arsenic through the KEN-box and ubiquitin-proteasome pathway, resulting in G2/M arrest.22 These studies suggest that even though arsenic itself

is not mutagenic, arsenic may interfere with DNA damage responses, such as

abolishment of checkpoint responses resulting in sensitization of cells to genotoxic agents.

Our observations that cell cycle profiles in cells treated with UV alone differed greatly

from those in cells treated with UV in the presence of arsenic raised our interest to look at

arsenic’s effect on cell cycle responses in greater detail. Previous studies have focused on

the effect of arsenic itself on the cell cycle, but few studies have examined alterations of

checkpoints activated by DNA damaging agents in the presence of arsenic. In this paper,

we examined the effect of arsenic on cell cycle progression under normal conditions, and

more importantly, following UV-induced DNA damage. We found that arsenic alone at

5 µM did not markedly alter cell cycle progression in HeLa cells other than initiating an

M-phase arrest. In contrast, following UV irradiation arsenic restored the G1 checkpoint

and enhanced the S and G2 checkpoints in HeLa cells.

REPORT-2 MATERIALS AND METHODS

Cell lines and treatments

HeLa cells were purchased from Coriel Cell Repository (Camden, NJ) and maintained in

Dulbecco’s Modified Eagle Media (DMEM; Invitrogen, Carlsbad, CA) with 10% fetal

bovine serum (FBS; Hyclone, Logan, UT) and 1% penicillin-streptomycin (Invitrogen).

HCT116 p53 (+/+) and HCT116 p53 (-/-) were obtained from Dr. Bert Vogelstein (Johns

Hopkins University, Baltimore, MD) and maintained in McCoy’s 5A medium (Invitrogen)

with 10% FBS. Primary human keratinocytes obtained from neonatal foreskins were

grown in keratinocyte growth medium MCDB153 (Invitrogen), supplemented with 1

ng/ml epidermal growth factor, 0.5 µg/ml hydrocortisone, 5 µg/ml insulin, and 0.5%

bovine pituitary extract, as described by Boyce and Ham.23 These cells have been proven

to have a normal p53 response after UV treatment. Arsenic treatment was done by adding

sodium arsenite from a stock solution into the cell medium to the desired concentration.

For UV irradiation, cells were washed with PBS once and irradiated in PBS at a rate of 1

J/m2/s using a low pressure mercury lamp (Mineralight lamp; model UVG-11; UVP, Inc.,

San Gabriel, CA) with a maximal output at 254 nm. After UV exposure, PBS was

removed and replaced with the original growth medium, and cells were incubated at 37°C until harvesting.

Antibodies

Anti-p53 (ab-1) was purchased from Oncogene (San Diego, CA) and used at a dilution of

1:1000 for western blotting. Anti-phospho-histone H3 was bought from Upstate

REPORT-2 Biotechnology (Lake Placid, NY) and used at a dilution of 1:200 in Fluorescence-

Activated Cell Sorting (FACS) analysis and immunostaining. FITC-conjugated anti- bromodeoxyuridine (BrdU), used for the detection of BrdU labeling in FACS analysis, was purchased from Becton Dickinson (San Jose, CA) and used at a 1:10 dilution.

Secondary antibodies used were FITC-conjugated anti-rabbit (Sigma-Aldrich, St. Louis,

MO) at a 1:100 dilution, and horseradish peroxidase-linked anti-mouse antibody

(Amersham, Piscataway, NJ) at a 1:3000 dilution.

Cell cycle analysis by FACS

For cell cycle profiling, cells were washed with cold phosphate-buffered saline (PBS), fixed with 70% ethanol, stained with 20 µg/ml propidium iodide (PI; Sigma-Aldrich) in

PBS, and subjected to FACS analysis on a FACScalibur (Becton & Dickinson, Franklin

Lakes, NJ) flow cytometer.

For BrdU pulse-chase labeling experiments, 70%-confluent cells were treated with 5 µM arsenic 3 h before UV treatment, then labeled with 10 µM BrdU for 20 min before UV treatment. After UV treatment, cells were incubated with fresh medium for different time periods. For the BrdU pulse-labeling experiment, HeLa cells were synchronized at the

G1/S boundary with 3 µM aphidicolin for 17 h. Cells were then released from the aphidicolin block by replacing the medium with fresh DMEM. At the time of release, 5

µM arsenic and 1.2 µM nocodazole were added to the medium, and the cells were incubated for another 2 h in order to allow entry into S phase. S phase cells were then treated with 10 J/m2 UV irradiation, pulse-labeled with 10 µM BrdU for 20 min, and

REPORT-2 collected at the indicated time points. To prepare the BrdU-labeled cells for FACS analysis, cells were washed with PBS and fixed with 70% ethanol, treated with 2 N

HCl/Triton X-100 to denature the DNA, and then neutralized with 1 ml 0.1 M Na2B4O7

(pH 8.5). Cells were then incubated with 100 µl anti-BrdU FITC, washed with 0.5%

Tween-20 and 1% BSA in PBS, stained with 20 µg/ml of PI, and then analyzed by flow cytometery.

Histone H3 phosphorylation as a mitotic marker

For measuring phospho-specific histone H3 phosphorylation, we followed the procedure of Xu et al.24 with slight modifications. Briefly, the cells were fixed with 70% ethanol, incubated with anti-phospho-histone H3 for 1 h, washed 3 times with 0. 5% Tween-20 and 1% BSA in PBS, and incubated with goat anti-rabbit FITC for 1 h. Finally, the cells

were stained with 20 µg/ml PI, dropped onto a slide and examined by fluorescent

microscopy, or analyzed by flow cytometery.

Western blotting

Whole cell lysates were solubilized in Laemmli sample loading buffer, placed at 100°C

for 3 min, and then separated on a 12% denaturing SDS–polyacrylamide gel. Proteins

were transferred to Immobilon-P polyvinyl-divinyl fluoride (PVDF) transfer membranes

(Millipore, Bedford, MA) using a semidry transfer cell apparatus (Bio-Rad, Hercules, CA)

at a maximum of 150 mA and 20 V for 1.5–2 h. The membranes were blocked for 0.5–1

h with TTBS (100 mM Tris–HCl, pH 7.5; 0.9% NaCl; 0.3% Tween-20) containing 5%

non-fat dry milk, and then probed with anti-p53 antibodies for 1–2 h. After washing 4

REPORT-2 times with TTBS, membranes were incubated in secondary antibody for 1 h, washed 3 times with TTBS, and the proteins were then visualized using chemiluminescence.

REPORT-2 RESULTS

Overall assessment of the effect of arsenic on cell cycle responses induced by UV

To investigate the effects of arsenic on UV-induced cell cycle responses, we pulse-

labeled cells with BrdU and then irradiated with UV. At different time points after UV

irradiation (Figure 1A), cells were immunostained by FITC-anti-BrdU and analyzed by

flow cytometry to determine the percentage of cells within the total population in each

stage of the cell cycle. In Figure 1B, the upper three boxes in each panel (R5, R2 and R6)

represent cells labeled with BrdU, indicating they were in S phase at the time of

irradiation. At the 0 h time point, cells in R5 are in early S phase and cells in R6 are in

late S phase. Cells in R3 are in G1, cells in R4 are in M, and cells in R7 are those that

entered S phase after BrdU pulse-labeling. Since cells continued to progress through the

cell cycle, the make-up of cells within the boxes changes over time. At the later time

points (8 h, 16 h and 24 h), cells in the R5 now include cells in early S phase and cells

that were in S phase during the irradiation but have divided and returned to G1. Cells in

R6 now include late S and G2/M cells.

UV treatment led to a slow-down in the progression through S phase (R2 + R6) as

expected, indicating the activation of the S phase checkpoint. Furthermore, there was no significant difference in R7 UV-treated cells at 8h after UV irradiation when compared to control cells, suggesting cells in both groups entered from G1 to S phase with similar kinetics. This is consistent with previous reports that demonstrate HeLa cells do not have a functional G1/S checkpoint due to accelerated p53 proteolysis directed by oncoprotein

REPORT-2 E6 from HPV.25 Arsenic treatment alone led to fewer cells progressing through G2/M to

G1 as seen by an accumulation of cells in R2+R6 and fewer cells in R5, suggesting that

cells were arrested somewhere in G2/M by arsenic. However, this arrest or pause is

temporary as demonstrated by the ability of the cells to return to G1 at the 16 hour time

point. Following UV irradiation in the presence of arsenic, the cells were blocked from

entering S phase as seen by fewer cells in R7 at 8 and 16 hours, suggesting that a G1/S

checkpoint was initiated, which is in contrast to UV alone or arsenic alone. Additionally,

cells which were at S phase during UV irradiation did not return to G1 as demonstrated

by fewer cells in R5 at 8 h, 16 h, and 24 h time point, suggesting the presence of a

permanent G2/M arrest induced by the arsenic and UV combined treatment.

Effect of arsenic on the G1 checkpoint

Results in Figure 1 suggested that UV in combination with arsenic induced a G1/S

checkpoint or at least a G1-S transition delay. To examine the G1-S transition in more

detail, we focused on cell cycle changes within the first 8 hours after UV irradiation.

Using the same pulse-labeling technique previously described, we looked at the

proportion of cells that entered S phase from G1 (R7), and plotted it against the length of

time after UV irradiation. As shown in Figure 2A, with UV irradiation alone, cells in G1

at the time of irradiation started entering S phase 4-5 h after UV treatment. However, in

the presence of arsenic, UV treated cells did not enter S-phase for at least 8 h after

irradiation.

REPORT-2 To confirm this result, we synchronized HeLa cells at the G2/M boundary with nocodazole and then released them into G1 by incubating in fresh medium. Two hours after release, at which time about 75% cells were in G1, cells were treated with UV. Cell cycle profiles were then obtained for the indicated time points using FACS analysis.

Consistent with previous results, cells treated with both UV and a combination of UV and arsenic were delayed in G1 (Figure 2B). Interestingly, when cells were synchronized at the G1/S boundary with aphidicolin, an inhibitor of DNA polymerase α, the presence of arsenic did not affect cellular progression into S phase after UV (Figure 2C), suggesting that the effect of arsenic on the G1/S transition occurs before the initiation of replication, the point at which aphidicolin acts.

The core molecular events controlling the G1 checkpoint are the induction and stabilization of p53, followed by the trans-activation of p21, an inhibitor of cyclin- dependent kinases (CDKs). The binding of p21 to CDKs causes deactivation of CDKs, resulting in cell cycle arrest.26 HeLa cells have been reported to have an abnormal G1

checkpoint due to the fact that they express the human papillomavirus (HPV) 18 protein

E6. E6 is able to bind to p53 and accelerate its degradation,25,27 resulting in unchecked

CDK function and progression of the cell cycle. In order to understand better the role p53

plays in the activation of a G1 checkpoint by combined arsenic and UV treatment, we

used a pair of isogenic cell lines, HCT116 (p53+/+) and HCT116 (p53-/-), and carried out

cell cycle analysis. In Figure 3A, we see that compared with HCT116 (p53-/-), HCT116

(p53+/+) had a smaller percentage of cells entering S phase from G1 at 8 h when UV

irradiation was given in the absence of arsenic, suggesting the G1/S checkpoint was

REPORT-2 activated by UV irradiation and dependent on p53. The p53 dependence of the UV- induced G1 checkpoint was further confirmed by the observation that no difference on the G1-S transition was observed in HCT116 (p53-/-), with or without 5 µM arsenic treatment. Since arsenic restored the G1 checkpoint, we expected to see arsenic increase p53 level following UV, which was later confirmed by western blot in Figure 3B.

Interestingly, we failed to see any difference on G1 arrest following UV irradiation in

HCT116 (p53+/+) cells. Considering both HeLa cells and HCT116 (p53+/+) carry a wild- type p53 but HeLa cells were HPV-18 transformed, these observations may be explained by the fact that arsenic can inhibit HPV E6 expression,28,29 which will indirectly restore

p53 function in HeLa cells. Thus, the G1 checkpoint which is normally inactivated in

HeLa cells is intact following UV treatment in the presence of arsenic, most likely due to

increased stabilization of p53.

Effect of arsenic on the S-phase checkpoint

When DNA is damaged in S phase, an intra-S phase checkpoint is activated, which

stabilizes replication forks and inhibits late origin firing, delaying progression through S

phase.30 In order to evaluate the S phase checkpoint after UV irradiation, we measured

the proportion of cells in S phase that were actively synthesizing DNA. HeLa cells were pulse labeled with BrdU at various times after UV radiation and harvested at the end of the BrdU pulse (Figure 4A). In order to focus on just the intra-S checkpoint, cells were synchronized at the G1/S boundary with aphidicolin, and nocodazole was added to prevent cells from progressing through M phase. In Figure 4B, cells in S phase and actively synthesizing DNA are represented by the R2+R5+R6 population (BrdU staining);

REPORT-2 cells in S phase but not synthesizing DNA are represented by R7 (DNA content 2N-4N, no BrdU staining). In Figure 4C, we plotted the percentage of S phase cells actively synthesizing DNA (S-active %; (R2+R5+R6)/total) at the indicated time points following

UV treatment. In Figure 4D, we plotted the percentage of S phase cells not synthesizing

DNA (S-arrested %; R7/total) at the indicated time points after UV treatment. With arsenic alone, the cells showed no difference from the control, either in active or arrested

S phase cells, indicating that no S phase checkpoint was activated by arsenic treatment alone. In Figure 1, we saw that cells irradiated with UV while in S phase progressed through S phase at a much slower rate, independent of the presence or absence of arsenic.

Here we found that even though actively synthesizing S phase cells (Figure 4C) did not differ much between the UV and the UV plus arsenic groups, the arrested S phase cells

(Figure 4D) increased dramatically when treated with UV in the presence of arsenic, with a magnitude greater than the sum of arsenic alone plus UV alone, suggesting that arsenic does not inhibit the activation of the S checkpoint, but instead caused a prolongation or an enhancement of the S-phase checkpoint.

Effect of arsenic on the G2 checkpoint

Before cells enter M phase, they pass through the G2 checkpoint, which ensures that

DNA replication is complete and that there is no residual or lingering DNA damage.

Cells with damaged DNA or incomplete DNA replication are stopped in G2 phase until the problem is resolved, or the cells are eliminated via apoptosis. There are two types of

G2 checkpoints. One is activated in cells in which the DNA is damaged during G2 phase.

This is demonstrated by activation of a G2 checkpoint immediately after ionizing

REPORT-2 radiation (IR). This G2 checkpoint is transient and dependent on ATM. The other type of

G2 checkpoint occurs when DNA damage is induced in earlier phases of the cell cycle and not resolved prior to cells entering G2.24 In order to evaluate the effect of arsenic on the G2 checkpoint initiated by DNA damage in G2, we irradiated asynchronous HeLa cells with UV and monitored the progression of cells from G2 into M phase. Cells were treated with 0.3 µM nocodazole to prevent cells that entered M phase from exiting, so that any change in the percentage of M phase cells would be exclusively due to entry of cells from G2 and not affected by exit into G1. We limited our time points to 1 and 3 h, so that cells entering M phase within this period must have been in G2 during UV irradiation (Figure 5A). The anti-phospho-histone H3 antibody, a mitotic marker,24 was

used to distinguish M phase cells from G2 cells (Figure 5B). In Figure 5C, the slopes of

the lines represent the rate of the cells entering M phase after UV irradiation. All the

treatment groups, even the control, have similar rates within the first hour after UV

irradiation. However, between 1 and 3 h, there is a significant change in rates of entry into M phase between the different experimental conditions. The rates of entry were not significantly different for control and arsenic-treated cells (p=0.065), whereas the rate of entry of UV-treated cells was decreased both in the presence and the absence of arsenic

(p<0.05). This implies that fewer cells were entering M phase due to activation of the G2 checkpoint by UV irradiation. Interestingly, the presence of arsenic almost completely blocked M phase entry even at 3 h after irradiation (p=0.0087, compared with UV alone).

Thus, arsenic enhances the UV-induced G2 checkpoint.

Effect of arsenic on M phase

REPORT-2 Due to the difficulty of distinguishing G2 cells from M cells, most previous studies have reported that arsenic treatment alone leads to arrest at G2/M,18,19 but did not pinpoint

more precisely the phase of the cell cycle that was affected. Here we used a mitotic-

specific marker to distinguish M phase cells from the G2 cells. As shown in Figure 6A,

untreated cycling HeLa cells had 21% of cells in G2/M and 5.9% cells in M phase. Cells

treated with 5 µM arsenic alone for 24 h had 59% of cells arrested at G2/M, but only 4%

were in M phase as distinguished by the mitotic marker anti-phospho-histone H3. Cells

treated with 10 J/m2 UV alone had 51% arrested in G2/M, and 36% were in M phase. In

cells that received UV and arsenic combined, the proportion of G2 and M phase cells was

similar to that in untreated cells. In the case of combined UV and arsenic treatment, the

cells may not have progressed to M phase because of the enhanced UV-induced S phase

and G2 arrest. These data demonstrated that the reported G2/M arrest induced by arsenic

alone is more specifically an M phase arrest.

Since microtubule-disrupting agents also induce an M phase arrest, we wanted to

compare cells arrested by arsenic with those arrested with nocodazole to determine if

these compounds work via a similar mechanism. To compare the cytogenetic changes

induced by arsenic or nocodazole treatment, we used fluorescent microscopy (Figure 6B).

Five hours after treatment, cells treated with nocodazole were halted at prophase as

expected31 and were demonstrated by condensed chromosomes that appear like a ball, and almost no metaphase cells were observed. However, cells treated with arsenic showed 36% of M phase cells at metaphase (arrow), suggesting that arsenic might affect

REPORT-2 the progression from metaphase to anaphase, which is distinctly different from the effect of nocodazole treatment.

We next wanted to determine if the effect of arsenic leading to M-phase arrest occurred in M phase or prior to M phase. Cells synchronized in M phase (prophase) by nocodazole were treated with 5 µM arsenic at the time of release. We observed that arsenic treatment of cells in M phase still led to the retention of cells in M phase (Figure 6C), suggesting that the M-phase effect of arsenic is related to the effect of arsenic on a protein and/or process specific to M phase. We cannot exclude the possibility that changes in earlier phases may also influence this effect.

Most cancer cell lines have abnormal checkpoint functions. Studies have shown that in response to DNA damage, lack of a checkpoint often leads to the enhancement of a later checkpoint.24 Since low concentrations of arsenic appear to cause little or no DNA damage, we suspect that the M phase arrest induced by arsenic treatment in HeLa cells might be partially due to the absence of a functional G1 checkpoint. In order to test this hypothesis, primary human keratinocytes from a patient with normal p53 function were treated with arsenic and M-phase cells were examined. As expected, primary keratinocytes did not show a marked M-phase arrest by arsenic (Figure 6D), while they seemed to arrest somewhere before M phase. This suggests that lack of a functional G1 checkpoint in HeLa cells may contribute to the observed arsenic-induced M-phase arrest.

REPORT-2 DISCUSSION:

In order to explain the carcinogenicity of arsenic in the absence of genotoxic potential at low concentrations, many investigators have hypothesized that arsenic may play a co- mutagenic or co-genotoxic role by inhibiting DNA repair.15,17,32 Here we tested an alternative hypothesis that arsenic might exert a co-genotoxic activity by affecting DNA damage-induced cell cycle checkpoints. In this study, we have investigated the effect of arsenic at low concentrations on normal cell cycle progression and on cell cycle checkpoints in response to a model DNA-damaging agent, 254 nm UV. Arsenic alone at

5 µM did not cause a marked arrest in G1, S or G2 phases of the cell cycle in HeLa cells, but caused a pronounced, albeit temporary, M-phase arrest. Contrary to our hypothesis, arsenic did not inhibit the activation of UV-induced DNA damage checkpoints in HeLa cells; instead, it seemed to activate or enhance these checkpoints. For convenience, we will discuss our results in the following sections according to the natural order of the cell cycle.

Arsenic does not alter cell cycle checkpoints by direct DNA damage

Arsenic is a human carcinogen, but due to arsenic’s negative results in carcinogenesis assays in experimental models, the genotoxicity potential of arsenic has been controversial. We showed that no DNA damage was detected using an alkaline comet assay in HeLa cells treated with 5 µM arsenic.33 A similar result was obtained by Bau et al.34 using the same cell line. However, Lynn et al.35 showed that 10 µM arsenic was able to induce DNA strand breaks (DSBs) in human vascular smooth muscle cells. Mre11, a

REPORT-2 DSB sensor, was found to be phosphorylated in a human bladder carcinoma cell line after treatment with 40 µM arsenic.36 Taken together, these studies indicate that arsenic

induces marked genotoxicity only at relatively high concentrations, but not at lower

concentrations. A single DSB if left unrepaired can activate a checkpoint response.3 Our results showed that 5 µM arsenic alone did not elicit any of the 3 major DNA damage checkpoints (G1, S, and G2), suggesting that 5 µM arsenic is not genotoxic, or at least does not induce sufficient damage to activate cell cycle checkpoints. The changes in UV- induced cell cycle checkpoints following arsenic treatment that we observed here are probably not due to an additive effect of arsenic-induced and UV-induced DNA damage.

Mechanism(s) of initiation of a UV-induced G1 checkpoint by arsenic in HeLa cells

The function of the G1 checkpoint is to prevent cells with damaged DNA from entering S phase. Normally UV irradiation elicits the G1 checkpoint in a p53-dependent manner.

HeLa cells, which are HPV-18 transformed, continuously express the viral oncoprotein

E6. E6 is able to bind to p53 and accelerate its degradation through the ubiquitin- mediated proteolysis pathway, preventing DNA damage-induced p53 activation.27

Consistent with this, we saw barely detectable levels of p53 and no G1 delay after HeLa

cells were treated with UV. When we used MCF-7, a cell line with normal p53, to assess

the UV-induced G1 checkpoint, the G1 checkpoint was robust. No cells entered S phase

even at 32 hours after UV (data not shown). These data support the notion that lack of

p53 stabilization contributes to the lack of a UV-induced G1 checkpoint in HeLa cells.

REPORT-2 When HeLa cells were pre-treated with arsenic before UV irradiation, the G1 checkpoint was activated and cells were delayed entering S phase by 8 h. Furthermore, the level of p53 protein was increased after UV in the presence of arsenic, compared with UV treatment alone. In cells with a normal p53 function, arsenic was reported to increase p53 expression and DNA damage induced by arsenic was thought to be the direct cause of this p53 upregulation.21,37,38 Our western blot data showed that the p53 level decreased in

cells treated with arsenic alone, especially at the time activation of the UV-induced G1

checkpoint was observed (around 4-8 h). Two previous studies have reported that arsenic

treatment restored p53 levels in HeLa cells due to its ability to downregulate the E6

oncoprotein.28,29 Both studies showed concomitant repression of E6 with the upregulation

of p53, suggesting that arsenic could upregulate p53 without inducing DNA damage.

However, Mantovani and Banks have shown that inhibition of E6-mediated p53

degradation is not sufficient to stabilize p53 in HeLa cells, suggesting that an additional

signal required for p53 stabilization is still absent in these cells.39 This may explain why

we did not observe p53 accumulation with arsenic treatment alone. The HCT116 p53+/+

and p53-/- isogenic cell lines, which do not contain the E6 oncoprotein, did not show an

altered G1 checkpoint following UV plus arsenic treatment. This suggests that the

activation of the G1 checkpoint in HeLa cells by UV plus arsenic co-treatment may be

through arsenic’s action on the E6 oncoprotein, and indirectly through p53.

Enhancement of UV-induced S-phase checkpoint in the presence of arsenic

In response to UV irradiation, but not arsenic treatment, HeLa cells showed a pronounced

S-phase checkpoint, demonstrated as a prolonged S phase and reduced BrdU

REPORT-2 incorporation. This effect was enhanced synergistically in the presence of arsenic. The S- phase checkpoint can be activated by two categories of agents: those that generate aberrant DNA templates, such as UV and IR, and those that inhibit the replication machinery, such as hydroxyurea (HU, inhibiting ribonucleotide reductase), aphidicolin

(inhibiting DNA polymerase α and δ), and inhibitors of topoisomerase I or II. The synergistic effect of arsenic on the UV-induced S-phase checkpoint could theoretically work through induction of more photoproducts, inhibition of repair of photoproducts, or interference with the replication machinery. We have demonstrated that arsenic treatment alone neither alters S phase progression nor inhibits BrdU incorporation, consistent with a previous report that arsenic does not affect the fidelity of DNA synthesis in vitro.40

Thus, the synergism of arsenic with UV on the S-phase checkpoint is not due to the

inhibition of DNA replication machinery itself.

Normally, the UV-induced S-phase checkpoint is thought to be initiated upon replication

fork stalling at sites of photoproducts.41 This process has been reported to be ATR- and

Chk1-dependent, but p53-indedendent.42 Neecke et al.43 reported that the S-phase

checkpoint was activated in rad14∆ (Saccharomyces cerevisiae homolog of human XPA)

cells, indicating that the stalled replication fork, not the ssDNA generated in the repair

process, is the essential event necessary to activate the S checkpoint. In mammalian cells,

replication protein A (RPA), a protein involved in both DNA repair and checkpoint

signaling, is also a substrate of ATR. The p34 subunit of RPA is hyper-phosphorylated in

response to DNA damage and stalled replication forks.44,45 The persistence of RPAp34

hyper-phosphorylation correlates well with persistent DNA damage, and hyper-

REPORT-2 phosphorylation after UV was observed in cells lacking global genomic repair (GGR) and/or transcription-coupled repair (TCR), suggesting that the stalled replication forks, but not NER intermediates, are the essential signals for RPAp34 hyper-phosphorylation.46

The persistence of RPAp34 hyper-phosphorylation is therefore indicative of the persistence of stalled replication forks. We have previously reported arsenic treatment alone did not induce RPAp34 hyper-phosphorylation, but RPAp34 hyperphosphorylation was very robust after UV irradiation. The amount of hyper-phosphorylated RPAp34 and the duration of phosphorylation were greatly increased in the presence of arsenic, suggesting arsenic treatment led to increased number and/or duration of stalled replication forks.33 At the same time, more TT dimers were detected in cells after UV in the presence of arsenic, possibly due to inhibition of nucleotide excision repair (NER)

processes by arsenic. Overall these data suggest that the inhibition of UV-induced

photoproduct removal by arsenic could be one of the mechanisms underlying the

synergism of arsenic with UV on the S-phase checkpoint.

Enhancement of the UV-induced G2 checkpoint in the presence of arsenic

Unlike the G1 checkpoint, the G2 checkpoint induced by UV is thought to be p53-

independent because cells lacking p53 are arrested in G2/M after UV irradiation.47-49 The

Chk1-Cdc25C-Cdk1 pathway is thought to be responsible for the UV-induced G2 checkpoint. In essence, Chk1, which is activated by DNA damage, phosphorylates and inhibits Cdc25C, which then fails to remove the inhibitory phosphates from cyclin- dependent kinase Cdk1, leading to cell cycle arrest at G2.50 A bona fide G2 checkpoint

occurs when cells are damaged after DNA replication is complete. In our control BrdU

REPORT-2 labeling experiment, cells labeled with BrdU appeared in G1 phase 6 hours after labeling, suggesting that G2 phase lasts about 5 hours if M phase lasts about 1 hour.51 This

suggests that cells entering M phase 3 h after UV irradiation must have been in G2 phase

during irradiation. While we observed that every treatment group delayed entry into M

within the first hour following UV irradiation, this is probably a transient stress response.

After a certain point in G2, UV irradiation does not activate the G2 checkpoint at all,52 which may explain why we observed G2 delay only at later time points.

We observed a UV-induced delay of cells entering M phase and arsenic enhanced this delay. Similar to the situation in G1 phase, repair of UV-induced photoproducts in G2 also depends on NER and the activation of the G2 checkpoint requires a normal NER process to generate ssDNA. Thus, it is conceivable that inhibition of DNA repair by arsenic would contribute to the enhancement of the G2 checkpoint. However, it has been reported that only about 20% of UV-induced thymine dimers can be removed within the first 3 hours after UV.53 Therefore, by the time cells reach M phase, the remaining dimers

are probably sufficient to elicit a G2 checkpoint. Chen et al.54 reported that arsenic alone

is sufficient to induce a G2 checkpoint due to the arsenic-directed degradation of Cdc25C.

Although we did not observe a G2 checkpoint in HeLa cells treated with 5 µM arsenic

alone, we did observe downregulation of Cdc25C in cells irradiated with UV at the G1/S

boundary. The Cdc25C levels were restored 18 hours after UV, a time coinciding with

normal cell cycle restoration, while Cdc25C levels were not restored in the presence of

arsenic even at 24 hours. These data suggest that arsenic’s enhancement of the UV-

REPORT-2 induced G2 checkpoint may be through the prolonged decrease in Cdc25C levels, but further studies are needed to confirm this.

Arsenic-induced M-phase arrest in HeLa cells

With arsenic treatment alone, the only effect on cell cycle progression that we observed in HeLa cells was an M-phase arrest. Previously, arsenic was reported to induce a G2/M arrest, and efforts were made to identify the mechanism(s) of this arrest, such as inhibition of Cdk1.19,55,56 Activation of Cdk1 is a prerequisite for M phase entry,

condensation of chromosomes, and the breakdown of nuclear envelopes.57 Here we

demonstrated with cytogenetic analysis that a substantial portion of cells were arrested in

metaphase with condensed chromosomes and no intact nuclear envelopes, suggesting that

cells had entered M phase. Thus, inhibition of Cdk1 activity probably does not account

for the arsenic-induced M-phase arrest we observed in HeLa cells.

In addition to causing an M-phase arrest, arsenic treatment also leads to aneuploidy and

polyploidy.58 Since these two characteristics are also common to microtubule poisons

such as nocodozole, taxol, and colchicine, it has been hypothesized that arsenic may have

a similar mechanism of action as these agents. Ramirez et al.58 reported arsenic could

inhibit microtubule assembly and induce tubulin depolymerizaition in vitro. However, intact mitotic spindles have been detected in arsenic-treated cells.59 Alignment of chromosomes in the metaphase plate depends on the formation of an intact microtubule spindle, and our observation of the accumulation of metaphase cells suggests that arsenic does not interfere with spindle assembly. Furthermore, we did not see an M-phase arrest

REPORT-2 effect of arsenic in primary keratinocytes. If arsenic interfered with microtubule assembly, we would expect to see an effect in keratinocytes as well as HeLa cells. Our study showed that even in cells in M phase at the time of treatment, arsenic still induced an M- phase arrest within 2 hours, suggesting the M-phase arrest induced by arsenic is at least partially due to its interaction with some target(s) within M phase.

The metaphase-anaphase transition within M phase is initiated by the activation of the anaphase-promoting complex/cyclosome (APC/C) by Cdk1.60 APC/C is a

multicomponent E3 ubiquitin ligase responsible for the degradation of many regulatory

proteins involved in M-phase progression. Previously, arsenic was demonstrated to

inhibit ubiqutin ligase E3 proteins by interacting directly with the vicinal SH- groups of

these proteins.61 We suspect that arsenic may interact with APC/C and affect its activity.

Consistent with this hypothesis, cyclin B, which is degraded by APC/C to facilitate M-

phase exit,62 was found to persist at high levels in arsenic-arrested cells.63 Additionally

dithiothreitol (DTT) was reported to inhibit arsenic-induced M-phase arrest in NB4 cells, which may involve its ability to maintain the thiol groups in an active state.64 Recently, arsenic treatment has been shown to increase the level of ubiquitinated proteins, implying arsenic may indeed affect the protein ubiquitination process.65

Although M-phase arrest by arsenic seems, at least partially, due to a direct interaction

with target(s) within M phase, other mechanisms may also be involved. DNA damage

can inhibit M phase progression from metaphase to anaphase. We cannot exclude the

possibility that minor DNA damage before M phase may also contribute to the M-phase

REPORT-2 arrest by arsenic, especially in cells with defective checkpoints. It has been postulated that lack of a cell cycle checkpoint may be compensated for by the initiation of a later checkpoint.24 Since HeLa cells lack a G1 checkpoint, if arsenic induced minor DNA damage, this damage may accumulate to an extent severe enough to activate an M-phase arrest in HeLa cells. Taken together, we conclude the M phase effect by arsenic is at least partially due to its effect on M phase-specific proteins/processes, but it is possible that arsenic-induced DNA damage in an earlier phase of the cell cycle may also contribute to the M-phase arrest.

Summary

We have shown here that the major DNA damage checkpoints are intact in the presence of arsenic. Arsenic treatment does not inhibit their activation. On the contrary, arsenic

seems to enhance UV-induced cell cycle checkpoints, possibly through inhibition of

DNA repair. We have also shown that the arsenic-induced G2/M checkpoint activation

that has been reported previously is more specifically an M-phase checkpoint, probably at

the transition from metaphase to anaphase. Together our data suggest that alteration of

genotoxin-induced checkpoints by arsenic does not contribute to arsenic co-

carcinogenesis and/or co-mutagenesis.

Acknowledgements:

This work was supported by the NIEHS Superfund Basic Research grant ES04908. We

thank Dr. Bert Vogelstein in Johns Hopkins University for providing the HCT116 p53

(+/+) and HCT116 p53 (-/-) cell lines, Daniel Marmer and Sue Vergamini in Cincinnati

REPORT-2 Children’s Hospital flow cytometry lab for help with FACS analysis. We would also like to thank Ms. Elizabeth Kopras, Ms. Rebecca Nebert, Dr. Nika van Tilburg and Dr.

Michael Thomas for editorial suggestions on the manuscript.

REPORT-2 References

1. Petrini JH, Stracker TH. The cellular response to DNA double-strand breaks:

defining the sensors and mediators. Trends Cell Biol. 2003;13:458-62.

2. Elledge SJ. Cell cycle checkpoints: preventing an identity crisis. Science

1996;274:1664-72.

3. Weinert T. DNA damage checkpoints update: getting molecular.

Curr.Opin.Genet.Dev. 1998;8:185-93.

4. Beamish H, Williams R, Chen P, Lavin MF. Defect in multiple cell cycle

checkpoints in ataxia-telangiectasia postirradiation. J.Biol.Chem. 1996;271:20486-

93.

5. Lim DS, Kim ST, Xu B, Maser RS, Lin J, Petrini JH et al. ATM phosphorylates

p95/nbs1 in an S-phase checkpoint pathway. Nature 2000;404:613-17.

6. Taniguchi T, Garcia-Higuera I, Xu B, Andreassen PR, Gregory RC, Kim ST et al.

Convergence of the fanconi anemia and ataxia telangiectasia signaling pathways.

Cell 2002;109:459-72.

7. Chan PC, Huff J. Arsenic carcinogenesis in animals and in humans: mechanistic,

experimental, and epidemiological evidence. Environ.Carcino.& Ecotox.Revs.,

1997;C15 (2):83-122.

8. Rossman TG. Molecular and genetic toxicology of arsenic. Environ Toxicol

1998;17:171-87.

REPORT-2 9. Rossman TG, Uddin AN, Burns FJ, Bosland MC. Arsenite is a cocarcinogen with

solar ultraviolet radiation for mouse skin: an animal model for arsenic

carcinogenesis. Toxicol.Appl.Pharmacol. 2001;176:64-71.

10. Soignet SL, Maslak P, Wang ZG, Jhanwar S, Calleja E, Dardashti LJ et al.

Complete remission after treatment of acute promyelocytic leukemia with arsenic

trioxide. N.Engl.J.Med. 1998;339:1341-48.

11. Shen ZX, Chen GQ, Ni JH, Li XS, Xiong SM, Qiu QY et al. Use of arsenic trioxide

(As2O3) in the treatment of acute promyelocytic leukemia (APL): II. Clinical

efficacy and pharmacokinetics in relapsed patients. Blood 1997;89:3354-60.

12. Novick SC, Warrell RP, Jr. Arsenicals in hematologic cancers. Semin.Oncol.

2000;27:495-501.

13. Chen Z, Chen GQ, Shen ZX, Sun GL, Tong JH, Wang ZY et al. Expanding the use

of arsenic trioxide: leukemias and beyond. Semin.Hematol. 2002;39:22-26.

14. Uslu R, Sanli UA, Sezgin C, Karabulut B, Terzioglu E, Omay SB et al. Arsenic

trioxide-mediated cytotoxicity and apoptosis in prostate and ovarian carcinoma cell

lines. Clin.Cancer Res. 2000;6:4957-64.

15. Hartwig A, Groblinghoff UD, Beyersmann D, Natarajan AT, Filon R, Mullenders

LH. Interaction of arsenic(III) with nucleotide excision repair in UV-irradiated

human fibroblasts. Carcinogenesis 1997;18:399-405.

REPORT-2 16. Tran HP, Prakash AS, Barnard R, Chiswell B, Ng JC. Arsenic inhibits the repair of

DNA damage induced by benzo(a)pyrene. Toxicol Lett. 2002;133:59-67.

17. Okui T, Fujiwara Y. Inhibition of human excision DNA repair by inorganic arsenic

and the co-mutagenic effect in V79 Chinese hamster cells. Mutat.Res. 1986;172:69-

76.

18. Park WH, Seol JG, Kim ES, Hyun JM, Jung CW, Lee CC et al. Arsenic trioxide-

mediated growth inhibition in MC/CAR myeloma cells via cell cycle arrest in

association with induction of cyclin-dependent kinase inhibitor, p21, and apoptosis.

Cancer Res. 2000;60:3065-71.

19. Zhao S, Tsuchida T, Kawakami K, Shi C, Kawamoto K. Effect of As2O3 on cell

cycle progression and cyclins D1 and B1 expression in two glioblastoma cell lines

differing in p53 status. Int.J.Oncol. 2002;21:49-55.

20. Vogt BL, Rossman TG. Effects of arsenite on p53, p21 and cyclin D expression in

normal human fibroblasts -- a possible mechanism for arsenite's comutagenicity.

Mutat.Res. 2001;478:159-68.

21. Hamadeh HK, Vargas M, Lee E, Menzel DB. Arsenic disrupts cellular levels of p53

and mdm2: a potential mechanism of carcinogenesis.

Biochem.Biophys.Res.Commun. 1999;263:446-49.

22. Chen F, Castranova V, Li Z, Karin M, Shi X. Inhibitor of nuclear factor kappaB

kinase deficiency enhances oxidative stress and prolongs c-Jun NH2-terminal

kinase activation induced by arsenic. Cancer Res. 2003;63:7689-93.

REPORT-2 23. Boyce ST, Ham RG. Calcium-regulated differentiation of normal human epidermal

keratinocytes in chemically defined clonal culture and serum-free serial culture.

J.Invest Dermatol. 1983;81:33s-40s.

24. Xu B, Kim ST, Lim DS, Kastan MB. Two molecularly distinct G(2)/M checkpoints

are induced by ionizing irradiation. Mol.Cell Biol. 2002;22:1049-59.

25. Athanassiou M, Hu Y, Jing L, Houle B, Zarbl H, Mikheev AM. Stabilization and

reactivation of the p53 tumor suppressor protein in nontumorigenic revertants of

HeLa cervical cancer cells. Cell Growth Differ. 1999;10:729-37.

26. Lakin ND, Jackson SP. Regulation of p53 in response to DNA damage. Oncogene

1999;18:7644-55.

27. Kessis TD, Slebos RJ, Nelson WG, Kastan MB, Plunkett BS, Han SM et al. Human

papillomavirus 16 E6 expression disrupts the p53-mediated cellular response to

DNA damage. Proc.Natl.Acad.Sci.U.S.A 1993;90:3988-92.

28. Um SJ, Lee SY, Kim EJ, Myoung J, Namkoong SE, Park JS. Down-regulation of

human papillomavirus E6/E7 oncogene by arsenic trioxide in cervical carcinoma

cells. Cancer Lett. 2002;181:11-22.

29. Chou RH, Huang H. Restoration of p53 tumor suppressor pathway in human

cervical carcinoma cells by sodium arsenite. Biochem.Biophys.Res.Commun.

2002;293:298-306.

REPORT-2 30. Tercero JA, Diffley JF. Regulation of DNA replication fork progression through

damaged DNA by the Mec1/Rad53 checkpoint. Nature 2001;412:553-57.

31. Dynek JN, Smith S. Resolution of sister telomere association is required for

progression through mitosis. Science 2004;304:97-100.

32. Li JH, Rossman TG. Inhibition of DNA ligase activity by arsenite: a possible

mechanism of its comutagenesis. Mol.Toxicol 1989;2:1-9.

33. Liu S, Patrick S, Robison JG, Medvedovic M, Oakley GG, Dixon K. Inhibition of

DNA repair as a mechanism of arsenic carcinogenesis. DNA Repair

2005;Submitted.

34. Bau DT, Gurr JR, Jan KY. Nitric oxide is involved in arsenite inhibition of

pyrimidine dimer excision. Carcinogenesis 2001;22:709-16.

35. Lynn S, Gurr JR, Lai HT, Jan KY. NADH oxidase activation is involved in

arsenite-induced oxidative DNA damage in human vascular smooth muscle cells.

Circ.Res. 2000;86:514-19.

36. Yuan SS, Su JH, Hou MF, Yang FW, Zhao S, Lee EY. Arsenic-induced Mre11

phosphorylation is cell cycle-dependent and defective in NBS cells. DNA Repair

(Amst) 2002;1:137-42.

37. Yih LH, Lee TC. Arsenite induces p53 accumulation through an ATM-dependent

pathway in human fibroblasts. Cancer Res. 2000;60:6346-52.

REPORT-2 38. Salazar AM, Ostrosky-Wegman P, Menendez D, Miranda E, Garcia-Carranca A,

Rojas E. Induction of p53 protein expression by sodium arsenite. Mutat.Res.

1997;381:259-65.

39. Mantovani F, Banks L. Inhibition of E6 induced degradation of p53 is not sufficient

for stabilization of p53 protein in cervical tumour derived cell lines. Oncogene

1999;18:3309-15.

40. Tkeshelashvili LK, Shearman CW, Zakour RA, Koplitz RM, Loeb LA. Effects of

arsenic, selenium, and chromium on the fidelity of DNA synthesis. Cancer Res.

1980;40:2455-60.

41. McGowan CH. Running into problems: how cells cope with replicating damaged

DNA. Mutat.Res. 2003;532:75-84.

42. Heffernan TP, Simpson DA, Frank AR, Heinloth AN, Paules RS, Cordeiro-Stone M

et al. An ATR- and Chk1-dependent S checkpoint inhibits replicon initiation

following UVC-induced DNA damage. Mol.Cell Biol. 2002;22:8552-61.

43. Neecke H, Lucchini G, Longhese MP. Cell cycle progression in the presence of

irreparable DNA damage is controlled by a Mec1- and Rad53-dependent

checkpoint in budding yeast. EMBO J. 1999;18:4485-97.

44. Robison JG, Elliott J, Dixon K, Oakley GG. Replication protein A and the

Mre11.Rad50.Nbs1 complex co-localize and interact at sites of stalled replication

forks. J.Biol.Chem. 2004;279:34802-10.

REPORT-2 45. Oakley GG, Loberg LI, Yao J, Risinger MA, Yunker RL, Zernik-Kobak M et al.

UV-induced hyperphosphorylation of replication protein A depends on DNA

replication and expression of ATM protein. Mol.Biol.Cell 2001;12:1199-213.

46. Rodrigo G, Roumagnac S, Wold MS, Salles B, Calsou P. DNA replication but not

nucleotide excision repair is required for UVC-induced replication protein A

phosphorylation in mammalian cells. Mol.Cell Biol. 2000;20:2696-705.

47. Pavey S, Russell T, Gabrielli B. G2 phase cell cycle arrest in human skin following

UV irradiation. Oncogene 2001;20:6103-10.

48. Herzinger T, Funk JO, Hillmer K, Eick D, Wolf DA, Kind P. Ultraviolet B

irradiation-induced G2 cell cycle arrest in human keratinocytes by inhibitory

phosphorylation of the cdc2 cell cycle kinase. Oncogene 1995;11:2151-56.

49. Gabrielli BG, Clark JM, McCormack AK, Ellem KA. Ultraviolet light-induced G2

phase cell cycle checkpoint blocks cdc25-dependent progression into mitosis.

Oncogene 1997;15:749-58.

50. Bunz F, Dutriaux A, Lengauer C, Waldman T, Zhou S, Brown JP et al.

Requirement for p53 and p21 to sustain G2 arrest after DNA damage. Science

1998;282:1497-501.

51. Wang XQ, Ellem KA. Heterogeneity in the HeLa cell cycle response to UVC

analyzed by the BrdUrd two-parameter method. Exp.Cell Res. 1994;212:176-89.

REPORT-2 52. Orren DK, Petersen LN, Bohr VA. A UV-responsive G2 checkpoint in rodent cells.

Mol.Cell Biol. 1995;15:3722-30.

53. Nakagawa A, Kobayashi N, Muramatsu T, Yamashina Y, Shirai T, Hashimoto MW

et al. Three-dimensional visualization of ultraviolet-induced DNA damage and its

repair in human cell nuclei. J.Invest Dermatol. 1998;110:143-48.

54. Chen F, Zhang Z, Bower J, Lu Y, Leonard SS, Ding M et al. Arsenite-induced

Cdc25C degradation is through the KEN-box and ubiquitin-proteasome pathway.

Proc.Natl.Acad.Sci.U.S.A 2002;99:1990-95.

55. Chen F, Zhang Z, Leonard SS, Shi X. Contrasting roles of NF-kappaB and JNK in

arsenite-induced p53-independent expression of GADD45alpha. Oncogene

2001;20:3585-89.

56. Chen F, Lu Y, Zhang Z, Vallyathan V, Ding M, Castranova V et al. Opposite effect

of NF-kappa B and c-Jun N-terminal kinase on p53-independent GADD45

induction by arsenite. J.Biol.Chem. 2001;276:11414-19.

57. Nigg EA. Mitotic kinases as regulators of cell division and its checkpoints.

Nat.Rev.Mol.Cell Biol. 2001;2:21-32.

58. Ramirez P, Eastmond DA, Laclette JP, Ostrosky-Wegman P. Disruption of

microtubule assembly and spindle formation as a mechanism for the induction of

aneuploid cells by sodium arsenite and vanadium pentoxide. Mutat.Res.

1997;386:291-98.

REPORT-2 59. Halicka HD, Smolewski P, Darzynkiewicz Z, Dai W, Traganos F. Arsenic trioxide

arrests cells early in mitosis leading to apoptosis. Cell Cycle 2002;1:201-09.

60. Murray AW. Recycling the cell cycle: cyclins revisited. Cell 2004;116:221-34.

61. Berleth ES, Kasperek EM, Grill SP, Braunscheidel JA, Graziani LA, Pickart CM.

Inhibition of ubiquitin-protein ligase (E3) by mono- and bifunctional

phenylarsenoxides. Evidence for essential vicinal thiols and a proximal nucleophile.

J.Biol.Chem. 1992;267:16403-11.

62. Peters JM. The anaphase-promoting complex: proteolysis in mitosis and beyond.

Mol.Cell 2002;9:931-43.

63. Huang S, Huang CF, Lee T. Induction of mitosis-mediated apoptosis by sodium

arsenite in HeLa S3 cells. Biochem.Pharmacol. 2000;60:771-80.

64. Cai X, Yu Y, Huang Y, Zhang L, Jia PM, Zhao Q et al. Arsenic trioxide-induced

mitotic arrest and apoptosis in acute promyelocytic leukemia cells. Leukemia

2003;17:1333-37.

65. Bredfeldt TG, Kopplin MJ, Gandolfi AJ. Effects of arsenite on UROtsa cells: low-

level arsenite causes accumulation of ubiquitinated proteins that is enhanced by

reduction in cellular glutathione levels. Toxicol Appl.Pharmacol. 2004;198:412-18.

94 REPORT-2

FIGURE LEGENDS

Figure 1. A. Protocol for arsenic, UV and BrdU treatment. HeLa cells were pre-treated with 5 µM sodium arsenite for 3 h; 20 minutes before UV irradiation, they were pulse- labeled with 10 µM BrdU for 20 minutes. The cells were irradiated with 10 J/m2 UV, collected at the indicated time points, and then subjected to FACS. B. Overall assessment of the effects of arsenic, UV, and a combination of arsenic plus UV on FACS profiles. C.

Graphic illustration of the results in Figure 1B.

Figure 2. A. Unlabeled cells in S phase (R7). Cells were treated as described in Figure 1, except that cells were harvested at the indicated time points for FACS analysis. B. FACS profiles of propidium iodide-labeled cells and time course of cell cycle progression of cells irradiated in G1. HeLa cells were synchronized at M phase with nocodazole, then released and incubated in fresh medium for another 3 h. The cells were irradiated with 10

J/m2 UV, and then incubated in either the presence or absence of 5 µM arsenic, until harvested at the indicated time points. C. Time course of cell cycle progression of cells irradiated at the G1/S border. HeLa cells were synchronized at the G1/S boundary with 3

µM aphidicolin for 17 hours. 5 µM arsenic was added at the time of release, and the cells were incubated for an additional 2 h before irradiation with 10 J/m2 UV irradiation. After irradiation, cells were collected at the indicated time points.

95 REPORT-2

Figure 3. A. FACS profiles of cell cycle progression following UV irradiation in the presence and absence of p53. HCT116 cells were synchronized at M phase with nocodazole, then released and incubated in fresh medium for another 3 h. The cells were irradiated with 10 J/m2 UV, and then incubated in either the presence or absence of 5

µM arsenic until harvested at the indicated time points. B. Western immunoblot demonstrating the effect of arsenic on the expression of p53 in HeLa cells. HeLa cells were pretreated with 5 µM arsenic for 3 h, then irradiated with 10 J/m2 UV. Cells were incubated in fresh medium in the presence or absence of 5 µM arsenic and harvested at the indicated time points. The positive control was cell lysate from MCF-7 cells treated with the same dose of UV for 8 h.

Figure 4. A. Treatment protocol. HeLa cells were synchronized at the G1/S boundary with aphidicolin; 5 µM arsenic and 1.2 µM nocodazole were added after the aphidicolin release, and the cells were incubated for another 2 h before UV treatment. At various times after 10 J/m2 UV irradiation, the cells were pulse-labeled with 10 µM BrdU for 20 mins, and then collected for analysis. B. An image from FACS analysis with anti-BrdU

FITC. C&D. Effect of arsenic on S-phase checkpoint. S-active represents cells that are in S phase and actively synthesizing DNA (R2+R5+R6 population; with BrdU staining);

S-arrested represents cells that are in S phase but not synthesizing DNA (R7; DNA content 2N-4N, but no BrdU staining).

96 REPORT-2

Figure 5. A. Treatment protocol. HeLa cells were treated with 0.6 µM nocodazole throughout the time course to trap any cells that entered M phase in M phase. They were treated with 5 µM arsenic added 1 hour after the nocodazole, and 3 h later, the cells were irradiated with 10 J/m2 UV, and then collected at the indicated time points. B. An image from the FACS analysis with anti-phospho-histone H3 immunostaining. C. Effect of arsenic on the G2 checkpoint.

Figure 6. A. Arsenic alone arrests cells at M phase, not G2. Cells were collected 24 h after incubation with 5 µM arsenic, then fixed with 70% ethanol, stained with anti- phospho-Histone H3, and subjected to FACS analysis. B. Cytogenetic analysis of cells arrested by nocodazole and arsenic. HeLa cells were treated with either 0.3 µM nocodazole or 5 µM arsenic for 5 h. The cells were collected, fixed with 100% methanol, then stained sequentially with anti-phospho-histone H3 and anti-rabbit-Alexa Fluor 488

(green), and counterstained with 0.15 µg/ml DAPI, and then examined under a fluorescent microscope. C. Effect of arsenic on nocodazole-arrested M phase cells. HeLa cells were synchronized in M phase by nocodazole, then released and incubated in fresh medium, either with or without 5 µM arsenic, until they were harvested at the indicated time points. D. Effect of arsenic on M-phase arrest in primary human keratinocytes.

Primary human keratinocytes were treated with 5 µM arsenic and harvested at indicated time points for FACS with anti-phospho-histone H3.

97 REPORT-2 Figure 1

A As UV(0h) 8h 16h 24h 3hr

BrdU pulse-labeled 20 min before UV

B 0h 8h 16h 24h

R2 R2 R2 R2

R6 R6 R6 R6 Control R5 R5 R5 R5 R3 R3 R3 R3 R4 R4 R4 R4 R7 R7 R7 R7

R2 R2 R2 R2 B r

R6 R6 R6 R6 d R5 R5 R5 UV R5 U 2 10 J/m R3 R4 R3 R4 R3 R4 R3 R4 R7 R7 R7 R7

R2 R2 R2 R2

R6 R6 R6 R6 As R5 R5 R5 R5

5 uM R4 R4 R3 R4 R3 R4 R3 R3 R7 R7 R7 R7

R2 R2 R2 R2

R6 R6 R6 R6 R5 R5 R5 R5 UV+As R3 R3 R3 R3 R4 R4 R4 R4 R7 R7 R7 R7

DNA content

98 Figure 1 C

R5, early S phase/ G1 R2, S phase R6, late S phase/G2/M

50 30 40

40 30 s

20 s s l l l l l l e e e 30 c c c f f f 20 o o o

20 % % BrdU labeled % 10 10 10

0 0 0 0 8 16 24 hr 0 8 16 24 hr 0 8 16 24 hr 9 9 R3, G1 R7, S phase R4, G2/M 50 60 50 50 50 40 40 40 s s

40 l l s s l l l l l l e 30 e

30 e 30 c e c c f f 30 c f o o f o

o 20 % % 20 Unlabeled 20 20 % % 10 10 10 10 0 0 0 0 8 16 24 hr 0 0 8 16 24 hr 0 8 16 24 hr

0 8 16 24 hr R E P

Control UV As UV+As O R T - 2 Figure 2

A ) 7 R

( 14 s l l

e 12 c e 1 10 s 0 0 a

h 8 p UV S

d 6 UV +A s e l

e 4 b a l

n 2 u f

o 0 %

0 2 4 6 8 10 R E P

Time after UV irradiation (hour) O R T - 2 Figure 2 B

t s UV + As 1 0 1

DNA content

0h 5h 9h 13h 18h 24h

80 80 80 90 70 70 70 80

60 60 % 60 70 R % % M E / 50 60

1 50 50 S 2 G P n G 40 50 UV 40 i 40 n i O n s i l s 40 l UV+As l 30

30 s

l 30 R e l e l C

e 30 T C 20 20 20 C 20 - 10 10 10 2 10 0 0 0 0 0 5 10 15 20 25 0 5 10 15 20 25 0 5 10 15 20 25 0 5 10 15 20 25 30 35 Time after UV (hr) Time after UV (hr) Time after UV (hr) Time (hr) Time (hr) Time (hr) Figure 2 C

t s 1

0 UV + As 2

DNA content 0h 3h 6h 10h 15h 24h 34h

90 80 60 90 80 70 80 70 50 60 %

% 70

60 %

1 40 50 M

60 R / S

G 50 2 n E i n 40 50 UV G i 40 30 P s l s n l l i 40 UV+As

30 O l 30 e s e

l 20 C R

l 30

C 20

20 e T

C 20

10 10 10 - 10 2 0 0 0 0 0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 Time after UV (hr) Time after UV (hr) 0 5 10 15 20 25 30 35 Time after UV (hr) REPORT-2 Figure 3

0 hr 8 hr 16 hr 24 hr 36 hr 48 hr

-As HCT116 +UV p53(+/+) +As +UV

-As HCT116 +UV p53(-/-) +As +UV

103 Figure 3

10 J/m 2 UV 5 uM As 10 J/m 2 UV + 5 uM As

1 Positive 0 4 Time (hr) 0 4 8 13 18 0 4 8 13 18 24 0 4 8 13 18 24 control

P53 R E P O R T - 2 REPORT-2 Figure 4

Aph Aph release UV (0h) 5h 10h 15h 20h 17h 2h

As Noc BrdU pulse-label 20 min before harvest

R6 R5

R3 R4 R7

105 Figure 4

C D

90 40 control control 80 5uM As 35 5uM As 70 10J UV 10J UV As+UV 30 As+UV 60 % 1 25 % d 0 e e 6 50 t v i s

t 20 e c r r

a 40 - a -

S 15 30 S 10 20 10 5 0 0 0 5 10 15 20 25 0 5 10 15 20 25 Time after UV (hr) Time after UV (hr) R E P O R T - 2 REPORT-2 Figure 5

Noc As UV 1h 3h 1h 3h 1h 2h

107 Figure 5

l 20 1

e 0 8 %

c l

l e e 15

s C

a M

p 10 Noc As UV

M Noc UV 5 Noc As Noc

0 R E P 0 1 2 3 4 O R T -

Time after UV (hr) 2 Figure 6

Control UV As UV+As

G1 64% G1 18% G1 37% G1 37% S 15% S 23% S 12% S 38% G2/M 21% G2/M 59% G2/M 51% G2/M 25% 1 0

9 PI Staining

5.9% 4.0% 36% 4.0%

R2 R2 R2 Anti-Histone R2 H3 Staining

(M phase cell) R E P O R T - 2 REPORT-2 Figure 6

Noc 0.6 uM As 5 uM

DAPI

p-His H3 (mitotic Marker)

110 REPORT-2 Figure 6

0h 2h 4h

Noc

Noc+As

111 Figure 6

6 60 control control arsenic arsenic 5 nocodazole 50 nocodazole 1 1

2 4 40 % % M

3 / 30 M 2 G 2 20

1 10

0 0 0 6 12 18 24 30 0 6 12 18 24 30 Hours Hours R E P O R T - 2

DISCUSSION & FUTURE DIRECTIONS 3. DISSCUSSION AND FUTURE DIRECTIONS

3.1 Summary

Arsenic is a widely distributed human carcinogen associated with a variety of cancers. In the United States, arsenic has been found in half of the superfund sites, where it usually co-exists with other genotoxic pollutants. The possibility of arsenic’s co-mutagenicity and co-carcinogenicity has raised the concern that it may be necessary to take into account a possible synergy between arsenic and other genotoxic agents in risk assessment.

Currently, exposure risk is assessed for compounds independently. Lack of an appropriate animal carcinogenesis model makes arsenic risk assessment even more difficult. Furthermore, the uncertainty about mechanism(s) of arsenic carcinogenesis and co-carcinogenesis has hindered the development of definitive arguments for setting a drinking water standard for arsenic. Thus, a call for more research on the mechanisms of action of arsenic has been put on the national agenda.

In order to understand the mechanisms of arsenic co-mutagenicity and co-carcinogenicty, we have tested two hypotheses: (1) arsenic inhibits DNA repair; (2) arsenic suppresses

DNA damage-induced cell cycle checkpoints.

To test hypothesis (1), we have completed multiple experiments using low concentrations of arsenite in cultured human cells. Our results showed that at a concentration of 2.5 µM, arsenic induced insertion/deletion mutations in a mutagenesis model using a shuttle vector pZ189, and synergistically enhanced UV mutagenicity in the same model, while it

113

DISCUSSION & FUTURE DIRECTIONS did not alter the UV mutation spectra. In cell culture, arsenic did not increase the generation of UV-induced thymine dimers, but did inhibit the removal of the dimers. In addition, arsenic at 1-5 µM enhanced and prolonged RPA-p34 hyper-phosphorylation induced by UV irradiation, suggesting the persistence of DNA damage. Arsenic did not alter expression of several critical NER proteins or inhibit the incision of UV-induced photoproducts in an in vitro incision assay. Together, these results suggest that arsenic inhibits NER, and more specifically, probably only one subtype of NER, transcription- coupled repair.

To test hypothesis (2), we studied the ability of arsenic treatment alone to activate/deactivate checkpoints, as well as arsenic’s ability to modulate checkpoints induced by ultraviolet light (UV) irradiation. Our study showed that sodium arsenite alone at 5 µM did not markedly alter cell cycle progression in HeLa cells other than initiating an M-phase arrest. In contrast, following UV irradiation arsenite restored the

G1 checkpoint and enhanced the S and G2 checkpoints in HeLa cells. These results suggest that arsenic does not inhibit the activation of DNA damage checkpoints after UV, and therefore its function as a co-mutagen or co-genotoxin most likely does not occur via cell cycle checkpoint suppression. Instead, the observed enhancement of cell cycle checkpoints suggests an increase in DNA damage signaling, probably due to the inhibition of DNA repair.

3.2 Cell cycle checkpoints and DNA repair

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DISCUSSION & FUTURE DIRECTIONS 3.2.1 Enhanced DNA damage checkpoint may be due to the inhibition of DNA repair by arsenic.

The UV-induced S phase checkpoint is activated by replication fork stalling at sites of photoproducts in the DNA templates. Tercero et al. demonstrated that cells with replication forks stalled by MMS or HU could resume normal cell cycle progression when MMS/HU was removed from the medium, as evidenced by dephosphorylation of

RAD53, a hallmark of deactivation of S-phase checkpoint [36;37]. We observed by FACS analysis that UV-treated cells resumed normal cell cycle progression in the absence of arsenic, but in the presence of arsenic more UV-treated cells stopped DNA synthesis and they arrested longer or even permanently in S phase, reflecting inability to remove UV- induced damage. This is consistent with our data on DNA repair demonstrating that arsenic inhibits the removal of UV-induced photoproducts. Arsenic-induced DNA repair inhibition may also play a role in the restoration of the UV-induced G1 checkpoint and the arsenic-induced M-phase arrest. However, these effects may be masked by the indirect stabilization of p53 via inhibition of expression of viral protein E6 and by an effect that arsenic may have on some M-phase specific target.

3.2.2 Prolonged cell cycle arrest does not facilitate DNA repair in the presence of arsenic.

The function of the G1/S checkpoint is to ensure DNA integrity before the DNA is replicated in S phase. Normally the arrest would allow the cells to have more time to repair the DNA damage. A study in budding yeast indicated NER could be transcriptionally enhanced within the G1 phase in response to UV irradiation [38]. It has

115

DISCUSSION & FUTURE DIRECTIONS been suggested that p53 plays a role in NER, facilitating removal of damage, mainly by

GGR. Our studies showed that arsenic restored an 8-hr G1 checkpoint and elevated p53 levels after UV in HeLa cells. However, the removal of UV-induced thymidine dimers was not increased but it was rather inhibited. One possible explanation is that arsenic may inhibit only TCR as supported by a reporter gene assay we carried out that showed that arsenic seems to generally inhibit gene transcription (data not shown). Another line of evidence consistent with the current observation came from Linke et al., who observed that in cells with a normal p53, G1 delay did not seem to lead to the recovery of viability.

They argued that G1 checkpoint may not necessarily facilitate the repair, but just serve the purpose of eliminating damaged cells [39]. This evidence supports the notion that upregulation of p53 may not necessarily facilitate DNA repair or that it just facilitates

global genomic repair.

We also observed prolonged UV-induced S-phase arrest and G2 arrest in the presence of

arsenic. In response to DNA damage, normal cells activate cell cycle checkpoints, which slow down cell cycle progression in S phase or arrest cells in G2, depending on in which phase of the cell cycle the DNA damage occurred. At the same time, cell cycle checkpoints also signal to activate the corresponding DNA repair pathways to repair the damage, such as upregulation of XPC and p48 to facilitate NER [40]. However, in the

presence of arsenic, DNA repair is inhibited, so even though cells have gained extra time

and elevated expression of NER proteins, if the inhibition imposed by arsenic is not

removed, the longer time and elevated NER proteins will be futile for DNA repair. For

example, if the inhibition of DNA repair by arsenic is due to arsenic’s ability to inhibit

116

DISCUSSION & FUTURE DIRECTIONS histone acetylation, resulting in a more compact chromatin and thereby hindering access of repair proteins to the damaged DNA, as long as histone deacetylaton persists, the longer arrest and elevated NER proteins would be useless. In the situation where UV- induced damage can not be removed, cells either choose to bypass the damage or trigger apoptosis to eliminate the damaged cells.

Arsenic alone induces an M-phase arrest, which seems to act through target(s) within M phase. However, minor DNA damage and loss of checkpoint may also be involved. Cells are under constant attack by endogenous and exogenous insults. When the DNA repair function is intact, cells can handle most of these endogenous insults without detectable consequences. In the presence of arsenic, if DNA repair is inhibited, endogenous DNA damage or even minor DNA damage induced by arsenic itself could persist and accumulate because of the inhibition of DNA repair, ultimately inducing a DNA damage- induced arrest in M phase.

3.3 Consistency between the experiments

The results of all four assays we used to test the inhibition of DNA repair by arsenic suggest that arsenic at the concentration tested does not produce detectable DNA damage.

Thus, the enhanced DNA damage signal, as demonstrated by RPA-p34 hyper- phosphorylation and the potentiation of UV-induced mutation in the mutagenesis assay, can not be explained by an additive effect of arsenic and UV. Furthermore, the reduced removal of UV-induced photoproducts suggests that DNA repair is inhibited by arsenic.

Consistent with that, no DNA damage checkpoints (except the M-phase arrest) were

117

DISCUSSION & FUTURE DIRECTIONS activated with arsenic treatment alone. However, prolonged cell cycle arrest in S phase and G2 in cells treated with both arsenic and UV seem to be the result of inhibited DNA repair, leading to persistent DNA damage signals due to inability to remove DNA damage in the presence of arsenic.

In the two assays we used to measure the removal of UV-induced photoproducts, the patterns of dimer removal were slightly different. The comet assay showed differences in dimer removal only at later time points and the difference was large, as demonstrated by almost no type III to type II switch occurring in the presence of arsenic. In contrast,

FACS analysis showed differences in dimer removal throughout the time course we investigated and the difference was relatively small, about 20%. This could be due to the difference of these two assays. The comet assay used the T4 endonuclease to incise on the CPDs sites and generates single strand breaks. The length of comet tails correlate with the size of DNA fragments, which depend on the distance between two CPDs. If

DNA repair occurs randomly, we would expect to see a uniform lengthening of DNA fragments due to the removal of CPDs. If DNA repair does not occur randomly, say,

damage in transcribable genes would never get repaired due to the inhibition by arsenic,

the migration of the DNA fragment will lose correlation with the amount of

photoproducts, as reflected by lack of switch from type III damage to type II damage in

the presence of arsenic even though damage in non-transcribled area of DNA did get

repaired. FACS analysis using the thymine dimer-specific antibody measures the

photoproducts directly. The intensity of the fluorescence correlates with the amount of

photoproducts in DNA. In addition, T4 endonuclease recognizes CPDs including TT, CT,

118

DISCUSSION & FUTURE DIRECTIONS TC and CC dimers, while the antibody we used in FACS reacts specifically with only thymine dimers (TT). TT dimers only account for about 50% of CPDs [41]. Thus, T4

endonuclease detects more UV-induced photoproducts than the thymine dimer-specific

antibody, which could result in detection of larger differences on removal of

photoproducts by comet assay.

The difference we observed at DNA repair with and without arsenic with FACS analysis

seemed relatively small, while the difference on the duration of cell cycle checkpoints

seemed larger, as demonstrated by much longer cell cycle arrest, even permanent arrest,

for example, in S phase. This raised the question if the difference in cell cycle

checkpoints could be explained by the difference at DNA repair resulting from the effect

of arsenic. Cell cycle checkpoint are activated and maintained by DNA damage and

restored by the repair of DNA damage. How much UV-induced DNA damage is

sufficient to maintain a checkpoint is not yet clear, but in the situation of double strand

breaks, a single double strand break is enough to activate a checkpoint. The nature of

checkpoint is a signal transduction cascade. A weak signal can be amplified in the

cascade and become strong enough when it reaches the effectors. Thus, a small amount of

persistent DNA damage may be enough to hold cells in cell cycle arrest.

3.4 A possible unified explanation for both DNA repair inhibition and cell cycle

alteration by arsenic

We observed in this study an enhanced cell cycle arrest and reduced DNA repair

following UV irradiation in the presence of arsenic. The enhanced cell cycle arrest seems

119

DISCUSSION & FUTURE DIRECTIONS to be a secondary effect of inhibition of DNA repair by arsenic. While the mechanism of inhibition of DNA repair by arsenic is not clear, our study demonstrated that it is not through reducing the expression of NER proteins or interacting directly with NER proteins. Mutation spectra analysis suggests that the damage recognition/excision step is inhibited. One possibility is that arsenic prevents the access of repair proteins to the damaged DNA.

DNA is packed into nucleosomes. In all the cellular processes that require DNA as a substrate, this compact structure has to be loosened up to allow protein access to DNA, which is mainly achieved by histone modification, such as acetylation and methylation.

This process is called chromatin remodeling [42]. Arsenite treatment has been reported to

induce a severe deacetylation and demethylation of core histones [43]. It has been shown

that deacetylation of core histones will result in more compact chromatin and less

accessibility by proteins. If arsenic induces deacetylation, it will hinder the access of

NER proteins to the damaged DNA. Histone acetylation by acetyltransferase also serve

as a mechanism for regulation of gene transcription [42], arsenic’s effect on chromatin

remodeling may also affect gene transcription, which is consistent with our preliminary

observation that arsenic inhibits transcription of a reporter gene [data not shown]. The transcription-coupled repair pathway shares some components with the transcription machinery; thus, the way arsenic interferes with gene transcription may also apply to transcription-coupled repair. Either of these could explain the inhibition of DNA repair despite the negative results we observed in the in vitro incision assay. Chromatin changes caused by arsenic can also affect the interaction between chromosomes and spindle

120

DISCUSSION & FUTURE DIRECTIONS microtubules, which may underlie the M phase arrest. Thus, there is a possibility that both DNA repair inhibition and cell cycle alteration by arsenic could be due to the effect of arsenic on chromatin remodeling. More work is needed to confirm this hypothesis.

3.5 Relevance in human health effects: carcinogenesis and chemotherapy

We have examined the effect of arsenic on DNA repair and cell cycle checkpoints.

Arsenic alone at 1-5 µmol does not induce point mutations, while arsenic inhibits the removal of UV-induced DNA damage, supporting the hypothesis that arsenic might work as a co-mutagen or co-carcinogen. Similar to XP group cells with defects in NER, impaired DNA repair in arsenic-treated cells can result in increased cancer susceptibility.

Arsenic also enhances S and G2 checkpoints, probably due to the inhibition of DNA repair. The enhancement of cell cycle checkpoints seems to lead to the activation of the apoptotic pathway. Arsenic has been used as a chemotherapeutic agent, and the synergism of arsenic with other chemotherapeutic agents accelerates the apoptosis of cancer cells, which could favor cancer treatment.

Arsenic alone induces an M phase arrest, which seems to occur only in cells with defects in cell cycle checkpoints but not in normal cells. Arsenic-induced M phase arrest has been related to the ensuing apoptosis. This provides selective elimination of cancer cells since most cancer cells have defects in at least one or more checkpoints and have a rapid turnover rate. On the other hand, cells that arrested in M phase and later escaped the

121

DISCUSSION & FUTURE DIRECTIONS apoptosis accidentally, failed to segregate their chromosomes and could result in multi- nuclei cells (polyploidy), which could become a source of carcinogenesis.

3.6 Future directions

Although our results and previous studies have demonstrated arsenic’s effect on DNA repair and cell cycle checkpoints, there are still some unanswered questions that need follow-up studies.

1. Arsenic and DNA repair:

We observed the potentiation of UV-induced mutagenicity, persistence of DNA damage, and reduced removal of UV-induced photoproducts in the presence of arsenic, while we failed to see any changes in expression of DNA repair proteins and on incision activity in vitro. However, more would have to be done to identify the actual target and mechanism(s) for inhibition of DNA repair by arsenic.

1a. Interpretation of comet assay results and TCR:

Due to the limitation of available image analysis software, the comet assay was analyzed by a semi-quantitative method, which seemed to work better for analysis of this kind of atypical comet image. It would be useful to re-analyze the comet assay results in a more quantitative way that would allow an estimate of the extent of dimer removal and the determination of possible asymmetrical removal which could be indicative of TCR.

122

DISCUSSION & FUTURE DIRECTIONS Based on results of the comet and incision assays, we proposed that only transcription-

coupled repair may be inhibited by arsenic. A way to confirm that is to compare the

effect of arsenic on TCR in cells with selective defects in these two subtypes of NER. A

classic way to measure TCR is to monitor the removal of dimers on a specific target gene

using strand-specific probes [44].

1b. Repair complex formation:

In response to different types of DNA damage, cells activate corresponding repair

pathways and form different types of foci (repair complexes). The successful formation

of proper foci can be used as an indicator of repair function. Using different fluorescence-

tagged antibodies, now it is possible to visualize and examine these foci, associated with

double strand breaks, replication fork stalling, TCR, GGR and BER in the cells, allowing

us to monitor the possible effects of arsenic on the complex formation and association

with sites of DNA damage.

1c. Chromatin structure:

Our failure to observe any effect of arsenic on incision of adducts in the in vitro incision

assay could be due to an intrinsic limitation of the in vitro assay, which tests the action of

repair proteins on naked DNA. A chromatin binding assay could be used to test the

accessibility of damaged DNA in arsenic-treated cells to repair proteins. There have been

studies showing that arsenic has an effect on some of the proteins involved in maintaining

the structure of chromatin. For example, arsenic was found to stimulate histone H3

phosphoacetylation, which indicates an effect on chromatin proteins [45;46]. Furthermore,

123

DISCUSSION & FUTURE DIRECTIONS we have preliminary data showing that arsenic reduces gene transcription measured by a reporter gene system. This effect on transcription could be due to arsenic’s effect on chromatin as well.

2. Arsenic and cell cycle checkpoints:

Arsenic restores the UV-induced G1 checkpoint in HeLa cells, perhaps by inhibiting the expression of HPV viral protein E6 [47]. Um et al. also speculated that the AP-1 sites

located proximal to the upstream regulatory region of the E6 gene could be the target of

arsenic suppression of gene expression. Since arsenic has been known to induce oxidative

stress [48] and the activity of the AP-1 transcription factor is affected by the cellular

oxidative status [49], it would be worth following up this logic to see how arsenic exerts

its effect on E6 viral gene expression. This may have a general significance for HPV-

induced cancers, and the mechanism of E6 gene suppression may be generalizable to

more global effects of arsenic on gene expression in other human cells. Considering the

possible effect of arsenic on chromatin remodeling, one possibility is that arsenic might

inhibit E6 expression by remodeling chromatin structure. At the same time, fluorescent

probes can be used to monitor cellular oxidative status.

Arsenic alone induced a temporary but marked M-phase arrest, specifically at the

metaphase-anaphase transition. Although our data suggest that some M-phase specific

target must have been involved, the actual signal triggering the arrest and the direct target

upon which arsenic acts is unknown. Several mechanisms are possible to halt metaphase-

anaphase transition, including direct inhibition of APC/C and activation of spindle

124

DISCUSSION & FUTURE DIRECTIONS assembly checkpoint [50;51]. The mechanism by which APC/C activity is inhibited by

arsenic need further study. The activity of APC/C can be assessed by in situ detection of

its specific substrates. For metaphase-anaphase transition, cyclin A has to be destroyed

before the onset of anaphase [52]. The presence of cyclin A in arrested metaphase cells is

an indicator of activation of spindle assembly checkpoint. Another indicator of activation

of spindle checkpoint is the presence of the phosphorylated Mad2 at the kinetochore [50].

If all the abovementioned experiments can be completed, that will probably lead us to a much better understanding of arsenic’s effect on DNA repair and cell cycle checkpoints.

125

REFERENCES References

[1] P.C.Chan, J.Huff. Arsenic carcinogenesis in animals and in humans: mechanistic,

experimental, and epidemiological evidence, Environ.Carcino.& Ecotox.Revs.,

C15 (2), (1997) 83-122.

[2] T.G.Rossman. Molecular and genetic toxicology of arsenic, Environ Toxicol, 17,

(1998) 171-187.

[3] T.G.Rossman. Arsenic, in: Rom WN (Ed.), Environmental and Occupational

Medicine, 1998, pp. 1011-1019.

[4] H.Y.Chiou, S.T.Chiou, Y.H.Hsu, Y.L.Chou, C.H.Tseng, M.L.Wei, C.J.Chen.

Incidence of transitional cell carcinoma and arsenic in drinking water: a follow-up

study of 8,102 residents in an arseniasis-endemic area in northeastern Taiwan,

Am.J.Epidemiol., 153, (2001) 411-418.

[5] J.P.Buchet, R.Lauwerys. Study of inorganic arsenic methylation by rat liver in

vitro: relevance for the interpretation of observations in man, Arch.Toxicol, 57,

(1985) 125-129.

[6] H.V.Aposhian, R.A.Zakharyan, M.D.Avram, M.J.Kopplin, M.L.Wollenberg.

Oxidation and detoxification of trivalent arsenic species, Toxicol

Appl.Pharmacol., 193, (2003) 1-8.

[7] M.Vahter. What are the chemical forms of arsenic in urine, and what can they tell

us about exposure?, Clin.Chem., 40, (1994) 679-680.

126

REFERENCES [8] S.Lin, Q.Shi, F.B.Nix, M.Styblo, M.A.Beck, K.M.Herbin-Davis, L.L.Hall,

J.B.Simeonsson, D.J.Thomas. A novel S-adenosyl-L-methionine:arsenic(III)

methyltransferase from rat liver cytosol, J.Biol.Chem., 277, (2002) 10795-10803.

[9] A.Hartmann, G.Speit. Effect of arsenic and cadmium on the persistence of

mutagen-induced DNA lesions in human cells, Environ Mol.Mutagen., 27, (1996)

98-104.

[10] A.Maier, B.L.Schumann, X.Chang, G.Talaska, A.Puga. Arsenic co-exposure

potentiates benzo[a]pyrene genotoxicity, Mutat.Res., 517, (2002) 101-111.

[11] J.H.Li, T.G.Rossman. Mechanism of comutagenesis of sodium arsenite with N-

methyl-N-nitrosourea, Biol.Trace Elem.Res., 21, (1989) 373-381.

[12] J.H.Li, T.G.Rossman. Inhibition of DNA ligase activity by arsenite: a possible

mechanism of its comutagenesis, Mol.Toxicol, 2, (1989) 1-9.

[13] T.G.Rossman, A.N.Uddin, F.J.Burns, M.C.Bosland. Arsenite is a cocarcinogen

with solar ultraviolet radiation for mouse skin: an animal model for arsenic

carcinogenesis, Toxicol.Appl.Pharmacol., 176, (2001) 64-71.

[14] T.Okui, Y.Fujiwara. Inhibition of human excision DNA repair by inorganic

arsenic and the co-mutagenic effect in V79 Chinese hamster cells, Mutat.Res.,

172, (1986) 69-76.

127

REFERENCES [15] A.Hartwig, U.D.Groblinghoff, D.Beyersmann, A.T.Natarajan, R.Filon,

L.H.Mullenders. Interaction of arsenic(III) with nucleotide excision repair in UV-

irradiated human fibroblasts, Carcinogenesis, 18, (1997) 399-405.

[16] A.S.Andrew, M.R.Karagas, J.W.Hamilton. Decreased DNA repair gene

expression among individuals exposed to arsenic in United States drinking water,

Int.J.Cancer, 104, (2003) 263-268.

[17] Y.Hu, L.Su, E.T.Snow. Arsenic toxicity is enzyme specific and its effects on

ligation are not caused by the direct inhibition of DNA repair enzymes,

Mutat.Res., 408, (1998) 203-218.

[18] T.Schwerdtle, I.Walter, A.Hartwig. Arsenite and its biomethylated metabolites

interfere with the formation and repair of stable BPDE-induced DNA adducts in

human cells and impair XPAzf and Fpg, DNA Repair (Amst), 2, (2003) 1449-

1463.

[19] J.H.Petrini, T.H.Stracker. The cellular response to DNA double-strand breaks:

defining the sensors and mediators, Trends Cell Biol., 13, (2003) 458-462.

[20] S.J.Elledge. Cell cycle checkpoints: preventing an identity crisis, Science, 274,

(1996) 1664-1672.

[21] T.Weinert. DNA damage checkpoints update: getting molecular,

Curr.Opin.Genet.Dev., 8, (1998) 185-193.

128

REFERENCES [22] H.Beamish, R.Williams, P.Chen, M.F.Lavin. Defect in multiple cell cycle

checkpoints in ataxia-telangiectasia postirradiation, J.Biol.Chem., 271, (1996)

20486-20493.

[23] D.S.Lim, S.T.Kim, B.Xu, R.S.Maser, J.Lin, J.H.Petrini, M.B.Kastan. ATM

phosphorylates p95/nbs1 in an S-phase checkpoint pathway, Nature, 404, (2000)

613-617.

[24] T.Taniguchi, I.Garcia-Higuera, B.Xu, P.R.Andreassen, R.C.Gregory, S.T.Kim,

W.S.Lane, M.B.Kastan, A.D.D'Andrea. Convergence of the fanconi anemia and

ataxia telangiectasia signaling pathways, Cell, 109, (2002) 459-472.

[25] S.L.Soignet, P.Maslak, Z.G.Wang, S.Jhanwar, E.Calleja, L.J.Dardashti, D.Corso,

A.DeBlasio, J.Gabrilove, D.A.Scheinberg, P.P.Pandolfi, R.P.Warrell, Jr.

Complete remission after treatment of acute promyelocytic leukemia with arsenic

trioxide, N.Engl.J.Med., 339, (1998) 1341-1348.

[26] Z.X.Shen, G.Q.Chen, J.H.Ni, X.S.Li, S.M.Xiong, Q.Y.Qiu, J.Zhu, W.Tang,

G.L.Sun, K.Q.Yang, Y.Chen, L.Zhou, Z.W.Fang, Y.T.Wang, J.Ma, P.Zhang,

T.D.Zhang, S.J.Chen, Z.Chen, Z.Y.Wang. Use of arsenic trioxide (As2O3) in the

treatment of acute promyelocytic leukemia (APL): II. Clinical efficacy and

pharmacokinetics in relapsed patients, Blood, 89, (1997) 3354-3360.

[27] S.C.Novick, R.P.Warrell, Jr. Arsenicals in hematologic cancers, Semin.Oncol.,

27, (2000) 495-501.

129

REFERENCES [28] Z.Chen, G.Q.Chen, Z.X.Shen, G.L.Sun, J.H.Tong, Z.Y.Wang, S.J.Chen.

Expanding the use of arsenic trioxide: leukemias and beyond, Semin.Hematol., 39,

(2002) 22-26.

[29] R.Uslu, U.A.Sanli, C.Sezgin, B.Karabulut, E.Terzioglu, S.B.Omay, E.Goker.

Arsenic trioxide-mediated cytotoxicity and apoptosis in prostate and ovarian

carcinoma cell lines, Clin.Cancer Res., 6, (2000) 4957-4964.

[30] H.P.Tran, A.S.Prakash, R.Barnard, B.Chiswell, J.C.Ng. Arsenic inhibits the

repair of DNA damage induced by benzo(a)pyrene, Toxicol Lett., 133, (2002) 59-

67.

[31] W.H.Park, J.G.Seol, E.S.Kim, J.M.Hyun, C.W.Jung, C.C.Lee, B.K.Kim, Y.Y.Lee.

Arsenic trioxide-mediated growth inhibition in MC/CAR myeloma cells via cell

cycle arrest in association with induction of cyclin-dependent kinase inhibitor,

p21, and apoptosis, Cancer Res., 60, (2000) 3065-3071.

[32] S.Zhao, T.Tsuchida, K.Kawakami, C.Shi, K.Kawamoto. Effect of As2O3 on cell

cycle progression and cyclins D1 and B1 expression in two glioblastoma cell lines

differing in p53 status, Int.J.Oncol., 21, (2002) 49-55.

[33] B.L.Vogt, T.G.Rossman. Effects of arsenite on p53, p21 and cyclin D expression

in normal human fibroblasts -- a possible mechanism for arsenite's

comutagenicity, Mutat.Res., 478, (2001) 159-168.

130

REFERENCES [34] H.K.Hamadeh, M.Vargas, E.Lee, D.B.Menzel. Arsenic disrupts cellular levels of

p53 and mdm2: a potential mechanism of carcinogenesis,

Biochem.Biophys.Res.Commun., 263, (1999) 446-449.

[35] F.Chen, V.Castranova, Z.Li, M.Karin, X.Shi. Inhibitor of nuclear factor kappaB

kinase deficiency enhances oxidative stress and prolongs c-Jun NH2-terminal

kinase activation induced by arsenic, Cancer Res., 63, (2003) 7689-7693.

[36] J.A.Tercero, M.P.Longhese, J.F.Diffley. A central role for DNA replication forks

in checkpoint activation and response, Mol.Cell, 11, (2003) 1323-1336.

[37] J.A.Tercero, J.F.Diffley. Regulation of DNA replication fork progression through

damaged DNA by the Mec1/Rad53 checkpoint, Nature, 412, (2001) 553-557.

[38] A.D.Scott, R.Waters. Inducible nucleotide excision repair (NER) of UV-induced

cyclobutane pyrimidine dimers in the cell cycle of the budding yeast

Saccharomyces cerevisiae: evidence that inducible NER is confined to the G1

phase of the mitotic cell cycle, Mol.Gen.Genet., 254, (1997) 43-53.

[39] S.P.Linke, K.C.Clarkin, G.M.Wahl. p53 mediates permanent arrest over multiple

cell cycles in response to gamma-irradiation, Cancer Res., 57, (1997) 1171-1179.

[40] M.E.Fitch, I.V.Cross, J.M.Ford. p53 responsive nucleotide excision repair gene

products p48 and XPC, but not p53, localize to sites of UV-irradiation-induced

DNA damage, in vivo, Carcinogenesis, 24, (2003) 843-850.

131

REFERENCES [41] D.E.Brash, W.A.Haseltine. UV-induced mutation hotspots occur at DNA damage

hotspots, Nature, 298, (1982) 189-192.

[42] P.Cheung, C.D.Allis, P.Sassone-Corsi. Signaling to chromatin through histone

modifications, Cell, 103, (2000) 263-271.

[43] A.P.Arrigo. Acetylation and methylation patterns of core histones are modified

after heat or arsenite treatment of Drosophila tissue culture cells, Nucleic Acids

Res., 11, (1983) 1389-1404.

[44] K.Kobayashi, M.O'Driscoll, P.Macpherson, L.Mullenders, M.Vreeswijk,

P.Karran. XPC lymphoblastoid cells defective in the hMutSalpha DNA mismatch

repair complex exhibit normal sensitivity to UVC radiation and normal

transcription-coupled excision repair of DNA cyclobutane pyrimidine dimers,

DNA Repair (Amst), 3, (2004) 649-657.

[45] J.Li, M.Gorospe, J.Barnes, Y.Liu. Tumor promoter arsenite stimulates histone H3

phosphoacetylation of proto-oncogenes c-fos and c-jun chromatin in human

diploid fibroblasts, J.Biol.Chem., 278, (2003) 13183-13191.

[46] J.Li, P.Chen, N.Sinogeeva, M.Gorospe, R.P.Wersto, F.J.Chrest, J.Barnes, Y.Liu.

Arsenic trioxide promotes histone H3 phosphoacetylation at the chromatin of

CASPASE-10 in acute promyelocytic leukemia cells, J.Biol.Chem., 277, (2002)

49504-49510.

132

REFERENCES [47] S.J.Um, S.Y.Lee, E.J.Kim, J.Myoung, S.E.Namkoong, J.S.Park. Down-regulation

of human papillomavirus E6/E7 oncogene by arsenic trioxide in cervical

carcinoma cells, Cancer Lett., 181, (2002) 11-22.

[48] Y.Qian, V.Castranova, X.Shi. New perspectives in arsenic-induced cell signal

transduction, J.Inorg.Biochem., 96, (2003) 271-278.

[49] M.Karin, T.Takahashi, P.Kapahi, M.Delhase, Y.Chen, C.Makris, D.Rothwarf,

V.Baud, G.Natoli, F.Guido, N.Li. Oxidative stress and gene expression: the AP-1

and NF-kappaB connections, Biofactors, 15, (2001) 87-89.

[50] A.Mikhailov, R.W.Cole, C.L.Rieder. DNA damage during mitosis in human cells

delays the metaphase/anaphase transition via the spindle-assembly checkpoint,

Curr.Biol., 12, (2002) 1797-1806.

[51] J.M.Peters. The anaphase-promoting complex: proteolysis in mitosis and beyond,

Mol.Cell, 9, (2002) 931-943.

[52] T.T.Su, B.Jaklevic. DNA damage leads to a Cyclin A-dependent delay in

metaphase-anaphase transition in the Drosophila gastrula, Curr.Biol., 11, (2001)

8-17.

* (not including the references in Report-1 and Report-2)

133