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: ______
Toxicogenetic Studies in Drosophila: Using Fruit Flies to
Study Arsenic Toxicity
A dissertation submitted to
The Graduate School of the University of Cincinnati
In partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY (Ph.D.)
In the Department of Molecular Genetics, Biochemistry, and Microbiology of the College of Medicine
2008
By Jorge Gerardo Muñiz Ortiz B.S., University of Dayton, 2002
Committee Chair: Iain L. Cartwright, Ph.D. Abstract
Arsenic contamination of drinking water supplies around the world is considered the worst environmental disaster of recent times. Chronic consumption of arsenic can lead to an array of serious pathological outcomes, in some of which methylation of the metal may be a crucial component in determining toxicity. Differential responsiveness within human populations suggests inter-individual genetic variation also plays an important role. We have used
Drosophila melanogaster as a model to study arsenic response pathways because of unrivalled access to varied genetic approaches and significant overlap with many aspects of mammalian physiology and disease phenotypes. Genetic analysis of various strains exhibiting relative susceptibility or resistance to arsenite toxicity resulted in the identification of a chromosomal region able to confer a differential response phenotype. We created fly lines harboring small, overlapping deficiencies in this region and found that relative arsenite sensitivity arose when the glutathione synthetase (GS) gene dose was reduced by half. Knock-down of GS expression by
RNA interference both in S2 cells and in vivo led to highly enhanced arsenite sensitivity. These analyses provide genetic proof that an optimally functioning glutathione (GSH) biosynthetic pathway is required for a robust defense against arsenite. Moreover, they unexpectedly highlight a step previously considered to be without regulatory significance; the implications of this are discussed in the context of GSH supply and demand under arsenite-induced stress. Recent work has shown that Drosophila does not possess an arsenic methylation pathway comparable to the human. Since methylated arsenicals (MAs) may be key players in the carcinogenic activity of arsenic, we have “humanized” Drosophila through the introduction of the human arsenic(III) methyltransferase (hAS3MT) gene expressed under easily manipulated regulatory control.
Transgenic flies can be induced to express an antigenically cross-reactive form with arsenic
iii methyltransferase activity of the hAS3MT enzyme and its expression does not affect the development or viability when exposed or unexposed to arsenic. Preliminary results using an in
vivo genotoxicity assay have shown that production of MAs induces tumorigenesis in
Drosophila. This model is ready for use in exploring mechanisms of arsenic genotoxicity and/or
carcinogenicity in many informative genetic backgrounds, as well as the effects of different
polymorphic variants of AS3MT found in human populations.
iv
v Dedication
To my late abuela Epi, the matriarch and rock behind the Ortiz family, for teaching me what the
term family really means and for teaching me how to make paper boats and paper airplanes.
And
To my wife, Deborah María Sánchez Aceituno, who taught me to stand up for what I believe in
and to surround myself with those who improve my quality of life.
And
To my dad, Papi, Viejo, Jorge Muñiz Morales, who taught me to never ever give up and how to
always be a gentleman.
And
To my mom, Mami, Carmen Belén Ortiz Rivera, who taught me that nobody is better than me,
but I’m not better than anybody and to shine by my own light.
And
To my brother, Javy, Jorge Javier Muñiz Ortiz, because even though we have been far apart for
so long, it is as if we are not.
And
Para mi Isla del Encanto, Puerto Rico, que esto sea una fuente de inspiración para los que
sueñan…
(For my Island of Enchantment, Puerto Rico, may this be a fountain of inspiration for those who
dream…)
vi Acknowledgements
First and foremost, I would like to thank my advisor, Dr. Iain L. Cartwright because even
when it seemed that no progress was being made, we were able to move forward because of the
psychological support he provided. I owe you everything I’ve learned throughout my graduate career, for challenging me and for always wanting what is best for me as a scientist. Thank you for your support in every endeavor I wanted to undertake, especially when pledging and when planning to go to conferences. Who thought that Sir Iain from England, now an American citizen, the foremost figure in the English language, would be so patient with me, a native
Spanish speaker who uses English to survive.
To my committee members, Drs. Gary E. Dean, thank you for challenging me, Dan J.
Hassett, thank you for expecting plenty of me, Anil G. Menon, thank you for your positive feedback, it made me feel good, and Alvaro Puga, thank you for your expertise on arsenic.
I want to thank the Shull lab for teaching me Northern analysis, the Miller lab for teaching me Western analysis, the DeKoter lab for teaching me real time PCR, Weiss, Stringer,
Thompson, Kovall, Herr, Shertzer, and Caruso labs for use of their equipment. Thanks also go to the Air Force Research Lab for use of their equipment and their friendship. I want to express my sense of gratitude to Dr. Richard M. Weinshilboum for providing the antibody against the human hAS3MT, Dr. David J. Thomas for technical advice on use of antibodies and Dr. James
Lessard for providing the antibodies against actin (there’s plenty for the whole department for an immeasurable amount of years). I want to thank Dr. Bryan Mackenzie for his help on statistical analyses.
I would like to thank my wife, Debbie, for lighting a match under me and helping me deal with the pressures of lab work and morally helping me push through the hard times when it
vii seemed that graduation was more than 10 years away. I can finally be grateful for receiving a
Ph.D., but without your love I have nothing. May the adventures we live from now on mark our
future and leave a mark for generations to come.
My family, Papi, Mami and Javy, for their great support even though sometimes it is hard when they are miles and miles away. For always being there since the day I left home to pursue
a career en el Norte. However, with the advancement of cellular phone communication devices,
just a second away, hopefully they feel as if they have personally witnessed every failure and
every glory. Los quiero mucho, les debo la vida que vivo y para ustedes le dejo la muestra de mi
esfuerzo y mis riesgos que nunca fueron en vano.
To my student colleagues, postdocs, faculty and office employees, past and present,
which helped me throughout my graduate career, from simple experiments to complicated
matters of life. I would also like to acknowledge Dan Kane for his summer work on the FLP-
FRT strains and Mayank Patel for the time he kept our flies well fed.
To the MolGen Softball Team, the Banana Slugs Softball Team and Los Caimanes de
Cincinnati Professional Softball Team, for providing me with an athletic avenue in order to vent
my frustrations. To my best friends outside of science, who gave me an escape from the daily
routine in the lab: Luis, Tommy, Millo y Pancho.
viii Table of Contents
Abstract...... iii
Dedication ...... vi
Acknowledgements ...... vii
Table of Contents ...... ix
List of Abbreviations and Symbols ...... xiii
List of Figures and Tables...... xvi
Chapter I. An Introduction to Arsenic and Drosophila ...... 1
History of Arsenic...... 1
Arsenic Worldwide and Health Effects...... 1
Toxicology and Mode of Action...... 3
1. Oxidative Stress...... 3
1.1.1. Production of ROS...... 3 1.1.2. ROS-induced DNA Damage...... 6 1.1.3. Interaction of Arsenic-induced ROS with Signaling Pathways ...... 7
1.2. Production of Reactive Arsenical Species...... 8
2. Inhibition of DNA Damage Repair...... 9
3. Epigenetic Alterations ...... 10
Metabolism and Genetic Susceptibility...... 11
1. Metabolic Pathway...... 11
1.1. Arsenite and Arsenate ...... 11
1.2. Influx...... 11
1.3.1. Methylation in Mammals...... 12
ix 1.3.2. Arsenic Methylation: Detoxification or Activation?...... 13 1.3.3. Absence of Methylation in Some Mammalian Species ..15
1.4. Efflux...... 16
1.5. Thio-Arsenicals...... 16
2. Genetics...... 17
Mammalian Models of Arsenic Induced Carcinogenicity...... 20
1. Hamsters...... 21
2. Rats...... 21
3. Mice...... 22
Drosophila as a Model for Chemical Genotoxicity...... 25
1. Why Drosophila?...... 25
2. Tests for Genotoxicity...... 26
2.1.1. mwh-flr SMART...... 26 2.1.2. lats SMART ...... 28
2.2. SRLRT...... 29
2.3. vermilion gene alkylation...... 30
2.4. SCLT...... 31
Arsenic and Drosophila ...... 31
Conclusion ...... 33
Chapter II. Investigating Arsenic Suceptibility from a Genetic Perspective in
Drosophila Reveals a Key Role for Glutathione Synthetase ...... 35
Introduction...... 35
Materials and Methods...... 37
Results ...... 45
x
Discussion...... 58
Chapter III. Establishment and Characterization of a Drosophila Model to Investigate the
Biological Effects of Arsenic Methylation In Vivo ...... 63
Introduction...... 63
Materials and Methods...... 67
Results ...... 74
Discussion...... 82
Chapter IV. Summary and Future Directions...... 88
Introduction...... 88
Population Toxicogenetics: Identification of GS Polymorphic Variants and Possible Susceptibility to Arsenic Health Effects in Exposed Individuals Worldwide ...... 88
Cytological Analysis of Arsenic Exposed AS3MT Transgenic Lines..90
Tissue Transplantation of Neuroblast Stem Cells from Arsenic- exposed AS3MT Transgenic Lines...... 91
Phenotypic Studies on Arsenic Exposure of hAS3MT Expression on Intersecting Biological Pathways...... 92
Intersection of Folate Metabolism and Arsenic Methylation ...... 93
Studies on the AS3MT M287T Variant ...... 94
References...... 96
Appendix I. Description of Unpublished Negative Results ...... 119
1. RNAi Approach to Identify Genes Required for a Robust Response to Arsenite...... 119
xi 2. Functional Characterization of the Role of GS Towards Arsenite Sensitivity Via P-element Mobilization...... 120
3. Rescue of Arsenite-exposed S2 cells Using Mannitol as an H2O2 Scavenger ...... 121
4. Identification of ROS in S2 Cells After Exposure to Sodium Arsenite ...... 122
5. Biochemical and Genetic Studies to Identify a Differential Response Mechanism to Arsenite Between PVM and Oregon-R 1970 Strains ...... 123
References...... 127
Appendix II. Standard Drosophila Procedures...... 128
Determination of hAS3MT Chromosomal Insertion ...... 128
Creation of hAS3MT; latsx1 Fly Line...... 129
Preparation of Fly Genomic DNA...... 130
RNA Isolation...... 130
xii List of Abbreviations and Symbols
8-OHdG 8-hydroxy-2’-deoxyguanosine APL Acute Promyelocytic Leukemia APP Amyloid Precursor Protein AQP Aquaporin ARE Antioxidant Response Element Arg Arginine AsIII Arsenite AsV Arsenate AS3MT Arsenic(III) Methyltransferase ATG Arseno-triglutathione ATO Arsenic Trioxide B[a]P Benzo-[a]-pyrene BBN N-butyl-N-(4-hydroxybutyl)nitrosamine BER Base Excision Repair BLAST Basic Local Alignment Search Tool BPDE Benzo-[a]-pyrene-7,8-diol 9,10-epoxide CHDH Choline Dehydrogenase Cys Cysteine DENA Diethylnitrosamine Df Deficiency DMAIII Dimethylarsinous Acid DMAV Dimethylarsinic Acid DMBA 9,10 dimethyl-1,2-benzanthracene DMDTAV Dimethyldithioarsinic Acid DMMTAV Dimethylmonothioarsinic Acid DMTAIII Dimethylthioarsinous Acid EGFR Epidermal Growth Factor Receptor EMS Ethyl Methansulfonate ER-α Estrogen Receptor-α ERK Extracellular Signal-regulated Kinase EZH2 Enhancer of Zeste Homolog 2 FLP Flippase FRT Flippase Recognition Target GCLM γ-glutamyl-cysteine Ligase Modifier Subunit GFR Growth Factor Receptor GLRX Glutaredoxin GLUT Glucose Permease GS Glutathione Synthetase GSH Glutathione GST Glutathione S-transferase HA Hemagglutinin HB High Bioactivation HPLC High-performance Liquid Chromatography
xiii iAs Inorganic Arsenicals ICPMS Inductively Coupled Plasma Mass Spectrometry IKK IκB Kinase i.p. Intra peritoneal i.v. Intra venous JNK c-Jun N-terminal Kinase Leu Leucine LOH Loss of Heterozygosity MAPK Mitogen-activated Protein Kinase MAs Methylated Arsenicals MADG monomethylarseno-diglutathione MAT Methionine Adenosyltransferase MCL Maximum Contaminant Limit Met Methionine MMAIII Monomethylarsinous Acid MMAV Monomethylarsinic Acid MMC Methylmercuric(II) Chloride MMS Methyl Methanesulfonate MRP Multidrug Resistance Associated-protein MTHFR Methyltetrahydrofolate-homocysteine Methyltransferase Reductase MPO Myeloperoxidase NF-κB Nuclear Factor-κ B NER Nucleotide Excision Repair NO Nitric Oxide ODC Ornithine Decarboxylase OMIM Online Mendelian Inheritance of Man PARP Poly(ADP-ribose) polymerase PCR Polymerase Chain Reaction PDGFR Platelet Derived Growth Factor Receptor PML Promyelocytic Leukemia p.o. Per os PRDX2 Peroxiredoxin 2 Pro Proline RAR-α Retinoic Acid Receptor-α RNAi RNA Interference ROS Reactive Oxygen Species SAM S-adenosyl methionine SCLT Sex-chromosome Loss Test SLRLT Sex-linked Recessive Lethal Mutation and Reciprocal Translocation Test SMART Somatic Mutation and Recombination Test SNP Single Nucleotide Polymorphism SOD Superoxide Dismutase Thio-DMA Dimethylated Thio-Arsenicals Thr Threonine TMAO Trimethylarsine Oxide TPA Tetradecanoyl Phorbol Acetate
xiv Trp Tryptophan UAS Upstream Activation Sequence UVR Ultraviolet Radiation VEGF Vascular Endothelial Growth Factor VEGFR Vascular Endothelial Growth Factor Receptor XPG Xeroderma Pigmentosum Complementation Group G Protein bp Basepairs °C Degrees Celsius Ct Threshold Cycle Δ Delta g Grams γ Gamma h Hours l Liter μ Micro M Molar m Milli mm Millimeter min Minutes n Nano U Units s Seconds x g Times the Force of Gravity
xv List of Figures and Tables
Figure 1.1 Chemical structures of arsenic compounds detected in the urine of some
mammals...... 4
Figure 1.2 Suggested metabolic pathway schemes of arsenic in methylation-efficient
cells...... 13
Figure 1.3 Suggested metabolic pathway of methylated thio-arsenicals in humans and
rats...... 17
Figure 2.1 Viability of D. melanogaster strains in sodium arsenite...... 46
Figure 2.2 Relative adult eclosion percentages for male and female progeny resulting
from reciprocal crosses of the Oregon R and PVM strains...... 47
Figure 2.3 Recombination mapping between Oregon R 1970 & PVM using strain-
specific microsatellite markers on the X chromosome to locate an arsenite
tolerance/sensitivity locus...... 49
Figure 2.4 X chromosomal overlapping deficiencies (Df) created using the FLP-FRT
recombination system to aid in identification of an arsenite response locus.
...... 50
Figure 2.5 Arsenite sensitivity of X chromosomal deficiency lines...... 52
Figure 2.6 A Gclm mutant line is highly sensitive to arsenite exposure compared to a
wild type revertant...... 53
Figure 2.7 RNAi induced knock down of glutathione synthetase expression in S2 cells.
...... 55
Figure 2.8 Effects of RNAi-induced knock-down of glutathione synthetase in flies. ...57
Figure 2.9 GAL4 expression per se is not overtly toxic to Drosophila either in the
xvi presence or absence of arsenite...... 57
Figure 3.1 Molecular characterization of AS3MT transgene insertion and expression.76
Figure 3.2 Functional enzymatic analysis of hAS3MT activity via HPLC/ICPMS...... 78
Figure 3.3 Relative viability of varying genotypic configurations tested by adult eclosion
AsIII-free or AsIII-containing food...... 79
Table 3.1 Frequency of tumor formation per fly from various genotypic crosses and arsenic
treatments...... 81
Table 3.2 Frequency of tumor formation in non-lats and lats adults irrespective of AsIII
treatment ...... 81
Figure 3.4 Tumors observed in different fly tissues using the lats-SMART genotoxic assay
...... 81
Figure 4.1 Folate metabolic pathway ...... 94
Table A.1 Primer set used to identify potential imprecise excision of EP1322...... 120
Table A.2 Primer sets used to produced mRNA probes for Northern analysis...... 125
Table A.3 Primer sets used to target GS (CG6835) and CG32495 transcripts...... 126
xvii Chapter I.
An Introduction to Arsenic and Drosophila
History of Arsenic
Although arsenic (meaning “yellow orpiment” in Greek, arshenikon), the 33rd element in the periodic table, was not conclusively identified as an element until 1649 (reviewed in [1]), it has been used throughout ancient and in present times as a death-inducing poison when administered in large doses (60,000 parts per billion (ppb)). Hippocrates and Galen popularized the use of arsenic as a therapeutic and healing agent and, interestingly, there is evidence of a population in Styria, Austria called the “Arsenic Eaters” who consumed arsenic for various health purposes [2] (reviewed in [3]). Arsenic came in the form of tablets, pastes, solutions, and injectable chemical forms of 1% arsenic trioxide (ATO) called Fowler’s solution and was widely used in the 19th century. Arsenic has been used to treat skin conditions, stomatitis, and Vincent’s angina, a microbial-induced condition of the gums that leads to necrotizing ulcerative gingivitis.
Arsenic was also used to treat syphilis during World War II, and a compound by the name of arsphenamine, which contained 30% arsenic, was used as an injectable form to treat syphilis, yaws, and infections by different protozoans [4]. More recently, arsenic trioxide has been used as a therapeutic agent to treat acute promyeloctyic leukemia [5].
Arsenic Worldwide and Health Effects
Arsenic is a common contaminant of drinking water that is found in many underdeveloped and developed parts of the world (reviewed in [6]) such as Bangladesh [7-9],
India [10], Mexico [11, 12], Argentina [13-15], Chile [16, 17] (reviewed in [18]), Taiwan [19],
1 and the United States [14, 20-24]. Contamination of the drinking water supply has been
considered the worst environmental calamity of human kind, especially in Bangladesh, where
millions of tube wells were bored to curb the use of water from alternate sources like rivers and
open wells, common locations for cholera and other water-borne diseases [25]. In Bangladesh,
arsenic in contaminated tube-well water is used by approximately 80 million people [26] and can
be released from river sediments through the reduction of arsenic-associated ferric (hydr)oxides
after the seasonal addition of organic carbon, which can occur upon irrigation pumping [27] and
river flood waters [28, 29]. The presence of arsenic in water used for subsistence and its chronic
consumption can lead to a variety of malignancies (reviewed in [30]) of the bladder [14], kidney
[13], liver [31], lung [13], and skin [32], as well as vascular and metabolic diseases such as
Blackfoot disease [19] and diabetes mellitus [33] and other non carcinogenic diseases affecting
the respiratory [34] and peripheral nervous systems [35]. Recently, a study from a population in
the United States demonstrated that a high incidence in type 2 diabetes was due to inorganic
arsenicals (iAs) found in drinking water [36]. These carcinogenic and non carcinogenic
conditions are mostly due to the chronic consumption of arsenic present in the diet of exposed
individuals. However, acute consumption of inorganic arsenic (reviewed in [4]) can cause
vomiting, diarrhea, and gastrointestinal hemorrhage leading to death from fluid loss and
circulatory collapse [37]. Due to the worldwide health effects related to the consumption of
arsenic in drinking water, the World Health Organization [38] and the United States
Environmental Protection Agency [39] have set the maximum contaminant limit (MCL) at 10 ppb (10 µg/L).
2 Toxicology and Mode of Action
The carcinogenic mode of action of arsenic has yet to be fully understood, but many
potentially causative mechanisms have been proposed, such as the formation of reactive oxygen
species (ROS) that can oxidize DNA and proteins, inhibition of DNA repair mechanisms, or via
interaction with the epigenetic machinery.
1. Oxidative Stress
1.1.1 Production of ROS
Several possible mechanisms of ROS production via arsenic have been described. ROS can be
formed by the release of iron from horse spleen or human liver ferritin after exogenous exposure
to the methylated metabolites of arsenic, dimethylarsinic acid (DMAV) and dimethylarsinous
acid (DMAIII) (further discussed below) (for chemical structures of arsenic compounds see
Figure 1.1) [40]. The iron released from ferritin reacts with hydrogen peroxide (H2O2) via a
Fenton reaction producing the reactive oxygen species, hydroxyl radical (HO·), which reacts and
- damages DNA [41] and a hydroxyl anion (OH ). It has been suggested that H2O2 is produced by
oxidation of AsIII to AsV [42, 43], leading to activation of the NF-E2-related transcription factor 2
(Nrf2) that is known to be involved in the expression of antioxidant and phase 2 drug-
metabolizing enzymes via their antioxidant response elements (AREs) [44] (reviewed in [45]).
III V Conversely, H2O2 has been shown to react with arsenite (As ) to produce arsenate (As ), thus detoxifying the trivalent inorganic form to the less toxic pentavalent form [46].
3
Figure 1.1. Chemical structures of arsenic compounds detected in the urine of some mammals.
4 In some cell types arsenic exposure can induce the production of superoxide (O2·) by activating the multi-membrane associated and cytosolic subunits of the NADPH oxidase complex [47]. DNA damage occurs via NADH oxidase-mediated ROS produced by arsenic exposure in vascular smooth muscle cells [48], and a study suggests that the ultimate ROS
- inducing apoptosis in leukemic cells exposed to ATO is O2· , formed via arsenic activation of
NADPH oxidase [49, 50] or p38 [51], (discussed below in terms of arsenic inhibition of signaling cascades). However, other studies have suggested that H2O2 can also induce apoptosis of acute promyelocytic leukemia (APL) cells after treatment with ATO [52], and the inherent levels of ROS in leukemic cells may render such cells highly susceptible to ATO treatment [53].
APL (reviewed in [54]) results from a translocation between chromosomes 15 and 17 (t(15:17)), which fuses the retinoic acid receptor-α (RAR-α) gene with the promyelocytic leukemia (PML)
gene, leading to a disrupted RAR-α enzyme fused to PML. ATO has not only been used to treat
APL but has been shown to induce apoptosis of esophageal [50], gastric [55] liver [56], cervical
[57], and colon [58] cancer cells. Other mechanisms suggested for ATO-induced APL cell death
include reduction of the mitochondrial membrane potential [59], which leads to activation of the
apoptotic pathway at high concentrations of ATO [60]. In fact, it has been suggested that
another mechanism for build-up of ROS via arsenic is by its interaction with complex I of the
mitochondrial respiratory chain [61]. Further mechanisms proposed for ATO-induced APL
remediation in which ROS may not play a substantial role, include cleavage of the PML-RAR-α
fusion protein [62], which could induce partial differentiation of APL to myelocyte- or
metamyelocyte-like cells at a dose dependent manner [63], or by inducing the translocation of
the PML-RAR-α fusion protein, via the PML moiety, to nuclear bodies where degradation occurs
[64].
5 An indirect pathway that could lead to ROS production is by the sequestration of the ultimate cellular redox factor glutathione (GSH) (reviewed in [65]). Studies have shown that arsenic forms a triglutathione complex that can be effluxed from the cell [66-68]. This may decrease the intracellular levels of GSH, leading to an increase in the levels of ROS formed in cells via metabolic activity. Interestingly, the methylated metabolites of arsenic, MMA and
DMA, have been shown to decrease the levels of hepatic GSH in rats [69] and mice [70] with concomitant DNA damage (discussed below).
1.1.2 ROS-induced DNA damage
Individuals that show arsenicosis, defined as health effects due to chronic arsenic consumption, such as hyperpigmentation, hyperkeratosis or ulcerative lesions, also frequently show DNA damage such as chromosomal aberrations and sister chromatid exchanges [71].
Studies have shown that arsenic consumption induces oxidative DNA damage by inducing the formation of DNA adducts mediated by ROS such as HO·, peroxynitrite (a reactive nitrogen
- species formed by the reaction of nitric oxide (NO) and O2· ), and hypochlorous acid, formed
III from the reaction of H2O2 and myeloperoxidase [72, 73]. These As -induced ROS can oxidize
DNA bases, forming formamidoprymidine and/or 8-oxoguanine [73]. In fact, the presence of 8- hydroxy-2’-deoxyguanosine (8-OHdG) adducts in DNA has been used as a marker associated with DNA damage-induced malignancies caused by arsenic exposure [74, 75]. The hypothesis that oxidative DNA damage is induced by arsenic-generated ROS is supported by the fact that treatment with free radical scavengers such as Tiron, Trolox [76] and DMSO [77] reduces the mutagenicity.
6 1.1.3 Interaction of Arsenic-induced ROS with Signaling Pathways
ROS production from arsenic exposure has been suggested to be involved in the
activation of specific signal transduction pathways (reviewed in [78]). Direct activation of
signaling cascades such as the mitogen-activated protein kinase (MAPK) pathway (which includes the extracellular signal-regulated kinases (ERKs) [79], c-Jun N-terminal kinases (JNKs)
[80], and p38 protein kinases [81]) or via growth factor receptors (including the epidermal
growth factor receptor (EGFR), platelet derived growth factor receptor (PDGFR) or vascular
endothelial growth factor receptor (VEGFR)), could contribute to the carcinogenic mode of action of arsenic (reviewed in [82]). It has been suggested that p38 plays a role in the arsenic- induced expression of the pro-angiogenic vascular endothelial growth factor (VEGF) [83].
Robust MAPK signaling or growth factor signaling depends on a phosphorylation/dephosphorylation balance of serine/threonine (MAPK) or tyrosine (GFR) residues produced by numerous kinases and phosphatases (reviewed in [84]). Inhibition of phosphatase via oxidation of essential thiols in theses enzymes by arsenic-generated ROS activity would lead to constitutive activation of such signaling cascades (reviewed in [78]). For example, the AsIII-induced activation of the AKT pathway, which in turn activates NF-κB [85],
has been suggested to occur via oxidative-stress inactivation of a phosphatase [86], while
activation of the stress activated transcription factor AP-1, which is regulated by the MAPK
pathway [87], can be induced by the AsIII-specific inhibition of a JNK phosphatase [88]. The
proto-oncogenes that encode the components of AP-1, c-fos and c-jun, have been shown to be
activated in WI-38 human diploid fibroblast cells exposed to arsenic by stimulating the
phosphoacetylation of histone H3 at Ser10 [89], in contrast to non-treated cells, suggesting a role
for arsenic in epigenetic changes that can induce transcriptional activation and carcinogenesis
7 (discussed below). NF-κB is a stress activated transcription factor involved in cell-cell
interaction, cell proliferation, pathogenic signaling, and tumorigenesis (reviewed in [90]).
Currently, AsIII-induced activation of NF-κB, suggested to occur via ROS generation, is one of
the more hotly debated processes, since other studies have suggested that arsenic actually inhibits
NF-κB activity [91-94]. A potential toxic mechanism postulated to occur after cellular influx of
AsIII could be induced by its tight binding to thiols of proteins in affected tissues [95] (reviewed
in [4]). For example, AsIII has been suggested to bind to the cysteine 179 residue of the
activation loop of IκB kinase (IKK) inhibiting its activity, thus inhibiting the activity of the pro-
inflammatory NF-κB protein [93]. Along with AsIII [96], MMAIII and DMAIII have been shown
to bind metallothionein [97], a protein involved in a cellular response to heavy metal exposure such as arsenic [98]. When exposed to iAs, metallothionein knockout mice have been shown to
be more susceptible to liver and kidney damage, by increasing the frequency and severity of
lesions compared to wild-type mice [99].
Tyrosine phosphorylation of membrane growth factor receptors induces their activation,
leading to activation of genes required for cell proliferation. Studies have shown that AsIII treatment stimulates phosphorylation of the EGFR and the proto-oncoprotein Shc [79], with the arsenic-induced activation of EGFR and the ERKs being dependent upon activation of the non- receptor tyrosine kinase c-Src [100] (reviewed in [101]).
1.2 Production of Reactive Arsenical Species
Even though many studies have suggested that ROS are the main factors in arsenic- induced carcinogenesis, other studies have shown that reactive arsenical species could also lead to tumorigenesis. After administering DMA to male mice, lung-specific DNA single stranded breaks were observed, but the levels of free radicals did not decrease when treated with the ROS
8 scavengers superoxide dismutase (SOD) or catalase, leading to the speculation that other types of reactive species could be responsible for induction of lung-specific DNA damage. Additional experiments identified the formation of the dimethylarsenic peroxyl radical [(CH3)2AsOO·]
[102]. The DMA peroxyl radical, as well as the DMA radical [(CH3)2As·], are thought to be
produced by the reaction of molecular oxygen (O2) with dimethylarsine, a further reductive
metabolite of DMA [103].
2. Inhibition of DNA Damage Repair
Defects in the DNA repair machinery would cause replication of damaged DNA, which in turn may ultimately induce cellular transformation and carcinogenesis. Nucleotide excision repair (NER) has been shown to be inhibited by AsIII or MMAIII in human cells treated with
either UV irradiation [104] or with the DNA adduct benzo-[a]-pyrene-7,8-diol 9,10-epoxide
(BPDE) [105]. Additionally, it has been demonstrated that MMAIII and DMAIII can inhibit the
base excision repair (BER) pathway [105]. One particular enzyme involved in the DNA repair
pathway that has been suggested to be inhibited by arsenic is poly(ADP-ribose) polymerase
(PARP) [106, 107]. The PARP protein is one of the first enzymes to respond to DNA damage
induced by arsenic exposure (reviewed in [108]). The inhibitory activity of arsenic towards
DNA repair enzymes was thought to occur via interactions with the sulfhydryl groups of the
proteins [109]; however, one study showed that this inhibition was independent of such
interaction and that other factors involved in protein phosphorylation/dephosphorylation or
signaling pathways deregulated by arsenic exposure could be responsible for the down regulation
in DNA repair [110]. Other studies have shown that AsIII can inhibit DNA ligase II activity
[111] and/or delay the activity of DNA ligase III [109], causing the cell to progress through cell
9 division regardless of DNA damage, and thereby suggesting a potential mechanism for arsenic- induced carcinogenicity.
3. Epigenetic Alterations
A number of studies have suggested that the carcinogenic mode of action of arsenic could be induced by epigenetic changes, such as DNA hypomethylation or hypermethylation [112], an increase in dimethylation of lysine residue 9 and tri- and dimethylation of lysine residue 4, a decrease in trimethylation of lysine residue 27 of histone H3 [113], and/or decreased acetylation of histone H3 [114]. An increase in H3K9 dimethylation, which induces silencing, might be explained by the fact that the mRNA and protein levels of the G9a histone methyltransferase, previously shown to methylate H3K9 [115], were increased after arsenic-treatment of the lung carcinoma cell line A549 [113]. It was further suggested that H3K9 and H3K4 dimethylation could be induced by the interaction and interference of arsenic with LSD1, a protein involved in histone demethylation [113]. Methylation of H3K4 occurs via the COMPASS-like methyltransferase complex [116], however as of yet no mechanisms on its possible interaction with arsenic have been proposed. Trimethylation of H3K27, which silences gene transcription, is catalyzed by enhancer of zeste homolog 2 (EZH2), a methyltransferase phosphorylated by
Akt. Akt signaling, which is activated by arsenic [86], leads to the phosphorylation of EZH2 inhibiting its catalytic activity [113]. Arsenic-induced DNA hypomethylation has been observed genome-wide but specific studies were performed on the promoter region of the estrogen receptor-α (ER-α) in rodent liver cells [117, 118] due to the fact that ER-α hypomethylation can lead to its increased expression, which has been associated with arsenic-induced hepatocarcinogenesis by altering estrogen signaling [119]. S-adenosyl methionine (SAM), used as a cofactor to methylate DNA, is also used by a different methyltransferase to methylate
10 inorganic arsenic (explained in more detail below), such that if the SAM pool decreases it may
lead to DNA hypomethylation and subsequent aberrant gene expression [120]. In contrast, after treatment with AsIII or AsV, hypermethylation of the p53 promoter, which suppresses gene
transcription, was observed in a human lung adenocarcinoma cell line [121]. Suppression of p53
expression could lead to cellular division with unrepaired DNA damage and thus promotion of
tumorigenesis. DNA hypermethylation occurs with concomitant increase in histone H3
deacetylation [114], which is induced by the recruitment of deacetylases via the transcriptional
repressor MeCP2, which binds to methylated DNA [122, 123].
Metabolism and Genetic Susceptiblity
1. Metabolic Pathway
1.1 Arsenite and Arsenate
In nature, arsenic is present in two different oxidative states, AsIII or AsV. AsV can
substitute for phosphate in ATP synthesis thereby uncoupling oxidative phosphorylation
(reviewed in [4, 124]), and as described above, AsIII can induce its toxicity via different
mechanisms, such as its ability to induce production of ROS, which could damage DNA or
oxidize signaling pathway proteins, alter the epigenome leading to aberrant gene expression, by
interfering with the DNA damage repair mechanism, and/or by tightly binding to thiols of
proteins inhibiting their enzymatic activity.
1.2 Influx
After consumption, AsIII can be taken up by cells via the aquaglyceroporin channels
AQP7 and AQP9 [125, 126] or via the glucose permease GLUT1 [127], whereas AsV can be
taken up by cells via phosphate transporters [128, 129] and is quickly reduced to AsIII [130]. As
11 reviewed above, trivalent arsenicals bind metallothionein, however it is not known whether this
interaction affects transport of iAs across the plasma membrane.
1.3.1 Methylation in mammals
Once inside mammalian cells, inorganic arsenicals (iAs) can be metabolized to produce
mono or dimethylated and trivalent or pentavalent forms (Figure 1.2). Different reaction
schemes by which this is accomplished have been proposed: (1) iAs undergo a series of successive reduction and oxidative methylation reactions [131] or (2) the reduced inorganic
form, AsIII, is conjugated to GSH and undergoes a series of successive methylations to form
monomethylarsonous acid (MMAIII) and DMAIII, each conjugated to GSH and providing a
substrate for oxidation to the +5 state to form monomethylarsonic acid (MMAV) and DMAV
[132, 133], these being the most prevalent species in urinary excretions [134] (schemes reviewed in [135]). In scheme (1) an AsV reductase identified as glutathione S-transferase omega (GSTO)
has been implicated by in vitro studies [136, 137], however 20% of the arsenic-reducing activity was retained in GSTO knock-out mice compared to wild-type [138]. Either these results suggested the presence of another enzyme capable of reducing the pentavalent arsenicals to trivalent forms or that residual GST enzymatic activity could catalyze the reaction. The enzyme arsenic(III) methyltransferase (AS3MT) [139] can provide the methylating activity necessary in scheme (2), and it has been suggested that it could also function in scheme (1) by linking oxidative methylation with reduction of arsenic metabolites, using GSH as a cofactor, thereby obviating the need for GST enzymatic activity [132, 140, 141]. The capacity to methylate iAs and produce DMA and MMA is strongly dependent on the AS3MT enzyme as shown by performing RNA interference studies in HepG2 human hepatic cells and demonstrating that the levels of methylation decreased by 70% compared to WT cells [142]. The authors point out that
12 production of methylated arsenicals (MAs) might be independent of AS3MT, albeit at a low
level, based on their percentages of mRNA knockdown (by 83%) and decrease in protein
expression (by 88%) after shRNA treatment and the methylation capacity (30%) [142].
However, since the knock-down of the mRNA transcripts is not 100%, the remaining enzymatic
levels might be enough to produce methylated arsenicals (MAs).
Figure 1.2. Suggested metabolic pathway schemes of arsenic in
methylation-efficient cells.
1.3.2 Arsenic Methylation: Detoxification or Activation?
Methylation of arsenic was previously thought to be a detoxification process based on
LD50 determinations of acute treatment with iAs, which can be considered unsophisticated [143].
Additionally, it had been suggested that methylation of iAs is a detoxification process because it
leads to higher levels of excretion of MAs due to the fact that MAs are more water soluble than iAs [144-146]. An important reason as to why it was previously thought that methylation of
arsenic functioned as a detoxification process was that once iAs were ingested, and underwent
metabolism, they were mostly excreted as DMAV and to a lesser extent as MMAV, as detected in
the urine of exposed individuals [147] and experiments performed using MMAV and DMAV
13 showed that they were less acutely toxic than iAs [69]. The detection methods of arsenicals in
human urine used earlier were less sophisticated than the methods currently used, therefore
MMAIII and DMAIII were not observed in the urine of exposed individuals and thus toxicity
studies were not performed using the trivalent organic arsenical forms. However, once the
methodology and the technology used for arsenical detection improved and the trivalent MMA
[148] and DMA [149] were detected studies were performed to determine their toxic potential.
III The LD50 of hamsters exposed to acute treatments of As was 4-fold higher than the LD50 of
MMAIII, suggesting that MMAIII is more toxic than AsIII and suggesting that methylation of
arsenic is normally not a detoxification process [150]. Studies have shown that the methylated trivalent forms of arsenic are more genotoxic by their ability to nick DNA in vitro, whereas the
methylated pentavalent and iAs did not [151]. Additionally, when human hepatocytes were
exposed to MMAIII they became more susceptible to cell death than when exposed to AsIII, proving that MAs are more cytotoxic than their inorganic counterparts [152, 153]. MMAIII and
DMAIII were shown to be more active in inducing chromosomal aberrations, such as DNA strand
breaks, than AsIII, AsV, MMAV and DMAV in human lymphocytes [154]. Strand breaks
observed after exposure to DMAIII appear to occur spontaneously, however results showed that
DMAIII and DMAV exposure blocked cells at metaphase, suggesting that DNA strand breaks
could also occur during DNA replication leading to aneuploidy or polyploidy [154].
Chromosomal damage induced by MAs has been attributed to the production of ROS (mainly
HO·) and their reaction with DNA bases, as described above [74]. Additionally, rat heart
microvessel endothelial cells and HL60 cells were more susceptible, as demonstrated via a
cytotoxic assay measuring cell viability, to the metabolic intermediate dimethylarsino-
glutathione (DMAG) than to iAs [155]. Studies on individuals from southwestern Taiwan
14 chronically exposed to arsenic-contaminated drinking water have correlated the efficiency of the
first iAs methylation step and low efficiency of the second methylation step, or the ratio of
MMA to DMA in their urine, with skin [156] and bladder [157] cancer risk, and also showed that
men were at a higher risk than women [157, 158]. Identical observations have been made in
populations from Argentina and the United States regarding high risk for bladder cancer [14] and
from Bangladesh with respect to high risk of developing skin lesions [159].
In conclusion, the higher the efficiency for the first methylation step of iAs, which
produces MMAIII, the higher the risk to develop either arsenic-related carcinogenic or non-
carcinogenic health effects. The higher MMAV:DMAV ratio in the urine of individuals exposed
to arsenic through their diet could be due to a lower affinity of the AS3MT enzyme for MMAIII, which is quickly oxidized to MMAV [148], as the studies show that high cumulative arsenic exposure leads to higher MMAV:DMAV ratios, which could potentially lead to higher levels of free MMAIII and, thus higher cancer risk [156].
1.3.3 Absence of Methylation in Some Mammalian Species
As described above it is well known that humans methylate inorganic arsenic once
transported into cells [147]. However, studies on the enzymatic activity of arsenic
methyltransferase identified several mammals that lack the ability to methylate iAs, e.g. great
apes such as the chimpanzee [160], gorilla [161] and orangutan [161], New World monkeys such
as the marmoset [162] and tamarin [162], and prosimians such as lemurs [161]. Old World
monkeys such as the rhesus, pig-tailed and long-tailed macaques have the ability to methylate
iAs [161]. Rodents that can methylate iAs include hamsters [163], mice [164], rats [165], and
rabbits [143] but not guinea pigs [166]. The inability of chimpanzees to methylate iAs is due to
the fact that a frameshift mutation in the AS3MT gene creates a premature stop codon that codes
15 for a 205 amino acid protein (compared to the 375 amino acid protein in humans) and also lacks
a well-defined cavity for Cys156, which has been suggested to be essential for the catalytic activity of the AS3MT enzyme [167].
1.4 Efflux
Efflux of xenobiotics from cells often consists of conjugation of glutathione (GSH) to
substrate via glutathione S-transferase (GST) and subsequent export by an ATP-dependent
transporter [168]. The multidrug resistance associated-protein 2 transporter (MRP2) has been
shown to transport arseno-triglutathione (ATG) and monomethylarseno-diglutathione (MADG),
and GSH is required for transport into rat bile after i.v. administration of AsIII [169]. GSTP1-1
has been suggested to play an important role in the secretion of a GSH-As complex via the ABC multidrug resistant transporter family member MRP1 [66]. Urinary profiles of mice administered sodium AsIII showed the presence of both ATG and MADG [67]. However, direct
data are lacking on whether a GST protein is essential for enzymatic GSH conjugation to
arsenicals for efflux, due to the fact that formation of ATG complexes can occur nonenzymatically [132].
1.5 Thio-Arsenicals
Recent studies of pregnant women chronically exposed to arsenic-contaminated tube-well water identified the presence of dimethylated thio-arsenicals (thio-DMA) in their urine, suggesting that an additional metabolic process occurs, at least in mammalian cells [170] (see
Figure 1 for chemical structures of thio-arsenicals). It has been suggested that inorganic arsenic is metabolized to DMAIII in hepatocytes, released into the bloodstream and taken up by red blood cells [171], where it is converted to dimethylmonothioarsinic acid (DMMTAV) by forming the
intermediate dimethylthioarsinous acid (DMTAIII) through a reaction with HS- [172]. DMMTAV
16 is then hydrolyzed into DMAV and HS-, which reacts with two molecules of DMMTAV to produce dimethyldithioarsinic acid (DMDTAV) and more DMAV, which can be reduced to
DMAIII [173] (Figure 1.3). Transformation of DMMTAV to DMDTAV can only occur in the presence of both GSH and Na2S [173]. Furthermore, thio-DMAs have been shown to be more cytotoxic than DMAs, and are capable of producing ROS [174]. For example, DMMTAV was
shown to be more toxic to human epidermoid carcinoma A431 cells than AsV and DMAV, but similar in toxicity to AsIII and DMAIII [174]. Using the hepatocarcinoma cell line HepG2,
cytotoxic effects of thio-DMAs, such as DNA fragmentation and cleavage of PARP, were
enhanced in the presence of 1 mM to 3 mM GSH [175].
Figure 1.3. Suggested metabolic pathway of methylated thio- arsenicals in humans and rats.
2. Genetics
It has been proposed that inter-individual variation in susceptibility occurring in populations chronically exposed to arsenic could have a genetic component (reviewed in [176]).
Differences in inorganic arsenic metabolism, identified after establishing a ratio between total levels of arsenic versus inorganic arsenicals and/or methylated arsenicals excreted in the urine of
17 individuals, have been observed between both children [15] and native Andean women [177] chronically exposed to arsenic-contaminated drinking water when compared to other populations. In patients with arsenic-correlated urothelial carcinoma, higher percentages of
MMAV compared to DMAV have been observed compared to healthy individuals with similar cumulative levels of As consumption [178]. A comparison of the ratios of total arsenic to arsenic metabolites (iAs and MAs) in the excreted urine of individuals in studied populations from
Mexico, China, and Chile has supported the idea that inter-individual differences in toxicity could be due to functional polymorphisms in genes whose enzymatic products are involved in metabolizing the metal [179]. Such studies are consistent with the idea that polymorphisms in the AS3MT gene that produce an enzyme with higher methylation efficiency could lead to higher susceptibility to arsenic-induced health effects (discussed below).
Studies have yet to identify potential polymorphisms in the AQP7, AQP9 transporter genes, which might lead to differential susceptibility to health effects from arsenic consumption.
Polymorphisms in GSTO [180] and in AS3MT [176, 181-184], specifically the M287T mutation in AS3MT, have been correlated with individual differences in urinary metabolic arsenic profiles. The AS3MT M287T mutant has been linked to higher enzymatic activity [185] leading to higher methylation efficiencies [186]. Higher methylation efficiencies could be detrimental to cells because of an intracellular increase in MAs, which, as discussed above, are more toxic than iAs. Although a required role for GSTP1-1 has been suggested in the cellular export of an As-
GSH complex [66], several studied polymorphisms in this enzyme were not associated with differences in As metabolism or incidence in skin lesions [187]. In addition, while susceptibility to arsenic exposure has not yet been correlated with polymorphisms in the ABC transporter genes, mdr1a/1b(-/-) double knockout mice were more sensitive than wild type mice due to
18 accumulation of arsenic in their tissues [188]. This suggests that individuals expressing a non-
functional (or reduced function) MDR transporter could potentially become susceptible to the
toxic effects of arsenic consumption.
Polymorphisms in enzymes involved in the production of cofactors required for arsenic metabolism (e.g. GSH and SAM) or in a robust response against the toxic effects of arsenic,
could also potentially render individuals chronically exposed to the toxicant susceptible to its
health effects. Thus, arsenic metabolism could be affected by polymorphisms in 5-
methyltetrahydrofolate-homocysteine methyltransferase reductase (MTHFR) [184, 189, 190], an
important enzyme involved in the early stages of biosynthesis of the AS3MT cofactor SAM.
Other candidates harboring relevant polymorphisms are methionine adenosyltransferase (MAT),
which joins methionine to the S-adenosyl moiety to produce SAM, and choline dehydrogenase
(CHDH) [191]. CDHD is an enzyme involved in an alternative pathway for methionine
synthesis (which occurs primarily by methylation of homocysteine into methionine using folate
as the methyl donor) by forming betaine from choline, with betaine providing the methyl group
to methylate homocysteine to methionine [192]. Polymorphisms in the glutaredoxin (GLRX)
gene, which uses GSH as a cofactor in an arsenic-coupled reduction system, which also includes
peroxiredoxin 2 (PRDX2), have also been associated with differential metabolism of arsenic
[141, 183, 192, 193]. Higher sensitivity towards AsIII and the induction of a variety of arsenic-
related conditions such as hyperkeratosis and hypertension have been correlated with polymorphisms in p53 [194] and NADPH oxidase [195]. Polymorphisms correlated with susceptibility to arsenic have also been identified in genes coding for the ROS scavengers manganese SOD [195], catalase [196], and myeloperoxidase (MPO) [196], and in three genes
involved in DNA damage response, the NER gene ERCC2 [197], the BER gene XRCC1 [198],
19 and ATM [199]. For example, higher risk in development of arsenic-induced premalignant
hyperkeratosis has been attributed to polymorphisms in ERCC2 [197], while individuals carrying
the Arg194Trp mutation in the XRCC1 enzyme were at a higher risk for urinary transitional cell
carcinoma [198].
In conclusion, observed inter-individual differences in the metabolism of arsenic leading to susceptibility can be associated with polymorphisms in genes involved in its metabolic pathway. Polymorphisms in genes involved in intersecting pathways, such as singaling cascades, DNA repair or redox maintenance could exacerbate the toxic potential of arsenic and its metabolites. Therefore, studies on arsenic-exposed populations should be undertaken to correlate potential genetic susceptibility to the toxic effects of arsenic that could lead to an increased risk for developing arsenic-related health conditions.
Mammalian Models of Arsenic Induced Carcinogenicity
The relative lack of understanding regarding the mechanism of arsenic carcinogenicity has not been helped by the absence of a good mammalian model that could be used to identify specific mechanisms for arsenic as a carcinogen (reviewed in [200, 201]). As described previously, many primates genetically similar to humans do not possess AsIII methyltransferase
activity (reviewed in [202]), thus, assuming that AS3MT enzymatic activity is relevant for As-
induced carcinogenicity, eliminates them from the possibility of establishing a model of arsenic
carcinogenicity that would mimic the human. However, studies in cynomolgus monkeys
(Maccaca fascicularis) fed AsV did not induce the development of malignant tumors even though
they possess arsenic methyltransferase activity [203]. Wildfang et al. point out that arsenic
studies in any of the great apes (gorilla, orangutan, and chimpanzee) would be controversial due to the fact that the apes are classified as endangered and/or threatened species by the U.S. Fish
20 and Wildlife Service and other international wildlife associations such as the United Nations
Environment Programme (UNEP) and the International Union for Conservation of Nature and
Natural Resources (IUCN) [161]. Therefore, attempts to establish hamster, rat and transgenic
and non-transgenic mice model systems have been made [204].
1. Hamsters
In one experiment, 2 out 10 Syrian golden hamsters and 2 out of 20 in a second
experiment showed growth of lung adenomas after intratracheal instillations of different concentrations of ATO once a week for 15 weeks [205]. The difference between the two experiments was the concentration of ATO used. In experiment one, hamsters were exposed to a total of 5.25 mg ATO throughout the 15 weeks and in experiment two hamsters were exposed to
3.75 mg, which could explain the lower totals of transformed hamsters [205]. More recently,
Syrian hamster embryonic cells treated chronically for 48 hours with 3-10 µM AsIII or 50-150
µM AsV were passaged and tested for neoplastic transformation by anchorage-independent
growth in semisolid agar and for malignant growth in newborn hamsters [206]. Some of the
isolated cell clones induced tumors in newborn hamsters and showed amplification and
hypomethylation of the c-myc and c-Ha-ras genes [206].
2. Rats
To test the carcinogenic potential of AsIII, Wistar rats were exposed to AsIII alone or pretreated with diethylnitrosamine (DENA) before being fed AsIII ad libitum [207]. Organ
analysis of cotreated Wistar rats did not show formation of hepatic tumors, but the kidney
showed tumor growth. Wistar rats treated only with AsIII did not develop tumors in the kidney
[207]. Wistar rats treated only with DMAV did not induce formation of tumors in the kidney,
21 however in rats pretreated with DENA, a tumor initiator, tumors were observed but the increase
in incidence was not statistically significant [208]. Liver analysis of Wistar rats cotreated with
DENA and DMAV showed the presence of lesions, although no hepatocellular carcinomas were
observed [208]. These studies suggested that DMAV only acts as a tumor promoter in the kidney and the liver. Similar results were observed when Fischer 344 (F344) and NCI-Black-Reiter rats
were pretreated with N-butyl-N-(4-hydroxybutyl)nitrosamine (BBN) and later treated with
DMAV [209, 210]. Both rat strains were exposed to DMAV via drinking water with or without
BBN and those who were treated with both DMAV and BBN showed a DMAV dose dependent
incidence and multiplicity of urinary bladder carcinomas, papillomas and preneoplastic lesions
[209, 210]. However, rats treated only with DMAV did not show any preneoplastic lesions or
tumors [209, 210]. F344/DuCrj rats pretreated with a variety of known carcinogens showed
enhanced tumor induction after being exposed to DMAV in their drinking water [211]. Although
the kidney, liver and the thyroid glands showed an effect, the bladder was more susceptible to
tumor induction at the lowest concentration of DMA exposure (50 ppm) [211]. Urinary bladder
carcinogenesis was observed in 2 F344 male rats out of 31 when administered high doses of
DMAV (200 ppm) via p.o. [212]. DMAV failed to promote lung carcinogenesis when F344 rats were initiated with N-bis(2-hydroxypropyl)nitrosamine [213]. Studies using MMAV and
trimethylarsine oxide (TMAO) also showed that only pretreatment with DENA induced hepatic preneoplastic lesion formation in F344 rats [214]. In contrast, a different study showed that at
200 ppm, TMAO alone could induce liver tumorigenicity in F344 rats [215].
3. Mice
In a skin cancer model, hairless mice (Skh1) fed AsIII for their first 21 days of life and exposed to UV radiation (UVR) after 42 days showed that AsIII enhanced the UVR-induced skin
22 carcinogenesis [216]. These studies suggest that arsenic acts as a cocarcinogen or a promoter, as
observed in rats, rather than an initiator. The non-carcinogenic potential of DMAV was
demonstrated when B6C3F1 mice did not show tumor formation in any of the organs studied
(kidneys, liver, lungs, thyroid, and urinary bladder) [217]. Recently mice have been used as a
model to study the possible transplacental carcinogenic potential of arsenic [218]. For example,
offspring from two different strains of pregnant mice (C3H and CD1) fed AsIII at concentrations
of up to 85 ppm were observed for up to two years. Female offspring from the C3H strain
showed ovarian tumors, lung carcinomas, and proliferative lesions of the uterus and oviduct and
offspring males from the same strain developed liver carcinoma and adrenal cortical adenoma
[218]. CD1 offspring male mice developed tumors of the liver and adrenal and renal hyperplasia
and female offspring, similar to C3H female offspring, developed tumors of the urogenital
system, ovary, uterus and adrenal hyperplasia of the oviduct [218]. These studies suggest that transplacental arsenic exposure can initiate tumors of the lungs, liver and reproductive organs, but cannot initiate skin tumors; rather, it acts as a promoter after exposure to other stressors like
UVR.
The importance of the p53 tumor suppressor function, especially during stress, has been widely studied in the context of carcinogenic potential after loss of function (reviewed in [219]).
Therefore, an established p53+/- mouse model was used to test the possible carcinogenic effects
of DMAV exposure via drinking water at concentrations of 0, 50 and 200 ppm [220]. Overall,
the results showed that p53+/- and wild type mice exposed to DMAV showed induction of tumors
significantly earlier than non-treated mice, and DMAV-treated p53+/- mice were more susceptible
to cancer induction at an earlier time point than treated wild type mice, further suggesting that
DMAV might induce DNA damage [220]. Transgenic mouse models for arsenic-induced skin
23 carcinogenesis have been established, for example Tg.AC mice (Tg.AC mice harbor the v-Ha- ras oncogene) topically exposed to the promoter tetradecanoyl phorbol acetate (TPA) showed a
marked increase in skin papillomas when fed AsIII compared to non-TPA treated transgenics,
transgenics unexposed to AsIII but TPA treated, and wild type mice treated with TPA and
exposed to AsIII [221]. Similar results in K6/ODC mice, which express a truncated form of
ornithine decarboxylase (ODC is a biomarker of cell proliferation, it is involved in polyamine
biosynthesis, and catalyzes ornithine from the Urea cycle to putrescine, which is required for cell
proliferation and cell differentiation), were observed when exposed to 100 ppm AsIII and DMAV via drinking water [222].
Even though many of the studies presented here using rodents have shown somewhat promising data on the carcinogenic potential of arsenic there are many limitations in establishing a carcinogenic model. For example, even though rats have been used frequently to study the carcinogenic potential of arsenic, their metabolic pathway yields TMAO [223], an arsenical rarely produced by humans, and one that could induce toxicity by unique means. In order to observe potential carcinogenic effects due to arsenic exposure, the doses (usually high), the mechanism of administration (i.p., i.v., or p.o.), and the chemical form (DMAV is widely used
because of the high levels found in human urine which were thought to be produced as a
detoxification mechanism after methylation of iAs) of arsenic that some of the experimental
animals are exposed to are usually different compared to the doses, mechanisms, or forms
humans are exposed to any given time point. The fact that arsenicals only act as promoters
confounds the research because which humans are rarely exposed to some of the chemical
initiators used in such studies. The carcinogenic effects observed in humans are usually due to
chronic lifetime exposure that could be potentiated by other environmental factors such as
24 smoking, diet, sunlight irradiation, and/or air pollutants. Laboratory animals have a shorter life
span compared to man, and they are usually maintained in a controlled environment. These are
problems scientists have faced in an attempt to establish a model system to study the
carcinogenic effects of arsenic and its synergy with other environmental stressors.
Drosophila as a Model for Chemical Genotoxicity
1. Why Drosophila?
Arsenic has been shown to be genotoxic in a variety of experimental in vitro and in vivo
models by inducing sister chromatid exchanges, DNA strand breaks, or oxidative DNA damage.
Drosophila offers a variety of advantages to study the effects induced by environmental
stressors, such as the ease and variety of genetic manipulations available that could help identify
genetic pathways involved (reviewed in [224-226]), e.g. the creation of genetic mutants via P-
element insertion [227] or deletion mutants via the FLP-FRT recombination system [228], as
well as in vivo functional studies employing RNAi-induced knock-down targeting of any gene in
the fly genome [229]. Additionally, the creation of transgenic fly strains utilizing the Gateway™
system [230] and the controlled expression of the inserted gene via the GAL4/UAS yeast expression system [231-233], provide the opportunity to study the effects observed in humans to
environmental stressors in the fly. A second major aspect of Drosophila as a model concerns the
high conservation of genes involved in many human disease pathways. A BLAST analysis of
929 human disease gene entries with at least one mutant allele in the Online Mendelian
Inheritance of Man (OMIM) database against the Drosophila genome identified 714 human genes (77%), formatted into a searchable database called Homophila, that matched 548 unique sequences in the fly genome [234, 235]. For example, the fly harbors genes encoding proteins homologous to genes correlated with neurodegenerative diseases (reviewed in [236]) including
25 Parkinson’s, for which fly models have been established [237, 238]. Fly models have also been
established to study the mechanisms involved in carcinogenesis [239, 240] (reviewed in [241,
242]). Additionally, many of the enzymes involved in neurodegeneration caused by Alzheimer’s
disease, such as presenilin [243, 244], nicastrin [245, 246] and the amyloid precursor protein
(APP) [247], have fly homologs and have been shown to function in a similar pathway. Human
congenital heart disease has been correlated to mutations in the transcription factor NKX2-5
[248], which has a fly homeobox orthologue called tinman that is involved in the formation of
the Drosophila heart [249]. According to the FlyAtlas database a number fly homologs of
human genes are expressed in fly tissues that are analogous to those of the affected human
tissues [250]. As far as practicalities go, the gestation period of the fly is shorter than the mouse,
as well as the age at which the fruit flies reach sexual maturity being much earlier. Moreover,
the cost of rearing and maintaining fly populations is relatively trivial compared to mouse
colonies, and there are no federal regulations or statutes that apply to experiments in Drosophila.
2. Tests for Genotoxicity
2.1.1 mwh-flr SMART
Drosophila has been used to study the genotoxic effects of known environmental
chemicals by employing the use of a loss of heterozygosity (LOH) technique called the Somatic
Mutation and Recombination Test (SMART) [251]. In fruit flies heterozygous for a visible
recessive marker mutation, certain mutagenic events induced by exposure to chemicals can lead
to the loss of the dominant wild-type allele opposite to the marker mutation and expression of the
recessive marker allele in a clone of the mutated cells. In the SMART assay the larval wing
imaginal discs are trans-heterozygous for the recessive markers multiple wing hairs (mwh), which is distal to the centromere, and flare (flr), which is proximal to the centromere of the left
26 arm of the 3rd chromosome. These markers are brought together by mating adult mwh virgin
females and flr/TM1 males [251]. The mwh enzyme is involved in imaginal disc derived wing
hair organization and biogenesis, wing morphogenesis, establishment of planar polarity and is
involved in Wnt receptor signaling [252]. The flr enzyme has no known molecular function or
known biological role [252], but homozygous mutants have distinct phenotypes of the chaetae
and trichomes in the abdomen, thorax, and wing surface [253, 254]. A variety of mechanisms
can lead to loss of heterozygosity (LOH), which can be observed phenotypically in the wings of
the eclosing adult. A deletion, recombination event, or point mutation of the distal marker
(mwh), can lead to a ‘single spot’ or a cell with 3 or more hairs (normal cells only have 1 or 2
hairs). A non-disjunction of the 3rd chromosome harboring the mwh or flr markers can also
exhibit ‘single spots’, as well as a point mutation in the flr gene. ‘Large single spots’ can be
induced by a large deletion of the distal mwh marker. ‘Twin spots’, or clones of cells with LOH in both markers, contain cells in the wings with three or more hairs and cells with shortened and thickened hairs or amorphic and balloon-like extrusions of melatonic chitinous material. ‘Twin spots’ arise from a mitotic recombination event proximal to the centromere generating cells homozygous recessive for mwh and cells homozygous recessive for flr after cell division. High bioactivation (HB) tester strains for the SMART test have been developed by substituting chromosomes 1 and 2 from the Oregon R(R) (ORR) strain into the original mwh female strain and crossing to an ORR flr male strain [255]. The ORR strain has increased cytochrome P-450
[256, 257] activity and was selected for its resistance to the pesticide DDT [258, 259]. An improved HB strain employs crossing an ORR flr female to an mwh male [260], which elminates the difficulty of spot classification due to irregular whorling (spiral hairs), high variations in
27 repeated experiments and low egg production of the ORR; mwh females and delay in development of the larvae of the HB cross [260].
Many chemicals have been tested and shown to be genotoxic by using the SMART. For example, chromium(IV) [261], cobalt, manganese, nickel, zinc [262], three polyaromatic hydrocarbons (PAHs), benzopyrene (B[a]P), benz[a]anthracene, and 7,12- dimethylbenz[a]anthracene [263], and several halogenated aliphatic compounds such as 1- bromo-2-chloroethane, 1,2-dichloroethane, 1,2-dibromoethane and 1-iodopropane [264], induced genotoxicity in the larval imaginal wing discs. In fact, B[a]P-DNA adducts have been identified in eclosing adult flies derived from the corresponding larvae utilized for the SMART [265]. It is important to note that neither of the heavy metals cadmium [266], mercury, nor methylmercuric(II) chloride (MMC) [267] induced genotoxicity in the SMART assay. The
SMART assay has also been employed for field risk assessment by testing the genotoxicity of pollutants present in surface waters, which are used for public and industrial supply, irrigation, animal feeding and recreation, in different urban and industrial sites of Brazil [268]. None of the samples from industrial sites showed genotoxicity; however urban samples from Montenegro and São Sebastião do Caí showed the presence of genotoxic compounds in surface waters [268].
2.1.2 lats SMART
A newly developed SMART system identifies LOH in a gene homologous to the human large tumor suppressor 1 (LATS1) gene, called warts (wts) in Drosophila [269]. The wts protein is a serine/threonine kinase tumor suppressor [269] involved in the Hippo pathway that controls organ size by regulating cell growth, proliferation, and apoptosis [270-273] via response to cell- cell contact [274]. In Drosophila, the Hippo-induced apoptotic pathway has been shown to be activated via p53 after DNA damage [275]. It would appear that this pathway is well conserved
28 among eukaryotes [276]. For example, Lats-/- mice develop soft-tissue sarcomas and ovarian
stromal cell tumors and are highly sensitive to 9,10 dimethyl-1,2-benzanthracene (DMBA) and
UVB cotreatment, expected to damage DNA [277]. Furthermore, ectopic expression of human
LATS1 in a Drosophila strain mutant for wts showed that tumor formation (see below) was
suppressed and flies developed normal adult structures [278]. Mutations in wts, induce dramatic overproliferated, outgrowth tissue phenotypes and a variety of developmental defects, such as early embryonic, larval, or pupal lethality, in mosaic and homozygous mutants [269]. In the lats-
SMART assay, if a mutation in the wild type allele of the wts gene occurs in a heterozygous
wts+/- 3rd instar larva exposed to a genotoxin, large tumorigenic clones can form and be observed
in the eclosed adult fly [279, 280]. This system was used to test the genotoxic potential of
several chemicals including B[a]P, pyrene [279], methyl methanesulfonate (MMS) and X-rays
[280] and showed that they were genotoxic.
2.2 SRLRT
Another test employed to identify the genotoxic potential of chemicals using Drosophila
as a model, specifically DNA damage to germ cells, is called the sex-linked recessive lethal
mutation and reciprocal translocation test (SRLRT) [281]. Using this test the genotoxic potential
of a chemical can be identified using this test by the induction of a lethal mutation in a wild type
X chromosome. Such a mutation would then be observed in surviving females due to the fact
that they would ‘balance’ the mutation over the inverted chromosome Basc [281]. The U.S.
National Toxicology Program has established a database of chemicals tested for their genotoxic potential using the SRLRT [282]. Two halogenated aliphatic compounds, 1,2-dibromo-3- chloropropane and 2,3-dibromo-1-propanol [283], and a compound formed by the heavy metal chromate, calcium chromate (CaCrO4), tested positive for the SRLRT [284]. Other chemicals
29 that have been tested but showed negative results for the SRLRT include the DNA adduct B[a]P
[285], the heavy metal mercuric chloride [286], and bisphenol A [287], a toxic chemical released
from polycarbonate bottles used to store drinking water [288] and may interfere with
chemotheraupetics used in cancer treatments [289]. The SRLRT has been used in combination
with repair-defective mutants, such as the NER-defective mei-9L1 or mus(2)201D1 strains, to test for the sensitization to effects of alkylation-induced genetic damage [290]. The mei-9 gene
expresses a protein homologous to the human XPF and has been suggested to be involved in
mismatch repair [291], NER [292, 293], and chromosomal resolution during meiotic
recombination [294].
2.3 vermilion gene alkylation
DNA adduct formation via alkylation can lead to genetic mutation(s) that can be
observed phenotypically. A repair deficient strain harboring the mus(2)201D1 allele, which
expresses a truncated form of the Drosophila homolog to human xeroderma pigmentosum (XPG)
protein [295], was used to test for the alkylation potential of ethyl methanesulfonate (EMS) on
the vermilion (v) locus and was shown to induce higher levels of GC to TA transversions than a
treated, but excision repair proficient, strain [296]. The v gene expresses a protein with
tryptophan 2,3-dioxygenase activity [252] that is involved in brown eye pigmentation [297]; thus
any genetic damage due to DNA alkylation and deficiency in DNA damage repair will be
observed by alterations in eye pigmentation. To identify a genetic mutation due to DNA
alkylation induced by a genotoxic chemical, exposed Drosophila mutants for the brown (bw)
gene (mutant flies lack red eye pigment, thus adult mutants have dark brownish wine eye color)
and wild type for the v gene are then mated to v heterozygous dominant mutants. Mutant v*/v; bw, where * denotes a severe, but non-lethal v phenotype, offspring are identified based on
30 differences in eye color pigmentation compared to the v; bw heterozygous dominant mutants, which are white eyed phenotypically [298]. The v system was also used to test the genotoxic potential of three different N-methyl-N-nitroso compounds, all of them inducing higher levels of
AT to GC transitions [299].
2.4 SCLT
Another test, called the sex-chromosome loss test (SCLT), characterizes the genotoxic potential of a chemical via chromosomal gain or loss (aneuploidy), rather than death, in the offspring derived from female germ cells [300]. The effects of aneuploidy are observed by the phenotypic yellow (y) body marker in females and the white (w) eye marker in males (termed
FIX system in which females carry a heterozygous inverted X chromosome), or the X-linked zeste (z) (termed ZESTE system in which females do not carry inversions) eye marker in females and the white (w) eye marker in males [300]. Cadmium chloride (CdCl2) induced chromosome gain in feeding 3rd instar larvae when the FIX system was employed, and chromosome loss and gain in the ZESTE system [301]. CdCl2 also induced chromosome loss in adults when the
ZESTE system was employed [301]. Another heavy metal compound, MMC, was negative for the FIX test in feeding larvae but positive in feeding adults [301]. These results suggest that chemicals can induce aneuploidy by disrupting meiotic division in germ cells and that
Drosophila can be used as an exceptional model system to observe these effects through easily identifiable phenotypes.
Arsenic and Drosophila
In order to learn more regarding the mechanisms and pathways affecting arsenic toxicity and genetic susceptibility, we have utilized Drosophila as our model system. Drosophila has been previously used as a model system to study the toxic effects of inorganic and organic
31 arsenicals, utilizing the SMART, SLRLT and SCLT assays. The earliest study using Drosophila
as a model was published in 1969 and showed that AsV increased the frequency of DNA
crossover, and that this was dependent on the concentration of selenocystine [302]. Later studies
demonstrated that when fed AsIII or AsV, flies were negative in the SMART assay [303, 304]. In
another SMART assay, using the improved HB strains, both AsIII and AsV were negative [305],
suggesting that neither AsIII nor AsV are genotoxic in flies. However, a similar approach
employing the SMART assay, using the w+/w eye markers, demonstrated that AsIII increased the levels of mosaic light spots, which are changes in the color composition of the eye ommatidia caused by genetic alterations in the eye pigment cells during larval development in both sexes
[306]. AsV induced a dose-response relationship for small and total light spots in the eyes of
females but not in males [305]. Two other genotoxic assays in germ cells showed that when
injected into males, AsIII, but not AsV, increased the frequency of SLRLT lethality, and both AsIII and AsV were negative in the SCLT assay [305]. The modulating effects of AsIII on other known
genotoxins have also been studied using Drosophila as a model and results have shown that AsIII does not increase the genotoxic potential of the heavy metal compound, potassium dichromate
(K2Cr2O7), or the alkylating agents EMS [304], ehtylnitrosourea, MMS, and ethylene oxide
[307]. More recently, studies of Drosophila larvae exposed to DMAV showed that eclosing adult
flies were positive for the SMART assay, and DMAV was observed in larval homogenates via
high-performance liquid chromatography (HPLC) followed by inductively coupled plasma mass spectrometry (ICPMS) [308]. Conversely, when fed iAs, the eclosing adults were negative for
the SMART assay and HPLC-ICMPS analyses of larval homogenates detected iAs, but not MAs
[308]. Taken together, these results suggest that Drosophila respond similarly to mammalian
cells in exhibiting genotoxic effects of MAs, but unlike mammalian cells, they do not methylate
32 inorganic arsenic, and generally speaking, iAs appear to have very weak genotoxic activity in flies. This makes sense because Drosophila appears not to harbor an arsenic methyltransferase gene, a notion reinforced when a BLAST analysis for a homolog to the human AS3MT gene was performed by our laboratory.
Conclusion
The massive poisoning of populations world wide due to the chronic consumption of arsenic-contaminated drinking water supplies has brought scientists from every part of the Earth together to answer questions and raise awareness about the health effects brought on by its systemic toxicity. Through many years of scientific research and advances in experimental approaches, we have been able to gain knowledge on the mechanism(s) of arsenic toxicity.
However, many questions still abound, especially how arsenic interacts with the complexity of the human cell and whether there is one single carcinogenic/toxic mode of action, exacerbating the problem due to the lack of a suitable mammalian model. Another scientific problem currently being researched is how inter-individual genetic variation, especially in genes coding for enzymes involved in the metabolic pathway of arsenic, could be correlated with differential responses, more importantly susceptibility, to the toxic effects of chronic arsenic consumption.
Establishment of a suitable model system to study arsenic toxicity and identification of genetic factors involved in susceptibility is important in risk assessment of arsenic exposure in populations.
Many differences between the human and Drosophila species abound. For example, genome complexity, organ/tissues used to maintain homeostatic balance, and the variations in metabolism of chemicals, could pose a problem for establishing a genotoxic mode of action.
However, the ease of genetic manipulation, the high level of genomic homology between the fly
33 and the human, the short time course required to obtain results from a given experiment, the
sample sizes that can be used and/or chromosomes that can be observed in a given experiment
make this system a quick and inexpensive system that could then be transferred to future human studies. For all these reasons, it seemed probable that we could use this experimental system to
explore some key questions in the toxicology of arsenic.
In this dissertation, data are presented that result from experiments designed to address
two key questions. First, using the natural variation available in strains derived from around the
globe, we sought to define a gene (or genes) that could be responsible for relative genetic
tolerance or susceptibility to the toxic effects of arsenic. Our experiments identified a key role
for the enzyme glutathione synthetase in a robust response towards AsIII, leading to the hypothesis that individuals expressing a polymorphic, and less functional, enzyme might be more
susceptible to arsenic exposure. Second, we have created and characterized transgenic lines that
express a functional human AS3MT enzyme in order to study the effects of MAs, which are
currently considered to be the primary genotoxic and/or carcinogenic species of arsenic. Our
goal with such an experimental resource is to study the effects of MAs in a variety of altered
genetic backgrounds that could shed light on the mechanisms and pathways affected in the
process of arsenic-mediated carcinogenesis.
34 Chapter II. (The text of this chapter has been published: Muñiz Ortiz et al., 2008, Tox Sci doi:10.1093/toxsci/kfn192)
Investigating Arsenic Susceptibility from a Genetic Perspective in Drosophila
Reveals a Key Role for Glutathione Synthetase
Introduction
Arsenic-contaminated drinking water is widely distributed throughout both the developed and developing world and represents an extremely serious public health issue in many locations, particularly in Bangladesh [309]. Here, millions of tube wells were bored to discourage use of water from ponds, rivers and open wells, often a source for cholera and other water-borne diseases [25]. Subsequently it was discovered that tube well water was frequently contaminated with arsenic [25] and thus tens of millions in Bangladesh have been, and still are, ingesting arsenic via drinking water. Chronic arsenic ingestion can induce a myriad of pathologies [310], including malignancies of the bladder, kidney, and lung, as well as diabetes mellitus, neuropathy, vascular disease, respiratory effects, and various types of skin lesions.
Owing to these worldwide health effects, a maximum contaminant limit for arsenic in water has been set to 10 µg/l [38].
It has been suggested that variation in susceptibility occurring in populations chronically exposed to arsenic could have a genetic component. Differences in the metabolism of inorganic arsenic have been observed between native Andean women [177] and children [15] chronically exposed to arsenic-contaminated drinking water when compared to other populations.
Comparing total arsenic and arsenic metabolites in urine of individuals from Mexico, China, and
35 Chile has supported the idea that inter-individual differences in toxicity could be due to
functional polymorphisms in genes involved in metabolizing the metal [179].
There are many studies of the cellular processes affected when arsenic interacts with
biological tissues (reviewed in [78], but it is germane to consider genes in three particular categories: those that affect arsenic uptake, cellular elimination, and metabolism within the cell.
Arsenite can be transported into mammalian cells via the aquaglyceroporin channels AQP7 and
AQP9 [125], but there are no data as yet regarding functional polymorphisms affecting individual sensitivity. Elimination of xenobiotics from cells frequently employs conjugation of glutathione (GSH) to substrate (via glutathione S-transferase (GST)), followed by ATP- dependent transporter-mediated export. Thus, a GSH-arsenic complex is secreted via the ABC multidrug resistant transporter family member MRP1 [66]. Differences in arsenic efflux might be associated with polymorphisms in GST-M1 and -T1 [311], and a GST-P1 polymorphism was suggested to increase the odds of arsenic-induced skin lesions [312]. Knockout of the ABC transporter MDR1 in mice produced greater susceptibility to arsenic and higher arsenic accumulation in their tissues [188].
When inside mammalian cells inorganic arsenicals can undergo a series of metabolic reactions leading to mono or dimethylated products of either the +3 or +5 oxidation state.
Different reaction schemes have been proposed: (1) inorganic arsenicals undergo a series of successive reduction and oxidative methylation reactions [131], or (2) inorganic arsenicals
(typically in the reduced +3 state) are conjugated to GSH and undergo a series of successive methylations, each intermediate providing a substrate for oxidation to the +5 state [132]. A
GSH-requiring arsenate reductase (identical to glutathione S-tranferase omega (GSTO)) has been implicated in scheme (1) [136], while an arsenic(+3) methyltransferase (AS3MT) [139] can
36 provide the methylating activity necessary in scheme (2) and could also function in scheme (1)
by linking oxidative methylation with reduction of arsenic metabolites using GSH as a cofactor
[132, 140]. Polymorphisms in GSTO [180] and in AS3MT [181, 182] have been correlated with
individual differences in urinary metabolic arsenic profiles.
Such studies reinforce the notion that susceptibility to arsenic will have a strong genetic basis. To explore this further, we have taken an unbiased approach to the genetics of arsenic susceptibility through examination of natural variation present in geographically distinct populations. We have utilized Drosophila as our experimental organism owing to the ease and variety of genetic manipulations available (reviewed in [313], as well as the high representation of genes homologous to those involved in many human disease pathways [235], including cancer
[242]. This approach has revealed that, surprisingly, optimal activity of the glutathione synthetase gene is required for an effective physiological defense towards arsenite. The data obtained reinforce the notion that organismal tolerance towards long-term arsenic exposure requires a robust anti-oxidant system based on the glutathione biosynthetic pathway, and imply that allelic variation affecting the activity of any genes contributing to this pathway will likely be pertinent to individual susceptibility and risk.
Materials and Methods
(For additional methods, see Appendix II)
Flies
Flies were maintained on standard cornmeal medium at room temperature. Most wild type strains were obtained from the Indiana University Stock Center at Bloomington, Indiana, although several strains came from the now defunct stock center at Bowling Green State
University, Ohio. For generation of deficiency lines, isogenic flies harboring inserted FRT
37 elements that flank regions to be deleted were obtained from either the Exelixis collection at
Harvard University Medical School or the Bloomington Drosophila Stock Center. FLP-induced
X chromosomal deletions were generated as previously described [228]. Stocks were maintained
by balancing over Binsinscy. Glutathione synthetase (GS) RNAi lines were obtained from the
Vienna Drosophila RNAi Center [229]. Since these lines were designed to target sequences
towards the 5’ end (VDRC49801) or the 3’ end (VDRC49719) of the GS genes CG6835 and
CG32495 we took advantage of their separate 2nd and 3rd chromosome insertion sites to create
(via the use of chromosomal balancers) a compound line (designated GSRNAi[5’/3’]) containing both RNAi inserts in homozygous condition. The da-GAL4 line was obtained from the
Bloomington stock center, and we received the e4 GclmL0580 line from Dr. Robert Saunders [314].
The e4 GclmL0580 stock contains a P-element insertion in the 5’-untranslated region of the gene
encoding the modifier subunit of glutamate cysteine ligase (Gclm). Wild type revertants were
induced by crossing to a stock containing the P{ry+ Δ2-3} element, which produces
constitutively active P-transposase. Revertants were selected on the basis of loss of eye
pigmentation, associated with loss of the w+-bearing P-element. Precise removal was confirmed
via PCR and homozygous lines were then established.
Arsenite sensitivity assays
Wild-type strains
Embryos were collected essentially as previously described [315]. Briefly, flies were allowed to lay eggs on a small grape juice agar plate, seeded with yeast paste, inserted into the neck of an inverted culture bottle. 150 to 200 embryos (0-8 h old) were transferred to a piece of sterilized grape juice-soaked filter paper. Filter paper with embryos was then laid down on 5 g of Instant
™ Drosophila Medium (Carolina Biological, Burlington, NC) hydrated with 30 ml of H2O or
38 sodium m-arsenite (Sigma, St. Loius, MO) solutions of various concentrations. Emerging adults
were counted, and eclosion data compared on AsIII-supplemented and non-supplemented food.
Microsatellite-based recombination mapping
A variety of microsatellite markers covering the X chromosome were identified either from
previous literature descriptions (e.g., [316] or from BLAST analysis of the relevant Drosophila
genomic sequence. Markers useful for further analysis were chosen by their size heterogeneity
when comparing PCR products (using unique sequence primers flanking the particular repetitive
regions) produced from Oregon R 1970 and PVM genomic DNA. For the recombination
analysis, F1 virgin females derived from an Oregon R-PVM cross were mated to PVM males, F2
embryos collected and placed on AsIII-free or AsIII-containing food as described, and individual
eclosing adult progeny collected for PCR analysis of specific microsatellite markers. PCR
products from experimental and control parental flies were sized on 3% MetaPhor agarose
(Cambrex, East Rutherford, NJ) gels run in 1X TAE at 4oC. For any given marker the
percentage of flies carrying one or the other parental allele was calculated.
X chromosome deficiency lines
Since most X-chromosomal deficiencies (Df) generated were lethal when homozygous, we maintained stocks as heterozygotes over the X chromosome balancer Binsinscy. In order to compare the AsIII sensitivity of deficiency lines with otherwise genetically identical control lines we had to create females heterozygous for the non-Df parental X chromosome (w1118) balanced
over Binsinscy. We crossed these w1118/Binsinscy females to Binsinscy/Y males and collected the
resulting embryos. These were placed on either AsIII-free or AsIII-supplemented food, and
eclosing female adults of the genotype w1118/Binsinscy were counted. We performed an AsIII dose-response assay to identify a threshold concentration where the control female flies
39 (w1118/Binsinscy) showed no obvious effects of AsIII on relative viability, and then used this
concentration (0.25 mM sodium AsIII) to assess the comparative viability of each deficiency line.
For these experiments isogenic deficiency females of the various lines (w1118, Df/Binsinsncy)
were crossed to Binsinscy/Y males, the resulting embryos were collected and exposed to AsIII- free and AsIII-supplemented food, and eclosing female adults of the genotype w1118, Df/Binsinscy
were counted.
Deficiency Line Data Analysis
A viability ratio was calculated for the average of eclosing w1118/Binsinscy females from three
bottles of 0.25 mM AsIII-supplemented food to that from three bottles of AsIII-free food. This
ratio was compared to the ratio of the average of w1118, Df/Binsinscy females eclosing from three
bottles of AsIII-supplemented food to that from three bottles of AsIII-free food. If the
chromosome deficiency produced sensitivity towards AsIII then the viability ratio of deficiency
lines should be significantly lower than that of control lines. On the other hand, if there is no effect of the chromosomal deficiency towards AsIII the ratios should not be significantly
different.
e4 GclmL0580 and wild type revertant lines
In order to compare the AsIII sensitivity of e4 GclmL0580 and wild type revertant lines we collected
embryos from each cross individually since they are homozygous viable. Embryo collection and
data analysis was performed as described above for X chromosome deficiency lines although
embryos were exposed to 0.125 mM AsIII.
GSRNAi[5’/3’] lines
GSRNAi[5’/3’] flies were crossed to a strain expressing Gal4 under the control of the daughterless
regulatory element. As controls, non-transgenic w1118 parental flies were crossed to the Gal4-
40 expressing strain and GSRNAi[5’/3’] flies were crossed to a non-GAL4 expressing w1118 line. In all
cases, progeny embryos were collected and the relative adult eclosion on AsIII-supplemented or
AsIII-free food was determined. Quantitative RT-PCR was performed on adults as described
below to demonstrate knock down of GS transcripts via RNAi.
PCR Confirmation of FLP-FRT-based Deletion and Gclm Revertant Lines
To confirm predicted recombination events occurred between FRT sites for the particular
Exelixis line combinations used, we performed PCR on genomic DNA as described previously, with some adjustments [228]. Each 50 µl PCR reaction included 1 µl of genomic DNA, 5 µl
® 10X ThermoPol reaction buffer (NEB , Ipswich, MA), 2 µl 25 mM MgCl2, 1 µl 10 mM dNTP
mix (NEB®, Ipswich, MA), 1 µl forward primer (1 µg/µl), 1 µl reverse primer (1 µg/µl), and 2.5
units (0.5 µl) Taq polymerase (NEB®, Ipswich, MA). Reaction conditions were 95° C for 10
min; 94° C for 30 s, 52.4° C for 1 min, 72° C for 2 minutes, repeated 30 times; and 72° C for 10
min. Agarose gel analysis was then performed to identify appropriate hybrid elements. For
determination of P-element excision from the e4GclmL0580 strain PCR conditions were identical
except the annealing temperature was 61° C.
PCR primer combinations for FLP induced recombinants:
XP(+) & WH(-):
F – 5’-AATGATTCGCAGTGGAAGGCT
R – 5’-GACGCATGATTATCTTTTACGTGAC
XP(+) & RB(+):
F – 5’-AATGATTCGCAGTGGAAGGCT
R – 5’-TGCATTTGCCTTTCGCCTTAT
RB(-) & XP(-):
41 F – 5’-TGCATTTGCCTTTCGCCTTAT
R – 5’-AATGATTCGCAGTGGAAGGCT
PCR primer combinations for Gclm mutant and wild type revertant:
F – 5’-AGCTGTGTAATCTGCTGCTTGAG
R – 5’-TCATTTGGATCTAGTACCCCTGG
Cell Culture
Schneider’s S2 cells were maintained at 25C in Schneider’s Drosophila Medium (1X) (Gibco,
Carlsbad, CA) supplemented with 10% FBS (Gibco, Carlsbad, CA) and 1%
Antibiotic/Antimycotic mix (Gibco, Carlsbad, CA).
Production of double stranded (ds) RNA for RNAi in tissue culture
Production of dsRNA was performed as described previously [317] with some adjustments. A
~900-1000 bp fragment corresponding to a segment of the target gene was amplified from fly
genomic DNA via PCR. A second round of PCR was performed to add the T7 promoter
sequence to either end of a ~500-700 bp segment within this fragment. Each T7 fragment was
produced individually. PCR products were purified by using the MinElute™ PCR Purification
Kit (Qiagen, Valencia, CA). 2 μg of each T7 fragment was used as template to produce dsRNA using the MEGAscript® RNAi Kit (Ambion, Austin, TX). Primers used in the generation of T7
constructs are described in the Supplementary Data section.
Conditions for RNAi in Drosophila Cell Culture
We followed the basic RNAi conditions, with some modifications, of those described previously
[317]. Drosophila S2 cells were diluted to a final concentration of 6.75 x 105 cells/ml in medium
containing 10% FBS and 1% antibiotic/antimycotic. dsRNA (1 μg) was added directly to
42 corresponding wells of a 96-well plate. Aliquots of cells (15 μl, 1 x 104 cells) were pipetted into
wells either containing dsRNA or not. FBS-free Schneider’s medium (50 μl) was added to the
wells and plates were then shaken vigorously at RT for 30 min, followed by addition of 100 μl of
AsIII-free or AsIII-supplemented (35, 45 or 80 µM) Schneider’s medium supplemented with 15%
FBS. Cells were incubated for 0, 24, 48 and 72 h at 25° C prior to testing for AsIII sensitivity by
cell viability. Quantitative RT-PCR was performed as described below to confirm knockdown of
GS mRNA transcripts.
Cell Viability Assay
Cell viability was determined using the CyQUANT® NF Cell Proliferation Assay kit
(Invitrogen™, Carlsbad, CA) following the manufacturer’s directions. After reagent addition the
microplate was covered and incubated at 25° C for 30 min. Fluorescence was measured with
excitation at 485 nm and emission at 530 nm using a BioTek® FL600 Microplate Fluorescence
Reader, and data were collected using the BioTek® KC4 version 3.01 computer software
program.
Quantitative Reverse Transcriptase-PCR
First strand synthesis was performed using the iScript cDNA synthesis kit using 1 µg total RNA
according to manufacturer’s directions (Bio-Rad). Real Time PCR was performed as previously
described, with variations [318]. Briefly, 2 µl cDNA was used as template in a 25 µl reaction
including 100 ng of primers, 1mM MgCl2, 0.2 mM dNTP’s (Fisher, Pittsburgh, PA), 1X
ThermoPol Taq Buffer (NEB®, Ipswich, MA), and 1U Taq polymerase (NEB®, Ipswich, MA) in
® filter-sterilized MilliQ H2O. SYBR green (Invitrogen™, Carlsbad, CA) was used at a final concentration of 0.5X from a 10,000X stock. All amplification protocols used a 2 min melting
43 step at 95°C followed by 40 cycles of amplification. Each cycle used a 15 s melting step at
95°C, an annealing step of 15 s at 62.1°C for actin, 61.4°C and 61.7°C for the 5’ and 3’ region
of CG6835 and CG32495, respectively, followed by an extension step at 72°C for 15 s. Each
cycle ended with measurement of fluorescence at 85°C for 10 s. Primers used in RT-PCR are described in the Supplementary Data section. Data analysis was performed using a relative
quantification of Ct values to calculate the expression of GS (CG6835 and CG32495) relative to actin control expression using the 2-ΔΔCt method [319].
Primers used in generation of T7 constructs
1149 bp fragment derived from 5’ region of GS genes (CG6835 and CG32495):
F – 5’-AGTTTCCAGCGATACCCAGCAG
R – 5’-AGCGACTACAGCCGACTTCTGG
Individual 641 bp T7 fragments derived from 5’ fragment of GS (incorporated T7 sequence is underlined):
F – 5’-GAATTAATACGACTCACTATAGGGAGATGTCCAGCGACGCCAATACG
R – 5’-ACAGATTGGCCGTGAACTCG
and
F – 5’-TGTCCAGCGACGCCAATACG
R – 5’-GAATTAATACGACTCACTATAGGGAGAACAGATTGGCCGTGAACTCG
885 bp fragment derived from 3’ region of GS gene:
F – 5’-TCGTACAACATTTGCGACCAG
R – 5’-TGAGAGCTTTGTGCGCAGCATG
Individual 664 bp T7 fragments derived from 3’ fragment of GS gene:
F – 5’GAATTAATACGACTCACTATAGGGAGATGTGCTCAACTCACACGCTTGC
44 R – 5’-ACAATGTGCTCCGCATCTCCGA and
F – 5’-TGTGCTCAACTCACACGCTTGC
R – 5’GAATTAATACGACTCACTATAGGGAGAACAATGTGCTCCGCATCTCCGA
Primers used in quantitative RT-PCR experiments actin (control):
F – 5’-ACCTTCTACAATGAGCTGCGTGTG
R – 5’-AGTCCAGAACGATACCGGTGGT
5’ Region of CG6835 and CG32495:
F – 5’-TGCCCCTTTTCTAGACGCAGCT
R – 5’-AATGGCATAATCCTTGGCCTTGG
3’ Region of CG6835 and CG32495:
F – 5’-TCATCTACTTCCGAGCTGGCT
R – 5’-AGACCCGTGAAGATCTTGCCCAC
Results
Identification of “arsenite-sensitive” and “arsenite-tolerant” Drosophila strains
Several dozen geographic variants of Drosophila melanogaster were obtained from stock centers and individual investigators and examined for their ability to eclose successfully as adults after seeding of embryos on to food containing sodium AsIII at a variety of concentrations. The average percent survivability of a representative sample of 35 such strains (over 60 were tested) as compared to those reared on control food is depicted graphically in Figure 2.1. On the basis of such data we decided to further test Oregon R 1970 and PVM as examples of strains that showed relative tolerance and relative susceptibility to AsIII respectively.
45
Figure 2.1. Viability of D. melanogaster strains in sodium arsenite. Seeded embryos were scored for percent adult eclosion on arsenite-containing food and normalized to values obtained when seeded on arsenite-free food. Two strains that showed relative resistance (R) and sensitivity (S) to arsenite are
marked by arrows and were selected for further analysis (though others could also have been investigated).
Contribution of an X-linked component to differential arsenite sensitivity
Embryonic offspring of reciprocal crosses between Oregon R 1970 and PVM adults were collected, reared on AsIII-free or AsIII-containing food as described above, and scored for percent
adult eclosion relative to their male or female siblings at that AsIII concentration (Figure 2.2).
The most notable result from these studies was that relative tolerance to dietary AsIII segregated
to a remarkable extent with parental origin of the X chromosome. Hence, males hemizygous for
the Oregon R-derived X (XR) were quite tolerant to AsIII, whereas those males that carried the
PVM-derived X chromosome (XS) were rather sensitive (see segregation scheme in Figure 2.2,
46 lower part). While this differential tolerance/sensitivity phenotype was clearly apparent in this first generation cross, its magnitude and X chromosome segregation rapidly diminished upon subsequent crosses into F2 and later generations (data not shown). We infer that (a) quantitatively significant component(s) related to the observed differential AsIII sensitivity is/are
encoded on the X chromosome, but that other segregating loci contributing to the phenotype are
present on the autosomes such that quantifiable Mendelian segregation is apparently lost in
subsequent generations.
Figure 2.2. Relative adult eclosion percentages for male and female progeny resulting from reciprocal crosses of the Oregon R and PVM strains. Embryos were seeded on arsenite-free or arsenite-containing food and the sex of eclosing adults scored as a percentage of the total flies hatching at that concentration. Strains are designated as resistant (R) or sensitive (S) based on the data in Figure 1; the crossing scheme shown below represents the expected genotype of progeny from the reciprocal crosses if the
arsenite tolerance/sensitivity gene(s) were X-
linked.
Mapping an X-linked arsenite-tolerance component to subdivision 16
Since it seemed clear that at least one region of the X chromosome was associated with a differential AsIII sensitivity phenotype, we attempted to gain some preliminary genomic location
information. To achieve this we made use of a variety of microsatellite markers mapped to different locations throughout the X chromosome (see Materials & Methods) which showed size
47 heterogeneity in direct comparisons between the Oregon R 1970 and PVM strains. In these
experiments the size of microsatellite-specific PCR fragments obtained from AsIII-tolerant F2
progeny of an Oregon R-PVM F1 heterozygous female crossed back to PVM males (see Figure
2.3A) was compared with the parental PVM and Oregon R fragments on high resolution agarose
gels. This procedure allows for recombination occurring on the X chromosome in the F1 hybrid
female to be detected in the F2 progeny. With the crossing scheme employed, only F2 progeny
that receive the relevant AsIII tolerance-encoding X chromosomal region from the F0 Oregon R
1970 parent would be expected to survive the selection on high arsenite concentration food (0.5-
1 mM). As the percentage of surviving flies that carry individual Oregon R-specific
microsatellite marker fragments increased towards 100%, we could infer increasingly tight linkage of the AsIII-tolerance region to the particular marker being scored. A representative
example of such data for the tightly linked marker 16DF[AGC9] is shown for surviving male F2
progeny (Figure 2.3B). Note that on non-selecting, AsIII-free food, eclosing flies have an approximately 1:1 distribution of the parental marker, as would be expected from randomly located recombination events in the F1 maternal X chromosome. The sum of these studies
(Figure 2.3C) led us to identify chromosome subdivision 16 as a likely location for an AsIII- tolerance region, and subsequent experiments were aimed at more closely defining (via deficiency analysis) a region where potential candidate genes could be identified and subsequently tested for their involvement in AsIII sensitivity.
48
Figure 2.3. Recombination mapping between Oregon R 1970 & PVM using strain-
specific microsatellite markers on the X chromosome to locate an arsenite
tolerance/sensitivity locus. A. Crossing scheme shows X chromosomal constitution of F1 heterozygous females backcrossed to PVM males – recombination on the female X leads
to male progeny that contain either the resistant (R) or sensitive (S) arsenite-response
allele. B. Microsatellite mapping in the 16DF region of the X chromosome shows mobility difference depending on parental source (R or S), allowing genotypic frequency to be scored in F2 males either selected for survival on 1 mM arsenite, or not selected (i.e.,
raised on normal food). C. “R”-derived allele frequencies were scored for a variety of
markers located along the X chromosome. As expected, when not selected for arsenite resistance, markers segregated in an approximately 1:1 ratio. When the “R” allele approaches 100% representation in resistant males, very close linkage to the arsenite-
responsive allele(s) is anticipated.
Creation of targeted X chromosomal deficiency lines
To create chromosomal deficiencies in this region we took advantage of the Exelixis library of
FRT-transformed lines that allow easy production (via FLP-FRT recombination) of a series of overlapping deficiencies of varying sizes [228]. Using this approach we were able to produce numerous deficiency lines within the chromosomal subdivision; several that contain overlapping
49 deficiencies spanning the 16C10 to 17A3 region of the X chromosome are depicted in Figure
2.4). All deficiency lines were confirmed by PCR analysis of genomic DNA using primers flanking the predicted deficiency limits (as deduced from coordinates available in FlyBase - data not shown). Most of these deficiencies were homozygous lethal and so were maintained as heterozygotes over the X chromosome balancer Binsinscy. Interestingly, a region towards the proximal end of that investigated here displays haploinsufficiency, while a region situated just beyond this could be deleted and homozygotes remained viable and fertile (Df #15).
Figure 2.4. X chromosomal overlapping deficiencies (Df) created using the FLP- FRTrecombination system to aid in identification of an arsenite response locus. Recovered
deficiencies are shown both above and below a physical map of the 16C-17A chromosomal region, while the annotated genetic organization of the region from FlyBase is shown directly
below it. *Haploinsufficient; ^Homozygous viable.
Arsenite sensitivity of X chromosomal deficiency lines
To determine the appropriate AsIII concentration for use in comparative sensitivity studies, we
exposed non-deficiency (but otherwise isogenic) embryos to various concentrations, ranging
from 0 to 0.625 mM, of sodium AsIII (Figure 2.5A). In this way we chose an experimental
toxicity testing threshold of 0.25 mM AsIII, a concentration at which the relative viability of
emerging w1118/Binsinscy female adults compared to those not exposed to AsIII was still one, but
50 a concentration beyond which the relative survival rate fell substantially. In order to conclude
that a particular X chromosomal deficiency line was sensitive to AsIII compared to its non-
deficient, but otherwise isogenic, parent, the ratio of emerging female adults exposed to 0.25 mM
AsIII versus those non-exposed should be significantly less than one. When tested in this way the
analysis showed that deficiency lines #9 and #11 were significantly more sensitive than all others
tested (Figure 2.5B). Inspection of the sequences removed in these lines, using annotated data
derived from FlyBase, disclosed that both lines are deficient in five genes located at cytological
subdivision 16F1, two of which, CG32496 and CG32495, appear to have resulted from an ancestral genomic duplication of sequences encompassing CG6788 and CG6835 respectively.
Intriguingly, given previous indications of the importance of glutathione (GSH) to arsenic
metabolism, the duplicated genes CG6835 and CG32495 were found to encode glutathione synthetase (GS), an enzyme which condenses γ–glutamylcysteine with glycine to create GSH in the second (and terminal) step of its biosynthesis.
51 Figure 2.5. Arsenite sensitivity of X chromosomal deficiency lines. A. Arsenite dose-response assay on parental w1118/Binsinscy strain to identify an experimental concentration threshold for testing toxic effects on Df lines. We chose to use 0.25 mM arsenite for the sensitivity assays. Each bar represents the relative viability of emerging w1118/Binsinscy adults exposed to the specified concentration of arsenite- supplemented food when compared to emerging w1118/Binsinscy adults exposed to non-supplemented food. * P < 0.01, ** P < 0.005. B. Viability ratio of various Df lines compared to that of the isogenic parental strain (w1118/Binsinscy) tested on 0.25 mM arsenite- supplemented food. * P < 0.05, ** P < 0.01. C. Deficiency #11 encompasses a region containing 5 annotated genes: CG6835 and CG32495 encode the enzyme glutathione synthetase, CG6788 and CG32496 are genes encoding cell adhesion molecules, and
CG7772 encodes a protein with carbonate
dehydratase activity.
Genetic manipulation of the GSH biosynthetic pathway drastically alters arsenite sensitivity
Previous work in mammalian tissue culture has supported a role for GSH in the cellular response to arsenic administration. GSH is synthesized in a two-step pathway that involves the initial
52 condensation of glutamic acid with cysteine by the rate-limiting enzyme glutamate-cysteine
ligase (GCL) to produce γ–glutamylcysteine, followed by the action of GS as described above.
In order to determine if this biosynthetic pathway plays a critical role in AsIII sensitivity we
decided to test the effects of genetic manipulation of its components. Fraser et al. (2003) described a viable line containing a P element insertion in the 5’ untranslated region of the gene encoding the modifier subunit of GCL (GclmL0580) that reduced cellular GSH levels to approximately 50% of wild type [314]. We induced precise excision of the P element in this line
(Figure 2.6A) and then measured the AsIII sensitivity of both. The analysis showed that the
GclmL0580 mutant line is substantially more sensitive to AsIII than the wild-type revertant (Figure
2.6B), and confirms data previously obtained in mouse cells that have sustained a Gclm knock-
out [320].
Figure 2.6. A Gclm mutant line is highly sensitive to arsenite exposure compared to a wild type revertant. A. (Top) Agarose gel analysis of a PCR fragment spanning the 5’ end of the Gclm gene in the mutant line (GclmL0580), an isogenic reference line (w1118), and several independently recovered putative revertants (Gclmrev). (Bottom) Amplification of a Gapdh1 fragment was used as a positive control. NT: no template control. B. Embryos of either the Gclm mutant line (GclmL0580) or the wild type revertant (Gclmrev) were tested for their ability to eclose as adults on arsenite- supplemented (0.125 mM) or regular food.
* P < 0.05, ** P < 0.01, *** P < 0.0001.
53 Since it is generally accepted that GCL-catalyzed synthesis of γ–glutamylcysteine
represents the rate-limiting step in GSH biosynthesis it might be expected that a reduction in activity of GS to 50% of wild-type levels, as anticipated for heterozygous deficiency lines #9 and
#11, would have little overall consequence for GSH levels in the cell. To confirm whether GS activity could be important for AsIII sensitivity, as suggested by the chromosomal deficiency
results, we initiated a series of RNA interference (RNAi) studies, directed towards GS, to independently confirm the importance of its quantitative levels of activity in AsIII toxicity. In
initial studies conducted in Drosophila S2 tissue culture cells we targeted both an upstream
(Figure 2.7A) and a downstream (Figure 2.7C) exon of the CG6835 and CG32495 genes via
transfection of appropriately located double-stranded RNA (dsRNA) oligomers. These GS-
compromised S2 cells showed heightened sensitivity towards AsIII as compared to cells exposed
to the same AsIII concentration but expressing wild type amounts of the enzyme. To determine the magnitude of RNAi-induced silencing of GS we performed quantitative real time RT-PCR of
GS transcripts. Those transcripts targeted by the upstream region dsRNA were knocked down by approximately 50% (Figure 2.7B), whereas transcripts targeted by the downstream region dsRNA were knocked down by approximately 80% (Figure 2.7D).
54
Figure 2.7. RNAi induced knock down of glutathione synthetase expression in S2 cells. A.
Double stranded (ds) RNA was targeted to the 5’ region of CG6835 and CG32495 and cell viability measured under differing concentrations of arsenite-supplemented growth medium. * P -6 < 1.0 x 10 for 35 and 45 µM As WT vs. 35 and 45 µM As GS RNAi. B. RT-PCR analysis of GS transcript levels after targeting the 5’ region. Results have been normalized to actin. * P <
0.01. C. dsRNA was targeted to the 3’ region of CG6835 and CG32495 and viability measured as described. * P < 1.0 x 10-7 for 35 and 45 µM As WT vs. 35 and 45 µM As GS RNAi. D. Real time RT-PCR analysis of GS transcript levels after targeting the 3’ region. Results have been normalized to actin. * P < 0.01.
55 To test whether similar effects could be seen in the whole organism, we procured two
lines of transgenic Drosophila engineered to inducibly express (under GAL4 regulatory control) dsRNA hairpin transcripts derived from either the 5’ or the 3’ regions of the two GS genes [229].
In order to more efficiently target GS transcripts in vivo we combined these two independent
transgenes into a single homozygous line, GSRNAi[5’/3’] (see Materials and Methods). After
crossing these compound RNAi flies to a line expressing the GAL4 protein under control of the
ubiquitously expressed daughterless (da) regulatory element, we tested progeny for their
sensitivity to food-borne AsIII, as previously described. Animals expressing the GAL4-induced
dsRNA GS hairpins displayed no adult hatching as compared to control non-GAL4-expressing
GSRNAi[5’/3’] animals when tested on food containing as little as 0.025 mM AsIII (Figure 2.8A). In
fact, we failed to observe any feeding third instar larvae under these conditions indicating a very
strong developmental toxicity had been induced. In contrast, control non-Gal4 expressing
GSRNAi[5’/3’] embryos not only formed larvae and pupae (tested at AsIII concentrations up to 0.2
mM), but developed into viable adults at high frequency. Quantitative real time RT-PCR data
showed that GS transcripts were indeed significantly reduced in the Gal4-expressing GSRNAi[5’/3’]
larvae compared to those in the non-GAL4 GSRNAi[5’/3’] larvae when tested on AsIII-free food
(Figure 2.8B). Additional control experiments demonstrated that expression of the yeast Gal4
protein by itself does not affect the development of embryos into adulthood under AsIII-free or
AsIII-supplemented conditions (Figure 2.9). We therefore conclude that, somewhat surprisingly,
the sensitivity of whole organisms to ingested AsIII depends critically on the level and/or activity
of the glutathione synthetase enzyme and not solely on that of the supposedly rate-limiting
enzyme glutamate-cysteine ligase.
56
Figure 2.8. Effects of RNAi-induced knock-down of glutathione synthetase in flies. A.
Embryos containing the GAL4-inducible GSRNAi[5’/3’] transgene were tested for their ability to eclose as adults in the presence or absence of ubiquitously expressed GAL4 on differing concentrations of arsenite-supplemented food. Survival is expressed relative to the non-
GAL4 expressing GSRNAi[5’/3’] line on control (0 mM arsenite) food. # P = 0.05, * P < 0.05,
** P < 0.01. B. Real time RT-PCR analysis of GS transcription in GSRNAi[5’/3’] flies in the presence or absence of a Gal4 transgene. * P < 0.01.
Figure 2.9. GAL4 expression per se is not overtly toxic to Drosophila either in the presence or absence of arsenite. Embryos containing the da-GAL4 transgene were tested for their ability to eclose as adults on differing concentrations of arsenite- supplemented food as a function of the presence or absence of the GAL4-inducible
GSRNAi[5’/3’] transgene. # P = 0.05, * P <
0.05, ** P < 0.01.
57 Discussion
The long-term ingestion of arsenic-contaminated ground water represents one of the
worst environmental calamities in history. Not only is such water extensively consumed in
developing nations, such as Bangladesh [309] and parts of India [321] but also in more highly developed countries such as China [322] and the United States [14]. Its health effects range widely, from conditions such as diabetes mellitus, peripheral vascular disease (e.g. Blackfoot disease) and neuropathy, to a large variety of cancers. Inter-individual genetic variations have been proposed as contributing to differences in susceptibility and response - for example, polymorphisms in either the AS3MT [182] or GSTO [180] genes have been associated with
increased likelihood in developing arsenic-related diseases. Given the pleiotropic nature of
disease outcomes upon arsenic exposure and the large number of pathways implicated in its
biological interactions (reviewed in [78, 323], an unbiased approach to determining genes and/or
pathways involved in differential susceptibility to its toxic effects at the whole organismal level
seems warranted. Here, we have used natural geographic variants of Drosophila melanogaster
as a model system to test for genetic factors present in wild type populations that may possibly
predispose to arsenic susceptibility. Historically, Drosophila has been used to shed light on
many fundamental biological processes, such as organismal development, owing to its ease of
genetic manipulation and analysis. However, it has become an increasingly used model system
to study human disease processes and signal transduction, owing to its unexpectedly high content
of cognates to many genes involved in human genetic disease and metabolic pathways (see
Introduction).
By employing a combination of classical chromosomal segregation and microsatellite
marker-based recombination analyses, together with the creation of a series of overlapping
58 deficiencies in the inferred region of interest, we identified a small region of the X chromosome
(defined by deficiency #11 and encompassing cytological location 16F1) as being of particular
interest with regard to arsenic susceptibility. The genomic sequence information available in
FlyBase for this region, together with the inferred annotation of its genetic function, shows that a
direct sequence duplication has apparently occurred, such that two copies of the glutathione
synthetase (GS) gene are present, along with two copies of an adjacent gene encoding a cell
adhesion protein and a single gene encoding a protein with carbonate dehydratase activity.
Given well-documented observations that GSH appears to play a role in the defense of cells against arsenic toxicity [324], it was obviously of interest that an enzyme involved in the biosynthesis of GSH was implicated by our genetic analysis. On the other hand, that this enzyme
might be GS seemed somewhat surprising in light of the prevailing view that it is the enzyme
that precedes GS in the two-step GSH biosynthetic pathway, namely glutamate-cysteine ligase
(GCL), that provides the rate-limiting step (reviewed in [325]. Indeed, when we examined a fly
line with a P-element insert in the 5’ UTR of the Gclm gene (encoding the regulatory subunit of
the heterodimeric GCL enzyme) that causes a ~2-fold reduction in GSH levels, it displayed very
high sensitivity to AsIII, confirming previous studies performed in Gclm knockout mouse embryo
fibroblasts [320]. Thus, one prediction would be that moderately reduced GS expression, as
anticipated in deficiency #11 owing to its 50% reduction in GS gene dose, would not affect
overall GSH levels in the cell, nor its AsIII sensitivity, as long as its substrate (γ-glutamyl
cysteine derived from the GCL-catalyzed step) was present at normal levels. The first part of
this prediction was true, since we found that GSH levels in deficiency line #11 appeared similar
to those of its non-deficient parent (data not shown). However, such deficient flies showed
59 distinct AsIII sensitivity, and encouraged us to investigate the role of GS in AsIII sensitivity in
greater detail.
RNA interference-based knockdown analysis in S2 tissue culture cells amply confirmed
that reduction in the expression of GS produced sensitivity to AsIII. Most strikingly, however,
whole organism knockdown of GS (to approximately 30% of normal levels) induced extreme
sensitivity, with complete developmental toxicity occurring at up to 10-fold lower concentrations
(and potentially even less) than those at which the first conspicuous effects on adult eclosion
typically start to occur. Particularly noticeable was the fact that this toxicity appeared to occur
very early in development (presumably shortly after embryo hatching), since very few active
larvae could be observed under these conditions. Though these data seemed highly contradictory
based on the prediction outlined above (in support of which Drosophila GCL has been shown to be rate-limiting [326] as in other organisms), they make a good deal more sense when the pathway for glutathione biosynthesis is viewed in a broader context. The key to this is understanding that GSH is not a static component in the cell under conditions of AsIII -induced
III stress. This is because it is actively bound by As in a stable As(GSH)3 complex [68], which not
only ties up free GSH from participating in its role as an antioxidant and regulator of the redox
state of the cell (many studies have shown high levels of ROS in the presence of arsenic – see
Kumagai and Sumi, 2007), but which also provides the substrate for active transport of arsenic
out of the cell by the multidrug resistance proteins [66]. It is in this situation of both synthesis
and active consumption of GSH that the rate-limiting properties of GCL are likely to be
compromised, because changes in flux through the pathway (as would be produced under AsIII
stress conditions when GSH is being consumed at a much higher rate) become sensitive to other
steps in both the supply and demand pathways. Furthermore, though GCL is feedback-inhibited
60 by GSH under zero or low GSH consumption conditions (forming the basis for its reported rate-
limiting behavior), this inhibition will be substantially relieved in a high GSH consumption
situation, allowing other control points (such as the GS-catalyzed step) to contribute to a
correspondingly greater extent. Such supply and demand considerations, inherent in the
biochemical approach known as Metabolic Control Analysis, have been recently discussed in
great detail, both for the GSH pathway [327] and for metabolic pathways in general [328]. In the
present case, it provides an extremely plausible rationale for why reduced GS activity sensitizes
cells that are experiencing chronic stress from AsIII exposure. The fact that this effect is
particularly strong when considering developmental susceptibility as compared to cultured cell
susceptibility emphasizes the importance of a whole animal model in studying mechanisms and
pathways of toxicity.
These results on the effects of reduced GS expression suggested that the differential
sensitivity displayed by the PVM and Oregon R 1970 strains towards AsIII might be due to
sequence polymorphisms in the CG6835 and/or CG32495 genes between the two strains. Such
polymorphisms could lead to differences in levels of the enzyme, differences in the levels of
transcripts, differences in transcript splice variants, or differences in enzyme activity, any or all
of which might then lead to differential availability of GSH under the sustained stress of AsIII
intoxication. While it is clear that both strains do contain the duplicated GS genes, we have not yet sequenced the genes and their flanking regions in the two strains in order to address these possibilities; preliminary analyses of multiple GS transcripts (i.e., splice variants) and their levels
have shown a good deal of complexity is present (data not shown). We have also measured GSH
levels in these two strains under both control and AsIII-stressed conditions and have not found
obvious differences (data not shown). However, these data might easily be compromised by the
61 fact that typical GSH assays [329] cannot distinguish between GSH and the substrate for the GS
reaction (i.e., -glutamyl cysteine), so this distinction needs to be made in further investigations.
According to the results described here, even though the GCL heterodimer is the rate-
limiting enzyme in the production of GSH under normal conditions, in the presence of AsIII (and
potentially other heavy metal toxicants) optimal GS activity is required to sustain high enough levels of bioavailable GSH to protect cells, and thus an organism, against the effects of the chronically ingested toxicant. The HapMap consortium has reported single nucleotide polymorphisms (SNPs) in the GS gene of individuals from different regions worldwide [330], and GS deficiency (whole or partial) is a well-described inherited autosomal recessive human condition (reviewed in [331]. Since it is clear that the synthesis and use of GSH in defense against arsenic intoxication is a common feature of both the invertebrate model studied here and the mammalian situation, we suggest that future studies of genetic polymorphism in human populations exposed to arsenic should consider potential associations between variant alleles of
genes in the GSH biosynthetic pathway (such as GCLM and GS) and disease susceptibility.
62 Chapter III. (All or part of the following is currently being prepared for submission to Environmental and Molecular Mutagenesis)
Establishment and Characterization of a Drosophila Model to Investigate the
Biological Effects of Arsenic Methylation In Vivo
Introduction
Arsenic contamination of drinking water supplies in both developed and underdeveloped countries, especially Bangladesh, may be the worst environmental disaster many parts of the world currently face. Chronic consumption of arsenic-contaminated water can induce a myriad of cancerous and non-cancerous conditions [310]. Owing to the worldwide health effects reported from consumption of arsenic-contaminated water, the World Health Organization [38] and the United States Environmental Protection Agency [39] have set the maximum contaminant limit for arsenic in water to 10 µg/L (10 ppb).
Arsenic in drinking water is present in two oxidative states, arsenite (AsIII) and arsenate
(AsV). Although the exact toxic mechanism of long term, low level exposure to arsenic is not known, it has been suggested that AsIII can induce the generation of reactive oxygen species
(ROS) that can oxidize DNA bases [73] and/or proteins required for signaling cascade pathways
[78]. AsIII can also interfere with the DNA damage repair machinery [104, 105], which may occur through its tight binding to dithiols in proteins [95, 109, 332]. It has also been suggested that arsenic can disrupt the epigenetic machinery by altering the acetylation and methylation status of histones [113, 114] and methylation of DNA [112, 120].
Once inside mammalian cells inorganic arsenicals (iAs) can undergo a series of successive methylation reactions, via the arsenic(III) methyltransferase (AS3MT) enzyme [139],
63 leading to the formation of monomethylarsonous acid (MMAIII) and dimethylarsinous acid
(DMAIII), which can be oxidized to form monomethylarsonic acid (MMAV) and dimethylarsinic
acid (DMAV) [132, 133]. Alternatively, AsIII and MMAIII may undergo a series of oxidative methylation reactions, producing MMAV and DMAV, respectively [131, 333], which are
potential reduction substrates of the enzyme glutathione S-transferase omega (GSTO), as in vitro
studies have shown [136, 137]. However, arsenic-reducing activity was retained in GSTO
knock-out mice [138] and the AS3MT enzyme can provide the methylating activity necessary by
linking oxidative methylation with reduction of arsenic metabolites, using GSH as a cofactor,
thereby obviating the need for GST enzymatic activity [132, 140, 141].
Metabolism of arsenic to produce methylated arsenicals (MAs) was previously thought to
be a detoxification process based on unsophisticated whole animal LD50 determinations of acute
treatment with various arsenicals, and because the MAs are more water soluble than iAs and
therefore more readily excreted [144-146]. Moreover, experiments performed using the
predominant methylated species detected in urine, namely DMAV and MMAV [147], showed that
they were less acutely toxic than iAs [69]. However, when improved technology detected both
DMAIII [149] and MMAIII [148] in the urine of exposed individuals, additional studies were
performed to determine their toxic potential. Experiments performed in vitro [152] and in vivo
[150, 151, 154] have demonstrated that MMAIII are more genotoxic than iAs and pentavalent
MAs. The efficiency of the initial methylation step has been directly correlated to a high risk of
developing arsenic-related carcinogenic health effects, presumably because of the high levels of
MMAIII produced [14, 156, 157].
The current relative lack of understanding regarding the mechanisms of arsenic-induced
carcinogenicity is, in part, due to the absence of a single mammalian model that can recapitulate
64 typical human outcomes or pathology (reviewed in [200, 201]). Many mammals genetically
similar to humans do not possess AS3MT activity (reviewed in [202]). Attempts to establish
hamster, rat, mouse and transgenic mouse model systems have been made [204], however the data obtained and the techniques used often do not translate to the effects observed and the methods of exposure in human populations. Several studies using rodents as a model system have concluded that arsenic is not a complete carcinogen [209, 210]; the evidence for or against this view in humans is currently lacking.
While clearly not a mammalian system, Drosophila offers a variety of advantages to study the biological pathways through which environmental stressors (e.g. arsenic), exert their pleiotropic, often pathologic, effects. Foremost among these are the variety of genetic manipulations available that help shed light on specific molecular processes involved (reviewed
in [224-226]). A second major aspect of Drosophila as a model concerns its high conservation
of genes that are involved in many human disease pathways, e.g. pathologies as assorted as
Alzheimer’s [243-247] and congenital heart disease [248, 249]. A BLAST analysis against the
Drosophila genome of 929 human disease genes with at least one mutant allele in the Online
Mendelian Inheritance of Man (OMIM) database identified 714 human genes (77%), formatted
into a searchable database called Homophila, that matched 548 unique sequences in the fly
genome [234, 235]. According to the FlyAtlas database many fly homologs of human genes are
expressed in fly tissues that are analogous to those of the affected human tissues [250]. As a
result of these insights Fly models have been established to study the mechanisms involved in
diseases as diverse as Parkinson’s [237, 238], and cancer [239, 240] (reviewed in [241, 242]).
As far as practicalities go, raising multiple generations of flies is relatively quick and cheap,
65 particularly when compared to mouse colonies, nor are there federal regulations or statutes that apply to most experiments in Drosophila.
The Drosophila system has previously been used to assess the genotoxic potential of a wide variety of environmental chemicals and stressors [261, 282, 334]. Previous studies employing a loss of heterozygosity (LOH) assay called the Somatic Mutation and Recombination
Test (SMART) [251] demonstrated that Drosophila adults derived from larvae fed iAs did not show any obvious genotoxic effects. However, MAs were not detected in larval homogenates, suggesting that methylation was not occurring in Drosophila [308]. In contrast, when larvae
were actually fed DMAV, adults scored strongly positive for SMART. This not only showed that
Drosophila was responsive to MAs, but that the nature of the response was fully in accord with the suggested higher genotoxicity of such methylated arsenic species seen previously in mammalian cell culture studies [152]. A BLAST search for the hAS3MT gene sequence in the
Drosophila genome showed that flies apparently do not harbor a homolog of the gene, providing
a compelling reason as to why larvae fed iAs alone did not produce MAs.
We have taken advantage of the fact that Drosophila does not methylate inorganic
arsenic (presumably because it does not harbor the AS3MT gene), and have created a series of
transgenic fly lines that carry a functional form of the human enzyme. We have characterized
the expression of this enzyme in the fly in terms of its effects on viability and development and
its genotoxic potential in the presence or absence of AsIII using a sensitive LOH SMART assay,
based on the lats/wts gene that can provide readily scorable phenotypes. The wts protein is a
serine/threonine kinase tumor suppressor [269] involved in the Hippo pathway that controls
organ size by regulating cell growth, proliferation, and apoptosis [270-273] via response to cell-
cell contact [274]. In the lats-SMART assay, if a mutation in the wild type allele of the wts gene
66 occurs in a heterozygous wts+/- 3rd instar larva exposed to a genotoxin, large dramatic
overproliferated tumorigenic clones can form and be observed in the eclosed adult fly [269, 279,
280]. Such a model will potentially allow us to more easily dissect the mechanisms involved in
the genotoxicity and/or carcinogenicity of MAs using the battery of genetic approaches for
which Drosophila is renowned.
Materials and Methods
(For additional methods, see Appendix II)
hAS3MT cloning
The hAS3MT sequence was isolated from a human kidney cDNA library kindly provided by Dr.
William Miller at the University of Cincinnati. The sequence was amplified in a 25 µl reaction
using Phusion™ Polymerase, (NEB®, Ipswich, MA), the forward primer 5’-
CACCATGGCTGCACTTCGTGACGCTG and the reverse primer 5’-
GCAGCTTTTCTTTGTGCCACAGCAG. The underlined sequence in the forward primer
allowed for unidirectional cloning into the pENTR™/D-TOPO® vector (Invitrogen™, Carlsbad,
CA). The PCR conditions were 2 min at 98C, 30 cycles of 30 s at 98C, 30 s at 65C and 1 min
at 72C, and an extension for 5 min at 72C. The reaction product was analyzed on an agarose gel and purified using the MinElute™ Gel Extraction Kit (Qiagen, Valencia, CA) according to
the manufacturer’s directions. To confirm the correct PCR product, a restriction endonuclease digest reaction was performed with XbaI. The hAS3MT PCR product (1 μl, 2.5 ng) was mixed with 1 μl salt solution (1.2 M NaCl, 0.06 M MgCl2), dH2O to a to a total volume of 5 µl and 15-
20 ng/μl of the TOPO® vector. The reaction was incubated at RT for 5 min, placed on ice and
transformed into One Shot® TOP10 chemically competent E. coli according to manufacturer’s
directions (Invitrogen™). A plasmid preparation of a 5 ml liquid culture was performed using
67 the QIAprep® Spin Miniprep Kit (Qiagen, Valencia, CA). Sequencing of purified DNA was
performed by Genewiz®, Inc. (South Plainfield, NJ). Positive transformants were confirmed via restriction endonuclease analysis using PvuII.
Site-directed mutagenesis
Nucleotide mutagenesis was performed using the QuikChange® II Site-Directed mutagenesis kit
according to the manufacturer’s directions (Stratagene®, La Jolla, CA). Primers used to mutate
C620 to T: F 5’-TTTATGGGGTGAGTGTCTGGGTGGTGCCTTTATACT and R 5’-
AGTATAAAGCACCACCCAGACACTCACCCCATAAA. Primers used to mutate C860 to T:
F 5’- AATTACAGGACATGAAAAAGAACTAATGTTTGATGCCAATTTTACATTTAAGG
and R 5’-
CCTTAAATGTAAAATTGGCATCAAACATTAGTTCTTTTTCATGTCCTGTAATT.
pENTR/D-TOPO-AS3MT and UAS LR recombination reaction
Each Drosophila specific Gateway® transformation vector, obtained from the Drosophila
Genomics Resource Center (Indiana University, Bloomington, IN), consists of a GAL4-specific
UAS sequence, which drives expression of an epitope-tagged protein-encoding gene, P-element
sequences allowing for genomic insertion of the vector, and a w+ marker to identify positive
transformants [230]. We used the pTHW (1099) vector for expression of an N-terminal HA-
tagged hAS3MT, the pTWH (1100) vector to express a C-terminal HA-tagged hAS3MT and the
pTFW (1115) vector to express an N-terminal FLAG-tagged hAS3MT. A mixture consisting of
150 ng of the pENTR-hAS3MT construct, 150 ng Gateway® vectors (pTHW, pTWH, or pTFW),
LR clonase II (2 μl) was incubated at 25C for 1 h. Subsequently, Proteinase K (1 µl) was added to the recombination reaction, incubated at 37C for 10 minutes, and the mixture used to
68 transform DH5α competent cells following standard procedures (Invitrogen™, Carlsbad, CA).
Colonies were grown in 5 ml liquid cultures and preparation of DNA was performed using
QIAfilter Plasmid Purification Midi Kit (Qiagen, Valencia, CA). Positive recombinants and transformants were confirmed via restriction endonuclease analysis using PvuII. Verified
recombinant DNA constructs were provided to Rainbow Transgenic Flies, Inc. (Newbury Park,
CA) for production of transformed Drosophila lines.
Flies
Flies were maintained on standard cornmeal medium at room temperature. Eclosing adults from
injected w1118 embryos were individually collected and crossed to w1118 males or female. We used standard procedures to identify the chromosomal insertion of the hAS3MT construct. The
GAL4 expressing lines y1 w*; Act5C-GAL4 and w1118; da-GAL4 were obtained from the Indiana
University Stock Center at Bloomington, IN. The w; latsx1/TM6B, Tb line was kindly provided
by Dr. Tian Xu [269]. We created the hAS3MT; latsx1/TM3, Sb line following standard
procedures using a variety of balancer lines.
Genomic analysis of transgenic flies
PCR for hAS3MT
Each 50 µl PCR reaction included 1 µl of genomic DNA, 5 µl 10X ThermoPol reaction buffer
® ® (NEB , Ipswich, MA) 2 µl 25 mM MgCl2, 1 µl 10 mM dNTP mix (NEB ), 1 µl forward primer
(1 µg/µl), 1 µl reverse primer (1 µg/µl), and 2.5 units (0.5 µl) Taq polymerase (NEB®). Reaction conditions were 95° C for 10 min; 94° C for 30 s, 52.4° C for 1 min, 72° C for 2 minutes, repeated 30 times; and 72° C for 10 min. Primers used for PCR are as described above for cloning the hAS3MT cDNA. PCR products were analyzed via agarose gel electrophoresis.
69
Analysis of hAS3MT mRNA transcription
Reverse Transcriptase PCR (RT-PCR)
First strand synthesis was performed using the iScript cDNA synthesis kit using 1 µg total RNA according to manufacturer’s directions (Bio-Rad, Hercules, CA). Each 50 µl PCR reaction
® included 2 µl of cDNA, 5 µl 10X ThermoPol reaction buffer (NEB ), 2 µl 25 mM MgCl2, 1 µl
10 mM dNTP mix (NEB®), 1 µl forward primer (1 µg/µl), 1 µl reverse primer (1 µg/µl), and 2.5
units (0.5 µl) Taq polymerase (NEB®). Primers used for PCR: F 5’-
ATTGAGAAGTTGGGAGAGGCTGGA and R 5’- TCTTCTGGCAGTTCAAGGCTCGTA.
PCR conditions were as follows: 2 min at 94°C, 30 cycles of 1 min at 94 °C, 1 min at 62.6°C,
and 30 s at 72°C, and an additional elongation step for 10 min at 72°C. PCR products were
visualized via agarose gel electrophoresis.
Western Blot analysis
Three adult flies were sonicated in 200 µl SDS loading buffer, and boiled for 5 minutes and
centrifuged at 15,800 x g for 3 minutes. The homogenates (20 µl) were separated by SDS-PAGE
and transferred to a nitrocellulose membrane following standard procedures. Immunoblotting
was performed using antibody dilutions for an HA epitope of the fused HA-hAS3MT at 1:1,000
(provided by Dr. William Miller), an epitope of hAS3MT at a dilution of 1:1,000 (provided by
Dr. Richard Weinshilboum) and Actin (provided by Dr. James Lessard) as a loading control at
1:10,000. Horseradish peroxidase conjugated anti-rabbit (HA and hAS3MT) or anti-mouse
(Actin) secondary antibody was used at a 1:1,000 dilution and SuperSignal West Pico
Chemiluminescent Substrate (Pierce, Rockford, IL) was used for signal amplification. Signals
were visualized by exposure to Cole Parmer Blue-Sensitive X-ray film.
70 Arsenic speciation
Drosophila exposure and sample preparation
Embryos from relevant crosses were collected as described previously [335] and allowed to
develop in AsIII-free or 0.1 mM AsIII-supplemented instant Drosophila Medium™ (Carolina
Biological, Burlington, NC). Sample preparation for analysis was performed as previously reported [308]. Briefly, thirty 3rd instar larvae were collected and homogenized in 2 mL
homogenizing buffer (250 mM sucrose, 1 mM MgCl2, and 10 mM Tris-HCl (pH 7.4)) using a
glass homogenizer with Teflon pestle. The homogenates were centrifuged at 105,000 x g for 90 minutes using a Beckman L7-55 ultracentrifuge, transferred to a 1.5 mL microcentrifuge tube, and stored at -80C until use.
Speciation analysis
Arsenic speciation via ion pairing reversed phase liquid chromatography followed by inductively coupled plasma mass spectrometry (ICPMS) for detection was performed at the University of
Cincinnati/Agilent Technologies Metallomics Center of the Americas following the methods previously described [336] with some adjustments.
Reagents and standards
Mobile phases and standards were prepared in 18 MΩ cm-1 doubly deionized water (DDW)
purified by cartridges from Sybron/Barnstead (Boston, MA). The mobile phases for the
chromatographic separation were as follows: mobile phase A consisted of 2.5 mmol l-1
tetrabutylammonium hydroxide from Fluka (Milwaukee, WI) and 2.5 mmol l-1 ammonium phosphate from Sigma-Aldrich (St.Louis, MO) at pH 6.0 and mobile phase B contained 10 mmol l-1 ammonium sulfate from Sigma-Aldrich at pH 6.0. Phosphoric acid and ammonium hydroxide
were used for pH adjustment for mobile phase A and mobile phase B, respectively. Standards
71 included methanearsonate hexahydrate (MMA) purchased from Chem Service (West Chester,
PA), dimethylarsinic acid (DMAV) from Fluka and AsIII as well as AsV, acquired from Sigma-
Aldrich (St. Louis, MO).
High Performance Liquid Chromatography
Chromatographic separations were performed with an Agilent 1100 liquid chromatograph
(Agilent Technologies, Santa Clara, CA) equipped with a binary HPLC pump, an autosampler, a vacuum de-gasser system and a thermostated column compartment. Reverse phase chromatography was performed with a ZORBAX Eclipse XDB-C18 column (5 µm x 4.6 mm id x
250 mm) (Agilent Technologies). The ion pairing reversed phase separation was accomplished utilizing the following gradient: 0-1 min 0% B, 1-2 min 0-100% B, 2-6 min 100% B, 6-7 min
100-0% B and 7-20 min 0% B with 1 ml min-1 flow and 100 µL injection.
Inductively Coupled Plasma Mass Spectrometry
The ICPMS used for arsenic detection at m/z 75 was an Agilent 7500ce. The instrument was equipped with a microconcentric nebulizer made by Glass Expansion (Pocasset, MA), a Scott double channel spray chamber (2C), a shielded torch, a CE lens stack, an octopole collision/reaction cell with hydrogen gas pressurization (purity of 99.999%), a quadrupole mass analyzer and an electron multiplier. 75As detection was accomplished at the following conditions: quadrupole bias -16, octopole bias -18 (a net + 2 volt energy discrimination barrier), and collision gas flow rate at 3.5 ml/min.
Arsenite sensitivity of wild type and transgenic lines
The response to arsenicals on the development and viability of hAS3MT-expressing and non- expressing lines was performed as previously described [335]. Briefly, flies were allowed to lay eggs on a small grape juice agar plate, seeded with yeast paste, inserted into the neck of an
72 inverted culture bottle. 150 embryos (0-16 h old) were transferred to a piece of sterilized filter
paper, which was then placed on 5 g of Instant Drosophila Medium™ hydrated with 30 ml of
III H2O or As solutions of various concentrations. Emerging adults were counted, and eclosion
data compared on AsIII-supplemented and non-supplemented food.
Data analysis
Data analysis was performed as previously described [335] with some adjustments. Briefly, an
eclosion average was calculated from three bottles of a control cross (w1118, da-GAL4) without
AsIII. This average was compared to the average of three bottles of eclosing w1118, da-GAL4
adults exposed to 0.1 mM AsIII, w1118; hAS3MT adults eclosing from three bottles with 0.1 mM
AsIII-supplemented or AsIII-free food and w1118; hAS3MT, da-GAL4 adults eclosing from three
bottles of AsIII-supplemented food and three bottles of AsIII-free food. If the genomic insertion
or induced expression of hAS3MT in the presence or absence of AsIII produced sensitivity then
the viability ratio of the induced or uninduced transgenic lines should be significantly lower than
that of control lines. On the other hand, if there is no effect on hAS3MT enzymatic expression and function the ratios should not be significantly different.
Arsenite genotoxicity assay
To determine if arsenite is genotoxic in Drosophila the w; hAS3MT; latsx1/TM3, Sb line or the w;
latsx1/TM6B, Tb were crossed to the w1118; da-GAL4 line, and females allowed to lay eggs on
AsIII-free or AsIII-supplemented (0.1 mM) instant Drosophila Medium. Eclosing adults were
observed under a stereo microscope at a magnification of 40X and scored for tissue overgrowth.
We analyzed and compared the data by dividing the amount of tumors observed by the amount
of adult flies scored from each treatment and genotype.
Statistical analysis
73 Data were analyzed by Chi-square test with Yate’s correction, using SigmaPlot v. 11 (Systat
Software, Inc.) and critical significance level α = 0.05.
Results
Sequencing and transformation of hAS3MT cDNA
Sequencing of the cloned hAS3MT cDNA construct in the pENTR vector revealed two single nucleotide polymorphisms (SNPs), one that has not been reported in the literature (T620C) and one that has been reported in populations (T860C) currently exposed to arsenic in drinking water [182] (data not shown). These two polymorphic variants are considered mutant forms of the hAS3MT gene due to the fact that other alleles are found at higher frequencies in human populations [185]. The T620C SNP leads to a Leu to Pro change in amino acid residue 207 and the T860C SNP leads to a Met to Thr change in amino acid residue 287. We performed site- directed mutagenesis to reverse the observed SNPs to the ‘wild-type’ form (most common allele reported by [185]) prior to in vitro recombination of the hAS3MT cDNA construct into three different Drosophila-specific Gateway™ P element vectors. These vectors, when present as transgenic constructs in flies, allow expression of N- or C-terminal tagged proteins under inducible UAS/GAL4 control (see Materials and Methods). Each of these was injected into recipient Drosophila embryos and transformants selected by their wild-type (red) eye color.
Several lines were established for each of the three Gateway™ constructs injected. These were designated as 1099 (N-terminal HA tag), 1100 (C-terminal HA tag), and 1115 (N-terminal
FLAG). Lines were checked for chromosomal location of the inserted transgene by standard balancer chromosome crossing.
74 Characterization of transgenic flies
Integration and expression of hAS3MT
After establishing several lines with a red eye phenotype suggesting positive integration
of the hAS3MT construct, we isolated whole fly DNA to confirm its presence via PCR using
hAS3MT specific primers. Agarose gel analysis of products from two independent transgenic
lines, 1099-2E and 1099-2F, demonstrated positive genomic integration of the hAS3MT construct
when compared to a non-transgenic w1118 fly line (Figure 3.1A).
As illustrated in Figure 3.1B, the integrated hAS3MT gene from the 1099-2E and 1099-2F
transgenics was transcriptionally induced in progeny obtained from crosses to two different lines
expressing the GAL4 protein under the control of the ubiquitous regulatory elements
daughterless (da) (w1118; da-GAL4) or Actin 5C (Act5C) (y1 w*; Act5C-GAL4). As expected,
little hAS3MT transcription was seen in adults emerging from a cross of either of the two
transgenic lines with a w1118 line that does not express the GAL4 protein.
Biochemical characterization
Even though low hAS3MT mRNA levels are observed in non-induced adults, Western
analysis showed no detectable hAS3MT protein expression (Figure 3.1C, top and middle panels).
In contrast, the hAS3MT protein is highly expressed in adult flies emerging from the transgenic
(1099-2E and 1099-2F) cross with either da-GAL4 or Act5C-GAL4, as observed when either an antibody against the HA fusion epitope (Figure 3.1C, top panel) or an antibody raised against a recombinant human hAS3MT protein [185] were used for detection (Figure 3.1C, middle panel).
75 Figure 3.1. Molecular characterization of hAS3MT transgene insertion and expression. The designations 1099-2E and 1099-2F are two independent pTHW transgenic lines. A. Agarose gel analysis of PCR products derived from hAS3MT insertion into w1118 genome. Gapdh1 was used as a loading control. B. Agarose gel analysis of reverse transcriptase PCR for hAS3MT mRNA. Transgenic lines were crossed into either a non-GAL4 or a GAL4-expressing background. Gapdh1 was used as a loading control. RT = reverse transcriptase. C. Western analyses for the expression of the AS3MT enzyme. Actin was used as a loading control. Ab = antibody used.
hAS3MT enzymatic function
In order to determine whether the cloned hAS3MT displayed any functional enzymatic activity of in Drosophila, detectable as production of methylated arsenic species, we collected 3rd instar larvae derived from each of three independent crosses of the 1099-2E transgenic line and raised on 0.1 mM AsIII-containing food. We chose to study the 1099-2E transgenic line for this analysis based on the fact that it has lower levels of constitutive hAS3MT mRNA transcription when crossed to a non-GAL4 inducing line compared to the 1099-2F transgenic line according to
76 RT-PCR results. Thus, the 1099-2E line was crossed to the non-GAL4-containing w1118 line
(negative control) or to the GAL4-expressing line w1118; da-GAL4. In addition, to eliminate the
possibility that GAL4 expression by itself could lead to production of MAs, w1118 flies were
mated to the da-GAL4 line. Homogenates of the different larvae were analyzed for the presence
of various arsenic species using an HPLC-linked inductively coupled mass spectrometry
approach [336].
An HPLC/ICP-MS-derived chromatogram produced using arsenic standards shows
separation of inorganic AsIII from AsV, as well as the ability to distinguish MMA and DMA species from the inorganic precursors (Figure 3.2A). When samples derived from larvae were analyzed, neither the separate presence of GAL4 (Figure 3.2B) nor the uninduced hAS3MT
transgene (Figure 3.2C) led to the production of methylated arsenic species. However, 3rd instar larvae in which induction of the hAS3MT enzyme was obtained via GAL4-regulated expression were able to produce high levels of DMA and lower levels of MMA species (Figure 3.2D). As expected 3rd instar larvae from the same crosses not exposed to AsIII produced no trace of MAs
(data not shown).
77
Figure 3.2. Functional enzymatic analysis of hAS3MT activity via HPLC/ICPMS.
A. Chromatogram of homogenizing buffer spiked with arsenic standards. B. Chromatogram of larval extracts derived from a w1118 cross and da-GAL4 and exposed to 0.1 mM AsIII. C. Chromatogram of larval extracts derived from a 1099-2E hAS3MT cross to w1118 exposed to 0.1 mM AsIII. D. Chromatogram of larval extracts III from a 1099-2E hAS3MT cross to da-GAL4 and exposed to 0.1 mM As . Each peak in B.-D. represents the abundance of a specific arsenic species in larval homogenates.
78 Arsenite sensitivity assays
To determine if expression of hAS3MT, and thus production of MAs, induces effects on the developmental cycle or the ability of adults to eclose (i.e. viability), we exposed embryos derived from each of the three crosses described above to AsIII-free food or food supplemented with 0.1 mM AsIII. We neither observed any significant (albeit qualitative) delay in the developmental cycle nor any significant quantitative effects on viability of the hAS3MT- expressing line as compared to the two non-hAS3MT expressing lines (Figure 3.3). Such data are consistent with the idea that neither the presence nor the expression of the hAS3MT transgene compromises normal physiological processes occurring in Drosophila growth and development.
Figure 3.3. Relative viability of varying genotypic configurations tested by adult
eclosion on AsIII-free or AsIII-containing food. Individual genotypes are grouped in pairs from left to right (0 mM or 0.1 mM AsIII) and are equivalent to those tested in Figure 3.2, B.-D. respectively. Viability is expressed relative to that seen for da-GAL4 adults on 0 mM AsIII.
79 Arsenite genotoxicity assay
To determine if the MAs produced in the hAS3MT transgenic Drosophila line are
genotoxic, we performed an LOH assay using the lats-SMART system, in which evidence of
induced chromosomal instability in imaginal disk cells can be monitored by the formation of
epithelial tumor outgrowths [269, 279, 280]. Table 3.1 shows the frequencies of tumor
formation per fly derived from various genotypes and arsenic treatments. Tumors were not
observed in the absence of AsIII in “control” (+/+) and hAS3MT eclosing adults, as would be
expected. The frequency of tumor development in lats and the compound hAS3MT; lats adults
unexposed to AsIII did not differ significantly. When exposed to 0.1 mM AsIII the frequency of
tumor formation in lats adults did not increase significantly (~1.3-fold), however the frequency
of tumor formation in the compound hAS3MT; lats adults increased by more than three-fold.
The frequency of tumor formation in the hAS3MT; lats adults exposed to 0.1 mM AsIII was two- fold higher than lats adults also exposed to 0.1 mM AsIII. Compared to non-arsenite exposed
eclosing adults, the frequency of tumor formation increased significantly in adult flies eclosing
from arsenite-supplemented food, irrespective of phenotype (P = 0.038). The level of tumor
formation frequency also increased significantly, irrespective of AsIII exposure in eclosing adults
harboring the heterozygous lats mutation, compared to the non-lats adults (wild-type and hAS3MT; P < 0.001) (Table 3.2). However, no statistical significance could be established in the lats; hAS3MT adults emerging from arsenite-supplemented food compared to control (0 mM
AsIII) food, due to the low numbers of adults scored. Future experiments will take into account
the numbers of adults to be scored in order to obtain statistically significant differences. We
observed tumor outgrowths of varying size, in various parts of the body, including the head,
wing, abdomen, thorax, and leg (see Figure 3.4 for examples).
80
Table 3.1. Frequency of MAs-induced tumor formation using lats-SMART assay. All genotypes expressed the GAL4 inducer. * P = 0.038
III Table 3.2. Frequency of tumor formation in non-lats and lats adults irrespective of As treatment. * P < 0.001
Figure 3.4. Examples of tissue outgrowths in a variety of Drosophila tissues, for example the leg (L), the thorax (T), and the wing (W), induced by LOH of the lats gene via genotoxic injury through production of MAs (H = head).
81 Discussion
The chronic consumption of arsenic-contaminated water for consumption is a world wide problem encountered in both developed (e.g. United States) [14] and underdeveloped countries
(e.g. Bangladesh) [337]. A variety of health effects such as bladder [14], kidney [13], and skin
[158] cancer, as well as non-cancerous conditions such as diabetes mellitus [36], Blackfoot disease [338], and peripheral neuropathy [35], have been attributed to chronic consumption of arsenic-contaminated drinking water. Upon ingestion of inorganic arsenic and its uptake into human cells, arsenic is methylated to produce MMAIII, MMAV, DMAIII, and DMAV [135] via either a series of successive reductive-methylations of the pentavalent forms followed by oxidation of the trivalent forms [132, 133], or a series of oxidative-methylations of the trivalent forms followed by reduction of the pentavalent forms [137, 143, 333].
The ability to methylate arsenic and produce MAs is strongly dependent on the AS3MT enzyme, as shown by RNA interference-mediated AS3MT knock-down studies in HepG2 human hepatic cells demonstrating that levels of methylation decreased by 70% compared to WT cells
[142]. The authors stress that production of methylated arsenicals (MAs) might be independent of hAS3MT, however since the knock-down of the mRNA transcripts is not 100%, the remaining enzymatic levels might be enough to produce MAs. While the methylation of arsenic was previously deemed a detoxification process, recent in vivo and in vitro studies have shown that the methylated forms of arsenic, especially the trivalent forms, are more toxic than the inorganic forms [151, 152, 154]. These methylated arsenic metabolites have now been suggested to be among the primary causative agents of arsenic-related health effects [157, 158].
Attempts to develop a mammalian model system to study the basis of carcinogenesis induced by methylated arsenicals have been made with only limited success [134]. Studies have
82 shown that rats exposed only to DMAV do not develop tumors, but when co-exposed to an
initiator and DMAV, the risk of developing tumors increases [208]. These results suggest that
arsenic is a tumor promoter, rather than an initiator. The use of rats as a model to study arsenic induced carcinogenesis has been hampered due to the fact that they can further methylate DMAIII
to trimethylarsine oxide (TMAO) [223], a compound not commonly found in human urine.
Other disadvantages of using rodent systems to study the carcinogenic potential of MAs are the
high cost to benefit ratio in maintenance of such mammals and the amount of samples that could
be studied at any given time point.
Studies performed by others and in our laboratory have suggested that Drosophila do not harbor an equivalent arsenic methylation to human cells, as previously observed in chimpanzees and some other primates [160]. Therefore, this presents an opportunity to use Drosophila with
its innate inability to methylate arsenic, to create a series of hAS3MT transgenic lines to study the
carcinogenic and/or toxic potential and of MAs and establish Drosophila as a viable model in
which to study the possible effects on and interactions with cellular pathways involved in arsenic
metabolism, DNA repair, and/or maintenance of the cellular redox state compared to a non-
methylation background.
Sequence analysis of the hAS3MT gene cloned here revealed two SNPs, one of which has
not been previously reported in the literature. Thus, there is no information on what effects the
T620C polymorphism might have on the function of the hAS3MT enzyme. The M287T hAS3MT polymorphic form of the enzyme has been described previously and it has been suggested that individuals harboring such a variant have a higher risk of developing arsenic-
related health effects [182] due to the higher methylation efficiency [185, 186] of the enzyme.
We performed site-directed mutagenesis on the identified SNPs in order to revert the sequence to
83 the allelic form found at the highest frequency in a studied human population [185]. Although
we have not yet created fly transgenics harboring these mutant hAS3MT alleles, we have their
cDNAs in the pENTR/D-TOPO vector and future studies using Drosophila transgenics harboring such a mutant will characterize its function and how it potentially correlates with arsenic toxicity.
The Gateway™ vectors used for expression of the hAS3MT enzyme in Drosophila allow
for a variety of epitope tags to be incorporated at the N- or C-terminus of the enzyme [230]. Our molecular and biochemical analyses of hAS3MT transgene insertion and mRNA transcription were performed on two transgenic lines (1099-2E and 1099-2F, see Figure 1), both expressing an
N-terminal HA tagged hAS3MT inserted at different locations on chromosome 2. Interestingly, the intensity of the red eyes (w+, marker for transgene insertion) correlates with the levels of
‘leaky’ RNA expression of the hAS3MT transgene, i.e. prior to GAL4 induction. Western
analysis using anti-HA antibodies shows that a protein of the expected size is translated in these
transgenic lines only when GAL4 is present. Thus, although its mRNA is expressed at low
levels in non-GAL4 adults, this may result from non-promoter based transcription of small RNA
species lacking the translation start site, since the protein is not translated at detectable levels
(Figure 3, lane 1, top and middle panels). This result was further confirmed by using enzyme-
specific antibodies raised against amino acids 341-360 of the hAS3MT enzyme [185].
The potential for enzymatic activity of the expressed hAS3MT enzyme was tested in the
1099-2E hAS3MT transgenic line crossed to the da-GAL4 inducer line by performing
HPLC/ICPMS analysis on larval homogenates exposed to 0.1 mM AsIII [336]. MMA is detected
at higher levels (~1.7-fold) in GAL4-induced hAS3MT expressing larval extracts than non-
induced extracts. DMA is only detected in transgenic larvae when the GAL4 activator is present.
84 Future experiments can help shed light on the biological effects of altering the DMA:MMA or
MAs:total arsenic ratios, which can be manipulated by altering the levels of folate in the diet fed
to larvae. GAL4 expression by itself does not lead to DMA generation as it is not detected in
non-transgenic larvae expressing the GAL4 activator alone. Nor is significant DMA seen in
transgenic hAS3MT larvae not expressing GAL4. Thus, we have created a genetically-
manipulable system that allows control of the expression of the hAS3MT enzyme in a variety of
tissue- or developmental-specific contexts, depending on the nature of the GAL4-driven
enhancer. Such flexibility in controlling the expression of the hAS3MT enzyme will allow for a
myriad of studies to be performed to help shed light on the mode of action of MAs.
We have shown that a variety of Drosophila lines unexposed or exposed to 0.1 mM AsIII and harboring (and expressing) the wild type form of the hAS3MT transgene are not compromised in either viability or developmental period, suggesting that insertion of the transgene did not affect normal physiology. Thus, production of DMA in vivo does not appear to cause any effects on the developmental fitness and/or ability to eclose of embryos exposed to 0.1 mM AsIII, suggesting that arsenic biotransformation in Drosophila is relatively benign at the
microscopic level. The concentration of AsIII used in these studies was near the upper threshold
in terms of observable effects on the ability to eclose in these lines. The absence of any obvious
phenotypic effects might also be attributed to efficient export and excretion of the toxic metal.
Future studies to investigate this process will be performed by creating flies harboring the UAS-
AS3MT transgene and mutations in or RNAi-inducible harpins for the MDR or MRP transporter
genes. Higher concentrations of AsIII have yet to be tested and it could be expected that if production of DMA is toxic to Drosophila, the effects on viability and development should be obvious between AS3MT non-expressing and expressing larvae exposed to AsIII. In addition, the
85 function of the AS3MT enzyme appears not be affected by the presence of the HA epitope. The
fact that we have shown that the enzyme is functional in Drosophila suggests that the enzyme is
correctly folded in fly cells.
Drosophila has frequently been used to study the genotoxic effects of known environmental chemicals by employing a SMART LOH technique [251, 334]. A newly developed SMART system identifies LOH in a gene homologous to the human large tumor suppressor 1 (LATS1) gene, called warts (wts) in Drosophila [269]. The wts protein is a serine/threonine kinase tumor suppressor [269] involved in the Hippo pathway that controls organ size by regulating cell growth, proliferation, and apoptosis [270-273] via response to cell- cell contact [274]. In the lats-SMART assay, if a mutation in the wild type allele of the wts gene occurs in a heterozygous wts+/- 3rd instar larva exposed to a genotoxin, large dramatic
overproliferated tumorigenic clones can form and be observed in the eclosed adult fly [269, 279,
280]. We have used the LOH lats-SMART assay in order to determine if MAs are genotoxic (as
in mammalian cells) by inducing the formation of tumorigenic outgrowths in hAS3MT transgenic
Drosophila. Adults harboring the lats gene have a low inherent frequency for tumor development, presumably due to spontaneous background mutation, but this frequency increased, albeit not-significantly, in the compound hAS3MT; lats flies, in the presence of AsIII.
We observed that the frequency of tumor formation increased significantly in adults exposed to
AsIII as well as adults harboring the lats gene, but not in flies harboring both the lats and the hAS3MT transgene suggesting that production of MAs is not genotoxic. However, the data obtained from the LOH lats-SMART analysis suffered from low statisitical power due to the low
numbers of emerging adults scored from any treatment and genotype. A solution to this pitfall is
to perform similar experiments scoring larger numbers of eclosing adults and using higher levels
86 of AsIII, which could potentially increase the frequency of tumor formation. Additionally, we
observed that the size of the tumor outgrowth increased as the time span after eclosion increased,
suggesting that in fact the outgrowths were made up of transformed mutant clones.
Therefore, this suggests that production of MAs has genotoxic consequences in
Drosophila, and that we are now in a position to exploit this system more fully using the range of genetic techniques and resources that are available in this higher eukaryotic model. For example, studies can now be envisaged where the effects of intersecting pathways on methylated arsenic and its toxicity are studied by crossing suitable transgenes into flies harboring mutations in genes
(or compromised for expression of) required for DNA repair, folate metabolism, GSH biosynthesis, arsenic cellular efflux and nutritional metabolism. Although we have not yet created fly transgenics harboring any polymorphic mutant hAS3MT alleles, we already have their
cDNAs cloned into the pENTR/D-TOPO vector and future studies using Drosophila harboring
such mutant(s) will characterize its function and how it relates to arsenic toxicity. Furthermore,
and of great interest to human arsenic susceptibility issues, studies on the potential relationship
between polymorphic gene alleles and susceptibility to arsenic exposure can be performed in
future endeavors using the basic transgenic model we have characterized herein.
87 Chapter IV.
Summary and Future Directions
Introduction
We have shown that strongly increased susceptibility to arsenic toxicity occurs as a result of reducing GS activity in vivo, since this profoundly affects the standard GSH supply and demand flux. Thus, we hypothesize that arsenic-exposed individuals harboring a GS polymorphic variant with reduced enzymatic activity might confer a substantially higer risk of developing arsenic-related health effects. Therefore, studies that would correlate GS polymorphisms with arsenic susceptibility in humans are warranted. In further studies we have established and characterized a Drosophila transgenic line capable of expressing the hAS3MT enzyme in order to study the effects of MAs in a non-methylation background and help shed light on the possible carcinogenic mechanisms induced by of chronic arsenic exposure and its relationship to genetic susceptibility. In the following discussion we propose various experiments to address some of these issues.
Population Toxicogenetics: Identification of GS Polymorphic Variants and Possible
Susceptibility to Arsenic Health Effects in Exposed Individuals Worldwide
Hereditary GS deficiency is an autosomal recessive disease characterized by hemolytic anemia, metabolic acidosis, 5-oxoprolinuria, central nervous system damage and/or recurrent bacterial infections [339]. Based on their clinical phenotypes, patients with these health effects attributed to GS deficiency have been divided into three groups. Mildly affected patients show hemolyitic anemia, moderately affected patients, along with hemolytic anemia show metabolic acidosis, and severely affected patients also show neurological defects and susceptibility to
88 bacterial infections [174]. A variety of polymorphisms causing missense, splice, deletion,
insertion and nonsense mutations have been associated with the disease [340, 341]. Single
nucleotide polymorphisms (SNPs) 656A G (D219G), 857T A (L286Q), and 988C T
(R339C) have been correlated with the mild form of GS deficiency [340, 342]. Mutations
causing frameshifts, premature stop codons or aberrant splicing have been correlated with the
moderate or severe phenotypes [341]. Many of the mutations in GS affect ligand binding or
catalysis: for example two SNPs, 373C T (R125C) and 799 C T (R267W) have been
correlated with reduced Vmax of 0.2% and 0.1% of wild type activity, respectively [343],
suggesting that these residues are important for active site catalysis. The R267W mutation was
observed in a newborn who died at 5 days of age [342]. A SNP, 1391G T (G464V) increases
the Km of the enzyme for glycine by approximately 100-fold [343] and has been correlated with the severe form of the disease [342].
Other polymorphisms in GS have been identified that do not show any obvious chemical
or pathological phenotypes but might render individuals susceptible to arsenic exposure via
drinking water. The international HapMap Consortium has characterized more than 3.1 million
human SNPs genotyped in 270 individuals from geographically diverse populations and includes
~25 to 35% of common SNP variation in the populations surveyed [344]. They have a identified
a total of 11 SNPs in the GS gene among 11 different individuals from various regions of the
world (available online at: http://www.hapmap.org) [344]. Furthermore, the Coriell Insititute for
Medical Research of the National Human Genome Research Institiute, offers the opportunity to
obtain cell lines donated from individuals who have participated in the HapMap project and these could be used to study the potential correlations of GS SNPs with susceptibility with environmental stressors such as arsenic. Preliminary experiments could be done on such cell
89 lines to test for susceptibility to arsenic exposure, including genotoxicity, cytotoxicity and
carcinogenic potential. Additionally, in order to determine if GS polymorphisms in individuals
chronically exposed to arsenic via drinking water are more susceptible to its toxic effects,
correlation studies could be performed in populations that have already been studied, such as the
ones in Bangladesh [345] and in the Taiwanese blackfoot disease-endemic area [338].
Cytological Analysis of AS3MT Transgenic Lines Exposed to Arsenic
Chromosomal aberrations such as hyperploidy, aneuploidy, sister chromatid exchanges
and other structural defects have been observed in arsenic-exposed cultured cells, for example
V79 Chinese hamster lung fibroblasts [346]. Human populations exposed to arsenic have also
shown increased chromosomal abnormalities, such as micronuclei formation, which are
extranuclear organelles separated from the main nucleus, generated by genotoxic exposition
during cellular division by a whole lagging chromosome or by acentric chromosome fragments
[347]. Other chromosomal aberrations observed in individuals showing signs of arsenic-related
skin conditions include chromatid breaks, gaps, and dicentrics [71]. Additionally and
importantly, recent studies have implicated MAs as responsible for the induction of such
chromosomal damage [154], but whether such abnormalities play a role in arsenic-induced
carcinogenesis is a topic of current debate [348, 349].
Future studies could be performed to observe chromosomal aberrations potentially
induced by arsenic exposure in neuroblasts isolated from the brain ganglions of 3rd instar
Drosophila larvae. The basis for this experimental approach stems from the fact that normal
Drosophila diploid neuroblasts undergo asymmetric division, which produces both another stem
cell and a daughter cell that can differentiate into other observable neural fates. Assymetric division is governed by polarity of the spindles at mitosis and, when disrupted, causes fate
90 destabilization of the daughter cells as cytoplasmic determinants are incorrectly distributed,
owing to a disruption in the cytokinetic plane. Techniques for neuroblast chromosome
preparations have been described and could easily be used for microscopic and
immunofluorescent visualization [350] to study the in vivo effects of AsIII in the hAS3MT
transgenic model at the cellular level. Studies should include karyotypic analysis using DAPI,
anti-H2Av (Drosophila equivalent of H2AX found at sites of double strand break repair) to
assess chromosomal breakage, anti-phosphorylated H3 antibody to identify mitotic
chromosomes, and anti-β tubulin or anti-γ tubulin to monitor centrosome number and
morphology, spindle morphology and polarity in mitotic cells.
Tissue Transplantation of Neuroblast Stem Cells from Arsenic-exposed AS3MT
Transgenic Lines
Studies have shown that arsenic consumption can lead to cancerous malignancies.
However, a mammalian model system to study the carcinogenic potential of arsenic has not been fully established. Because of a variety of advantages, Drosophila could be used as an alternative system to study the carcinogenic potential of arsenic. One technique that could be especially useful to study the carcinogenicity of MAs is the transplantation of neuroblasts from treated animals into the abdomen of adult hosts, allowing prolonged culture of arsenic-exposed cells in vivo [351]. Malignant cellular transformation has been commonly observed when mutations have been created in genes responsible for arranging the correct spindle orientation [352]. When transplanted into the abdomen of a wild type host adult, these incorrectly divided cells can give rise to malignant tumors that fill the abdominal cavity, metastasize to other tissues or organs, and cause lethality [351]. This experimental approach should be undertaken by transplanting neuroblast cells derived from hAS3MT-expressing and non-expressing larvae exposed or
91 unexposed to arsenic into wild type adult host abdomens, monitoring for malignant growth via
sectioning of adult abdomens and monitoring for tissue overgrowth in tumor masses by DAPI
staining and/or in situ hybridization to the hAS3MT gene.
Phenotypic Studies on Arsenic Exposure of hAS3MT Expression on Intersecting Biological
Pathways
The creation of a hAS3MT fly transgenic line shortens the gap between the mammalian
and fruit fly systems in relation to the metabolic pathway of arsenic. Therefore, to shed more
light on the GSH biosynthetic pathway, the toxic effects of methylated metabolites of arsenic can
be further studied in the RNAi-induced GS deficient lines. We expect that if MAs are more toxic
than inorganic arsenicals, such fly lines should be more susceptible than the non-methylation GS
deficient line. Such studies would approximate the use of Drosophila to what would be observed
in populations chronically exposed to arsenic.
Additionally, a better understanding of mechanisms involved in the genotoxic mode of
action of arsenic could potentially be obtained by crossing the hAS3MT transgenic line into
various heterozygous mutant genetic backgrounds for the DNA repair and cell cycle checkpoint pathways, such as mei-41 (ATR homolog), grp (chk1 homolog), okra (rad54 homolog), dBlm
(Bloom’s helicase), and dOgg (oxidative DNA damage repair). A role in arsenic toxicity for
enzymes suggested to be involved in the metabolic pathway of arsenic, such as GSTO, the
MDR/MRP and AQP transporters, could also be targeted in the fly via RNAi [229] and
combined into flies harboring mutations in other genetic backgrounds.
92 Intersection of Folate Metabolism and Arsenic Methylation
Deficiencies in the enzymes involved in the folate one-carbon metabolic pathway, which
produces the methyl donor S-adenosylmethionine for methylation of inorganic arsenic, have been
implicated in a methylation phenotype of arsenic-exposed individuals (reviewed in [176]).
Folate deficiency has been shown to be a risk factor for arsenic-induced skin tumorigenesis by
exacerbating the altered expression of epidermal cell growth/proliferation genes in K6/ODC
mice [353]. During oncogenic transformation, human prostate epithelial cells acquire adaptive
arsenic efflux by altering the levels of substrates and enzymes involved in the folate and transulfuration pathways and increasing the expression of the efflux protein, ATP binding cassette protein C1 [120]. A search for genes involved in the folate one-carbon and transulfuration metabolic pathways in the Drosophila genome shows a high degree of conservation (Figure 4.1). Therefore, studies on the toxicity of MAs could be performed in backgrounds mutant for the CG7560 (methylene-tetrahydrofolate reductase, MTHFR), Dhfr
(dihydrofolate reductase, DHFR), CG3011 (serine hydroxyl methyltransferase, SHMT), CG2674
(methionine adenosyltransferase, MAT), CG14882 (methionine synthase, MTR), Ahcy13
(adenosylhomocysteinase, AHCY), CG1753 (cystathionine β-synthase, CSB), and Eip55E
(cystathionase, CTH) genes. Alternatively if lines harboring mutations in these genes cannot be identified, lines harboring GAL4-inducible dsRNA hairpins targeting specific transcripts could be obtained from the Vienna Drosophila RNAi Center (Vienna, Austria) [229] (or other sources) and crossed into the hAS3MT background. Observations of a higher incidence of tumor formation via the lats/wts SMART assay in any of these backgrounds would suggest a strong link between arsenic toxicity and a specific biological pathway. Larvae from these mutant strains and wild type strains could be exposed to folate-supplemented or folate-deficient food to
93 study the potential correlations of dietary intake and the toxic effects of MAs. Additionally, the folate concentrations to which larvae would be exposed could be altered in other experiments that would include arsenic speciation studies to identify potential dose-response relationships between levels of folate versus levels of MAs.
Figure 4.1. Metabolic pathway of folate. Folate metabolism leads to synthesis of SAM, the substrate for hAS3MT. Enzymes involved in the metabolism of folate and the transulfuration pathway are in bold and the Drosophila genes that express the homologous enzymes are italicized and in parenthesis.
Studies on the AS3MT M287T Variant
Previous studies have shown that the M287T genetic variant has increased activity compared to wild type leading to higher methylation efficiency of inorganic arsenic [185, 186].
The MAs MMA and DMA in both the pentavalent and trivalent forms have been shown to be
94 more toxic than their inorganic counterparts. Higher methylation efficiencies, which are based
on higher levels of excreted MMA in the urine, could render individuals highly susceptible to
arsenic-related health effects due to the high levels of MAs synthesized which could overwhelm the cell or organism. We have cloned the M287T variant from a human kidney cDNA library into the pENTR/D-TOPO vector, but have yet not incorporated it into the Drosophila specific
Gateway constructs [230]. After recombination, embryo microinjection, and establishment of an
AS3MT M287T transgenic line(s), gene expression studies (mRNA, and Western analyses), and
protein function analysis (arsenic speciation), and studies such as cytological analysis, tissue
transplantation of neuroblasts, and effects on intersecting pathways will be performed and
compared to wild type.
95 References
1. Bentley, R. and Chasteen, T.G. (2002) Arsenic Curiosa and Humanity. Chem. Educator. 7: 51-60. 2. Maclagan, R.C. (1864) On the arsenic eaters of Styria. Edinburgh Med. J. 10: 200-207. 3. Przygoda, G., Feldmann, J., and Cullen, W.R. (2001) The arsenic eaters of Styria: a different picture of people who were chronically exposed to arsenic. Appl. Organometal. Chem. 15: 457-462. 4. Ratnaike, R.N. (2003) Acute and chronic arsenic toxicity. Postgrad Med J. 79: 391-6. 5. Shen, Z.X., Chen, G.Q., Ni, J.H., Li, X.S., Xiong, S.M., Qiu, Q.Y., Zhu, J., Tang, W., Sun, G.L., Yang, K.Q., Chen, Y., Zhou, L., Fang, Z.W., Wang, Y.T., Ma, J., Zhang, P., Zhang, T.D., Chen, S.J., Chen, Z., and Wang, Z.Y. (1997) Use of arsenic trioxide (As2O3) in the treatment of acute promyelocytic leukemia (APL): II. Clinical efficacy and pharmacokinetics in relapsed patients. Blood. 89: 3354-60. 6. Nordstrom, D.K. (2002) Public health. Worldwide occurrences of arsenic in ground water. Science. 296: 2143-5. 7. Rahman, M., Vahter, M., Wahed, M.A., Sohel, N., Yunus, M., Streatfield, P.K., El Arifeen, S., Bhuiya, A., Zaman, K., Chowdhury, A.M., Ekstrom, E.C., and Persson, L.A. (2006) Prevalence of arsenic exposure and skin lesions. A population based survey in Matlab, Bangladesh. J Epidemiol Community Health. 60: 242-8. 8. Alam, M.G., Allinson, G., Stagnitti, F., Tanaka, A., and Westbrooke, M. (2002) Arsenic contamination in Bangladesh groundwater: a major environmental and social disaster. Int J Environ Health Res. 12: 235-53. 9. Bhattacharya, P., Jacks, G., Ahmed, K.M., Routh, J., and Khan, A.A. (2002) Arsenic in groundwater of the Bengal delta plain aquifers in Bangladesh. Bull Environ Contam Toxicol. 69: 538-45. 10. Guha Mazumder, D.N., Chakraborty, A.K., Ghose, A., Gupta, J.D., Chakraborty, D.P., Dey, S.B., and Chattopadhyay, N. (1988) Chronic arsenic toxicity from drinking tubewell water in rural West Bengal. Bull World Health Organ. 66: 499-506. 11. Cebrian, M.E., Albores, A., Aguilar, M., and Blakely, E. (1983) Chronic arsenic poisoning in the north of Mexico. Hum Toxicol. 2: 121-33. 12. Meza, M.M., Kopplin, M.J., Burgess, J.L., and Gandolfi, A.J. (2004) Arsenic drinking water exposure and urinary excretion among adults in the Yaqui Valley, Sonora, Mexico. Environ Res. 96: 119-26. 13. Hopenhayn-Rich, C., Biggs, M.L., and Smith, A.H. (1998) Lung and kidney cancer mortality associated with arsenic in drinking water in Cordoba, Argentina. Int J Epidemiol. 27: 561-9. 14. Steinmaus, C., Bates, M.N., Yuan, Y., Kalman, D., Atallah, R., Rey, O.A., Biggs, M.L., Hopenhayn, C., Moore, L.E., Hoang, B.K., and Smith, A.H. (2006) Arsenic methylation and bladder cancer risk in case-control studies in Argentina and the United States. J. Occup. Environ. Med. 48: 478-88. 15. Concha, G., Nermell, B., and Vahter, M.V. (1998) Metabolism of inorganic arsenic in children with chronic high arsenic exposure in northern Argentina. Environ Health Perspect. 106: 355-9.
96 16. Hopenhayn-Rich, C., Browning, S.R., Hertz-Picciotto, I., Ferreccio, C., Peralta, C., and Gibb, H. (2000) Chronic arsenic exposure and risk of infant mortality in two areas of Chile. Environ Health Perspect. 108: 667-73. 17. Marshall, G., Ferreccio, C., Yuan, Y., Bates, M.N., Steinmaus, C., Selvin, S., Liaw, J., and Smith, A.H. (2007) Fifty-year study of lung and bladder cancer mortality in Chile related to arsenic in drinking water. J Natl Cancer Inst. 99: 920-8. 18. Ferreccio, C. and Sancha, A.M. (2006) Arsenic exposure and its impact on health in Chile. J Health Popul Nutr. 24: 164-75. 19. Chen, C.J., Chuang, Y.C., Lin, T.M., and Wu, H.Y. (1985) Malignant neoplasms among residents of a blackfoot disease-endemic area in Taiwan: high-arsenic artesian well water and cancers. Cancer Res. 45: 5895-9. 20. Meliker, J.R., Wahl, R.L., Cameron, L.L., and Nriagu, J.O. (2007) Arsenic in drinking water and cerebrovascular disease, diabetes mellitus, and kidney disease in Michigan: a standardized mortality ratio analysis. Environ Health. 6: 4. 21. Knobeloch, L.M., Zierold, K.M., and Anderson, H.A. (2006) Association of arsenic- contaminated drinking-water with prevalence of skin cancer in Wisconsin's Fox River Valley. J Health Popul Nutr. 24: 206-13. 22. Lewis, D.R., Southwick, J.W., Ouellet-Hellstrom, R., Rench, J., and Calderon, R.L. (1999) Drinking water arsenic in Utah: A cohort mortality study. Environ Health Perspect. 107: 359-65. 23. Burgess, J.L., Meza, M.M., Josyula, A.B., Poplin, G.S., Kopplin, M.J., McClellen, H.E., Sturup, S., and Lantz, R.C. (2007) Environmental Arsenic Exposure and Urinary 8- OHdG in Arizona and Sonora. Clin Toxicol (Phila). 45: 490-8. 24. Moore, L.E., Lu, M., and Smith, A.H. (2002) Childhood cancer incidence and arsenic exposure in drinking water in Nevada. Arch Environ Health. 57: 201-6. 25. British Geological Survey, Groundwater studies for arsenic contamination in Bangladesh - summary of Phase I report. 1999, British Geological Survey. 26. Chowdhury, A.M. (2004) Arsenic crisis in Bangladesh. Sci Am. 291: 86-91. 27. Harvey, C.F., Swartz, C.H., Badruzzaman, A.B., Keon-Blute, N., Yu, W., Ali, M.A., Jay, J., Beckie, R., Niedan, V., Brabander, D., Oates, P.M., Ashfaque, K.N., Islam, S., Hemond, H.F., and Ahmed, M.F. (2002) Arsenic mobility and groundwater extraction in Bangladesh. Science. 298: 1602-6. 28. Polizzotto, M.L., Harvey, C.F., Sutton, S.R., and Fendorf, S. (2005) Processes conducive to the release and transport of arsenic into aquifers of Bangladesh. Proc Natl Acad Sci U S A. 102: 18819-23. 29. Polizzotto, M.L., Kocar, B.D., Benner, S.G., Sampson, M., and Fendorf, S. (2008) Near- surface wetland sediments as a source of arsenic release to ground water in Asia. Nature. 454: 505-8. 30. Yoshida, T., Yamauchi, H., and Fan Sun, G. (2004) Chronic health effects in people exposed to arsenic via the drinking water: dose-response relationships in review. Toxicol Appl Pharmacol. 198: 243-52. 31. Chen, C.J., Chen, C.W., Wu, M.M., and Kuo, T.L. (1992) Cancer potential in liver, lung, bladder and kidney due to ingested inorganic arsenic in drinking water. Br J Cancer. 66: 888-92.
97 32. Tseng, W.P., Chu, H.M., How, S.W., Fong, J.M., Lin, C.S., and Yeh, S. (1968) Prevalence of skin cancer in an endemic area of chronic arsenicism in Taiwan. J Natl Cancer Inst. 40: 453-63. 33. Lai, M.S., Hsueh, Y.M., Chen, C.J., Shyu, M.P., Chen, S.Y., Kuo, T.L., Wu, M.M., and Tai, T.Y. (1994) Ingested inorganic arsenic and prevalence of diabetes mellitus. Am J Epidemiol. 139: 484-92. 34. Mazumder, D.N., Haque, R., Ghosh, N., De, B.K., Santra, A., Chakraborti, D., and Smith, A.H. (2000) Arsenic in drinking water and the prevalence of respiratory effects in West Bengal, India. Int J Epidemiol. 29: 1047-52. 35. Mukherjee, S.C., Rahman, M.M., Chowdhury, U.K., Sengupta, M.K., Lodh, D., Chanda, C.R., Saha, K.C., and Chakraborti, D. (2003) Neuropathy in arsenic toxicity from groundwater arsenic contamination in West Bengal, India. J Environ Sci Health A Tox Hazard Subst Environ Eng. 38: 165-83. 36. Navas-Acien, A., Silbergeld, E.K., Pastor-Barriuso, R., and Guallar, E. (2008) Arsenic exposure and prevalence of type 2 diabetes in US adults. JAMA. 300: 814-22. 37. Levin-Scherz, J.K., Patrick, J.D., Weber, F.H., and Garabedian, C., Jr. (1987) Acute arsenic ingestion. Ann Emerg Med. 16: 702-4. 38. World Health Organization, Guidelines for drinking-water quality: incorporating first addendum. Vol. 1, Recommendations. Third ed. 2006, Geneva: World Health Organization. 39. Environmental Protection Agency, U.S., 66 FR 6975 (2001) National primary drinking water regulations; arsenic and clarifications to compliance and new source contaminants monitoring; Final Rule. Federal Register. 66: 6975-7066. 40. Ahmad, S., Kitchin, K.T., and Cullen, W.R. (2000) Arsenic species that cause release of iron from ferritin and generation of activated oxygen. Arch Biochem Biophys. 382: 195- 202. 41. Imlay, J.A., Chin, S.M., and Linn, S. (1988) Toxic DNA damage by hydrogen peroxide through the Fenton reaction in vivo and in vitro. Science. 240: 640-2. 42. Del Razo, L.M., Quintanilla-Vega, B., Brambila-Colombres, E., Calderon-Aranda, E.S., Manno, M., and Albores, A. (2001) Stress proteins induced by arsenic. Toxicol Appl Pharmacol. 177: 132-48. 43. Shi, H., Shi, X., and Liu, K.J. (2004) Oxidative mechanism of arsenic toxicity and carcinogenesis. Mol Cell Biochem. 255: 67-78. 44. Pi, J., Qu, W., Reece, J.M., Kumagai, Y., and Waalkes, M.P. (2003) Transcription factor Nrf2 activation by inorganic arsenic in cultured keratinocytes: involvement of hydrogen peroxide. Exp Cell Res. 290: 234-45. 45. Nguyen, T., Sherratt, P.J., and Pickett, C.B. (2003) Regulatory mechanisms controlling gene expression mediated by the antioxidant response element. Annu Rev Pharmacol Toxicol. 43: 233-60. 46. Aposhian, H.V., Zakharyan, R.A., Avram, M.D., Kopplin, M.J., and Wollenberg, M.L. (2003) Oxidation and detoxification of trivalent arsenic species. Toxicol Appl Pharmacol. 193: 1-8. 47. Smith, K.R., Klei, L.R., and Barchowsky, A. (2001) Arsenite stimulates plasma membrane NADPH oxidase in vascular endothelial cells. Am J Physiol Lung Cell Mol Physiol. 280: L442-9.
98 48. Lynn, S., Gurr, J.R., Lai, H.T., and Jan, K.Y. (2000) NADH oxidase activation is involved in arsenite-induced oxidative DNA damage in human vascular smooth muscle cells. Circ Res. 86: 514-9. 49. Chou, W.C., Jie, C., Kenedy, A.A., Jones, R.J., Trush, M.A., and Dang, C.V. (2004) Role of NADPH oxidase in arsenic-induced reactive oxygen species formation and cytotoxicity in myeloid leukemia cells. Proc Natl Acad Sci U S A. 101: 4578-83. 50. Shen, Z.Y., Shen, W.Y., Chen, M.H., Shen, J., and Zeng, Y. (2003) Reactive oxygen species and antioxidants in apoptosis of esophageal cancer cells induced by As2O3. Int J Mol Med. 11: 479-84. 51. Iwama, K., Nakajo, S., Aiuchi, T., and Nakaya, K. (2001) Apoptosis induced by arsenic trioxide in leukemia U937 cells is dependent on activation of p38, inactivation of ERK and the Ca2+-dependent production of superoxide. Int J Cancer. 92: 518-26. 52. Jing, Y., Dai, J., Chalmers-Redman, R.M., Tatton, W.G., and Waxman, S. (1999) Arsenic trioxide selectively induces acute promyelocytic leukemia cell apoptosis via a hydrogen peroxide-dependent pathway. Blood. 94: 2102-11. 53. Yi, J., Gao, F., Shi, G., Li, H., Wang, Z., Shi, X., and Tang, X. (2002) The inherent cellular level of reactive oxygen species: one of the mechanisms determining apoptotic susceptibility of leukemic cells to arsenic trioxide. Apoptosis. 7: 209-15. 54. Zhu, J., Chen, Z., Lallemand-Breitenbach, V., and de The, H. (2002) How acute promyelocytic leukaemia revived arsenic. Nat Rev Cancer. 2: 705-13. 55. Jiang, X.H., Wong, B.C., Yuen, S.T., Jiang, S.H., Cho, C.H., Lai, K.C., Lin, M.C., Kung, H.F., and Lam, S.K. (2001) Arsenic trioxide induces apoptosis in human gastric cancer cells through up-regulation of p53 and activation of caspase-3. Int J Cancer. 91: 173-9. 56. Kito, M., Akao, Y., Ohishi, N., Yagi, K., and Nozawa, Y. (2002) Arsenic trioxide- induced apoptosis and its enhancement by buthionine sulfoximine in hepatocellular carcinoma cell lines. Biochem Biophys Res Commun. 291: 861-7. 57. Kang, Y.H., Yi, M.J., Kim, M.J., Park, M.T., Bae, S., Kang, C.M., Cho, C.K., Park, I.C., Park, M.J., Rhee, C.H., Hong, S.I., Chung, H.Y., Lee, Y.S., and Lee, S.J. (2004) Caspase- independent cell death by arsenic trioxide in human cervical cancer cells: reactive oxygen species-mediated poly(ADP-ribose) polymerase-1 activation signals apoptosis-inducing factor release from mitochondria. Cancer Res. 64: 8960-7. 58. Nakagawa, Y., Akao, Y., Morikawa, H., Hirata, I., Katsu, K., Naoe, T., Ohishi, N., and Yagi, K. (2002) Arsenic trioxide-induced apoptosis through oxidative stress in cells of colon cancer cell lines. Life Sci. 70: 2253-69. 59. Woo, S.H., Park, I.C., Park, M.J., Lee, H.C., Lee, S.J., Chun, Y.J., Lee, S.H., Hong, S.I., and Rhee, C.H. (2002) Arsenic trioxide induces apoptosis through a reactive oxygen species-dependent pathway and loss of mitochondrial membrane potential in HeLa cells. Int J Oncol. 21: 57-63. 60. McCafferty-Grad, J., Bahlis, N.J., Krett, N., Aguilar, T.M., Reis, I., Lee, K.P., and Boise, L.H. (2003) Arsenic trioxide uses caspase-dependent and caspase-independent death pathways in myeloma cells. Mol Cancer Ther. 2: 1155-64. 61. Corsini, E., Asti, L., Viviani, B., Marinovich, M., and Galli, C.L. (1999) Sodium arsenate induces overproduction of interleukin-1alpha in murine keratinocytes: role of mitochondria. J Invest Dermatol. 113: 760-5. 62. Shao, W., Fanelli, M., Ferrara, F.F., Riccioni, R., Rosenauer, A., Davison, K., Lamph, W.W., Waxman, S., Pelicci, P.G., Lo Coco, F., Avvisati, G., Testa, U., Peschle, C.,
99 Gambacorti-Passerini, C., Nervi, C., and Miller, W.H., Jr. (1998) Arsenic trioxide as an inducer of apoptosis and loss of PML/RAR alpha protein in acute promyelocytic leukemia cells. J Natl Cancer Inst. 90: 124-33. 63. Chen, G.Q., Shi, X.G., Tang, W., Xiong, S.M., Zhu, J., Cai, X., Han, Z.G., Ni, J.H., Shi, G.Y., Jia, P.M., Liu, M.M., He, K.L., Niu, C., Ma, J., Zhang, P., Zhang, T.D., Paul, P., Naoe, T., Kitamura, K., Miller, W., Waxman, S., Wang, Z.Y., de The, H., Chen, S.J., and Chen, Z. (1997) Use of arsenic trioxide (As2O3) in the treatment of acute promyelocytic leukemia (APL): I. As2O3 exerts dose-dependent dual effects on APL cells. Blood. 89: 3345-53. 64. Zhu, J., Koken, M.H., Quignon, F., Chelbi-Alix, M.K., Degos, L., Wang, Z.Y., Chen, Z., and de The, H. (1997) Arsenic-induced PML targeting onto nuclear bodies: implications for the treatment of acute promyelocytic leukemia. Proc Natl Acad Sci U S A. 94: 3978- 83. 65. Meister, A. and Anderson, M.E. (1983) Glutathione. Annu Rev Biochem. 52: 711-60. 66. Leslie, E.M., Haimeur, A., and Waalkes, M.P. (2004) Arsenic transport by the human multidrug resistance protein 1 (MRP1/ABCC1). Evidence that a tri-glutathione conjugate is required. J. Biol. Chem. 279: 32700-8. 67. Kala, S.V., Kala, G., Prater, C.I., Sartorelli, A.C., and Lieberman, M.W. (2004) Formation and urinary excretion of arsenic triglutathione and methylarsenic diglutathione. Chem Res Toxicol. 17: 243-9. 68. Kobayashi, Y., Cui, X., and Hirano, S. (2005) Stability of arsenic metabolites, arsenic triglutathione [As(GS)3] and methylarsenic diglutathione [CH3As(GS)2], in rat bile. Toxicology. 211: 115-23. 69. Brown, J.L., Kitchin, K.T., and George, M. (1997) Dimethylarsinic acid treatment alters six different rat biochemical parameters: relevance to arsenic carcinogenesis. Teratog Carcinog Mutagen. 17: 71-84. 70. Ahmad, S., Anderson, W.L., and Kitchin, K.T. (1999) Dimethylarsinic acid effects on DNA damage and oxidative stress related biochemical parameters in B6C3F1 mice. Cancer Lett. 139: 129-35. 71. Mahata, J., Basu, A., Ghoshal, S., Sarkar, J.N., Roy, A.K., Poddar, G., Nandy, A.K., Banerjee, A., Ray, K., Natarajan, A.T., Nilsson, R., and Giri, A.K. (2003) Chromosomal aberrations and sister chromatid exchanges in individuals exposed to arsenic through drinking water in West Bengal, India. Mutat Res. 534: 133-43. 72. Bau, D.T., Wang, T.S., Chung, C.H., Wang, A.S., Wang, A.S., and Jan, K.Y. (2002) Oxidative DNA adducts and DNA-protein cross-links are the major DNA lesions induced by arsenite. Environ Health Perspect. 110 Suppl 5: 753-6. 73. Wang, T.S., Hsu, T.Y., Chung, C.H., Wang, A.S., Bau, D.T., and Jan, K.Y. (2001) Arsenite induces oxidative DNA adducts and DNA-protein cross-links in mammalian cells. Free Radic Biol Med. 31: 321-30. 74. Kinoshita, A., Wanibuchi, H., Wei, M., Yunoki, T., and Fukushima, S. (2007) Elevation of 8-hydroxydeoxyguanosine and cell proliferation via generation of oxidative stress by organic arsenicals contributes to their carcinogenicity in the rat liver and bladder. Toxicol Appl Pharmacol. 221: 295-305. 75. Matsui, M., Nishigori, C., Toyokuni, S., Takada, J., Akaboshi, M., Ishikawa, M., Imamura, S., and Miyachi, Y. (1999) The role of oxidative DNA damage in human
100 arsenic carcinogenesis: detection of 8-hydroxy-2'-deoxyguanosine in arsenic-related Bowen's disease. J Invest Dermatol. 113: 26-31. 76. Nesnow, S., Roop, B.C., Lambert, G., Kadiiska, M., Mason, R.P., Cullen, W.R., and Mass, M.J. (2002) DNA damage induced by methylated trivalent arsenicals is mediated by reactive oxygen species. Chem Res Toxicol. 15: 1627-34. 77. Liu, S.X., Athar, M., Lippai, I., Waldren, C., and Hei, T.K. (2001) Induction of oxyradicals by arsenic: implication for mechanism of genotoxicity. Proc Natl Acad Sci U S A. 98: 1643-8. 78. Kumagai, Y. and Sumi, D. (2007) Arsenic: signal transduction, transcription factor, and biotransformation involved in cellular response and toxicity. Annu. Rev. Pharmacol. Toxicol. 47: 243-62. 79. Chen, W., Martindale, J.L., Holbrook, N.J., and Liu, Y. (1998) Tumor promoter arsenite activates extracellular signal-regulated kinase through a signaling pathway mediated by epidermal growth factor receptor and Shc. Mol Cell Biol. 18: 5178-88. 80. Porter, A.C., Fanger, G.R., and Vaillancourt, R.R. (1999) Signal transduction pathways regulated by arsenate and arsenite. Oncogene. 18: 7794-802. 81. Liu, Y., Guyton, K.Z., Gorospe, M., Xu, Q., Lee, J.C., and Holbrook, N.J. (1996) Differential activation of ERK, JNK/SAPK and P38/CSBP/RK map kinase family members during the cellular response to arsenite. Free Radic Biol Med. 21: 771-81. 82. Qian, Y., Castranova, V., and Shi, X. (2003) New perspectives in arsenic-induced cell signal transduction. J Inorg Biochem. 96: 271-8. 83. Duyndam, M.C., Hulscher, S.T., van der Wall, E., Pinedo, H.M., and Boven, E. (2003) Evidence for a role of p38 kinase in hypoxia-inducible factor 1-independent induction of vascular endothelial growth factor expression by sodium arsenite. J Biol Chem. 278: 6885-95. 84. den Hertog, J., Groen, A., and van der Wijk, T. (2005) Redox regulation of protein- tyrosine phosphatases. Arch Biochem Biophys. 434: 11-5. 85. Romashkova, J.A. and Makarov, S.S. (1999) NF-kappaB is a target of AKT in anti- apoptotic PDGF signalling. Nature. 401: 86-90. 86. Souza, K., Maddock, D.A., Zhang, Q., Chen, J., Chiu, C., Mehta, S., and Wan, Y. (2001) Arsenite activation of P13K/AKT cell survival pathway is mediated by p38 in cultured human keratinocytes. Mol Med. 7: 767-72. 87. Karin, M. (1995) The regulation of AP-1 activity by mitogen-activated protein kinases. J Biol Chem. 270: 16483-6. 88. Cavigelli, M., Li, W.W., Lin, A., Su, B., Yoshioka, K., and Karin, M. (1996) The tumor promoter arsenite stimulates AP-1 activity by inhibiting a JNK phosphatase. Embo J. 15: 6269-79. 89. Li, J., Gorospe, M., Barnes, J., and Liu, Y. (2003) Tumor promoter arsenite stimulates histone H3 phosphoacetylation of proto-oncogenes c-fos and c-jun chromatin in human diploid fibroblasts. J Biol Chem. 278: 13183-91. 90. Chen, F., Castranova, V., and Shi, X. (2001) New insights into the role of nuclear factor- kappaB in cell growth regulation. Am J Pathol. 159: 387-97. 91. Mathas, S., Lietz, A., Janz, M., Hinz, M., Jundt, F., Scheidereit, C., Bommert, K., and Dorken, B. (2003) Inhibition of NF-kappaB essentially contributes to arsenic-induced apoptosis. Blood. 102: 1028-34.
101 92. Shumilla, J.A., Wetterhahn, K.E., and Barchowsky, A. (1998) Inhibition of NF-kappa B binding to DNA by chromium, cadmium, mercury, zinc, and arsenite in vitro: evidence of a thiol mechanism. Arch Biochem Biophys. 349: 356-62. 93. Kapahi, P., Takahashi, T., Natoli, G., Adams, S.R., Chen, Y., Tsien, R.Y., and Karin, M. (2000) Inhibition of NF-kappa B activation by arsenite through reaction with a critical cysteine in the activation loop of Ikappa B kinase. J Biol Chem. 275: 36062-6. 94. Roussel, R.R. and Barchowsky, A. (2000) Arsenic inhibits NF-kappaB-mediated gene transcription by blocking IkappaB kinase activity and IkappaBalpha phosphorylation and degradation. Arch Biochem Biophys. 377: 204-12. 95. Kitchin, K.T. and Wallace, K. (2008) The role of protein binding of trivalent arsenicals in arsenic carcinogenesis and toxicity. J Inorg Biochem. 102: 532-9. 96. Ngu, T.T. and Stillman, M.J. (2006) Arsenic binding to human metallothionein. J Am Chem Soc. 128: 12473-83. 97. Jiang, G., Gong, Z., Li, X.-F., Cullen, W.R., and Chris Le, X. (2003) Interaction of trivalent arsenicals with metallothionein. Chem Res Toxicol. 16: 873-880. 98. Kreppel, H., Bauman, J.W., Liu, J., McKim, J.M., Jr., and Klaassen, C.D. (1993) Induction of metallothionein by arsenicals in mice. Fundam Appl Toxicol. 20: 184-9. 99. Liu, J., Liu, Y., Goyer, R.A., Achanzar, W., and Waalkes, M.P. (2000) Metallothionein- I/II null mice are more sensitive than wild-type mice to the hepatotoxic and nephrotoxic effects of chronic oral or injected inorganic arsenicals. Toxicol Sci. 55: 460-7. 100. Simeonova, P.P., Wang, S., Hulderman, T., and Luster, M.I. (2002) c-Src-dependent activation of the epidermal growth factor receptor and mitogen-activated protein kinase pathway by arsenic. Role in carcinogenesis. J Biol Chem. 277: 2945-50. 101. Simeonova, P.P. and Luster, M.I. (2002) Arsenic carcinogenicity: relevance of c-Src activation. Mol Cell Biochem. 234-235: 277-82. 102. Yamanaka, K. and Okada, S. (1994) Induction of lung-specific DNA damage by metabolically methylated arsenics via the production of free radicals. Environ Health Perspect. 102 Suppl 3: 37-40. 103. Yamanaka, K., Kato, K., Mizoi, M., An, Y., Takabayashi, F., Nakano, M., Hoshino, M., and Okada, S. (2004) The role of active arsenic species produced by metabolic reduction of dimethylarsinic acid in genotoxicity and tumorigenesis. Toxicol Appl Pharmacol. 198: 385-93. 104. Hartwig, A., Groblinghoff, U.D., Beyersmann, D., Natarajan, A.T., Filon, R., and Mullenders, L.H. (1997) Interaction of arsenic(III) with nucleotide excision repair in UV- irradiated human fibroblasts. Carcinogenesis. 18: 399-405. 105. Schwerdtle, T., Walter, I., and Hartwig, A. (2003) 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: 1449-63. 106. Hartwig, A., Pelzer, A., Asmuss, M., and Burkle, A. (2003) Very low concentrations of arsenite suppress poly(ADP-ribosyl)ation in mammalian cells. Int J Cancer. 104: 1-6. 107. Yager, J.W. and Wiencke, J.K. (1997) Inhibition of poly(ADP-ribose) polymerase by arsenite. Mutat Res. 386: 345-51. 108. Hartwig, A., Asmuss, M., Ehleben, I., Herzer, U., Kostelac, D., Pelzer, A., Schwerdtle, T., and Burkle, A. (2002) Interference by toxic metal ions with DNA repair processes and cell cycle control: molecular mechanisms. Environ Health Perspect. 110 Suppl 5: 797-9.
102 109. Lynn, S., Lai, H.T., Gurr, J.R., and Jan, K.Y. (1997) Arsenite retards DNA break rejoining by inhibiting DNA ligation. Mutagenesis. 12: 353-8. 110. Hu, Y., Su, L., and Snow, E.T. (1998) Arsenic toxicity is enzyme specific and its affects on ligation are not caused by the direct inhibition of DNA repair enzymes. Mutat Res. 408: 203-18. 111. Li, J.H. and Rossman, T.G. (1989) Inhibition of DNA ligase activity by arsenite: a possible mechanism of its comutagenesis. Mol Toxicol. 2: 1-9. 112. Zhong, C.X. and Mass, M.J. (2001) Both hypomethylation and hypermethylation of DNA associated with arsenite exposure in cultures of human cells identified by methylation- sensitive arbitrarily-primed PCR. Toxicol Lett. 122: 223-34. 113. Zhou, X., Sun, H., Ellen, T.P., Chen, H., and Costa, M. (2008) Arsenite alters global histone H3 methylation. Carcinogenesis. 29: 1831-1836. 114. Jensen, T.J., Novak, P., Eblin, K.E., Gandolfi, A.J., and Futscher, B.W. (2008) Epigenetic remodeling during arsenical-induced malignant transformation. Carcinogenesis. 29: 1500-8. 115. Tachibana, M., Sugimoto, K., Fukushima, T., and Shinkai, Y. (2001) Set domain- containing protein, G9a, is a novel lysine-preferring mammalian histone methyltransferase with hyperactivity and specific selectivity to lysines 9 and 27 of histone H3. J Biol Chem. 276: 25309-17. 116. Krogan, N.J., Dover, J., Khorrami, S., Greenblatt, J.F., Schneider, J., Johnston, M., and Shilatifard, A. (2002) COMPASS, a histone H3 (Lysine 4) methyltransferase required for telomeric silencing of gene expression. J Biol Chem. 277: 10753-5. 117. Zhao, C.Q., Young, M.R., Diwan, B.A., Coogan, T.P., and Waalkes, M.P. (1997) Association of arsenic-induced malignant transformation with DNA hypomethylation and aberrant gene expression. Proc Natl Acad Sci U S A. 94: 10907-12. 118. Chen, H., Li, S., Liu, J., Diwan, B.A., Barrett, J.C., and Waalkes, M.P. (2004) Chronic inorganic arsenic exposure induces hepatic global and individual gene hypomethylation: implications for arsenic hepatocarcinogenesis. Carcinogenesis. 25: 1779-86. 119. Waalkes, M.P., Liu, J., Chen, H., Xie, Y., Achanzar, W.E., Zhou, Y.S., Cheng, M.L., and Diwan, B.A. (2004) Estrogen signaling in livers of male mice with hepatocellular carcinoma induced by exposure to arsenic in utero. J Natl Cancer Inst. 96: 466-74. 120. Coppin, J.F., Qu, W., and Waalkes, M.P. (2008) Interplay between cellular methyl metabolism and adaptive efflux during oncogenic transformation from chronic arsenic exposure in human cells. J Biol Chem. 283: 19342-50. 121. Mass, M.J. and Wang, L. (1997) Arsenic alters cytosine methylation patterns of the promoter of the tumor suppressor gene p53 in human lung cells: a model for a mechanism of carcinogenesis. Mutat Res. 386: 263-77. 122. Jones, P.L., Veenstra, G.J., Wade, P.A., Vermaak, D., Kass, S.U., Landsberger, N., Strouboulis, J., and Wolffe, A.P. (1998) Methylated DNA and MeCP2 recruit histone deacetylase to repress transcription. Nat Genet. 19: 187-91. 123. Nan, X., Ng, H.H., Johnson, C.A., Laherty, C.D., Turner, B.M., Eisenman, R.N., and Bird, A. (1998) Transcriptional repression by the methyl-CpG-binding protein MeCP2 involves a histone deacetylase complex. Nature. 393: 386-9. 124. Dixon, H.B.F., The biochemical action of arsonic acids especially as phosphate analogs., in Advances in Inorganic Chemistry, A.G. Sykes, Editor. 1997, Academic Press: Orlando. p. 191-227.
103 125. Liu, Z., Shen, J., Carbrey, J.M., Mukhopadhyay, R., Agre, P., and Rosen, B.P. (2002) Arsenite transport by mammalian aquaglyceroporins AQP7 and AQP9. Proc Natl Acad Sci U S A. 99: 6053-8. 126. Liu, Z., Carbrey, J.M., Agre, P., and Rosen, B.P. (2004) Arsenic trioxide uptake by human and rat aquaglyceroporins. Biochem Biophys Res Commun. 316: 1178-85. 127. Liu, Z., Sanchez, M.A., Jiang, X., Boles, E., Landfear, S.M., and Rosen, B.P. (2006) Mammalian glucose permease GLUT1 facilitates transport of arsenic trioxide and methylarsonous acid. Biochem Biophys Res Commun. 351: 424-30. 128. Villa-Bellosta, R. and Sorribas, V. (2008) Role of rat sodium/phosphate cotransporters in the cell membrane transport of arsenate. Toxicol Appl Pharmacol. 232: 125-34. 129. Kenney, L.J. and Kaplan, J.H. (1988) Arsenate substitutes for phosphate in the human red cell sodium pump and anion exchanger. J Biol Chem. 263: 7954-60. 130. Csanaky, I. and Gregus, Z. (2005) Role of glutathione in reduction of arsenate and of gamma-glutamyltranspeptidase in disposition of arsenite in rats. Toxicology. 207: 91- 104. 131. Cullen, W.R., McBride, B.C., and Reglinski, J. (1984) The reaction of methylarsenicals with thiols: some biological implications. J. Inorg. Biochem. 21: 179-193. 132. Hayakawa, T., Kobayashi, Y., Cui, X., and Hirano, S. (2005) A new metabolic pathway of arsenite: arsenic-glutathione complexes are substrates for human arsenic methyltransferase Cyt19. Arch. Toxicol. 79: 183-91. 133. Naranmandura, H., Suzuki, N., and Suzuki, K.T. (2006) Trivalent arsenicals are bound to proteins during reductive methylation. Chem Res Toxicol. 19: 1010-8. 134. Cohen, S.M., Arnold, L.L., Eldan, M., Lewis, A.S., and Beck, B.D. (2006) Methylated arsenicals: the implications of metabolism and carcinogenicity studies in rodents to human risk assessment. Crit Rev Toxicol. 36: 99-133. 135. Thomas, D.J. (2007) Molecular processes in cellular arsenic metabolism. Toxicol Appl Pharmacol. 222: 365-73. 136. Zakharyan, R.A., Sampayo-Reyes, A., Healy, S.M., Tsaprailis, G., Board, P.G., Liebler, D.C., and Aposhian, H.V. (2001) Human monomethylarsonic acid (MMA(V)) reductase is a member of the glutathione-S-transferase superfamily. Chem Res Toxicol. 14: 1051-7. 137. Zakharyan, R.A. and Aposhian, H.V. (1999) Enzymatic reduction of arsenic compounds in mammalian systems: the rate-limiting enzyme of rabbit liver arsenic biotransformation is MMA(V) reductase. Chem Res Toxicol. 12: 1278-83. 138. Chowdhury, U.K., Zakharyan, R.A., Hernandez, A., Avram, M.D., Kopplin, M.J., and Aposhian, H.V. (2006) Glutathione-S-transferase-omega [MMA(V) reductase] knockout mice: enzyme and arsenic species concentrations in tissues after arsenate administration. Toxicol Appl Pharmacol. 216: 446-57. 139. Lin, S., Shi, Q., Nix, F.B., Styblo, M., Beck, M.A., Herbin-Davis, K.M., Hall, L.L., Simeonsson, J.B., and Thomas, D.J. (2002) A novel S-adenosyl-L- methionine:arsenic(III) methyltransferase from rat liver cytosol. J. Biol. Chem. 277: 10795-803. 140. Thomas, D.J., Waters, S.B., and Styblo, M. (2004) Elucidating the pathway for arsenic methylation. Toxicol Appl Pharmacol. 198: 319-26. 141. Waters, S.B., Devesa, V., Del Razo, L.M., Styblo, M., and Thomas, D.J. (2004) Endogenous reductants support the catalytic function of recombinant rat cyt19, an arsenic methyltransferase. Chem Res Toxicol. 17: 404-9.
104 142. Drobna, Z., Xing, W., Thomas, D.J., and Styblo, M. (2006) shRNA silencing of AS3MT expression minimizes arsenic methylation capacity of HepG2 cells. Chem Res Toxicol. 19: 894-8. 143. Zakharyan, R.A., Ayala-Fierro, F., Cullen, W.R., Carter, D.M., and Aposhian, H.V. (1999) Enzymatic methylation of arsenic compounds. VII. Monomethylarsonous acid (MMAIII) is the substrate for MMA methyltransferase of rabbit liver and human hepatocytes. Toxicol Appl Pharmacol. 158: 9-15. 144. Gebel, T.W. (2002) Arsenic methylation is a process of detoxification through accelerated excretion. Int J Hyg Environ Health. 205: 505-8. 145. Buchet, J.P., Lauwerys, R., and Roels, H. (1981) Comparison of the urinary excretion of arsenic metabolites after a single oral dose of sodium arsenite, monomethylarsonate, or dimethylarsinate in man. Int Arch Occup Environ Health. 48: 71-9. 146. Hughes, M.F. and Kenyon, E.M. (1998) Dose-dependent effects on the disposition of monomethylarsonic acid and dimethylarsinic acid in the mouse after intravenous administration. J Toxicol Environ Health A. 53: 95-112. 147. Crecelius, E.A. (1977) Changes in the chemical speciation of arsenic following ingestion by man. Environ Health Perspect. 19: 147-50. 148. Le, X.C., Ma, M., Cullen, W.R., Aposhian, H.V., Lu, X., and Zheng, B. (2000) Determination of monomethylarsonous acid, a key arsenic methylation intermediate, in human urine. Environ Health Perspect. 108: 1015-8. 149. Le, X.C., Lu, X., Ma, M., Cullen, W.R., Aposhian, H.V., and Zheng, B. (2000) Speciation of key arsenic metabolic intermediates in human urine. Anal Chem. 72: 5172- 7. 150. Petrick, J.S., Jagadish, B., Mash, E.A., and Aposhian, H.V. (2001) Monomethylarsonous acid (MMA(III)) and arsenite: LD(50) in hamsters and in vitro inhibition of pyruvate dehydrogenase. Chem Res Toxicol. 14: 651-6. 151. Mass, M.J., Tennant, A., Roop, B.C., Cullen, W.R., Styblo, M., Thomas, D.J., and Kligerman, A.D. (2001) Methylated trivalent arsenic species are genotoxic. Chem Res Toxicol. 14: 355-61. 152. Petrick, J.S., Ayala-Fierro, F., Cullen, W.R., Carter, D.E., and Vasken Aposhian, H. (2000) Monomethylarsonous acid (MMA(III)) is more toxic than arsenite in Chang human hepatocytes. Toxicol Appl Pharmacol. 163: 203-7. 153. Styblo, M., Del Razo, L.M., Vega, L., Germolec, D.R., LeCluyse, E.L., Hamilton, G.A., Reed, W., Wang, C., Cullen, W.R., and Thomas, D.J. (2000) Comparative toxicity of trivalent and pentavalent inorganic and methylated arsenicals in rat and human cells. Arch Toxicol. 74: 289-99. 154. Kligerman, A.D., Doerr, C.L., Tennant, A.H., Harrington-Brock, K., Allen, J.W., Winkfield, E., Poorman-Allen, P., Kundu, B., Funasaka, K., Roop, B.C., Mass, M.J., and DeMarini, D.M. (2003) Methylated trivalent arsenicals as candidate ultimate genotoxic forms of arsenic: induction of chromosomal mutations but not gene mutations. Environ Mol Mutagen. 42: 192-205. 155. Hirano, S. and Kobayashi, Y. (2006) Cytotoxic effects of S-(dimethylarsino)-glutathione: a putative intermediate metabolite of inorganic arsenicals. Toxicology. 227: 45-52. 156. Yu, R.C., Hsu, K.H., Chen, C.J., and Froines, J.R. (2000) Arsenic methylation capacity and skin cancer. Cancer Epidemiol Biomarkers Prev. 9: 1259-62.
105 157. Chen, Y.C., Su, H.J., Guo, Y.L., Hsueh, Y.M., Smith, T.J., Ryan, L.M., Lee, M.S., and Christiani, D.C. (2003) Arsenic methylation and bladder cancer risk in Taiwan. Cancer Causes Control. 14: 303-10. 158. Chen, Y.C., Guo, Y.L., Su, H.J., Hsueh, Y.M., Smith, T.J., Ryan, L.M., Lee, M.S., Chao, S.C., Lee, J.Y., and Christiani, D.C. (2003) Arsenic methylation and skin cancer risk in southwestern Taiwan. J Occup Environ Med. 45: 241-8. 159. Lindberg, A.L., Rahman, M., Persson, L.A., and Vahter, M. (2008) The risk of arsenic induced skin lesions in Bangladeshi men and women is affected by arsenic metabolism and the age at first exposure. Toxicol Appl Pharmacol. 230: 9-16. 160. Vahter, M., Couch, R., Nermell, B., and Nilsson, R. (1995) Lack of methylation of inorganic arsenic in the chimpanzee. Toxicol Appl Pharmacol. 133: 262-8. 161. Wildfang, E., Radabaugh, T.R., and Vasken Aposhian, H. (2001) Enzymatic methylation of arsenic compounds. IX. Liver arsenite methyltransferase and arsenate reductase activities in primates. Toxicology. 168: 213-21. 162. Zakharyan, R.A., Wildfang, E., and Aposhian, H.V. (1996) Enzymatic methylation of arsenic compounds. III. The marmoset and tamarin, but not the rhesus, monkeys are deficient in methyltransferases that methylate inorganic arsenic. Toxicol Appl Pharmacol. 140: 77-84. 163. Wildfang, E., Zakharyan, R.A., and Aposhian, H.V. (1998) Enzymatic methylation of arsenic compounds. VI. Characterization of hamster liver arsenite and methylarsonic acid methyltransferase activities in vitro. Toxicol Appl Pharmacol. 152: 366-75. 164. Healy, S.M., Casarez, E.A., Ayala-Fierro, F., and Aposhian, H. (1998) Enzymatic methylation of arsenic compounds. V. Arsenite methyltransferase activity in tissues of mice. Toxicol Appl Pharmacol. 148: 65-70. 165. Styblo, M., Delnomdedieu, M., and Thomas, D.J. (1996) Mono- and dimethylation of arsenic in rat liver cytosol in vitro. Chem Biol Interact. 99: 147-64. 166. Healy, S.M., Zakharyan, R.A., and Aposhian, H.V. (1997) Enzymatic methylation of arsenic compounds: IV. In vitro and in vivo deficiency of the methylation of arsenite and monomethylarsonic acid in the guinea pig. Mutat Res. 386: 229-39. 167. Li, J., Waters, S.B., Drobna, Z., Devesa, V., Styblo, M., and Thomas, D.J. (2005) Arsenic (+3 oxidation state) methyltransferase and the inorganic arsenic methylation phenotype. Toxicol Appl Pharmacol. 204: 164-9. 168. Rappa, G., Lorico, A., Flavell, R.A., and Sartorelli, A.C. (1997) Evidence that the multidrug resistance protein (MRP) functions as a co-transporter of glutathione and natural product toxins. Cancer Res. 57: 5232-7. 169. Kala, S.V., Neely, M.W., Kala, G., Prater, C.I., Atwood, D.W., Rice, J.S., and Lieberman, M.W. (2000) The MRP2/cMOAT transporter and arsenic-glutathione complex formation are required for biliary excretion of arsenic. J Biol Chem. 275: 33404- 8. 170. Raml, R., Rumpler, A., Goessler, W., Vahter, M., Li, L., Ochi, T., and Francesconi, K.A. (2007) Thio-dimethylarsinate is a common metabolite in urine samples from arsenic- exposed women in Bangladesh. Toxicol Appl Pharmacol. 222: 374-80. 171. Naranmandura, H. and Suzuki, K.T. (2008) Formation of dimethylthioarsenicals in red blood cells. Toxicol Appl Pharmacol. 227: 390-9. 172. Suzuki, K.T., Iwata, K., Naranmandura, H., and Suzuki, N. (2007) Metabolic differences between two dimethylthioarsenicals in rats. Toxicol Appl Pharmacol. 218: 166-73.
106 173. Naranmandura, H., Suzuki, N., and Suzuki, K.T. (2008) Reaction mechanism underlying the in vitro transformation of thioarsenicals. Toxicol Appl Pharmacol. 231: 328-35. 174. Naranmandura, H., Ibata, K., and Suzuki, K.T. (2007) Toxicity of dimethylmonothioarsinic acid toward human epidermoid carcinoma A431 cells. Chem Res Toxicol. 20: 1120-5. 175. Ochi, T., Kita, K., Suzuki, T., Rumpler, A., Goessler, W., and Francesconi, K.A. (2008) Cytotoxic, genotoxic and cell-cycle disruptive effects of thio-dimethylarsinate in cultured human cells and the role of glutathione. Toxicol Appl Pharmacol. 228: 59-67. 176. Hernandez, A. and Marcos, R. (2008) Genetic variations associated with interindividual sensitivity in the response to arsenic exposure. Pharmacogenomics. 9: 1113-32. 177. Vahter, M., Concha, G., Nermell, B., Nilsson, R., Dulout, F., and Natarajan, A.T. (1995) A unique metabolism of inorganic arsenic in native Andean women. Eur. J. Pharmacol. 293: 455-62. 178. Huang, Y.K., Huang, Y.L., Hsueh, Y.M., Yang, M.H., Wu, M.M., Chen, S.Y., Hsu, L.I., and Chen, C.J. (2008) Arsenic exposure, urinary arsenic speciation, and the incidence of urothelial carcinoma: a twelve-year follow-up study. Cancer Causes Control. 19: 829-39. 179. Loffredo, C.A., Aposhian, H.V., Cebrian, M.E., Yamauchi, H., and Silbergeld, E.K. (2003) Variability in human metabolism of arsenic. Environ. Res. 92: 85-91. 180. Marnell, L.L., Garcia-Vargas, G.G., Chowdhury, U.K., Zakharyan, R.A., Walsh, B., Avram, M.D., Kopplin, M.J., Cebrian, M.E., Silbergeld, E.K., and Aposhian, H.V. (2003) Polymorphisms in the human monomethylarsonic acid (MMA V) reductase/hGSTO1 gene and changes in urinary arsenic profiles. Chem Res Toxicol. 16: 1507-13. 181. Meza, M.M., Yu, L., Rodriguez, Y.Y., Guild, M., Thompson, D., Gandolfi, A.J., and Klimecki, W.T. (2005) Developmentally restricted genetic determinants of human arsenic metabolism: association between urinary methylated arsenic and CYT19 polymorphisms in children. Environ Health Perspect. 113: 775-81. 182. Hernandez, A., Xamena, N., Surralles, J., Sekaran, C., Tokunaga, H., Quinteros, D., Creus, A., and Marcos, R. (2008) Role of the Met(287)Thr polymorphism in the AS3MT gene on the metabolic arsenic profile. Mutat. Res. 637: 80-92. 183. Engstrom, K.S., Nermell, B., Concha, G., Stromberg, U., Vahter, M., and Broberg, K. (2007) Arsenic metabolism is influenced by polymorphisms in genes involved in one- carbon metabolism and reduction reactions. Mutat Res/Fundamental and Molecular Mechanisms of Mutagenesis. doi:10.1016/j.mrfmmmm.2008.07.003. 184. Lindberg, A.L., Kumar, R., Goessler, W., Thirumaran, R., Gurzau, E., Koppova, K., Rudnai, P., Leonardi, G., Fletcher, T., and Vahter, M. (2007) Metabolism of low-dose inorganic arsenic in a central European population: influence of sex and genetic polymorphisms. Environ Health Perspect. 115: 1081-6. 185. Wood, T.C., Salavagionne, O.E., Mukherjee, B., Wang, L., Klumpp, A.F., Thomae, B.A., Eckloff, B.W., Schaid, D.J., Wieben, E.D., and Weinshilboum, R.M. (2006) Human arsenic methyltransferase (AS3MT) pharmacogenetics: gene resequencing and functional genomics studies. J Biol Chem. 281: 7364-73. 186. Hernandez, A., Xamena, N., Sekaran, C., Tokunaga, H., Sampayo-Reyes, A., Quinteros, D., Creus, A., and Marcos, R. (2008) High arsenic metabolic efficiency in AS3MT287Thr allele carriers. Pharmacogenet Genomics. 18: 349-55. 187. McCarty, K.M., Chen, Y.C., Quamruzzaman, Q., Rahman, M., Mahiuddin, G., Hsueh, Y.M., Su, L., Smith, T., Ryan, L., and Christiani, D.C. (2007) Arsenic methylation,
107 GSTT1, GSTM1, GSTP1 polymorphisms, and skin lesions. Environ Health Perspect. 115: 341-5. 188. Liu, J., Liu, Y., Powell, D.A., Waalkes, M.P., and Klaassen, C.D. (2002) Multidrug- resistance mdr1a/1b double knockout mice are more sensitive than wild type mice to acute arsenic toxicity, with higher arsenic accumulation in tissues. Toxicology. 170: 55- 62. 189. Ahsan, H., Chen, Y., Kibriya, M.G., Slavkovich, V., Parvez, F., Jasmine, F., Gamble, M.V., and Graziano, J.H. (2007) Arsenic metabolism, genetic susceptibility, and risk of premalignant skin lesions in Bangladesh. Cancer Epidemiol Biomarkers Prev. 16: 1270- 8. 190. Brouwer, O.F., Onkenhout, W., Edelbroek, P.M., de Kom, J.F., de Wolff, F.A., and Peters, A.C. (1992) Increased neurotoxicity of arsenic in methylenetetrahydrofolate reductase deficiency. Clin Neurol Neurosurg. 94: 307-10. 191. Engstrom, K.S., Nermell, B., Concha, G., Stromberg, U., Vahter, M., and Broberg, K. (2008) Arsenic metabolism is influenced by polymorphisms in genes involved in one- carbon metabolism and reduction reactions. Mutat Res.: Fundamental and Molecular Mechanisms of Mutagenesis. doi:10.1016/j.mrfmmmm.2008.07.003. 192. Chiuve, S.E., Giovannucci, E.L., Hankinson, S.E., Zeisel, S.H., Dougherty, L.W., Willett, W.C., and Rimm, E.B. (2007) The association between betaine and choline intakes and the plasma concentrations of homocysteine in women. Am J Clin Nutr. 86: 1073-81. 193. Engstrom, K.S., Broberg, K., Concha, G., Nermell, B., Warholm, M., and Vahter, M. (2007) Genetic polymorphisms influencing arsenic metabolism: evidence from Argentina. Environ Health Perspect. 115: 599-605. 194. De Chaudhuri, S., Mahata, J., Das, J.K., Mukherjee, A., Ghosh, P., Sau, T.J., Mondal, L., Basu, S., Giri, A.K., and Roychoudhury, S. (2006) Association of specific p53 polymorphisms with keratosis in individuals exposed to arsenic through drinking water in West Bengal, India. Mutat Res. 601: 102-12. 195. Hsueh, Y.M., Lin, P., Chen, H.W., Shiue, H.S., Chung, C.J., Tsai, C.T., Huang, Y.K., Chiou, H.Y., and Chen, C.J. (2005) Genetic polymorphisms of oxidative and antioxidant enzymes and arsenic-related hypertension. J Toxicol Environ Health A. 68: 1471-84. 196. Ahsan, H., Chen, Y., Kibriya, M.G., Islam, M.N., Slavkovich, V.N., Graziano, J.H., and Santella, R.M. (2003) Susceptibility to arsenic-induced hyperkeratosis and oxidative stress genes myeloperoxidase and catalase. Cancer. Lett. 201: 57-65. 197. Banerjee, M., Sarkar, J., Das, J.K., Mukherjee, A., Sarkar, A.K., Mondal, L., and Giri, A.K. (2006) Polymorphism in the ERCC2 codon 751 is associated with arsenic-induced premalignant hyperkeratosis and significant chromosome aberrations. Carcinogenesis. 198. Hsu, L.I., Chiu, A.W., Huan, S.K., Chen, C.L., Wang, Y.H., Hsieh, F.I., Chou, W.L., Wang, L.H., and Chen, C.J. (2008) SNPs of GSTM1, T1, P1, epoxide hydrolase and DNA repair enzyme XRCC1 and risk of urinary transitional cell carcinoma in southwestern Taiwan. Toxicol Appl Pharmacol. 228: 144-55. 199. Mei, N., Lee, J., Sun, X., Xing, J.Z., Hanson, J., Le, X.C., and Weinfeld, M. (2003) Genetic predisposition to the cytotoxicity of arsenic: the role of DNA damage and ATM. Faseb J. 17: 2310-2. 200. IARC, Monographs on the evaluation of carcinogenic risks to humans: Some drinking- water contaminants and disinfectants, including arsenic. Vol. 84. 2004, Lyon, France: International Agency for Research on Cancer.
108 201. Kitchin, K.T. (2001) Recent advances in arsenic carcinogenesis: modes of action, animal model systems, and methylated arsenic metabolites. Toxicol Appl Pharmacol. 172: 249- 61. 202. Vahter, M. (2002) Mechanisms of arsenic biotransformation. Toxicology. 181-182: 211- 7. 203. Thorgeirsson, U.P., Dalgard, D.W., Reeves, J., and Adamson, R.H. (1994) Tumor incidence in a chemical carcinogenesis study of nonhuman primates. Regul Toxicol Pharmacol. 19: 130-51. 204. Wanibuchi, H., Salim, E.I., Kinoshita, A., Shen, J., Wei, M., Morimura, K., Yoshida, K., Kuroda, K., Endo, G., and Fukushima, S. (2004) Understanding arsenic carcinogenicity by the use of animal models. Toxicol Appl Pharmacol. 198: 366-76. 205. Ishinishi, N., Yamamoto, A., Hisanaga, A., and Inamasu, T. (1983) Tumorigenicity of arsenic trioxide to the lung in Syrian golden hamsters by intermittent instillations. Cancer Lett. 21: 141-7. 206. Takahashi, M., Barrett, J.C., and Tsutsui, T. (2002) Transformation by inorganic arsenic compounds of normal Syrian hamster embryo cells into a neoplastic state in which they become anchorage-independent and cause tumors in newborn hamsters. Int J Cancer. 99: 629-34. 207. Shirachi, D.Y., Johansen, M.G., McGowan, J.P., and Tu, S.H. (1983) Tumorigenic effect of sodium arsenite in rat kidney. Proc West Pharmacol Soc. 26: 413-5. 208. Johansen, M.G., McGowan, J.P., and Tu, S.H. (1984) Tumorigenic effect of dimethylarsinic acid in the rat. Proc West Pharmacol Soc. 27: 289-291. 209. Wanibuchi, H., Yamamoto, S., Chen, H., Yoshida, K., Endo, G., Hori, T., and Fukushima, S. (1996) Promoting effects of dimethylarsinic acid on N-butyl-N-(4- hydroxybutyl)nitrosamine-induced urinary bladder carcinogenesis in rats. Carcinogenesis. 17: 2435-9. 210. Li, W., Wanibuchi, H., Salim, E.I., Yamamoto, S., Yoshida, K., Endo, G., and Fukushima, S. (1998) Promotion of NCI-Black-Reiter male rat bladder carcinogenesis by dimethylarsinic acid an organic arsenic compound. Cancer Lett. 134: 29-36. 211. Yamamoto, S., Konishi, Y., Matsuda, T., Murai, T., Shibata, M.A., Matsui-Yuasa, I., Otani, S., Kuroda, K., Endo, G., and Fukushima, S. (1995) Cancer induction by an organic arsenic compound, dimethylarsinic acid (cacodylic acid), in F344/DuCrj rats after pretreatment with five carcinogens. Cancer Res. 55: 1271-6. 212. Wei, M., Wanibuchi, H., Yamamoto, S., Li, W., and Fukushima, S. (1999) Urinary bladder carcinogenicity of dimethylarsinic acid in male F344 rats. Carcinogenesis. 20: 1873-6. 213. Seike, N., Wanibuchi, H., Morimura, K., Nishikawa, T., Kishida, H., Nakae, D., Hirata, K., and Fukushima, S. (2002) Lack of promoting effect due to oral administration of dimethylarsinic acid on rat lung carcinogenesis initiated with N-bis(2- hydroxypropyl)nitrosamine. Cancer Lett. 175: 113-9. 214. Nishikawa, T., Wanibuchi, H., Ogawa, M., Kinoshita, A., Morimura, K., Hiroi, T., Funae, Y., Kishida, H., Nakae, D., and Fukushima, S. (2002) Promoting effects of monomethylarsonic acid, dimethylarsinic acid and trimethylarsine oxide on induction of rat liver preneoplastic glutathione S-transferase placental form positive foci: a possible reactive oxygen species mechanism. Int J Cancer. 100: 136-9.
109 215. Shen, J., Wanibuchi, H., Salim, E.I., Wei, M., Kinoshita, A., Yoshida, K., Endo, G., and Fukushima, S. (2003) Liver tumorigenicity of trimethylarsine oxide in male Fischer 344 rats--association with oxidative DNA damage and enhanced cell proliferation. Carcinogenesis. 24: 1827-35. 216. Burns, F.J., Uddin, A.N., Wu, F., Nadas, A., and Rossman, T.G. (2004) Arsenic-induced enhancement of ultraviolet radiation carcinogenesis in mouse skin: a dose-response study. Environ Health Perspect. 112: 599-603. 217. Arnold, L.L., Eldan, M., Nyska, A., van Gemert, M., and Cohen, S.M. (2006) Dimethylarsinic acid: results of chronic toxicity/oncogenicity studies in F344 rats and in B6C3F1 mice. Toxicology. 223: 82-100. 218. Waalkes, M.P., Liu, J., and Diwan, B.A. (2007) Transplacental arsenic carcinogenesis in mice. Toxicol Appl Pharmacol. 222: 271-80. 219. Murray-Zmijewski, F., Slee, E.A., and Lu, X. (2008) A complex barcode underlies the heterogeneous response of p53 to stress. Nat Rev Mol Cell Biol. 9: 702-12. 220. Salim, E.I., Wanibuchi, H., Morimura, K., Wei, M., Mitsuhashi, M., Yoshida, K., Endo, G., and Fukushima, S. (2003) Carcinogenicity of dimethylarsinic acid in p53 heterozygous knockout and wild-type C57BL/6J mice. Carcinogenesis. 24: 335-42. 221. Germolec, D.R., Spalding, J., Yu, H.S., Chen, G.S., Simeonova, P.P., Humble, M.C., Bruccoleri, A., Boorman, G.A., Foley, J.F., Yoshida, T., and Luster, M.I. (1998) Arsenic enhancement of skin neoplasia by chronic stimulation of growth factors. Am J Pathol. 153: 1775-85. 222. Chen, Y., Megosh, L.C., Gilmour, S.K., Sawicki, J.A., and O'Brien, T.G. (2000) K6/ODC transgenic mice as a sensitive model for carcinogen identification. Toxicol Lett. 116: 27-35. 223. Yoshida, K., Inoue, Y., Kuroda, K., Chen, H., Wanibuchi, H., Fukushima, S., and Endo, G. (1998) Urinary excretion of arsenic metabolites after long-term oral administration of various arsenic compounds to rats. J Toxicol Environ Health A. 54: 179-92. 224. Adams, M.D. and Sekelsky, J.J. (2002) From sequence to phenotype: reverse genetics in Drosophila melanogaster. Nat Rev Genet. 3: 189-98. 225. Venken, K.J. and Bellen, H.J. (2005) Emerging technologies for gene manipulation in Drosophila melanogaster. Nat Rev Genet. 6: 167-78. 226. Ryder, E. and Russell, S. (2003) Transposable elements as tools for genomics and genetics in Drosophila. Brief Funct Genomic Proteomic. 2: 57-71. 227. Bellen, H.J., Levis, R.W., Liao, G., He, Y., Carlson, J.W., Tsang, G., Evans-Holm, M., Hiesinger, P.R., Schulze, K.L., Rubin, G.M., Hoskins, R.A., and Spradling, A.C. (2004) The BDGP gene disruption project: single transposon insertions associated with 40% of Drosophila genes. Genetics. 167: 761-81. 228. Parks, A.L., Cook, K.R., Belvin, M., Dompe, N.A., Fawcett, R., Huppert, K., Tan, L.R., Winter, C.G., Bogart, K.P., Deal, J.E., Deal-Herr, M.E., Grant, D., Marcinko, M., Miyazaki, W.Y., Robertson, S., Shaw, K.J., Tabios, M., Vysotskaia, V., Zhao, L., Andrade, R.S., Edgar, K.A., Howie, E., Killpack, K., Milash, B., Norton, A., Thao, D., Whittaker, K., Winner, M.A., Friedman, L., Margolis, J., Singer, M.A., Kopczynski, C., Curtis, D., Kaufman, T.C., Plowman, G.D., Duyk, G., and Francis-Lang, H.L. (2004) Systematic generation of high-resolution deletion coverage of the Drosophila melanogaster genome. Nat. Genet. 36: 288-92.
110 229. Dietzl, G., Chen, D., Schnorrer, F., Su, K.C., Barinova, Y., Fellner, M., Gasser, B., Kinsey, K., Oppel, S., Scheiblauer, S., Couto, A., Marra, V., Keleman, K., and Dickson, B.J. (2007) A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila. Nature. 448: 151-6. 230. Murphy, T. The Drosophila GatewayTM Vector Collection. 2003 August 14, 2003; Available from: http://www.ciwemb.edu/labs/murphy/Gateway%20vectors.html. 231. Duffy, J.B. (2002) GAL4 system in Drosophila: a fly geneticist's Swiss army knife. Genesis. 34: 1-15. 232. Klueg, K.M., Alvarado, D., Muskavitch, M.A., and Duffy, J.B. (2002) Creation of a GAL4/UAS-coupled inducible gene expression system for use in Drosophila cultured cell lines. Genesis. 34: 119-22. 233. Brand, A.H. and Perrimon, N. (1993) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development. 118: 401-15. 234. Chien, S., Reiter, L.T., Bier, E., and Gribskov, M. (2002) Homophila: human disease gene cognates in Drosophila. Nucleic Acids Res. 30: 149-51. 235. Reiter, L.T., Potocki, L., Chien, S., Gribskov, M., and Bier, E. (2001) A systematic analysis of human disease-associated gene sequences in Drosophila melanogaster. Genome Res. 11: 1114-25. 236. Driscoll, M. and Gerstbrein, B. (2003) Dying for a cause: invertebrate genetics takes on human neurodegeneration. Nat Rev Genet. 4: 181-94. 237. Haywood, A.F. and Staveley, B.E. (2004) Parkin counteracts symptoms in a Drosophila model of Parkinson's disease. BMC Neurosci. 5: 14. 238. Feany, M.B. and Bender, W.W. (2000) A Drosophila model of Parkinson's disease. Nature. 404: 394-8. 239. Tapon, N., Harvey, K.F., Bell, D.W., Wahrer, D.C., Schiripo, T.A., Haber, D.A., and Hariharan, I.K. (2002) salvador Promotes both cell cycle exit and apoptosis in Drosophila and is mutated in human cancer cell lines. Cell. 110: 467-78. 240. Pagliarini, R.A. and Xu, T. (2003) A genetic screen in Drosophila for metastatic behavior. Science. 302: 1227-31. 241. Vidal, M. and Cagan, R.L. (2006) Drosophila models for cancer research. Curr Opin Genet Dev. 16: 10-6. 242. Brumby, A.M. and Richardson, H.E. (2005) Using Drosophila melanogaster to map human cancer pathways. Nat. Rev. Cancer. 5: 626-39. 243. Boulianne, G.L., Livne-Bar, I., Humphreys, J.M., Liang, Y., Lin, C., Rogaev, E., and St George-Hyslop, P. (1997) Cloning and characterization of the Drosophila presenilin homologue. Neuroreport. 8: 1025-9. 244. Hong, C.S. and Koo, E.H. (1997) Isolation and characterization of Drosophila presenilin homolog. Neuroreport. 8: 665-8. 245. Chung, H.M. and Struhl, G. (2001) Nicastrin is required for Presenilin-mediated transmembrane cleavage in Drosophila. Nat Cell Biol. 3: 1129-32. 246. Hu, Y., Ye, Y., and Fortini, M.E. (2002) Nicastrin is required for gamma-secretase cleavage of the Drosophila Notch receptor. Dev Cell. 2: 69-78. 247. Rosen, D.R., Martin-Morris, L., Luo, L.Q., and White, K. (1989) A Drosophila gene encoding a protein resembling the human beta-amyloid protein precursor. Proc Natl Acad Sci U S A. 86: 2478-82.
111 248. Schott, J.J., Benson, D.W., Basson, C.T., Pease, W., Silberbach, G.M., Moak, J.P., Maron, B.J., Seidman, C.E., and Seidman, J.G. (1998) Congenital heart disease caused by mutations in the transcription factor NKX2-5. Science. 281: 108-11. 249. Bodmer, R. and Frasch, M., Genetic determination of Drosophila heart development, in Heart Development, R.P. Harvey and N. Rosenthal, Editors. 1999, Academic Press: San Diego, CA. p. 65-90. 250. Chintapalli, V.R., Wang, J., and Dow, J.A. (2007) Using FlyAtlas to identify better Drosophila melanogaster models of human disease. Nat Genet. 39: 715-20. 251. Graf, U., Wurgler, F.E., Katz, A.J., Frei, H., Juon, H., Hall, C.B., and Kale, P.G. (1984) Somatic mutation and recombination test in Drosophila melanogaster. Environ Mutagen. 6: 153-88. 252. Wilson, R.J., Goodman, J.L., Strelets, V.B., and The FlyBase Consortium. (2008) FlyBase: integration and improvements to query tools. Nucleic Acids Res. 36: D588-93. 253. Lindsley, D.L. and Zimm, G.G., The Genome of Drosophila melanogaster. 1992, San Diego: Academic Press. viii, 1133 p. 254. Garcia-Bellido, A. and Dapena, J. (1974) Induction, detection and characterization of cell differentiation mutants in Drosophila. Mol Gen Genet. 128: 117-30. 255. Frolich, A. and Wurgler, F.E. (1989) New tester strains with improved bioactivation capacity for the Drosophila wing-spot test. Mutat Res. 216: 179-87. 256. Hallstrom, I. (1986) Genetic regulation of the cytochrome P-450 system in Drosophila melanogaster. Prog Clin Biol Res. 209B: 419-25. 257. Hallstrom, I. and Blanck, A. (1985) Genetic regulation of the cytochrome P-450 system in Drosophila melanogaster. I. Chromosomal determination of some cytochrome P-450- dependent reactions. Chem Biol Interact. 56: 157-71. 258. Dapkus, D. and Merrell, D.J. (1977) Chromosomal analysis of DDT-resistance in a long- term selected population of Drosophila melanogaster. Genetics. 87: 685-97. 259. Hallstrom, I., Blanck, A., and Atuma, S. (1984) Genetic variation in cytochrome P-450 and xenobiotic metabolism in Drosophila melanogaster. Biochem Pharmacol. 33: 13-20. 260. Graf, U. and van Schaik, N. (1992) Improved high bioactivation cross for the wing somatic mutation and recombination test in Drosophila melanogaster. Mutat Res. 271: 59-67. 261. Graf, U., Heo, O.S., and Ramirez, O.O. (1992) The genotoxicity of chromium(VI) oxide in the wing spot-test of Drosophila melanogaster is over 90 percent due to mitotic recombination. Mutation Research. 266: 197-203. 262. Ogawa, H.I., Shibahara, T., Iwata, H., Okada, T., Tsuruta, S., Kakimoto, K., Sakata, K., Kato, Y., Ryo, H., Itoh, T., and Fujikawa, K. (1994) Genotoxic activities in vivo of cobaltous chloride and other metal chlorides as assayed in the Drosophila wing spot test. Mutation Research. 320: 133-140. 263. Frolich, A. and Wurgler, F.E. (1990) Drosophila wing-spot test - improved detectability of genotoxicity of polycyclic aromatic-hydrocarbons. Mutation Research. 234: 71-80. 264. Chroust, K., Pavlova, M., Prokop, Z., Mendel, J., Bozkova, K., Kubat, Z., Zajickova, V., and Damborsky, J. (2007) Quantitative structure-activity relationships for toxicity and genotoxicity of halogenated aliphatic compounds: Wing spot test of Drosophila melanogaster. Chemosphere. 67: 152-159.
112 265. Zordan, M., Osti, M., Pavanello, S., Costa, R., and Levis, A.G. (1994) Relationship between benzo(a)pyrene-DNA adducts and somatic mutation and recombination in Drosophila melanogaster. Environmental and Molecular Mutagenesis. 23: 171-178. 266. Rizki, M., Kossatz, E., Creus, A., and Marcos, R. (2004) Genotoxicity modulation by cadmium treatment: Studies in the Drosophila wing spot test. Environmental and Molecular Mutagenesis. 43: 196-203. 267. Carmona, E.R., Kossatz, E., Creus, A., and Marcos, R. (2008) Genotoxic evaluation of two mercury compounds in the Drosophila wing spot test. Chemosphere. 70: 1910-1914. 268. do Amaral, V.S., da Silva, R.M., Reguly, M.L., and de Andrade, H.H.R. (2005) Drosophila wing-spot test for genotoxic assessment of pollutants in water samples from urban and industrial origin. Mutation Research-Genetic Toxicology and Environmental Mutagenesis. 583: 67-74. 269. Xu, T., Wang, W., Zhang, S., Stewart, R.A., and Yu, W. (1995) Identifying tumor suppressors in genetic mosaics: the Drosophila lats gene encodes a putative protein kinase. Development. 121: 1053-63. 270. Harvey, K. and Tapon, N. (2007) The Salvador-Warts-Hippo pathway - an emerging tumour-suppressor network. Nat Rev Cancer. 7: 182-91. 271. Harvey, K.F., Pfleger, C.M., and Hariharan, I.K. (2003) The Drosophila Mst ortholog, hippo, restricts growth and cell proliferation and promotes apoptosis. Cell. 114: 457-67. 272. Justice, R.W., Zilian, O., Woods, D.F., Noll, M., and Bryant, P.J. (1995) The Drosophila tumor suppressor gene warts encodes a homolog of human myotonic dystrophy kinase and is required for the control of cell shape and proliferation. Genes Dev. 9: 534-46. 273. Pan, D. (2007) Hippo signaling in organ size control. Genes Dev. 21: 886-97. 274. Zhao, B., Wei, X., Li, W., Udan, R.S., Yang, Q., Kim, J., Xie, J., Ikenoue, T., Yu, J., Li, L., Zheng, P., Ye, K., Chinnaiyan, A., Halder, G., Lai, Z.C., and Guan, K.L. (2007) Inactivation of YAP oncoprotein by the Hippo pathway is involved in cell contact inhibition and tissue growth control. Genes Dev. 21: 2747-61. 275. Colombani, J., Polesello, C., Josue, F., and Tapon, N. (2006) Dmp53 activates the Hippo pathway to promote cell death in response to DNA damage. Curr Biol. 16: 1453-8. 276. Zhang, J., Smolen, G.A., and Haber, D.A. (2008) Negative regulation of YAP by LATS1 underscores evolutionary conservation of the Drosophila Hippo pathway. Cancer Res. 68: 2789-94. 277. St John, M.A., Tao, W., Fei, X., Fukumoto, R., Carcangiu, M.L., Brownstein, D.G., Parlow, A.F., McGrath, J., and Xu, T. (1999) Mice deficient of Lats1 develop soft-tissue sarcomas, ovarian tumours and pituitary dysfunction. Nat Genet. 21: 182-6. 278. Tao, W., Zhang, S., Turenchalk, G.S., Stewart, R.A., St John, M.A., Chen, W., and Xu, T. (1999) Human homologue of the Drosophila melanogaster lats tumour suppressor modulates CDC2 activity. Nat Genet. 21: 177-81. 279. Sidorov, R.A., Ugnivenko, E.G., Khovanova, E.M., and Belitsky, G.A. (2001) Induction of tumor clones in D. melanogaster wts/+ heterozygotes with chemical carcinogens. Mutat Res. 498: 181-91. 280. Eeken, J.C., Klink, I., van Veen, B.L., Pastink, A., and Ferro, W. (2002) Induction of epithelial tumors in Drosophila melanogaster heterozygous for the tumor suppressor gene wts. Environ Mol Mutagen. 40: 277-82. 281. Woodruff, R.C., Mason, J.M., Valencia, R., and Zimmering, S. (1984) Chemical mutagenesis testing in Drosophila: I. Comparison of positive and negative control data
113 for sex-linked recessive lethal mutations and reciprocal translocations in three laboratories. Environ Mutagen. 6: 189-202. 282. Tennant, R.W. (1991) The genetic toxicity database of the National Toxicology Program: Evaluation of the relationship between genetic toxicity and carcinogenicity. Environmental Health Perspectives. 96: 47-51. 283. Yoon, J.S., Mason, J.M., Valencia, R., Woodruff, R.C., and Zimmering, S. (1985) Chemical mutagenesis in Drosophila: 4. Results of 45 coded compounds tested for the National Toxicology Program. Environmental Mutagenesis. 7: 349-367. 284. Zimmering, S., Mason, J.M., Valencia, R., and Woodruff, R.C. (1985) Chemical mutagenesis testing in Drosophila: 2. Results of 20 coded compounds tested for the National Toxicology Program. Environmental Mutagenesis. 7: 87-100. 285. Valencia, R., Mason, J.M., and Zimmering, S. (1989) Chemical mutagenesis testing in Drosophila: 6. Interlaboratory comparison of mutagenicity tests after treatment of larvae. Environmental and Molecular Mutagenesis. 14: 238-244. 286. Foureman, P., Mason, J.M., Valencia, R., and Zimmering, S. (1994) Chemical mutagenesis testing in Drosophila: 9. Results of 50 coded compounds tested for the National Toxicology Program. Environmental and Molecular Mutagenesis. 23: 51-63. 287. Foureman, P., Mason, J.M., Valencia, R., and Zimmering, S. (1994) Chemical mutagenesis testing in Drosophila: 10. Results of 70 coded chemicals tested for the National Toxicology Program. Environmental and Molecular Mutagenesis. 23: 208-227. 288. Le, H.H., Carlson, E.M., Chua, J.P., and Belcher, S.M. (2008) Bisphenol A is released from polycarbonate drinking bottles and mimics the neurotoxic actions of estrogen in developing cerebellar neurons. Toxicol Lett. 176: 149-56. 289. LaPensee, E.W., Tuttle, T.R., Fox, S.R., and Ben-Jonathan, N. (2008) Bisphenol A at low nanomolar doses confers chemoresistance in estrogen receptor alpha positive and negative breast cancer cells. Environ Health Perspect. doi: 10.1289/ehp.11788. 290. Vogel, E.W., Dusenbery, R.L., and Smith, P.D. (1985) The relationship between reaction kinetics and mutagenic action of monofunctional alkylating agents in higher eukaryotic systems. IV. The effects of the excision-defective mei-9L1 and mus(2)201D1 mutants on alkylation-induced genetic damage in Drosophila. Mutat Res. 149: 193-207. 291. Bhui-Kaur, A., Goodman, M.F., and Tower, J. (1998) DNA mismatch repair catalyzed by extracts of mitotic, postmitotic, and senescent Drosophila tissues and involvement of mei-9 gene function for full activity. Mol Cell Biol. 18: 1436-43. 292. Boyd, J.B., Golino, M.D., and Setlow, R.B. (1976) The mei-9 alpha mutant of Drosophila melanogaster increases mutagen sensitivity and decreases excision repair. Genetics. 84: 527-44. 293. Sekelsky, J.J., McKim, K.S., Chin, G.M., and Hawley, R.S. (1995) The Drosophila meiotic recombination gene mei-9 encodes a homologue of the yeast excision repair protein Rad1. Genetics. 141: 619-27. 294. Radford, S.J., McMahan, S., Blanton, H.L., and Sekelsky, J. (2007) Heteroduplex DNA in meiotic recombination in Drosophila mei-9 mutants. Genetics. 176: 63-72. 295. Calleja, F.M., Nivard, M.J., and Eeken, J.C. (2001) Induced mutagenic effects in the nucleotide excision repair deficient Drosophila mutant mus201(D1), expressing a truncated XPG protein. Mutat Res. 461: 279-88.
114 296. Pastink, A., Heemskerk, E., Nivard, M.J., van Vliet, C.J., and Vogel, E.W. (1991) Mutational specificity of ethyl methanesulfonate in excision-repair-proficient and - deficient strains of Drosophila melanogaster. Mol Gen Genet. 229: 213-8. 297. Tearle, R. (1991) Tissue specific effects of ommochrome pathway mutations in Drosophila melanogaster. Genet Res. 57: 257-66. 298. Pastink, A., Vreeken, C., Nivard, M.J., Searles, L.L., and Vogel, E.W. (1989) Sequence analysis of N-ethyl-N-nitrosourea-induced vermilion mutations in Drosophila melanogaster. Genetics. 123: 123-9. 299. Nivard, M.J., Pastink, A., and Vogel, E.W. (1996) Mutational spectra induced under distinct excision repair conditions by the 3 methylating agents N-methyl-N-nitrosourea, N-methyl-N'-nitro-N-nitrosoguanidine and N-nitrosodimethylamine in postmeiotic male germ cells of Drosophila. Mutat Res. 352: 97-115. 300. Zimmering, S., Osgood, C., and Mason, J.M. (1990) Aneuploidy in Drosophila, I. Genetic test systems in the female Drosophila melanogaster for the rapid detection of chemically induced chromosome gain and chromosome loss. Mutat Res. 234: 319-26. 301. Osgood, C., Zimmering, S., and Mason, J.M. (1991) Aneuploidy in Drosophila, II. Further validation of the FIX and ZESTE genetic test systems employing female Drosophila melanogaster. Mutat Res. 259: 147-63. 302. Walker, G.W. and Bradley, A.M. (1969) Interacting effects of sodium monohydrogenarsenate and selenocystine on crossing over in Drosophila melanogaster. Can J Genet Cytol. 11: 677-88. 303. Tripathy, N.K., Wurgler, F.E., and Frei, H. (1990) Genetic toxicity of six carcinogens and six non-carcinogens in the Drosophila wing spot test. Mutat Res. 242: 169-80. 304. Rizki, M., Kossatz, E., Xamena, N., Creus, A., and Marcos, R. (2002) Influence of sodium arsenite on the genotoxicity of potassium dichromate and ethyl methanesulfonate: studies with the wing spot test in Drosophila. Environ Mol Mutagen. 39: 49-54. 305. Ramos-Morales, P. and Rodriguez-Arnaiz, R. (1995) Genotoxicity of two arsenic compounds in germ cells and somatic cells of Drosophila melanogaster. Environ Mol Mutagen. 25: 288-99. 306. Vogel, E.W. and Zijlstra, J.A. (1987) Mechanistic and methodological aspects of chemically-induced somatic mutation and recombination in Drosophila melanogaster. Mutat Res. 182: 243-64. 307. de la Rosa, M.E., Magnusson, J., Ramel, C., and Nilsson, R. (1994) Modulating influence of inorganic arsenic on the recombinogenic and mutagenic action of ionizing radiation and alkylating agents in Drosophila melanogaster. Mutat Res. 318: 65-71. 308. Rizki, M., Kossatz, E., Velazquez, A., Creus, A., Farina, M., Fortaner, S., Sabbioni, E., and Marcos, R. (2006) Metabolism of arsenic in Drosophila melanogaster and the genotoxicity of dimethylarsinic acid in the Drosophila wing spot test. Environ Mol Mutagen. 47: 162-8. 309. Parvez, F., Chen, Y., Argos, M., Hussain, A.Z., Momotaj, H., Dhar, R., van Geen, A., Graziano, J.H., and Ahsan, H. (2006) Prevalence of arsenic exposure from drinking water and awareness of its health risks in a Bangladeshi population: results from a large population-based study. Environ Health Perspect. 114: 355-9. 310. National Research Council, Arsenic in drinking water : 2001 update, ed. (U.S.). Subcommittee on Arsenic in Drinking Water. 2001, Washington, DC: National Academy Press. xiv, 225 p.
115 311. Chiou, H.Y., Hsueh, Y.M., Hsieh, L.L., Hsu, L.I., Hsu, Y.H., Hsieh, F.I., Wei, M.L., Chen, H.C., Yang, H.T., Leu, L.C., Chu, T.H., Chen-Wu, C., Yang, M.H., and Chen, C.J. (1997) Arsenic methylation capacity, body retention, and null genotypes of glutathione S- transferase M1 and T1 among current arsenic-exposed residents in Taiwan. Mutat. Res. 386: 197-207. 312. McCarty, K.M., Ryan, L., Houseman, E.A., Williams, P.L., Miller, D.P., Quamruzzaman, Q., Rahman, M., Mahiuddin, G., Smith, T., Gonzalez, E., Su, L., and Christiani, D.C. (2007) A case-control study of GST polymorphisms and arsenic related skin lesions. Environ. Health. 6: 5. 313. Bier, E. (2005) Drosophila, the golden bug, emerges as a tool for human genetics. Nat. Genet. Rev. 6: 9-23. 314. Fraser, J.A., Kansagra, P., Kotecki, C., Saunders, R.D., and McLellan, L.I. (2003) The modifier subunit of Drosophila glutamate-cysteine ligase regulates catalytic activity by covalent and noncovalent interactions and influences glutathione homeostasis in vivo. J. Biol. Chem. 278: 46369-77. 315. Polak, M., Opoka, R., and Cartwright, I.L. (2002) Response of fluctuating asymmetry to arsenic toxicity: support for the developmental selection hypothesis. Environ. Pollut. 118: 19-28. 316. Kauer, M., Zangerl, B., Dieringer, D., and Schlotterer, C. (2002) Chromosomal patterns of microsatellite variability contrast sharply in African and non-African populations of Drosophila melanogaster. Genetics. 160: 247-56. 317. Clemens, J.C., Worby, C.A., Simonson-Leff, N., Muda, M., Maehama, T., Hemmings, B.A., and Dixon, J.E. (2000) Use of double-stranded RNA interference in Drosophila cell lines to dissect signal transduction pathways. Proc Natl Acad Sci U S A. 97: 6499-503. 318. Schweitzer, B.L. and DeKoter, R.P. (2004) Analysis of gene expression and Ig transcription in PU.1/Spi-B-deficient progenitor B cell lines. J. Immunol. 172: 144-54. 319. Livak, K.J. and Schmittgen, T.D. (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods. 25: 402-8. 320. Kann, S., Estes, C., Reichard, J.F., Huang, M.Y., Sartor, M.A., Schwemberger, S., Chen, Y., Dalton, T.P., Shertzer, H.G., Xia, Y., and Puga, A. (2005) Butylhydroquinone protects cells genetically deficient in glutathione biosynthesis from arsenite-induced apoptosis without significantly changing their prooxidant status. Toxicol. Sci. 87: 365-84. 321. Guha Mazumder, D.N., Haque, R., Ghosh, N., De, B.K., Santra, A., Chakraborty, D., and Smith, A.H. (1998) Arsenic levels in drinking water and the prevalence of skin lesions in West Bengal, India. Int. J. Epidemiol. 27: 871-7. 322. Yu, G., Sun, D., and Zheng, Y. (2007) Health effects of exposure to natural arsenic in groundwater and coal in China: an overview of occurrence. Environ Health Perspect. 115: 636-42. 323. Rossman, T.G. (2003) Mechanism of arsenic carcinogenesis: an integrated approach. Mutat Res. 533: 37-65. 324. Brambila, E.M., Achanzar, W.E., Qu, W., Webber, M.M., and Waalkes, M.P. (2002) Chronic arsenic-exposed human prostate epithelial cells exhibit stable arsenic tolerance: mechanistic implications of altered cellular glutathione and glutathione S-transferase. Toxicol Appl Pharmacol. 183: 99-107. 325. Griffith, O.W. and Mulcahy, R.T. (1999) The enzymes of glutathione synthesis: gamma- glutamylcysteine synthetase. Adv. Enzymol. Relat. Areas Mol. Biol. 73: 209-67.
116 326. Fraser, J.A., Saunders, R.D., and McLellan, L.I. (2002) Drosophila melanogaster glutamate-cysteine ligase activity is regulated by a modifier subunit with a mechanism of action similar to that of the mammalian form. J. Biol. Chem. 277: 1158-65. 327. Mendoza-Cozatl, D.G. and Moreno-Sanchez, R. (2006) Control of glutathione and phytochelatin synthesis under cadmium stress. Pathway modeling for plants. J. Theor. Biol. 238: 919-36. 328. Moreno-Sanchez, R., Saavedra, E., Rodriguez-Enriquez, S., and Olin-Sandoval, V. (2008) Metabolic control analysis: a tool for designing strategies to manipulate metabolic pathways. J. Biomed. Biotechnol. 2008: 597913. 329. Senft, A.P., Dalton, T.P., and Shertzer, H.G. (2000) Determining glutathione and glutathione disulfide using the fluorescence probe o-phthalaldehyde. Anal Biochem. 280: 80-6. 330. The International HapMap Consortium (2003) The International HapMap Project. Nature. 426: 789-96. 331. Njalsson, R. (2005) Glutathione synthetase deficiency. Cell. Mol. Life Sci. 62: 1938-45. 332. Delnomdedieu, M., Basti, M.M., Otvos, J.D., and Thomas, D.J. (1993) Transfer of arsenite from glutathione to dithiols: a model of interaction. Chem Res Toxicol. 6: 598- 602. 333. Cullen, W.R., McBride, B.C., and Reglinski, J. (1984) The reduction of trimethylarsine oxide to trimethylarsine by thiols: a mechanistic model for the biological reduction of arsenicals. JIC. 21: 45-60. 334. Delgado Rodriguez, A., Ortiz Marttelo, R., Graf, U., Villalobos Pietrini, R., and Gomez Arroyo, S. (1995) Genotoxic activity of environmentally important polycyclic aromatic- hydrocarbons and their nitro-derivatives in the wing spot-test of Drosophila melanogaster. Mutation Research-Genetic Toxicology. 341: 235-247. 335. Muñiz Ortiz, J.G., Opoka, R., Kane, D., and Cartwright, I.L. (2008) Investigating arsenic susceptibility from a genetic perspective in Drosophila reveals a key role for glutathione synthetase. Toxicol Sci. doi:10.1093/toxsci/kfn192. 336. Afton, S., Kubachka, K., Catron, B., and Caruso, J.A. (2008) Simultaneous characterization of selenium and arsenic analytes via ion-pairing reversed phase chromatography with inductively coupled plasma and electrospray ionization ion trap mass spectrometry for detection Applications to river water, plant extract and urine matrices. J Chromatogr A. 337. Nickson, R., McArthur, J., Burgess, W., Ahmed, K.M., Ravenscroft, P., and Rahman, M. (1998) Arsenic poisoning of Bangladesh groundwater. Nature. 395: 338. 338. Yu, H.S. (1984) Blackfoot disease and chronic arsenism in southern Taiwan. Int J Dermatol. 23: 258-60. 339. Ristoff, E. and Larsson, A. (2007) Inborn errors in the metabolism of glutathione. Orphanet J Rare Dis. 2: 16. 340. Shi, Z.Z., Habib, G.M., Rhead, W.J., Gahl, W.A., He, X., Sazer, S., and Lieberman, M.W. (1996) Mutations in the glutathione synthetase gene cause 5-oxoprolinuria. Nat Genet. 14: 361-5. 341. Njalsson, R., Ristoff, E., Carlsson, K., Winkler, A., Larsson, A., and Norgren, S. (2005) Genotype, enzyme activity, glutathione level, and clinical phenotype in patients with glutathione synthetase deficiency. Hum Genet. 116: 384-9.
117 342. Dahl, N., Pigg, M., Ristoff, E., Gali, R., Carlsson, B., Mannervik, B., Larsson, A., and Board, P. (1997) Missense mutations in the human glutathione synthetase gene result in severe metabolic acidosis, 5-oxoprolinuria, hemolytic anemia and neurological dysfunction. Hum Mol Genet. 6: 1147-52. 343. Njalsson, R., Carlsson, K., Olin, B., Carlsson, B., Whitbread, L., Polekhina, G., Parker, M.W., Norgren, S., Mannervik, B., Board, P.G., and Larsson, A. (2000) Kinetic properties of missense mutations in patients with glutathione synthetase deficiency. Biochem J. 349: 275-9. 344. The International HapMap Consortium. (2007) A second generation human haplotype map of over 3.1 million SNPs. Nature. 449: 851-61. 345. Ahsan, H., Chen, Y., Parvez, F., Argos, M., Hussain, A.I., Momotaj, H., Levy, D., van Geen, A., Howe, G., and Graziano, J. (2006) Health Effects of Arsenic Longitudinal Study (HEALS): description of a multidisciplinary epidemiologic investigation. J Expo Sci Environ Epidemiol. 16: 191-205. 346. Ochi, T., Suzuki, T., Isono, H., Schlagenhaufen, C., Goessler, W., and Tsutsui, T. (2003) Induction of structural and numerical changes of chromosome, centrosome abnormality, multipolar spindles and multipolar division in cultured Chinese hamster V79 cells by exposure to a trivalent dimethylarsenic compound. Mutat Res. 530: 59-71. 347. Chakraborty, T., Das, U., Poddar, S., Sengupta, B., and De, M. (2006) Micronuclei and chromosomal aberrations as biomarkers: a study in an arsenic exposed population in West Bengal, India. Bull Environ Contam Toxicol. 76: 970-6. 348. Klein, C.B., Leszczynska, J., Hickey, C., and Rossman, T.G. (2007) Further evidence against a direct genotoxic mode of action for arsenic-induced cancer. Toxicol Appl Pharmacol. 222: 289-97. 349. Kligerman, A.D. and Tennant, A.H. (2007) Insights into the carcinogenic mode of action of arsenic. Toxicol Appl Pharmacol. 222: 281-8. 350. Gatti, M., Bonaccorsi, S., and Pimpinelli, S. (1994) Looking at Drosophila mitotic chromosomes. Methods Cell Biol. 44: 371-91. 351. Caussinus, E. and Gonzalez, C. (2005) Induction of tumor growth by altered stem-cell asymmetric division in Drosophila melanogaster. Nat Genet. 37: 1125-9. 352. Gonzalez, C. (2007) Spindle orientation, asymmetric division and tumour suppression in Drosophila stem cells. Nat Rev Genet. 8: 462-72. 353. Nelson, G.M., Ahlborn, G.J., Delker, D.A., Kitchin, K.T., O'Brien, T.G., Chen, Y., Kohan, M.J., Roop, B.C., Ward, W.O., and Allen, J.W. (2007) Folate deficiency enhances arsenic effects on expression of genes involved in epidermal differentiation in transgenic K6/ODC mouse skin. Toxicology. 241: 134-45.
118 Appendix I.
Description of Unpublished Negative Results
1. RNAi Approach to Identify Genes Required for a Robust Response to Arsenite.
In order to identify genes that could render Drosophila S2 cells susceptible to AsIII, we employed RNA interference (RNAi) to knock down the expression of genes that were hypothesized to be involved in maintaining the redox state of the cell, which has been shown to be deregulated in the presence of AsIII. We targeted thioredoxin peroxidase 1 (Jafrac1),
peroxiredoxin (Prx5037), thioredoxin 2 (trx-2), thioredoxin reductase 1 (Trxr-1) and 2 (Trxr-2),
uncoupler of phosphorylation 4A (Ucp4A), CG6788, CG32496, CG7772, and catalase genes.
We did not observe susceptibility to AsIII when any of these genes were knocked down.
The main reason for the observed results could be that transcript knock-down via RNAi
may not have occurred. Transcripts levels were only verified for CG6788, CG32496, and
CG7772 and, since along with GS they were the only genes absent in Df #11, the results showed
that the levels of supposedly knocked-down transcripts were actually higher than the wild type.
The levels of mRNA transcripts in the knock-down cells could have been higher than wild-type
(unexposed to RNAi) because the PCR primers used could be detecting the dsRNA construct
used for RNAi. Another reason could be the functional redundancy of the enzymes targeted.
Some knock-down experiments were performed in combinations in an attempt to reduce the
activities of several similar enzymes, but we obtained the same results as when knock-downs
were performed individually. Finally, the genes examined may not be required for a robust
response to AsIII exposure.
119 Alternative approach:
RNAi in Drosophila can be performed in vivo by obtaining adult flies from the Vienna
Drosophila RNAi Center that harbor a construct that expresses a dsRNA hairpin targeting any
gene in the fly genome under the control of the GAL4/UAS expression system. This system has been verified for all of our target genes for off-target effects.
2. Functional Characterization of the Role of GS Towards Arsenite Sensitivity Via P-
element Mobilization.
After identifying a role for GS in arsenic toxicity and susceptibility via RNAi, an in vivo
approach to further characterize the function of the gene with regards to arsenic was undertaken.
A fly line harboring a P-element insertion (EP1322) near the GS gene was obtained from the
Bloomington Stock Center (Bloomington, Indiana). We attempted to induce an imprecise
excision of the P-element, causing loss of genetic material around the GS gene, thus leading to a
mutation in the gene. After performing the necessary crosses to induce the imprecise excision,
PCR was performed across the deletion site. After screening seven putative mutants, we
observed a precise excision rather than an imprecise one. If more lines could be examined the
chances of identifying an imprecise excision should increase.
Alternative approach:
Two alternative approaches were taken by creating a FLP-FRT deletion mutant and using
a GS-RNAi targeted fly strain to assess the role of GS in arsenic toxicity.
Table A.1. Primer set used to identify potential imprecise excision of EP1322. Forward Primer AGTCACAACAATCGCTCGTTGAG
Reverse Primer TGGCAAACTGTTGCTCAAGCAGC Annealing Temp.(C) 60.6
120
3. Rescue of Arsenite-exposed S2 Cells Using Mannitol as an H2O2 Scavenger.
We performed knock-down experiments in S2 cells targeting MnSOD and CuSOD,
which scavenge for superoxide producing H2O2, which is then scavenged by catalase to produce
H2O and O2. Our initial experiments using the CellTiter-Glo luminescent (Promega) assay
showed that knock-down of MnSOD and CuSOD in S2 cells exposed to AsIII caused an increase
in cell growth when compared to cells expressing wild type levels of both SODs at the same
concentrations of AsIII. However, we were unable to reproduce these results using the fluorescent CyQuant-NF Cell Proliferation kit (Invitrogen) under the same conditions. RNAi- induced knock down of catalase gene transcription did not show any effects on cell growth with or without AsIII treatment. Thus, we hypothesized that the ultimate AsIII -induced ROS leading
III to cellular toxicity was H2O2. We treated S2 cells with non-toxic and highly toxic levels of As with or without mannitol in an attempt to rescue the cells from H2O2-induced cytotoxicity for 0-
72 hr and measuring cell density every 24 hrs or from 0-24 hrs measuring cell density every eight hours. Our results did not support to our hypothesis in that the cells were susceptible to
AsIII with our without mannitol. Next, we used sodium formate instead of mannitol but we
observed the same results. Lastly, we used deferoxamine to chelate iron in order to prevent a
Fenton reaction between iron and H2O2, which would produce a hydroxyl radical (HO·) and a
hydroxyl anion (OH-). Our results showed that deferoxamine was toxic to S2 cells; thus, further
experimentation using the chelator was not performed. Our experimental approach may have not
III produced the results we expected because H2O2 may not be the ultimate ROS leading to As -
induced cytotoxicity, the levels of mannitol used could have been too low for rescue, or the
stoichiometry between mannitol and AsIII might have to be adjusted.
121
Alternative approach:
Other ROS could be targeted for scavenging by using other chelators such as Tiron or
Tempol, which shed some light on an ultimate AsIII -induced oxidative stressor.
4. Identification of ROS in S2 Cells After Exposure to Sodium Arsenite.
Many studies have shown that a carcinogenic/genotoxic mode of action of AsIII occurs
via the production of ROS [1, 2]. Thus, we attempted to detect the production of ROS by
sodium AsIII in S2 cells incubated in 4-well plates, by using the Image-iT LIVE green ROS detection kit (Molecular Probes). We exposed S2 cells to 50 µM AsIII, a non-toxic
concentration, but could not detect any ROS at any time point up to three hours. We also
attempted to detect ROS after treatment with 100 µM and 500 µM TBHP, a positive control for
production of ROS, taking an image every 5 seconds for 25 seconds but could not observe a
difference in fluorescence between untreated and treated cells. We attempted to shorten the time
of AsIII exposure by exposing S2 cells to 60 µM AsIII for 15 seconds, taking an image every 5
seconds, but we could not see a difference in fluorescence compared to non-exposed cells. We observed similar results when S2 cells were exposed to 100 or 500 µM TBHP. Reasons for not observing the expected results could be because of a spike in ROS, which may occur earlier than our first time point; S2 cells might be scavenging for the ROS formed at a higher rate; the concentration of AsIII, though it approximated toxic levels, might be too low for production of
ROS; the fluorescent probe used was not at the optimal level required for S2 cells; and the
fluorescent probe might have undergone bleaching between exposure and visualization under the
fluorescent microscope; or simply that no ROS were produced.
122 Alternative approach:
As an alternative, another kit for detecting ROS could be used to detect specific forms of
ROS. S2 cells could be exposed to higher levels of AsIII and for shorter time points. The visualization of fluorescence could be performed using a different microscope at the College of
Medicine Center for Biological Microscopy. The S2 cells could be fixed in cover slips rather than using 4-well plates.
5. Biochemical and Genetic Studies to Identify Differential Response Mechanism to
Arsenite Between PVM and Oregon-R 1970 Strains.
Previous work identified a differential response towards AsIII between PVM (sensitive) and
Oregon-R 1970 (Ore-R; tolerant), but biochemical and genetic analysis detailing the reason for such contrasting differences have yet to be performed. We approached the study of biochemical and genetic differences in four ways.
A. As described previously, we identified GS as a candidate for susceptibility towards AsIII;
thus, we decided to verify the potential difference in levels of GSH in PVM and Ore-R [3].
Our results showed that PVM had slightly higher levels of GSH than Ore-R, contrary to what
we expected, since the PVM is more sensitive than Ore-R when exposed to AsIII. We also
measured levels of oxidized GSH, GSSG, and the results showed that Ore-R had slightly
higher levels than PVM whether exposed or unexposed to AsIII. These results suggest that
the differential response towards AsIII between PVM and Ore-R is not due to the levels of
GSH produced, but might be due to the rate of production in the presence of AsIII.
B. Because we could not identify a significant biochemical difference in the levels of
GSH/GSSG between PVM and Ore-R, we decided to look into genetic differences. The first
approach we undertook was identifying any potential differences in transcripts made by the
123 two strains via Northern analysis. Table 1 shows the primers used to produce specific probes
against the 5’ region of the GS (CG6835) and CG32495 genes, and RpL32 was used as a
loading control. Results were inconclusive because we could not identify the targeted
transcripts, except for those against RpL32.
C. The images from the Northern analysis were not conclusive enough to gain any knowledge
regarding the transcription of the GS and CG32495 genes. Therefore, we performed real
time-reverse transcriptase PCR targeting a individual and/or combinations of transcripts from
both genes (Table 2). The levels of each transcript were normalized to the levels of the
control gene alpha-tubulin (α-Tub84B). The only consistent results we obtained were the
higher levels of the GS RC and RD transcript levels in Ore-R compared to PVM. The other
PCRs we attempted were inconclusive in that they were not reproducible.
D. Having identified the GS gene as coding for the differential response observed between Ore-
R and PVM, we thought the activity of the enzyme could be different between the two
strains. We used a method previously described [4] to measure and compare the enzymatic
activity of GS between Ore-R and PVM. Our results were inconclusive because we could
not obtain consistent readings between samples and the levels of activity were lower than
expected.
Alternative approaches:
First, HPLC could be performed to identify possible differences in the levels of gamma- glutamyl cysteine (γ-GC), the precursor to GSH. The presence of γ-GC could interfere with the
GSH measurements because γ-GC also reacts with OPA. Therefore, our previous readings could be measuring of the total levels of GSH and γ-GC. HPLC could be used to determine the levels of GSH and γ-GC individually. Secondly, the Northern analysis could be performed utilizing
124 new larger probes. The genomic region between the GS (CG6835) and CG32495 in Ore-R and
PVM could also be sequenced to identify polymorphic differences that could be harbored
between the two strains. The new probes for the Northern analysis could subsequently be
derived from the sequence information. Our results from the GS activity enzymatic activity
could have been negative due to the preparation of the γ-GC we obtained from Bachem
Americas, which could have traces of GSH that would interfere with our readouts of GSH
production by the native GS enzyme. If another company could produce a better yield of γ-GC,
the experiment could be attempted and the protocol could be perfected. However, the results
obtained suggest that no biochemical or genetic difference exists between PVM and Ore-R in the
production of GSH via GS.
Table A.2. Primer sets used to produce mRNA probes for Northern analysis. Probe Forward Primer (5’ 3’) Reverse Primer (5’ 3’) Annealing Probe Target Temp (C) Size (bp) 5’ GS & CCAAGCAGAACGATCAT TGGGTGAATCCGTGCGC 65.6 386 CG32495 GTCCAGC CAGCA 3’ GS & TGGATCCCAAGAGGTGG AGCGCCATCTCGTAGCT 66.3 297 CG32495 CCGTCAT GGCA RpL32 GCACATGTTATCAATGG CACAAATGGCGCAAGCC 62.3 463 TGCTG CAAGGGT
125 Table A.3. Primer sets used to target GS (CG6835) and CG32495 transcripts. Transcript Forward Primer (5’ 3’) Reverse Primer (5’ 3’) Annealing Targeted Temp (C) GS RA AGAGTTTTCATTTGCTGACGCT CAGGATGCAGGATGCGGGATG 62.2 GC CCA GS RC CATAGTTATATTTTTGTCTGTG GCACTGTCAATTTGGACACC 60.5 GGC GS RC RD TGCAGGAGTCGGAACTGCCGC TGGTGACCCGGCTTAGCTGGA 63.5 T C CG32495 RB GCGATCCGACTATATGGCACA GGAGGCCACCGTGTTGATCTC 60.2 CGT GAC CG32495 ex. ACTCCGGAGAGATTCGTGCTG TAGTGCATCTGGTATATCCACG 62.2 6 RA RB ex. 4 AA CC RC 5’ GS & CGCCATGCGATCGAAGACGGC TCGAACTCCTTGCGCGGAAAC 62.4 CG32495 CTT G 3’ GS & TCGTGGACATGGTCTCCGAGC GCAGCATGTGTCCCGCCTGGT 62.7 CG32495 TGG A α-Tub84D CGATGAGGTCCGTACCGGAAC ACGGGCGTAGTTGTTGGCCGC 66.3 CTACCGTCAGCT ATCC
126 References
1. Hei, T.K. and Filipic, M. (2004) Role of oxidative damage in the genotoxicity of arsenic. Free Radic Biol Med. 37: 574-81. 2. Kitchin, K.T. and Ahmad, S. (2003) Oxidative stress as a possible mode of action for arsenic carcinogenesis. Toxicol Lett. 137: 3-13. 3. Senft, A.P., Dalton, T.P., and Shertzer, H.G. (2000) Determining glutathione and glutathione disulfide using the fluorescence probe o-phthalaldehyde. Anal Biochem. 280: 80-6. 4. Volohonsky, G., Tuby, C.N., Porat, N., Wellman-Rousseau, M., Visvikis, A., Leroy, P., Rashi, S., Steinberg, P., and Stark, A.A. (2002) A spectrophotometric assay of gamma- glutamylcysteine synthetase and glutathione synthetase in crude extracts from tissues and cultured mammalian cells. Chem Biol Interact. 140: 49-65.
127 Appendix II.
Standard Drosophila Procedures
Determination of hAS3MT Chromosomal Insertion
Eclosing adults from injected w1118 embryos were individually collected and crossed to w1118 males or female.
X-chromosomal insertion
If the AS3MT insertion lied on the X-chromosome of red-eyed (w*; hAS3MT, w+) males eclosing from the first cross, it was expected that only w*; hAS3MT, w+ females and w1118 males would
eclose. Each w*; hAS3MT, w+ female offspring from the first cross was then mated to an X-
chromosome balancer strain to homozygose the transgenic line. If w*; h-AS3MT, w+ female
offspring eclosed from the microinjected embryos, they were crossed to w1118 males and the w*; hAS3MT, w+ offspring males were then mated to w1118 females and homozygosed as described above. If the cross of a w*; hAS3MT, w+ male offspring from the first cross to a w1118 female
resulted in w* male and female offspring, as well as w*; hAS3MT, w+ males and females, we
could conclude that the insertion was harbored in any of the other two autosomal chromosomes.
2nd and 3rd chromosome insertion
To determine if the insertion lied either on the 2nd or 3rd chromosome a w*; hAS3MT, w+ male was crossed to 2nd balancer strain marked with the CyO gene. The offspring from this cross would consist of w*; hAS3MT, w+/CyO and w-/CyO, male and female genotypes. The w*;
hAS3MT, w+/CyO males and females were then intercrossed. If only w*; hAS3MT, w+ offspring
appear, it meant the insertion lied on the 2nd chromosome. These offspring were then
homozygosed by crossing w*; hAS3MT, w+ males to w*; hAS3MT, w+ females. If w*; CyO or
128 w*, along with the transgenic offspring emerged from the w*; hAS3MT, w+/CyO reciprocal cross,
it was inferred that the insertion then lied on the 3rd chromosome. To homozygose this line w*;
hAS3MT, w+/Cy males were crossed to a 3rd chromosome balancer strain harboring the TM6B,
Tb marker. TM6B, Tb pupae from this cross were collected and w*; hAS3MT, w+; TM6B, Tb
females and males were collected and intercrossed. w*; hAS3MT, w+ pupae (non-TM6B Tb)
were collected from this mating and w*; hAS3MT, w+ (non-TM6B, Tb) females and males were
intercrossed to homozygose the line.
Creation of hAS3MT; latsx1 Fly Line
Since the latsx1 allele is homozygous lethal, it is balanced over the TM6B, Tb balancer
chromosome. Therefore, in order to create a Drosophila line with both the hAS3MT the latsx1 mutation in the same fly we had to create a hAS3MT line with a 2nd chromosome balancer CyO
and a double 3rd chromosome balancer line heterozygote for the TM3, Sb and TM6B, Tb 3rd
chromosome balancers. We mated the 1099-2E transgenic line to a line harboring the TM3, Sb
3rd balancer chromosome and, in a parallel cross, the 1099-2E line was mated to a line harboring
the TM6B, Tb 3rd chromosome balancer. The offspring from these two parallel crosses showing
both CyO; Sb or CyO; Tb phenotypic markers were intercrossed and eclosed adults were selected
for by the Sb and Tb phenotypes and the loss of the CyO marker (maintained as a stable stock).
These were then crossed to the offspring of the latsx1/TM6B, Tb line mating with the TM6B, Tb
3rd chromosome balancer line that were Cy and Tb phenotypically. This cross yielded w; hAS3MT, w+/CyO; latsx1/TM3, Sb females and males that were then intercrossed to maintain a
UAS-AS3MT homozygote line harboring the latsx1 mutant marker balanced over the TM3, Sb 3rd
chromosome balancer denominated w; hAS3MT; latsx1/TM3, Sb.
129 Preparation of Fly Genomic DNA
Fly genomic DNA was isolated according to a protocol at the Berkeley Drosophila Genome
Project (http://www.fruitfly.org:9005/about/methods/inverse.pcr.html) with several
modifications. Briefly, to two flies in a microcentrifuge tube 100 μl of homogenization buffer
(1% SDS, 100 mM Tris-HCl, pH 9.0, 100 mM EDTA prepared in DEPC-treated deionized H2O)
was added, followed by homogenization using a plastic pestle (Fisher Scientific, Pittsburgh, PA).
The homogenate was incubated at 70° C for 30 min, 14 μl 8 M KOAc was added and the tube
placed on ice for 30 min. The contents were centrifuged at 10,000 x g at 4° C for 15 min and the
supernatant transferred to a fresh microcentrifuge tube to which 50 μl isopropanol was added,
followed by centrifugation at 14,000 x g for 5 minutes at RT. The supernatant was discarded,
100 μl of 70% ethanol (prepared with DEPC-dH2O) was added to the pellet, the contents mixed well, and again centrifuged at RT. The ethanol was discarded, the pellet was air dried, and then resuspended in 10 μl deionized H2O.
RNA Isolation
S2 Cells
Isolation of RNA from S2 cells was performed using the RNeasy® Mini Kit according to
manufacturer’s directions (Qiagen, Valencia, CA) for cultured cells, including the addition of 10
μl DNase I stock solution in 70 μl buffer RDD.
Third instar larvae/adults
Forty (40) third instar larvae or adults were snap-frozen in a cooled porcelain mortar in liquid
nitrogen and homogenized in 5 ml TRI Reagent® (Ambion®, Austin, TX). The homogenate was
centrifuged at 12,000 x g at 4° C for 10 min, the supernatant decanted, and chloroform (1 ml)
added, followed by vigorous shaking, incubation for 10 min at RT, and centrifugation (12,000 x
130 g) for 15 min at 4° C. The aqueous layer was withdrawn, further TRI Reagent (2 ml) added with
gentle mixing, followed by a further round of chloroform treatment (400 µl) as above. The
recovered aqueous layer was mixed with isopropanol (2.5 ml), incubated for 10 min at RT and
the precipitate recovered by centrifugation at 12,000 x g for 8 min at 4° C. After washing with
70% ethanol (5 ml, made with DEPC-treated dH2O) the pellet was dissolved in DEPC-treated
H2O (500 µl), transferred to a 1.5 ml tube, 3M sodium acetate added (50 μl) (made in milliQ
H2O and autoclaved) followed by 100% ethanol (1 ml) and incubation for 6 h to overnight at -
20°C. The pellet was recovered by centrifugation (12,000 x g for 5 min at 4° C) and subjected to a further round of dissolution and precipitation. The final pellet was dried at RT for 1 h and resuspended in milliQ H2O (100 µl).
131