Cadmium alters the formation of benzo[a]pyrene DNA adducts in the RPTEC/TERT1 human renal proximal...

Accepted Manuscript

Title: Cadmium alters the formation of benzo[a]pyrene DNAadducts in the RPTEC/TERT1 human renal proximal tubuleepithelial cell line

Author: Bridget R. Simon Mark J. Wilson Diane A. BlakeHaini Yu Jeffrey K. Wickliffe

PII: S2214-7500(14)00046-8DOI: http://dx.doi.org/doi:10.1016/j.toxrep.2014.07.003Reference: TOXREP 45

To appear in:

Received date: 14-6-2014Revised date: 7-7-2014Accepted date: 8-7-2014

Please cite this article as: B.R. Simon, M.J. Wilson, D.A. Blake, H. Yu, J.K.Wickliffe, Cadmium alters the formation of benzo[a]pyrene DNA adducts in theRPTEC/TERT1 human renal proximal tubule epithelial cell line, Toxicol. Rep. (2014),http://dx.doi.org/10.1016/j.toxrep.2014.07.003

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http://dx.doi.org/doi:10.1016/j.toxrep.2014.07.003

http://dx.doi.org/10.1016/j.toxrep.2014.07.003

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Title: Cadmium alters the formation of benzo[a]pyrene DNA adducts in the 1

RPTEC/TERT1 human renal proximal tubule epithelial cell line2

Bridget R. Simon a, b, Mark J Wilson b, Diane A. Blake a,c, Haini Yu c, and Jeffrey K. 3

Wickliffe a, b, d4

a Graduate Program in Biomedical Sciences, Tulane University School of Medicine, New 5

Orleans, LA 701126

b Department of Global Environmental Health Sciences, Tulane University School of 7

Public Health and Tropical Medicine, New Orleans, LA 701128

c Department of Biochemistry and Molecular Biology, Tulane University School of 9

Medicine, New Orleans, LA 7011210

d To whom correspondence should be addressed at 1440 Canal Street, Suite 2100, 11

New Orleans, LA, 70112. Telephone: 504-988-3910. E-mail: [email protected]

Running title: Co-exposure alters DNA adduct formation13

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Abstract15

Previously, we demonstrated the sensitivity of RPTEC/TERT1 cells, an 16

immortalized human renal proximal tubule epithelial cell line, to two common 17

environmental carcinogens, cadmium (Cd) and benzo[a]pyrene (B[a]P). Here, we 18

measured BPDE-DNA adducts using a competitive ELISA method after cells were 19

exposed to 0.01, 0.1, and 1 μM B[a]P to determine if these cells, which appear 20

metabolically competent, produce BPDE metabolites that react with DNA. BPDE-DNA 21

adducts were most significantly elevated at 1 μM B[a]P after 18 and 24 hours with 36.34 22

+/- 9.14 (n = 3) and 59.75 +/- 17.03 (n = 3) adducts/108 nucleotides respectively. For 23

mixture studies, cells were exposed to a non-cytotoxic concentration of Cd, 1 μM, for 24 24

hours and subsequently exposed to concentrations of B[a]P for 24 hours. Under these 25

conditions, adducts detected at 1 μM B[a]P after 24 hours were significantly reduced, 26

17.28 +/- 1.30 (n = 3) adducts/108 nucleotides, in comparison to the same concentration 27

at previous time points without Cd pre-treatment. We explored the NRF2 antioxidant 28

pathway and total glutathione levels in cells as possible mechanisms reducing adduct29

formation under co-exposure. Results showed a significant increase in the expression of 30

NRF2-responsive genes, GCLC, HMOX1, NQO1, after 1 μM Cd x 1 μM B[a]P co-31

exposure. Additionally, total glutathione levels were significantly increased in cells 32

exposed to 1 μM Cd alone and 1 μM Cd x 1 μM B[a]P. Together, these results suggest 33

that Cd may antagonize the formation of BPDE-DNA adducts in the RPTEC/TERT1 cell 34

line under these conditions. We hypothesize that this occurs through priming of the 35

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antioxidant response pathway resulting in an increased capacity to detoxify BPDE prior 36

to BPDE-DNA adduct formation. 37

Key words: Mixtures toxicology; BPDE-DNA adducts; renal cancer; RPTEC/TERT138

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Introduction40

Over 90% of kidney cancers originate in the renal proximal tubule epithelial cells 41

and are classified as renal cell carcinoma (RCC). However, only about 2% of kidney 42

cancer cases can be attributed to a genetic predisposition (Motzer et al., 1996; Polascik43

et al., 2002). The remaining cases occur in otherwise healthy individuals with no prior 44

familial history (Cancer Facts and Figures 2012, 2012). Substantial evidence of 45

environmental risk factors contributing to the development of RCC suggests that further 46

scrutiny of human mutagens and carcinogens on the cellular and molecular level is 47

warranted (USRDS 2013 Annual Data Report; Chow et al., 2010). 48

Exposure to polycyclic aromatic hydrocarbons (PAHs) has been associated with 49

an elevated risk of many cancers including skin, lung, bladder, liver, and stomach (Toxic 50

Substances Database, 2011). PAHs are formed as byproducts of incomplete 51

combustion and are ubiquitous in the environment. Major routes of exposure include52

inhalation and ingestion, which can result from cigarette smoking, consumption of grilled 53

or contaminated foods, and atmospheric pollution associated with the burning of fossil 54

fuels (IARC Working Group on the Evaluation of Carcinogenic Risks to Humans. and 55

International Agency for Research on Cancer., 2010). Increased consumption of 56

chargrilled meats has been shown to directly correlate with elevated PAH exposure and57

risk of RCC (Daniel et al., 2012; Daniel et al., 2011). Additional factors associated with 58

the development of RCC include obesity, smoking, and hypertension (Chow, et al., 59

2010; Jonasch et al., 2012; Ljungberg et al., 2011).60

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The ability of cells to repair bulky PAH-DNA adducts may be altered by the 61

presence of environmental contaminants such as cadmium (Cd). Cd has been shown to 62

substitute for zinc ion co-factors in many DNA repair proteins and enzymes specifically 63

responsible for recognizing and repairing DNA adducts (Hartwig and Schwerdtle, 2002). 64

Cd, a heavy metal and known nephrotoxicant, is present in the environment in food, 65

cigarettes, and contaminated water runoff. Human exposure to Cd occurs primarily 66

through inhalation of fine particulates (i.e. tobacco smoke) and consumption of foods 67

such as rice, cereal, and mollusks. (IARC Working Group on the Evaluation of 68

Carcinogenic Risks to Humans. and International Agency for Research on Cancer., 69

2012). Cd accumulates in the liver, kidneys, and bone and is suspected to promote 70

cancers in these organs as well as in the lungs. Cd lacks strong mutagenic properties 71

but may act as a co-carcinogen in the body by inhibiting DNA damage repair processes 72

and increasing oxidative stress in cells (Joseph, 2009; Waisberg et al., 2003).73

Because individuals are rarely exposed to single chemical agents or carcinogens74

in the environment, it is important to study these compounds as humans might 75

encounter them on a daily basis. Exposure to chemical mixtures can result in76

toxicological outcomes that substantially differ from the expected effects of each77

compound alone. Toxicants in mixtures may act through similar or distinctly different 78

mechanisms of action. Chemicals and compounds can act antagonistically, additively,79

synergistically, or one chemical may potentiate the effects of another (Klassen, 2008).80

Ultimately, these interactions may substantially alter toxicity to different and possibly 81

unexpected degrees. For example, in vitro studies have shown that exposure to binary 82

combinations of PAHs including benzo[a]pyrene (B[a]P) and benzo[b]fluoranthene 83

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(B[b]F) results in a significant increase in the formation of DNA adducts than exposure 84

to B[a]P alone. However, exposure to both B[a]P and benzo[k]fluoranthene (B[k]F), a 85

similarly structured PAH, results in a significant reduction in the formation of DNA 86

adducts than exposure to B[a]P alone (Sevastyanova et al., 2007; Staal et al., 2007; 87

Tarantini et al., 2011). The opposing results that occur even after exposure to 88

compounds of the same toxicant class emphasize the need for studies investigating 89

effects elicited after exposure to mixtures of toxicants from similar and different classes.90

In order to study the mechanisms of mixture exposure, which may promote RCC, 91

we utilized an immortalized human renal cell line, RPTEC/TERT1. The RPTEC/TERT1 92

cell line was derived from the renal proximal tubule epithelial cells (RPTEC) of a normal, 93

healthy male donor. These cells were immortalized with the catalytic subunit of the 94

human telomerase reverse transcriptase enzyme (TERT1) (Wieser et al., 2008). 95

Previously, we determined that RPTEC/TERT1 cells exhibit sensitivity and compound-96

specific responses to B[a]P and Cd treatment (Simon et al., 2014). Our results were 97

consistent with canonical biological responses to both environmental toxicants and 98

demonstrate metabolic competency of the RPTEC/TERT1 cell line. To test our 99

hypothesis that Cd may alter formation of adducts after B[a]P exposure, we have 100

explored concentration-dependent formation of BPDE-DNA adducts through cellular 101

bioactivation of B[a]P. We examined the persistence of those adducts under conditions 102

of pre-treatment with Cd. We intended to determine the effects of Cd on the persistence 103

of BPDE-DNA adducts as a function of time, co-exposure, and oxidative stress. 104

We hypothesize that exposure to a binary combination of the environmental 105

carcinogens, Cd, a heavy metal, and B[a]P, a representative PAH, acts to alter DNA 106

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adduct formation in comparison to levels found after B[a]P exposure alone. As Cd is 107

known to inhibit the recognition and/or repair of PAH-DNA adducts, it is plausible to find108

persistence of adducts under conditions of co-exposure (Kopera et al., 2004).109

Alternatively, co-exposure may result in an antagonistic response leading to the 110

formation of fewer DNA adducts through increased detoxification or inhibition of 111

bioactivation. However, our previous work in the RPTEC/TERT1 cell line suggests that112

the inhibition of bioactivation is unlikely (Simon, et al., 2014). The interaction of chronic, 113

low level exposure to both Cd and PAHs over a lifetime may provide support for 114

environmental contributions to the development of RCC in healthy individuals. 115

116Materials and Methods117

118Reagents119

120Chemicals121

122All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) unless noted 123

otherwise. Cadmium chloride (CdCl2, 202908) was dissolved in fresh complete medium 124

and delivered at 0.1% of the final culture volume to yield the appropriate target 125

concentrations. Benzo[a]pyrene (B[a]P, B1760) was dissolved in dimethyl sulfoxide 126

(DMSO, D8418) and delivered at 0.05% of the final culture volume to yield the 127

appropriate target concentrations. B[a]P preparations and exposures were carried out 128

under low light conditions. 129

DNA Isolation Reagents130

Enzymes used for DNA isolation including RNaseT1, mRNAse A, and proteinase 131

K were purchased from Sigma-Aldrich. Tris-buffered saturated phenol, 132

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phenol:chloroform:isoamyl (25:24:1), and 5 PRIME Phase Lock Gel, light, 15mL tubes 133

for DNA isolation were purchased from Fisher Scientific (Pittsburg, PA).134

BPDE-DNA Adduct ELISA Reagents135

Greiner Bio-One microplates (high-binding, white) were purchased from Fisher 136

Scientific. I-Block casein-based blocking solution and CPD-Star Substrate with Emerald-137

II Enhancer were purchased from Life Technologies™ (Grand Island, NY). Polyclonal 138

BPDE-DNA antiserum was kindly provided by Dr. Regina Santella. Biotin-labeled goat 139

anti-rabbit secondary antibody (Cat # 111-065-045) was purchased from Jackson 140

ImmunoResearch (West Grove, PA). Streptavidin-alkaline phosphatase conjugate (Cat 141

#21324) was a product of Pierce and purchased from Fisher Scientific. Standard BPDE-142

DNA adducts were prepared from highly purified calf thymus DNA (Sigma, St. Louis,143

MO) and benzo[a]pyrene-r-7,t-8-dihydrodiol-t-9,10-epoxide(),(anti) from MRIGLOBAL 144

Chemical Carcinogen Repository (Kansas City, MO) according to the procedures 145

described by (Jennette et al., 1977). 146

Cell Culture147

RPTEC/TERT1 cells and culture medium were purchased from Evercyte 148

Laboratories (Vienna, Austria), and grown according to Evercyte’s instructions. Cells 149

were cultured at 37oC in a humidified atmosphere containing 5% CO2. RPTEC/TERT1 150

cells were passaged approximately once or twice per week and subcultured at a 1:2 or 151

1:3 ratio. Cell culture vessels were purchased from Fisher Scientific and CellTreat®152

Scientific Products (Shirley, MA) and were tissue culture treated to promote adherent 153

cell growth.154

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Cell Exposure155156

Cd was dissolved in fresh complete medium and delivered at 0.1% of final 157

volume to give appropriate dose ranges. B[a]P was dissolved in DMSO and delivered at 158

0.05% final volume to give appropriate concentration ranges. B[a]P exposures were 159

conducted under low light conditions. Regardless of exposure format, final volume 160

percentage of each chemical was maintained. For co-exposure experiments, 1 µM Cd 161

was used to pre-treat cells for 24 hours before B[a]P exposure. The Cd pre-treatment162

concentration was determined based on previous characterization of the cell line’s 163

responses to various Cd concentrations. One micromolar Cd was the highest164

concentration tested that showed no significant cytotoxicity at 24 hours, 48 hours, or 1-165

week post exposure while demonstrating significantly increased cellular responses at 166

the level of the gene and protein (Simon, et al., 2014). 167

For DNA isolation, RPTEC/TERT1 cells were treated at confluence in T75cm2168

tissue culture treated flasks. After exposure time points, cells were washed twice with 169

cold 1X PBS, collected by centrifugation at 4oC, and stored at -80oC until DNA was 170

isolated. 171

Gene Expression 172

RPTEC/TERT1 cells were grown to confluence in 60mm dishes and exposed to 173

Cd or B[a]P as described above. Cells were exposed in triplicate for each concentration 174

and time point examined. Total RNA was isolated from cells after appropriate time 175

points using the QIAshredder (QIAGEN, 79656, Valencia, CA) and RNeasy extraction 176

kit (QIAGEN, 74136) following the manufacturer’s instruction. RNA concentration and 177

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purity was assessed using a Thermo Scientific Nanodrop 2000c spectrophotometer. 178

RNA samples were diluted to 0.5μg/μL in nuclease-free water. 179

Two microliters of each RNA sample were used for cDNA synthesis reactions to 180

deliver 1μg template in a 20μL total reaction volume. cDNA was synthesized using 181

iScript cDNA synthesis (BioRad, 170-8891, Hercules, CA) protocol as follows: 5 mins at 182

25oC, 30 mins at 42oC, and 5 mins at 85oC. RNA templates and cDNA were stored at -183

20oC until use. Gene expression was determined using primer-probe sets from Applied 184

Biosystems®TaqMan® Gene Expression Assays. Actin, beta (ACTB) was used as a 185

reference gene. Primers used are listed in Table 1. The thermal cycling protocol 186

followed the manufacturer’s instructions: 50oC for 2 min and 95oC for 10 min followed by 187

40 cycles of 95oC for 15s and 60oC for 1min. Reactions were conducted in 20μL 188

volumes with each sample being run in duplicate. All reactions were carried out using a 189

BioRad C1000™ thermal cycler equipped with a CFX96™ Real-Time PCR Detection 190

System.191

Table 1. Primer-probe sets used for RPTEC/TERT1 Gene Expression, Applied 192

Biosystems® TaqMan® Gene Expression Assays193

Gene IDGene

FunctionGene

LocationAssay ID

GCLC Antioxidant 6p12 Hs00155249_m1HMOX1 Antioxidant 22q13.1 Hs01110250_m1

NQO1Quinone

ReductionAntioxidant

16q22.1 Hs00168547_m1

ACTB Reference 7p22.1 Hs99999903_m1194

DNA Isolation195196

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Genomic DNA was isolated with a standard phenol chloroform extraction. 197

Briefly, cell pellets were thawed and incubated with 1X TE buffer, RNaseT1, mRNAse 198

A, and SDS for 45 minutes at 37oC. Pellets were incubated with proteinase K for 60 199

mins at 60oC and then overnight at 37oC. Deproteinized DNA was extracted using 5 200

PRIME Phase Lock Gel light, 15mL, tubes to increase yield from the aqueous phase. 201

Precipitated DNA was spooled onto a glass pipette, transferred to 70% ethanol, and 202

collected by centrifugation (18,000 rcf for 10 minutes). Ethanol was decanted and DNA 203

was allowed to dry completely before reconstituting in sterile, DNA grade water. DNA 204

concentration and purity was assessed using a Thermo Scientific Nanodrop 2000c 205

spectrophotometer.206

207

BPDE-DNA Adduct ELISA208

BPDE-DNA adducts were measured by a competitive ELISA method (Gammon209

et al., 2002; Mumford et al., 1996; Santella et al., 1988). Briefly, 96-well white210

microplates were coated by adding 50pg BPDE-substituted DNA in PBS to each 211

microwell. The DNA was sonicated and denatured in a boiling water bath for 5 min212

before coating. Plates were allowed to dry overnight and washed twelve times the next 213

day with washing buffer (1X PBS/0.05%Tween 20). All subsequent wash steps were 214

also performed twelve times. Plates were treated with I-Block (200 μL/well) for 90 min at 215

37oC to prevent non-specific binding. Standard curves and samples were prepared by 216

mixing and incubating with the previously characterized polyclonal BPDE-DNA antisera 217

at 1:3,000,000 in I-Block buffer (Mumford, et al., 1996). A 5-point standard curve was 218

used, in triplicate, to give a range of 0.312-10 fM adducts/well. Unknown samples were 219

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assessed at 10ug DNA per well in triplicate after sonication and denaturation. The plate 220

was washed after incubation with primary antibody, and a biotin-labeled goat anti-rabbit 221

secondary antibody (1:2,500 in I-Block) was incubated with in each well for 1 hour. After 222

an additional wash, the plate was incubated for one hour with streptavidin-alkaline 223

phosphatase conjugate (1:40,000 in I-Block). After one more wash step, the CPD-Star 224

Substrate with Emerald-II Enhancer was used to produce and amplify signal. 225

Luminescence was read with a Tecan Infinite® 200 PRO multimode reader (Tecan, San 226

Jose, CA). 227

Adducts were calculated for unknown samples based on percent inhibition of the 228

standard curve and expressed as average number of adducts per 108 nucleotides. Non-229

specific background signal detected in vehicle control groups was subtracted.230

Total Glutathione Assay231

After determined exposure time points, cells were trypsinized, collected, and 232

washed twice in 1X cold PBS. Total glutathione levels in cells were determined using 233

OxiSelect™ Total Glutathione (GSSG/GSH) Assay Kit (Cell Biolabs, Inc., San Diego, 234

CA) according to the manufacturer’s instructions. Cell isolates were diluted at 1:100 for 235

use within the linear range of the assay. 236

237Statistical Analysis 238

One- and two-way ANOVAs were performed using the GraphPad Prism 239

analytical software, version 6.0 (San Diego, CA). Data total glutathione assays were 240

analyzed using a one-way ANOVA and Dunnett’s multiple comparison tests. Data for 241

gene expression was analyzed using a two-way ANOVA and Tukey’s post hoc test. An 242

alpha of 0.05 was used as the criteria for determining significance. 243

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General linear models were used to test for differences among treatments, 244

treatment groups, and time points for the BPDE-DNA adduct ELISA. Where the initial 245

GLM analysis of variance (GLM-ANOVA) indicated a significant difference, post hoc246

mean comparisons were conducted using a Tukey correction. Statistical testing was 247

conducted using IBM SPSS Statistics version 19 software (Armonk, NY). An alpha of 248

0.05 was used as the criteria for determining significance.249

250251252

Results253254

BPDE-DNA adducts are formed and detected after exposure to B[a]P but altered after 255

co-exposure to B[a]P and Cd256

After 18 hours of exposure to B[a]P alone, BPDE-DNA adducts were detected in 257

RPTEC/TERT1 DNA samples. Although there appeared to be a dose-dependent 258

increase in adduct formation after 18 hours, exposure to 1 μM B[a]P was significantly 259

increased over DMSO vehicle control or lower concentrations, 0.01 and 0.1 μM B[a]P. 260

After 24 hours of exposure to B[a]P alone, adduct formation was most significantly 261

increased at 1 μM B[a]P in comparison to DMSO vehicle control, 0.01 and 0.1 μM B[a]P 262

at both 18 and 24 hours post exposure. Fewer adducts were detected after 24 hours of 263

exposure to 0.1 μM B[a]P in comparison to the same concentration at 18 hours although 264

the difference was not statistically significant (Figure 1, Table 2). 265

In order to assess the ability of Cd to alter adduct formation and persistence, 266

adducts were analyzed under conditions of Cd and B[a]P co-exposure. Cells were 267

exposed to Cd alone for 18 and 24 hours to verify the absence of adducts. There were 268

no BPDE-DNA adducts found above background at either time point after Cd exposure 269

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(data not shown). For co-exposure, cells were exposed to a non-cytotoxic concentration 270

of Cd, 1μM, for 24 hours. Cytotoxicity of each compound was based on previous work271

(Simon, et al., 2014). After 24 hours, cells were exposed to DMSO vehicle control or 272

appropriate concentrations of B[a]P for 24 hours. Adducts detected in groups exposed 273

to lower concentrations of B[a]P remained relatively unchanged between treatment 274

groups. However, cells exposed to 1 μM Cd x 1 μM B[a]P demonstrated significantly 275

reduced levels of adducts in comparison to 1 μM B[a]P alone at either time point (Figure 276

1, Table 2). 277

Figure 1 BPDE-DNA adducts are formed and detected in RPTEC/TERT1 cells after 278

exposure to B[a]P but reduced under co-exposure to 1 μM Cd and 1 μM B[a]P279

18 Hrs B[a]P 24 Hrs B[a]P 24 Hrs Cd x 24 Hrs B[a]P280

281

282

283

284

285

286

287

288

289

290

Figure 1. BPDE-DNA adducts are formed and detected in RPTEC/TERT1 cells after 291

exposure to B[a]P but reduced under co-exposure to 1 μM Cd and 1 μM B[a]P. After 18 292

DMSO

0.01

µM

0.1

µM1

µM

DMSO

0.01

µM

0.1

µM1

µM

DMSO

0.01

µM

0.1

µM1

µM0

20

40

60

80

B[a]P Concentration

Ad

du

cts

/10

^8

nu

cle

otid

es

*

Bars represent mean +/- SEM

* +

+

* +

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and 24 hours of treatment with B[a]P a significant increase in the number of adducts 293

was detected at 1 μM B[a]P. Treatment of cells for 24 hours with 1 μM Cd before a 24 294

hour exposure to concentrations of B[a]P showed a significant decrease in adducts 295

detected at 1 μM B[a]P in comparison to B[a]P alone. Bars represent average 296

adducts/108 nucleotides (n = 3) +/- SEM. * indicates significant difference from 1 μM 297

B[a]P at 18 hours, p < 0.01, + indicates significant difference from 1 μM B[a]P at 24 298

hours, p < 0.01. 299

300

Table 2. BPDE-DNA adducts formed after B[a]P and Cd exposure in RPTEC/TERT1 301

cells detected by ELISA302

303

Exposure Duration TreatmentAverage adducts/108

nucleotides +/- SEM, n = 3DMSO 0.0 +/- 0.15

0.01 μM B[a]P 0.58 +/- 0.370.1 μM B[a]P 9.92 +/- 2.66

18 hours B[a]P

1 μM B[a]P 36.34 +/- 9.14DMSO 0.0 +/- 0.21

0.01 μM B[a]P 0.0 +/- 0.820.1 μM B[a]P 5.72 +/- 1.43

24 hours B[a]P

1 μM B[a]P 59.75 +/- 17.03DMSO 0.0 +/- 0.16

0.01 μM B[a]P 1.18 +/- 0.140.1 μM B[a]P 7.88 +/- 1.33

24 hours 1 μM Cd x 24 hours B[a]P

1 μM B[a]P 17.28 +/- 1.30304305

Exposure to Cd increases expression of NRF2 responsive genes306

Gene expression changes of the NRF2 responsive genes, glutamate-cysteine 307

ligase, catalytic subunit (GCLC), heme oxygenase 1 (HMOX1), and NAD(P)H 308

dehydrogenase, quinone 1 (NQO1), were examined after exposure to determine if Cd309

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alone or Cd and B[a]P together appeared to induce an antioxidant response that may310

increase BPDE detoxification and reduce BPDE-DNA adduct formation under co-311

exposure conditions at 1 μM Cd x 1 μM B[a]P. While GCLC was detected, there was no 312

change among treatment groups after a 24-hour exposure to Cd (Figure 2A). After 24 313

hours of exposure to 0.1, 1, and 10 μM Cd, there was nearly a 3-fold increase in 314

HMOX1 at 10 μM Cd in comparison to untreated cells and all other concentrations 315

(Figure 2B). Additionally, all concentrations of Cd showed approximately a 2-3-fold 316

increase in NQO1 over that of untreated cells (Figure 2C). 317

Figure 2 RPTEC/TERT1 cells respond to 24 hour Cd exposure by upregulating HMOX1 318

and NQO1 but not GCLC. 319

Figure 2A320

0µM

0.1

µM1

µM

10µM

0

1

2

3

No

rma

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dF

old

Ex

pre

ss

ion

GCLC

Cd Concentration321

Figure 2B322

323324

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0µM

0.1

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10µM

0

1

2

3

HMOX1N

orm

aliz

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Fo

ldE

xp

res

sio

n

Cd Concentration

*

# ##

325326

Figure 2C327

0µM

0.1

µM

1µM

10µM

0

1

2

3

NQO1

No

rma

lize

dF

old

Ex

pre

ss

ion

Cd Concentration

+

328329

Figure 2. RPTEC/TERT1 cells respond to 24 hour Cd exposure by upregulating 330

HMOX1 and NQO1 but not GCLC. After 24 hours of treatment with Cd at various 331

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concentrations, RPTEC/TERT1 cells showed no change in (A) GCLC at any 332

concentration. There was a significant increase in gene expression at the highest 333

concentration, 10 μM Cd, of (B) HMOX1 and (C) NQO1. Bars represent mean fold 334

expression (n = 3) +/- SEM. All genes of interest were normalized to ACTB. 0 μM, 335

where denoted, was set as 1. * indicates significant difference from 0μM Cd, p < 0.01, # 336

indicates significant difference from 10μM Cd, p < 0.01, and + indicates significant 337

difference from 0μM Cd, p < 0.05.338

339

Twenty-four hours of B[a]P exposure did not increase gene expression of GCLC, 340

HMOX1, or NQO1 (Figures 3A, 3B, and 3C). All genes were detected at basal levels by 341

real time PCR. However, co-exposure significantly increased gene expression of all 342

three genes at the highest concentration of 1 μM Cd x 1 μM B[a]P over vehicle control 343

and other co-exposure groups. GCLC gene expression was increased by approximately 344

2-fold, HMOX1 gene expression was increased by approximately 3-fold, and NQO1345

gene expression was increased by approximately 4-fold (Figures 4A, 4B, and 4C). This 346

suggests that co-exposure, under these conditions, triggers a stronger transcriptional 347

antioxidant response than Cd alone.348

Figure 3. 24 hour B[a]P exposure does not induce changes in gene expression of 349

GCLC, HMOX1, or NQO1.350

351

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Figure 3A352

DMSO

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µM0

1

2

3

No

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dF

old

Ex

pre

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GCLC

B[a]P Concentration

Bars represent mean +/- SEM353354

Figure 3B355

DMSO

0.01

µM

0.1

µM1

µM0

1

2

3

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Figure 3C358

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µM0

1

2

3

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B[a]P Concentration


Figure 3. 24 hour B[a]P exposure does not induce changes in GCLC, HMOX1, or360

NQO1. None significantly differ. Bars represent mean fold expression (n = 3) +/- SEM. 361

All genes of interest were normalized to ACTB. DMSO, where denoted, was set as 1.362

Figure 4. Co-exposure conditions with Cd and B[a]P result in upregulation of GCLC, 363

HMOX1, and NQO1 in RPTEC/TERT1 cells.364

365366

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Figure 4A367

DMSO

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0.1

µM1

µM0

2

4

6

1 µM Cd x B[a]P Concentrations

No

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old

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GCLC

*# # #

368369

Figure 4B370

371

DMSO

0.01

µM

0.1

µM1

µM0

2

4

6

1µM Cd x B[a]P Concentrations

No

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Ex

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Figure 4C373

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µM1

µM0

2

4

6

NQO1

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1µM Cd x B[a]P Concentrations

Bars represent mean +/- SEM

*

# # #

374

Figure 4. Co-exposure conditions with Cd and B[a]P result in upregulation of GCLC, 375

HMOX1, and NQO1 in RPTEC/TERT1 cells. Cells were exposed to 1 μM Cd for 24 376

hours followed by a 24 hour exposure to B[a]P at different concentrations. 1 μM Cd was377

previously determined to be non-cytotoxic to RPTEC/TERT1 cells after 24 hours. 378

RPTEC/TERT1 cells demonstrated significant upregulation of (A) GCLC, (B) HMOX1, 379

and (C) NQO1 after exposure to 1 μM Cd x 1 μM B[a]P. Bars represent mean fold 380

expression (n = 3) +/- SEM. All genes of interest were normalized to ACTB. DMSO, 381

where denoted, was set as 1. * indicates significant difference from DMSO, p < 0.01, 382

and # indicates significant difference from 1μM Cd x 1μM B[a]P, p <0.01.383

384

Cd and Cd x B[a]P exposure increase total glutathione levels in RPTEC/TERT1 cells385

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Total glutathione was measured after Cd exposure for 24 hours and after co-386

exposure with B[a]P. Total glutathione levels were approximately double in cells treated 387

with 1 and 10 μM Cd in comparison to untreated groups after 24 hours. Cells treated 388

with 0.1 μM Cd exhibited a slight increase in total glutathione, but this increase was not 389

statistically significant. Total glutathione was also significantly increased in cells pre-390

treated with 1 μM Cd for 24 hours followed by exposure to 0.01, 0.1, and 1 μM B[a]P for 391

24 hours (Figure 5). This supports our hypothesis that Cd induces a biochemical 392

antioxidant response, and co-exposure to Cd and B[a]P results in a substantial increase 393

in reduced glutathione (GSH) levels possibly greater than those induced by Cd alone.394

Figure 5. Cd exposure increases total glutathione in RPTEC/TERT1 cells395

396397

0µM

0.1

µMCd

1µM

Cd

10µM

Cd

1µM

Cdx

0.01

µMB[a

]P

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]P

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Figure 5. Cd exposure increases total glutathione in RPTEC/TERT1 cells. Total 399

glutathione levels were significantly increased after 24 hours of exposure to 1 and 10 400

μM Cd. Total glutathione levels were also significantly increased after exposure to 1 μM 401

Cd for 24 hours followed by a 24 hour exposure to B[a]P concentrations. Bars represent 402

mean total glutathione levels (n = 3) +/- SEM. * indicates significant difference from 0 403

μM Cd, p < 0.05.404

405

406

Discussion407

The multifaceted effects that environmental mixtures have on the human body 408

have been notoriously problematic to resolve. For over a decade, scientists have faced 409

exceedingly difficult challenges in chemical toxicological research when studying 410

chemical mixtures designed to address gaps in our knowledge (Feron and Groten, 411

2002). However, the importance of pursuing chemical mixture experiments continues to 412

increase with the rise in diseases that have no clear genetic predisposition. 413

Assessments by the American Cancer Society credit heritable mutations in the 414

development of only 5% of all cancers (Cancer Facts and Figures 2012, 2012). 415

Likewise, current evidence suggests that an overwhelming 90% of human disease 416

burden, especially degenerative conditions, can be attributed to environmental factors 417

such as exposure, lifestyle, and diet (Anand et al., 2008; Rappaport, 2011; Rappaport, 418

2013; Sung et al., 2011). Humans encounter mixtures of chemical compounds daily 419

and throughout their lives; however, relatively little research to date has aimed to420

address the differential effects that mixtures have on molecular and mechanistic 421

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endpoints in comparison to studies focused on individual chemicals and compounds. A 422

recent review on PAH mixtures toxicology illustrates these complexities but also 423

provides strong rationale for approaching these issues (Jarvis et al., 2014). Biological 424

processes and molecular factors that counteract the development of cancer (e.g. 425

increased antioxidant or detoxification capacity) must be studied in the context of 426

exposure to these mixtures. Environmental toxicants that interfere with efficient 427

processing and accurate repair of DNA adducts may increase the mutagenicity of other 428

toxicants by decreasing DNA repair capacity. In this context, such environmental 429

toxicants function as co-carcinogens. 430

In an effort to characterize the effects of a simple binary mixture on renal 431

proximal tubule cells, we have examined cellular responses of the RPTEC/TERT1 432

immortalized cell line to B[a]P and Cd, two distinctly different carcinogens. Our previous 433

studies with this cell line have demonstrated its sensitivity to both B[a]P and Cd as well 434

as compound-specific responses (Simon, et al., 2014). Here, we confirm the 435

metabolism of B[a]P to metabolites which form DNA adducts under these conditions. 436

We detected BPDE-DNA adducts at 18 and 24 hours post exposure to B[a]P alone. At 437

24 hours post exposure, there were fewer adducts detected at intermediate 438

concentrations, 0.01 and 0.1 μM B[a]P, than at 18 hours post exposure. While the 439

numbers of adducts were not significantly reduced, the decrease at these 440

concentrations suggests that there may be some removal or repair of the initial adducts. 441

However, the limited sensitivity of the ELISA method at the lower concentrations of 442

B[a]P tested in these experiments makes these suggestions speculative. At the highest 443

concentration of B[a]P (1 μM) tested alone, we found that significantly greater adduct 444

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levels remained at both 18- and 24-hour time points. At this concentration of B[a]P, 445

detoxification and repair mechanisms may have been unable to process the amount of 446

B[a]P metabolites in the examined time period. There may a threshold effect, possibly 447

short-term but in excess of 24 hours, in which bioactivation exceeds detoxification and 448

repair, which generates BPDE-DNA adducts at a rate greater than the rate at which 449

DNA repair processes can remove them. This conclusion is warranted especially if 450

B[a]P treatments alone in this experiment do not induce an antioxidant response, 451

increase detoxification capacity, or increase DNA repair capacity. In contrast, when cells 452

were pre-treated with 1 μM Cd, BPDE-DNA adducts formed after 1 μM B[a]P exposure 453

were significantly reduced. This effect was not observed to this degree at other454

concentrations of B[a]P after Cd pre-treatment. It is possible that CYP-mediated 455

biotransformation at the lower concentrations of B[a]P was occurring without exceeding 456

detoxification and/or DNA repair capabilities. This would allow cellular mitigation of 457

BPDE-DNA adducts without an increase in induction as supported, in part, by our 458

previous work (Simon, et al., 2014). However, as mentioned previously, the sensitivity of 459

the ELISA method at lower concentrations of B[a]P used in these experiments may not 460

be adequate to distinguish statistically significant differences in BPDE-DNA adduct 461

levels among the treatment and co-treatment groups with adequate precision. We 462

suspect that B[a]P is metabolized at the lower concentrations, but more sensitive 463

analytical methods are necessary to discriminate significant differences in adduct 464

formation and persistence based on treatment regimen. We suggest that future 465

experiments be designed to address such experimental possibilities and statistical 466

power limitations. 467

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The reduction in adducts under co-exposure at the highest concentration of B[a]P 468

may be a function Cd x B[a]P priming the detoxification system through the NRF2 469

antioxidant pathway. These effects appear to result in increased levels of glutathione470

and increased inactivation or detoxification of BPDE prior to adduct formation. Future 471

measurements of BPDE-DNA conjugates in conjunction with glutathione levels should 472

be used to confirm this supposition. B[a]P exposure alone, under these conditions, does 473

not appear to induce any such response. While it appears that Cd alone induces a 474

significant antioxidant response resulting in increased levels of glutathione, Cd and 475

B[a]P together at the highest concentrations tested induce an even more robust 476

response. This response to both Cd and B[a]P in our experiments seems to mitigate 477

DNA adduction formation through enhanced detoxification capacity. Our previous work 478

does not support one alternative explanation that Cd inhibits the formation of BDPE 479

through CYP feedback inhibition (Simon, et al., 2014).480

The binding of B[a]P to the aryl hydrocarbon receptor increases the expression of 481

xenobiotic response element (XRE) genes and their encoded enzymes, which are 482

responsible for metabolizing B[a]P to reactive intermediates (Shimada and Fujii-483

Kuriyama, 2004; Xue and Warshawsky, 2005). These reactive intermediates, along 484

with reactive oxygen species (ROS) from heavy metals, can increase the transcription485

of antioxidant response element (ARE) genes through NRF2 binding (Aleksunes and 486

Manautou, 2007; Nguyen et al., 2009). Activation of this gene battery may be 487

responsible for metabolite detoxification. We suspect that this process, under our 488

experimental co-exposure conditions, reduced level of adducts detected at 1 μM B[a]P 489

following 1 μM Cd pre-treatment. We found the expression of NRF2-targeted genes, 490

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GCLC, NQO1, HMOX1, to be significantly increased under experimental conditions 491

coinciding with the most significant reduction in BPDE-DNA adducts under co-exposure. 492

Our results are similar to in vivo studies which have discovered that priming the NRF2 493

system decreases the levels of adducts formed after B[a]P exposure. Nrf2 knockout 494

mice develop more tumors than wild-type mice when treated with B[a]P alone. When 495

mice are given a Nrf2 activator with B[a]P, wild-type mice develop half as many tumors. 496

However, tumor reduction is not seen in Nrf2 knockout mice given a Nrf2 activator with 497

B[a]P exposure (Aleksunes and Manautou, 2007). In other studies including 498

transformed kidney cell lines from humans and rats, Cd has been shown to induce the 499

NRF2 pathway through an oxidative stress mechanism (Chen and Shaikh, 2009; He et 500

al., 2008; Wilmes et al., 2011). Future studies in the RPTEC/TERT1 cell line should 501

consider the application of a NRF2 inhibitor to mimic in vivo Nrf2 knockout conditions 502

and further verify the responses seen after B[a]P exposure. Several NRF2 activating 503

agents, both natural and synthetic, have been examined as chemoprotectives for 504

chronic disease and overall cell health. Flavonoids, for example, are naturally occurring 505

antioxidants found in cruciferous vegetables, apples, and onions. They have been 506

shown to increase NRF2 mediated expression of NQO1 and GST. Additionally, naturally 507

occurring phytochemicals such as chalcones and coumarins have been shown to act 508

similarly by inducing NRF2 expression of NQO1 and GST to act as anti-inflammatories509

and antioxidants (Kumar et al., 2014). Similarly, bardoxolone methyl, a synthetic NRF2 510

activator derived from natural antioxidants, has been successful in increasing kidney 511

function and halting the progression of renal injury in Phase 2 clinical trials in patients 512

with chronic kidney disease (Ruiz et al., 2013).513

https://www.researchgate.net/publication/234157678_Targeting_the_Transcription_Factor_Nrf2_to_Ameliorate_Oxidative_Stress_and_Inflammation_in_Chronic_Kidney_Disease?el=1_x_8&enrichId=rgreq-cc7f000a-faa0-4eb7-a1e7-21e56e371142&enrichSource=Y292ZXJQYWdlOzI2Mzg5MjE1ODtBUzoxMjUwMjIyNjE3NDc3MTJAMTQwNjgxOTAzODgxMQ==

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Our goals were to measure responses in RPTEC/TERT1 cells to defined, non-514

cytotoxic mixtures of two distinctly different toxicants. While our results suggest an 515

antagonistic, or perhaps a hormetic effect, on the endpoint examined, DNA adduct 516

formation and persistence, further studies are necessary to explore these results and 517

determine effects on other downstream biomarkers. Of particular interest is the 518

mutagenicity of BPDE-DNA adducts under such conditions of co-exposure especially at 519

lower, environmentally relevant concentrations. We hypothesize that increased 520

detoxification capacity is responsible for the reduced levels of BPDE-DNA adducts521

which may protect cells from these premutagenic lesions. This could be interpreted as a 522

hormetic effect (Calabrese, 2008). It is also possible that, while detoxification capacity is 523

increased, subsequent DNA repair is inhibited by the presence of Cd. Cd inhibition of 524

DNA repair may promote repair mistakes or error-prone translesion synthesis of the 525

remaining adducts leading to a relative increase in mutagenicity under these conditions526

(for a review, see Hartwig, 2013).527

Apparent NRF2 activation and the increased total glutathione levels found in Cd 528

and co-exposure groups are evidence of cellular oxidative stress. Cd and Cd 529

compounds are Group 1 carcinogens and are known to cause cancer in humans (IARC 530

Working Group on the Evaluation of Carcinogenic Risks to Humans. and International 531

Agency for Research on Cancer., 2012). DNA damage caused indirectly by Cd, such as 532

oxidative insult and repression of DNA damage repair, must be considered in mutational 533

investigations. Quantifying the levels of PAH-DNA adducts in human studies can serve 534

as a biomarker for exposure as well as provide information on an individual’s DNA 535

repair capacity and mutagenic risk (Gammon, et al., 2002). However, it would be ideal 536

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to also measure mutation frequency to confirm mutagenic potential as a function of 537

adduct formation under controlled conditions with in vitro and in vivo models to better 538

represent and understand mechanisms by which mixtures impact humans.539

We have conducted these studies in the RPTEC/TERT1 human immortalized cell 540

line because they were derived from a normal, healthy individual, have proven to be 541

metabolically competent, and exhibit canonical responses similar to human kidney cells 542

when exposed to the selected environmental toxicants. However, we acknowledge the 543

difficulties in carrying out robust, controlled experimentation on chemical mixtures. Due 544

to the complicated nature of mixtures toxicology, it remains challenging to extrapolate 545

the results obtained in this or any in vitro or in vivo model to actual human risk. 546

Nevertheless, in vitro models permit higher throughput screening of many compounds 547

and mixtures. As they improve to better model the tissue of interest, in vitro models will 548

prove extremely valuable in mixtures toxicology. Advancements in high throughput 549

screening have allowed scientists to begin to elucidate increasingly complex 550

mechanisms and interactions inherent to mixtures toxicology. As research continues on 551

both environmental and genetic components of disease, the causes of conditions such 552

as cancer are becoming more recognized as environmentally mediated. Thus, it can be 553

hypothesized that the majority of genetic changes resulting in cancer are acquired over 554

a lifetime through one’s interactions with the environment. Therefore, it is critical to 555

better understand the molecular processes that contribute to the initiation of cancer. 556

557

Funding558

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Funding was provided in part by a generous grant from the Baton Rouge Area 559

Foundation, Baton Rouge, LA. Funding was also provided in part by a grant and 560

cooperative agreement from the NIH/NIEHS 1U19ES20677-01. Its contents are solely 561

the responsibility of the authors and do not necessarily represent the official views of the 562

NIEHS or NIH. Funding and support was also provided by the Tulane Cancer Center 563

and the Louisiana Cancer Research Consortium.564

565

Acknowledgements566

We would like to thank Dr. Regina M. Santella, Professor of Environmental Health 567

Sciences, Mailman School of Public Health at Columbia University, New York, NY, for 568

training in the BPDE-DNA adduct ELISA method and for generously providing critical 569

reagents used in the assay.570

571

572

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Cadmium alters the formation of benzo[a]pyrene DNA adducts in the RPTEC/TERT1 human renal proximal...

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