Haloacetic Acid Water Disinfection Byproducts Affect Pyruvate

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Haloacetic Acid Water Disinfection By-Products Affect Pyruvate Dehydrogenase Activity and Disrupt Cellular Metabolism AZRA DAD, Clara Hyunhee Jeong, Elizabeth D. Wagner, and Michael J Plewa Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b04290 • Publication Date (Web): 20 Dec 2017 Downloaded from http://pubs.acs.org on December 24, 2017

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Environmental Science & Technology

HALOACETIC ACID WATER DISINFECTION BY-PRODUCTS AFFECT PYRUVATE DEHYDROGENASE ACTIVITY AND DISRUPT CELLULAR METABOLISM

Azra Dad1**,*, Clara H. Jeong2,3 , Elizabeth D. Wagner1, Michael J. Plewa1

1. Safe Global Water Institute, University of Illinois at Urbana-Champaign and the Department of Crop Sciences, University of Illinois at Urbana-Champaign. 2. Molecular and Environmental Toxicology Center, School of Medicine and Public Health, University of Wisconsin-Madison, Madison, WI, USA. 3. Department of Urology, University of Wisconsin-Madison, Madison, WI, USA.

**Current address: Division of Genetic and Molecular Toxicology, National Center for Toxicological Research, US Food and Drug Administration, Jefferson, AR, USA

*Corresponding Author: Azra Dad, [email protected]

ACS Paragon Plus Environment

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ABSTRACT

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The disinfection of drinking water has been a major public health achievement. However,

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haloacetic acids (HAAs), generated as by-products of water disinfection, are cytotoxic,

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genotoxic, mutagenic, carcinogenic and teratogenic. Previous studies of monoHAA-

5

induced genotoxicity and cell stress demonstrated that the toxicity was due to inhibition

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of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) leading to disruption of cellular

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metabolism and energy homeostasis. DiHAAs and triHAAs are also produced during

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water disinfection, and whether they share mechanisms of action with monoHAAs is

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unknown. In this study, we evaluated the effects of mono-, di-, and tri-HAAs on cellular

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GAPDH enzyme kinetics, cellular ATP levels, and pyruvate dehydrogenase complex

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(PDC) activity. Here, treatments conducted in Chinese hamster ovary (CHO) cells

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revealed differences among mono-, di-, and triHAAs in their molecular targets. The

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monoHAAs, iodoacetic acid and bromoacetic acid, were the strongest inhibitors of

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GAPDH and greatly reduced cellular ATP levels. Chloroacetic acid, diHAAs and

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triHAAs were weaker inhibitors of GAPDH and some increased the levels of cellular

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ATP. HAAs also affected PDC activity, with most HAAs activating PDC. The primary

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finding of this work is that mono- versus multihalo HAAs address different molecular

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targets and the results are generally consistent with a model in which monoHAAs

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activate the PDC through GAPDH inhibition-mediated disruption in cellular metabolites,

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including altering ATP:ADP and NADH:NAD ratios. The monoHAA-mediated

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reduction in cellular metabolites results in accelerated PDC activity by way of

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metabolite-ratio-dependent PDC regulation. DiHAAs and triHAAs are weaker inhibitors

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of GAPDH, but many also increase cellular ATP levels, and we suggest that they increase

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PDC activity by inhibiting pyruvate dehydrogenase kinase.


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INTRODUCTION Disinfection of drinking water was one of the major public health achievements of

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the 20th century, significantly reducing the outbreak of water-borne diseases including

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cholera, typhoid and dysentery.1 Disinfectants are oxidants that not only kill pathogenic

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microorganisms but also generate disinfection by-products (DBPs). 2 Currently over 600

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DBPs have been identified.3 In chlorinated water, the second largest DBP chemical class

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is the haloacetic acids (HAAs).4, 5 The U.S. Environmental Protection Agency (EPA)

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regulates five HAAs (chloroacetic acid (CAA), dichloroacetic acid (DCA),

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trichloroacetic acid (TCA), bromoacetic acid (BAA), and dibromoacetic acid (DBA)) to a

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total maximum contaminant level of 60 µg/L.6

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HAAs as a class of DBPs expressed different levels and types of toxicity based on

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their chemical structures.3, 7-11 DBP exposure is associated with increased risks of bladder

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and colorectal cancer,9 and spontenous abortion and other negative pregnancy

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outcomes.10 DCA, TCA, and DBA induced hepatocellular adenomas and hepatocellular

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carcinomas in B6C3F1 mice and F344 rats11, and gestational exposure of a mixture of

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regulated HAAs resulted in pregnancy loss and eye malformation in rats.8 HAAs were

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cytotoxic and genotoxic in Chinese hamster ovary (CHO) cells 12, mutagenic in

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Salmonella typhimurium and CHO cells,13-15 and cytotoxic and genotoxic in primary

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human cells.16 Results from in vitro studies have provided information on the molecular

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mode of toxicity mechanisms of monoHAAs.16-18

45 46

HAAs are SN2 alkylating agents15 and thus capable of binding cellular macromolecules. Previously, we demonstrated that monoHAAs do not directly damage


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DNA, but inhibit glyceraldehyde 3-phosphate dehydrogenase (GAPDH), by alkylating

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the sulfhydryl group of the cysteine residue in the active site, rendering the enzyme

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inactive.16, 18 GAPDH is a pivotal enzyme in glycolysis and the HAA-induced inhibition

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of GAPDH activity can affect pyruvate generation, which is required by mitochondrial

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metabolism. Alterations in mitochondrial metabolism can lead to elevated levels of

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reactive oxygen species (ROS) that can damage DNA.19, 20 Treating cells with exogenous

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pyruvate overcame the monoHAA –mediated reduction in cellular ATP and reduced

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genomic DNA damage.17 We hypothesized that di-, and triHAAs may follow the same

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toxicity pathway by inhibiting GAPDH and reducing cellular ATP levels. Previous

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studies demonstrated that DCA was used as a treatment for patients with lactic acidosis as

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a result of pyruvate dehydrogenase (PDH) deficiency. 21 DCA also has been used as a

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chemotherapeutic agent because it reverses the Warburg effect 22 by activating PDH

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through inhibition of PDH kinase. 21, 23-25

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The pyruvate dehydrogenase complex (PDC) is a multi-enzyme complex 26 that is

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responsible for catalyzing the conversion of pyruvate, NAD+, and coenzyme-A into

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acetyl-CoA, NADH and CO2 in the presence of thiamine pyrophosphate and Mg+2.27

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Mammalian PDC is made up of multiple isoforms of 6 different components: pyruvate

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dehydrogenase (E1), dihydrolipoamide acetyltransferase (E2), dihydrolipoamide

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dehydrogenase (E3), E1 specific kinases, phospho-E1 phosphotase, and X protein.26, 28

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E1, E2, and E3 are the major catalytic components, and are responsible for the stepwise

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decarboxylation and dehydrogenation of pyruvate into acetyl- CoA. 29

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The E1 component of PDC is a tetramer (α2β2) with two thiamine pyrophosphate (TPP) binding sites. The E1α component is inactivated/phosphorylated by pyruvate


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dehydrogenase kinase (PDK) and activated/dephosphorylated by phospho-E1-

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phosphatase. Dephosphorylation by phospho-E1-phosphatase leads to the activation of

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the enzyme.29, 30 E2, besides generating acetyl-CoA, also has L1 and L2 lipoyl domains

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that facilitate the binding of PDK and phospho-E1-phosphatase and thus plays a crucial

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role in the activation and inactivation of these enzymes. 29, 31

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Kato and colleagues found that rat PDK2 had a unique component known as the

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“ATP lid”. The binding of ATP lid to the L2 domain of PDC-E2 regulated PDK activity

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by inducing allosteric conformational changes in the ATP lid changing its affinity for

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ADP. They demonstrated that unbound ATP lid had a higher affinity for ADP, rendering

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PDK inactive, and that bound ATP lid promoted PDK activity. 32

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Kato and colleagues also found that DCA reduced PDK1 and PDK3 activity to

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4% and 17% of the control, respectively.33 Bonnet and colleagues demonstrated that

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DCA induced apoptosis in tumor cells by reversing the Warburg effect; DCA activated

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the PDC by inhibiting PDK and shifting the glycolytic-based metabolism to mitochondria

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in tumor cells.34

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These studies indicated that the PDC is regulated by PDK. However, the cellular

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ATP/ADP ratio also plays a significant role in regulating PDK, PDC and overall

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mitochondrial metabolism.35

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We propose that HAAs disrupt metabolism by activating PDC either by

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inhibiting GAPDH or PDK, depending on the HAA halogenation level (mono-, di-, and

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tri-HAAs). We also hypothesize that these two mechanisms of HAA-induced PDC

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activation, through either GAPDH inhibition or PDK inhibition, operate independently.


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To test this hypothesis, we conducted a series of studies in Chinese hamster ovary (CHO)

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cells to (i) measure GAPDH inhibition following HAA-treatment; (ii) determine the

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effect of HAA-treatment on PDC activity; and (iii) measure the impact of HAA exposure

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on cellular ATP levels. CHO cells were employed in this study since they were used in

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developing the largest database on chronic cell cytotoxicity and genomic DNA damage

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induced by DBPs12.36


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MATERIALS AND METHODS

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Reagents

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DBA, DCA, tribromoacetic acid (TBA), TCA, and CAA were purchased from

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Fluka (St. Louis, MO); iodoacetic acid (IAA) and BAA were obtained from Sigma (St.

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Louis, MO), dibromochloroacetic acid (BDCA) from Radian International (Austin, TX),

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and DBCA from Cerilliant (Round Rock, TX); bromochloroacetic acid (BCA) was from

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the U. S. EPA. Nutrient mixture F12 and fetal bovine serum (FBS) were purchased from

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Fisher Scientific (Itasca, IL). Cell Titer-Glo reagent was obtained from Promega

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(Madison, WI), and the Pyruvate Dehydrogenase (PDH) Activity Colorimetric Assay Kit

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was purchased from BioVision (San Francisco, CA). Glyceraldehyde-3-phosphate was

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purchased from Sigma-Aldrich (St. Louis, MO). Other reagents were obtained from

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either Fisher Scientific or Sigma in the highest purities available. Stock HAA solutions

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were dissolved in dimethylsulfoxide (DMSO) and stored at −20°C in sealed sterile glass

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vials (Sulpelco, Bellefonte, PA). Working HAA solutions were diluted in Hank’s

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balanced salt solution (HBSS) to the appropriate concentration.

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CHO Cell Culture and HAA Treatment The CHO cell line AS52, clone 11-4-8 (derived from CHOK1 cells) was used in

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all experiments. 37-39 Routine CHO cultures were maintained in 100-mm glass petri plates

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containing F12 medium supplemented with 5% FBS, 1% glutamine and 1% antibiotic-

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antimycotic (F12 complete medium) solution at 37°C in a humidified atmosphere of 5%

119

CO2 in air.


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Noncytotoxic concentrations of HAAs were used for treatments in order to

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measure endpoints without the confounding factor of overt cytotoxicity. The test doses

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were based on the dose response cytotoxicity data reported by Plewa et al. (2010)12 and

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preliminary range-finding experiments conducted for this study (data not shown). When

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the HAAs produced alterations in the endpoints that were measured (e.g., activation of

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PDC), the lowest noncytotoxic doses producing significant responses were chosen for

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making comparisons among HAAs.

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GAPDH Enzyme Kinetics Analysis

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The GAPDH enzyme kinetics analysis method was adapted from (Pals et

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al.2011).18 CHO cells were grown in F12 complete medium on 60-mm plastic cell culture

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plates until the cells reached confluency. The medium was removed and the cells were

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washed 3× with 2 mL cold phosphate-buffered saline (PBS) and once with 2 mL cold

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Buffer K (1 mM Tris-HCl, 1 mM EDTA, 1 mM MgCl2, pH 7.6). The cells then were

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tumefied with Buffer K for 2 min at ambient temperature, followed by homogenization

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with a sterilized cell scraper. The cell homogenates were collected in microfuge tubes,

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centrifuged at 16,100 × g for 3 min at room temperature to remove cellular debris, and

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the supernatants transferred to a fresh tube. Next, 854 µL of reaction buffer (0.1 M Tris-

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HCl (pH 8.5), 5 mM KH2PO4, 20 mM NaF, 1.7 mM NaAsO2, and 1 mM NAD), and 220

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µL of a 5× solution of the HAA test agents were added to 22 µL cell homogenates. The

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mixture was vortexed, and incubated for 2 min, and 996.3 µL of the assay mixture was

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transferred to a quartz spectrophotometer cuvette, and the instrument was zeroed at 340

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nm. Finally, 3.7 µL of 1 mM glyceraldyde-3-phospate (G3P) was added to the assay

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mixture to initiate the GAPDH-dependent NAD+ to NADH conversion; NADH


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absorption was measured at 340 nm every 10 s for a total of 60 s using a Beckman

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DU7400 UV-vis spectrophotometer. Each experiment included a negative control and

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three replicates of all HAA test agent concentrations. Absorbance values were plotted

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against time for each individual experiment and a linear regression was fitted to the data

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to calculate the GAPDH kinetic rates. Cell homogenate protein levels were obtained as

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described below. GAPDH activity was calculated as µMol NADH/min/µg protein. Data

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were compared to the concurrent negative control for each experiment and the percent

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reduction in the GAPDH activity per µM of each HAA was calculated. ANOVA with

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Holm-Sidak pairwise comparisons were performed to compare the reduction in the

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GAPDH activity among mono-, di- and triHAAs.

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ATP Analysis and Protein Quantification

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Parallel ATP and protein analyses were performed following HAA treatments.

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Cellular ATP levels were measured using the Promega Cell Titer-Glo reagent in a 96-

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well microplate assay and using a TUNE-SpectraMax Paradigm® Multi-Mode

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Microplate Detection Platform. The day before ATP analysis, 3×104 CHO cells/well were

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cultured in an opaque 96-well microplate in 200 µL of F12 plus 5% FBS. The next day,

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the cells were washed with 100 µL of HBSS and treated with monoHAAs (final

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concentrations of 3 µM IAA, 15 µM BAA, or 300 µM CAA); diHAAs (900 µM DBA,

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900 µM BCA, or 50 µM DCA); or triHAAs (300 µM TCA, 300 µM CDBA, 500 µM

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TBA, or 900 µM BDCA) in 50 µL HBSS (supplemented with 1.3 mM CaCl2 and 1.1 mM

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MgSO4). The microplate was covered with AlumnaSeal and incubated for 4 h at 37°C

164

with 5% CO2. Each experiment contained a concurrent negative control, a

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bioluminescence background control, HAA-treated cultures, and wells for an ATP


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standard curve. After 4 h incubation, the cells were washed with 100 µL of HBSS and

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equilibrated to room temperature for 30 min. The ATP content of the cells was measured

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according to the manufacturer’s protocol using 50 µL of Cell Titer-Glo ATP reagent.

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For protein quantification, 3×104 CHO cells/well were cultured in a 96-well flat-

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bottom clear microplate in 200 µL F12 complete media the day before the experiment.

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The next day, the cells were treated with the HAAs as described previously for the ATP

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analysis. After treatment, the protein was quantified in the cell lysates as described in

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Dad et al. (2013).17

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PDH Enzyme Kinetics Analysis

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CHO cells were seeded in 6-well plates at a concentration of 6×106 cells/2 mL of

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F12 complete medium/well and incubated overnight at 37°C, in a humidified atmosphere

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of 5% CO2 in air. The following day, the medium was aspirated and the cells were

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washed twice with 2 mL HBSS at 37°C. The cells were then treated with monoHAAs

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(final concentration of 3 µM IAA, 15 µM BAA, or 300 µM CAA); diHAAs (900 µM

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DBA, 900 µM BCA, or 50 µM DCA); or triHAAs (300 µM TCA, 300 µM CDBA, 500

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µM TBA, or 900 µM BDCA) in HBSS (supplemented with 1.3 mM CaCl2 and 0.5 mM

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MgCl2) and incubated for 4 h at 37°C. Following the treatment interval, the cells were

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washed once with HBSS at ambient temperature and harvested with 0.05% trypsin.

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Harvested cells from different wells treated with the same concentration of each HAA

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were pooled into a single tube and centrifuged at 16,100 ×g for 1 min; the supernatant

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was transferred to a new tube.


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PDH activity was measured using a BioVision PDH colorimetric assay kit. The

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cell pellets were washed with 500 µL HBSS and homogenized in 200 µL PDH buffer

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using a motorized tissue grinder (Fisher). Homogenates were incubated on ice for 10 min,

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followed by centrifugation at 10,000 × g for 5 min at 4°C. A 50 µL sample of the

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supernatant from each treatment group was transferred to a 96-well microplate, 50 µL of

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the kit reaction buffer were added to each well, and absorbance was measured

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immediately with a TUNE-SpectraMax Paradigm® Multi-Mode Microplate Detection

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Platform (Molecular Devices, Sunnyvale, CA). Measurements were made at 450 nm in

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kinetic mode for 60 min at 37 °C at 10 min intervals. Each treatment group had three

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replicates for every experiment and each experiment was repeated at least three times. An

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NADH standard curve was used to determine the PDH activity as nmol NADH/min/mL

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using the formula:

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Sample PDH Activity = B/(∆T × V) × D = nmol/min/mL

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Where: B = NADH calculated from the standard curve (nmol)

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∆T = reaction time (min)

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V = Sample volume in the reaction well (ml)

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D = Dilution factor

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The amount of protein measured in each sample was determined using the

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Bradford assay and was used to calculate nmol NADH/min/mg protein. ANOVA with a

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Holm-Sidak post-hoc test determined statistical significance among the HAA treatments.


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RESULTS AND DISCUSSION

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HAAs as a class of DBPs expressed different levels and types of toxicity,

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determined by their individual chemical structure.3, 7 The mechanisms responsible for the

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toxicity of HAAs, and the differences among the molecular targets and toxicity

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mechanisms of mono-, di-, and triHAAs, are not well understood. Previously, we found

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that monoHAAs disrupt cellular energy homeostasis by inhibiting GAPDH. It is

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unknown if di-, and triHAAs share this toxicity mechanism with monoHAAs. We believe

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that PDH plays a central role in the HAA mechanism of action that leads to disruption in

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metabolism and cellular energy homeostasis.

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PDH plays a pivotal role in mitochondrial metabolism as part of a multi-enzyme

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PDC and produces reducing power for oxidative phosphorylation, thereby linking

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glycolysis with the citric acid cycle (CAC). 27 The activity of the PDC is regulated by

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homeostasis in ATP:ADP, NADH:NAD, and Acetyl-CoA :CoA metabolic precursor

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ratios. 24 In addition to metabolic precursor ratios, PDK and phospho-E1-phosphatase

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also regulate PDC. 25, 32

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The E1α component of the tetrameric E1 component is inactivated/phosphorylated

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by PDK and activated/dephosphorylated by a specific phosphatase, phospho-E1-

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phosphotase.29, 30 In humans, mutations in the X-linked PDHA1 gene that encodes the

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E1α subunit result in PDH deficiency disease.40 PDH deficiency disease is characterized

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by reduced mitochondrial function and is associated with two forms of abnormalities: a

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metabolic form (lactic acidosis) and a neurological form (neurodevelopment delay and

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hypotonia). 41 Studies indicate that PDH is also involved in oncogene-induced


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senescence.34, 42 Activation of PDH enhances pyruvate utilization resulting in increased

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respiration and redox stress. 34

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Previous studies indicate that the di-HAA, DCA, inhibits PDK by inducing

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allosteric changes, resulting in increased PDC activity.32 Thus, DCA was used to activate

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the PDC to treat different disease states caused by PDH deficiency.43, 44 It was reported

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that the biotransformation of diHAAs to glyoxylate is dependent on cytosolic glutathione

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S-transferase-zeta (GST-zeta)45-47. Glyoxylate can undergo transamination,

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decarboxylation, and oxidation to form glycine, carbon dioxide and oxalate, respectively.

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DiHAAs can also induce toxicity by generating intermediate compounds. Glyoxylate can

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induce toxicity by covalently reacting with N-terminal amino groups of proteins or by

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acting as a suicide substrate for GST-zeta47, 48. Since GST-zeta is identical to

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maleylacetoacetate isomerase (the enzyme that catalyzes tyrosine catabolism) its diHAA-

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induced inactivation leads to the buildup of maleylacetoacetate and maleylacetone,

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which can cause tissue damage due to oxidative stress and by reacting with cellular

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nucleophiles. Therefore, the use of DCA to cure PDH deficiency potentially can induce

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toxicity by deactivating tyrosine catabolism, although it is generally regarded as well-

245

tolerated.49, 50

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When monoHAAs are involved in blocking glycolysis by inhibiting GAPDH,

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their cytotoxicity and genotoxicity occur through oxidative stress16 resulting from

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GAPDH inhibition. 18, 51 Addition of pyruvate or treatment with antioxidants relieves cell

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stress and reduces genotoxicity.17, 52 Stalter et al (2016) demonstrated that HAAs induced

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oxidative stress, depleted reduced glutathione (GSH) levels, and activated p53, thus

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confirming indirect HAA-induced DNA damage linked to GAPDH inhibition.53


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These mechanisms, however, were established with monoHAAs; any differences

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among the toxicity mechanisms for mono-, di-, and triHAAs are unknown. We postulate

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that di-, and triHAAs inhibit PDK and activate PDC leading to increased cellular ATP

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levels. This contrasts with the monoHAAs, IAA and BAA, which are direct inhibitors of

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GAPDH and greatly reduce ATP levels leading to ATP/ADP ratio disruption that require

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cells to accelerate the CAC by increasing PDC activity. To test this hypothesis, we

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analyzed the effect of a series of monoHAAs, diHAAs, and triHAAs on GAPDH

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kinetics, cellular ATP levels, and PDC activity.

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Effect of HAAs on GAPDH Kinetics

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GAPDH is pivotal in glycolysis and plays a crucial role in cellular energy

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homeostasis. A GAPDH kinetic study measuring NADH levels was performed on

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homogenates from CHO cells treated with HAAs (monoHAAs: 25 µM IAA, 150 µM

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BAA, or 5 mM CAA; diHAAs: 2 mM DBA, 2 mM BCA, or 2 mM DCA; or triHAAs: 2

265

mM TCA, 2 mM CDBA, 2 mM TBA, or 2 mM BDCA) for 2 min. The results indicated

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that the monoHAAs IAA and BAA are strong direct inhibitors of GAPDH (Figure 1).

267

The data in Figure 1 were plotted as the index values of percent of GAPDH inhibition per

268

µM of HAA. The data were normalized as percent reduction of the negative control.

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Since IAA and BAA were the strongest inhibitors of GAPDH, lower concentrations were

270

used for these two HAAs as compared to other HAAs. To make the comparison among

271

different HAAs more appropriate, the data were normalized based on per µM HAA-

272

induced inhibition. The pairwise comparisons among different HAAs were based on an

273

ANOVA test using the Holm-Sidak method and the analysis is presented in Table 1 of

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the Supporting Information.


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Among all the HAAs, IAA was the strongest inhibitor of GAPDH followed by

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BAA, while CAA was the weakest inhibitor of GAPDH (Figure 1). Comparisons

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between the alterations to GAPDH kinetics produced by different HAAs indicated that

278

the IAA and BAA induced interference with GAPDH activity was significantly greater

279

than that produced by any of the di- and tri-HAAs (Supporting Information Table 1). The

280

reactivity of IAA vs BAA vs CAA is due to the dissociation energy of the leaving group

281

(the halogen) generating an alkylating product that reacts with the cysteine at the active

282

site of GAPDH15,18. In addition, weaker inhibitory reponses produced by the di-, and

283

triHAAs indicate that they may not interfere with GAPDH, possibly due to the

284

structurally based steric hindrance. Pairwise comparison between most of the di- and tri-

285

HAAs revealed that the reduction in GAPDH activity by these HAAs was not

286

significantly different from each other (Supporting Information, Table 1).

287

The Effect of HAAs on ATP Levels in CHO Cells

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Our data demonstrated that structurally different HAAs inhibit GAPDH

289

production of NADH to varying degrees and that the monoHAAs, IAA and BAA, were

290

the strongest inhibitors of GAPDH. To study further the associations among structurally

291

different mono-, di-, and triHAAs on cellular energy depletion, we also evaluated the

292

effect HAAs have on cellular ATP levels. Initially, as preliminary experiments, CHO

293

cells were treated with individual HAAs for 4 h at high noncytotoxic concentrations in

294

order to normalize their biological activity on the basis of cytotoxicity (monoHAAs: 4

295

µM IAA, 15 µM BAA, or 700 µM CAA; diHAAs: 700 µM DBA, 700 µM BCA, or 2000

296

µM DCA; and triHAAs: 2000 µM TCA, 1000 µM CDBA, 1000 µM TBA, or 1000 µM

297

BDCA) .12


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Figure 1 in Supporting Information illustrates the effect of HAAs on ATP levels,

299

expressed as a percent of the concurrent negative control. MonoHAAs induced the

300

greatest reduction in ATP, whereas diHAAs induced a moderate reduction with higher

301

concentration ranges, while triHAAs, except for TCA, induced significant increases in

302

ATP levels. The weaker reduction in ATP levels by diHAAs was expected, as they did

303

not significantly inhibit GAPDH production of NADH, and likely induce energy

304

depletion through a different mechanism than monoHAAs. However, most of the

305

triHAAs (TBA, CDBA, DCBA, but not TCA) induced a significant increase in cellular

306

ATP levels. These observations suggest a general association between cellular ATP level

307

and the number of halogens in the HAA. The results indicated that monoHAAs, including

308

the marginally toxic CAA, significantly reduce ATP levels as compared to concurrent

309

negative controls, and as compared to di-, and triHAAs. The unexpected increase in ATP

310

level produced by most triHAAs suggested that they might have cellular targets involved

311

in the ATP regulatory pathways not shared with the monoHAAs (Figure 1 in

312

Supplemental Information). The association, however, is not absolute (e.g., the TCA

313

response), indicating that various factors, in addition to the number of halogen

314

substituents, influence the toxicity of HAAs.

315

Previous studies demonstrated that DCA and TCA inhibit PDK25, 32 and promote

316

activation of PDC. 25 The decarboxylation of pyruvate into acetyl-CoA by PDC is a rate-

317

limiting step connecting glycolysis to the CAC. Pyruvate decarboxylation by PDC

318

provides substrate (acetyl-CoA) for the CAC. 27 Reports indicated that 1 mM DCA

319

increased the activity of PDC in rat heart by inhibiting PDK, and also showed that there

320

was an inverse relation between the NADH:NAD+, ATP:ADP, and acetyl-CoA:CoA


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ratios and the activity of PDC.24 Consistent with previous studies, we observed that the

322

monoHAAs, with the exception of CAA, were the strongest inhibitors of GAPDH among

323

all HAAs. Furthermore, monoHAAs including CAA, greatly reduced ATP levels leading

324

to disruptions in ATP:ADP and NADH:NAD+ ratios. Di-, and triHAAs were relatively

325

weaker inhibitors of GAPDH in CHO cells and triHAAs increased ATP levels suggesting

326

a different underlying mechanism for inducing changes in overall cellular metabolism.

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The CAC and oxidative phosphorylation are pivotal processes in cellular energy

328

homeostasis. Since the CAC is dependent on PDC for its substrate, PDC regulation plays

329

an important role in overall energy homeostasis. Our data demonstrate a general

330

structure-activity relationship for HAA-induced GAPDH inhibition and overall disruption

331

of cellular energy homeostasis, suggesting that differences may exist in the mechanisms

332

exhibited by individual HAAs in modulating PDC activity.

333

Effect of HAAs on PDC Activity

334

Previous studies demonstrated that DCA-induced PDK inhibition activated

335

PDC.32,33 Kato and colleagues found that DCA prevented ADP dissociation from the

336

ATP lid of PDK resulting in PDK inhibition. The presence of DCA and ADP

337

synergistically decreased the affinity of PDK for the L2 domain by 130-fold as compared

338

to ADP alone and eventually affected the L2-induced activation of PDK. The inhibition

339

of PDK by the DCA-induced allosteric change kept the PDC in its active state and

340

accelerated the CAC. 33 Moreover, studies showed that ATP:ADP and NADH:NAD+

341

ratios are also involved in the PDC regulation .24 Therefore, based on these known

342

mechanisms of PDC regulation, we reasoned that IAA and BAA strongly inhibit GAPDH


17


Page 18 of 29

343

leading to reduced cellular ATP levels that disrupt the ATP:ADP ratio. This altered ratio

344

stimulates cells to activate PDC activity and eventually upregulate the CAC to produce

345

more energy and maintain the ATP:ADP balance. Di-, and triHAAs, however, are weaker

346

inhibitors of GAPDH and some triHAAs increase cellular ATP levels. We further

347

reasoned that di-, and triHAAs may increase ATP levels primarily via PDK inhibition.

348

PDK inhibition leads to increased PDC activity and generates elevated levels of ATP.

349

To test this hypothesis, PDC activity was measured in CHO cells after exposure to

350

structurally different HAAs (Figure 2). We found that most of the HAAs (except for

351

CDBA, BCA and BDCA) increased PDC activity. These data also demonstrated that

352

monoHAAs increased PDC activity to a greater degree than did di- or triHAAs. Among

353

monoHAAs, IAA induced the greatest increase in PDC activity (370% of the negative

354

control), followed by BAA (308%) and CAA (194%) (Figure 2). Previous studies

355

concluded that IAA was the strongest inhibitor of GAPDH and reduced ATP levels to the

356

greatest degree, followed by BAA and CAA.17, 18 It was also revealed that there was an

357

inverse relationship between NADH:NAD+, ATP:ADP, and acetyl-CoA:CoA ratios and

358

PDC activity.24 Our current results with monoHAAs are consistent with these previous

359

findings. The pattern for the increase in PDC activity (IAA > BAA >> CAA) also

360

correlated with the pattern found for other HAA toxicity metrics. 17, 18 Among diHAAs,

361

DCA and DBA increased PDC activity to 135% and 184% above the negative control,

362

respectively, while BCA had no significant effect on PDC activity (Figure 2). Among

363

triHAAs, TBA and TCA increased PDC activity to 255% and 148% of the negative

364

control, respectively. However, BDCA significantly decreased the PDC activity to

365

35.4%, while CDBA had no significant effect on PDC activity. Importantly, these data


18

Page 19 of 29


366

are generally consistent with our model in which increased PDC activity by the

367

monoHAAs, IAA and BAA is due to disturbances in the metabolite ratios (ATP:ADP,

368

NADH:NAD+) induced by GAPDH inhibition. In contrast, di- and tri-HAAs were weaker

369

inhibitors of GAPDH and had more modest effects on ATP levels (Figure 1 in

370

Supplemental Information), which suggests that the increased PDC activity produced by

371

di- and tri-HAAs is affected by an alternative mechanism, such as the inhibition of PDK.

372

Note that not all HAAs fit these patterns perfectly (e.g., CAA and GAPDH inhibition),

373

indicating that additional mechanisms are probably involved.

374

The purpose of the preliminary experiments indicated in Figure 1 of Supplemental

375

Information was to determine the relationship between the number of halogens in the

376

HAAs and alterations in ATP production. We tested several different concentrations for

377

individual HAAs and chose the lowest noncytotoxic concentration that induced a

378

significant change in ATP reduction for comparing the effects of the different HAAs on

379

ATP production. The lowest concentration of individual HAAs that induced cytotoxicity

380

was published previously. 12 Even though the specific percent ATP change varied by

381

individual HAAs, a general trend was resolved where the mono-HAAs reduced the

382

percent ATP production while tri-HAAs increased percent ATP production. This

383

observation led us to analyze the PDC activity and to repeat the ATP analysis in HAA-

384

treated cells. We demonstrated that the changes in PDC activity correlated with ATP

385

levels based on similar HAA concentrations (Figure 3). We normalized the percent ATP

386

levels per mg protein to serve as more robust values for comparing responses. Moreover,

387

our findings were consistent with our hypothesis regarding HAA-induced effects on PDC

388

activity.


19


389

Page 20 of 29

MonoHAAs significantly reduced cellular ATP, indicating an inverse relationship

390

between the ATP level and PDC activity. Among the diHAAs, DCA significantly

391

increased cellular ATP to 207.4% as compared to negative control (Figure 3); 50 µM

392

DCA significantly increased the PDC activity as well (Figure 2). Therefore, increases in

393

PDC activity and changes in energy homeostasis by DCA, DBA, TBA, and TCA are not

394

due to changes in the metabolite ratios induced by GAPDH inhibition, but more likely

395

due to allosteric changes induced in PDK by DCA and other di- and tri-HAAs. These

396

changes may lead to inhibition of PDK and activation of PDC, findings that are

397

consistent with previous studies. 24, 32 Furthermore, the relatively strong effect that DCA

398

has on ATP levels and PDC activity may be because it acts as a pyruvate analog, unlike

399

other HAAs54. Significant increases and decreases in the ATP content of HAA-treated

400

cells are shown in Figure 3.

401

In summary, these findings support a model in which the number and type of

402

halogens affect the mechanism and degree of toxicity of HAAs. Our primary finding was

403

that HAAs depending upon their chemical structure affect different molecular target

404

molecules that generate a toxic cascade. Two of the monoHAAs (IAA and BAA)

405

induced their toxicity by indirectly affecting mitochondrial metabolism due to

406

downstream effects of GAPDH inhibition. Di- and triHAAs appeared to affect

407

mitochondrial metabolism by inhibiting PDK, which enhanced PDC activation.

408

Moreover, we found that the monoHAAs strongly affected GAPDH inhibition, ATP

409

depletion and PDC activation whereas di-, and triHAAs produced their effects only at

410

much higher concentrations, which may make their toxicity less relevant to

411

environmental DBP exposures. Di-, and triHAA-induced weaker effects may be due to


20

Page 21 of 29


412

the steric hindrance induced by the multiple halogen groups in their structures. The

413

relationship between the number of halogens and HAA toxicity, however, was not a

414

perfect one (e.g., CAA, triHAAs). Thus, there are likely to be other factors affecting

415

HAA toxicity that will have to be determined in additional studies.

416

Acknowledgments

417

We appreciate the support of all the funding sources. This work was supported by

418

the EPA STAR Grant R834867. We appreciate support by the Center of Advanced

419

Materials for the Purification of Water with Systems (WaterCAMPWS), a National

420

Science Foundation Science and Technology Center, under Award CTS-0120978. C.J.

421

was supported by a NIEHS Pre-doctoral Fellowship under Grant No. T32 ES007326.

422


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References 1. Cutler, D.; Miller, G., The role of public health improvements in health advances: The twentieth-century United States. Demography 2005, 42, (1), 1-22. 2. Richardson, S. D.; Postigo, C., Formation of DBPs: State of the Science. In Recent Advances in Disinfection By-Products, American Chemical Society: 2015; Vol. 1190, pp 189-214. 3. Richardson, S. D.; Plewa, M. J.; Wagner, E. D.; Schoeny, R.; Demarini, D. M., Occurrence, genotoxicity, and carcinogenicity of regulated and emerging disinfection byproducts in drinking water: a review and roadmap for research. Mutat. Res. Rev. Mutat. Res. 2007, 636, (1-3), 178-242. 4. Krasner, S. W.; Weinberg, H. S.; Richardson, S. D.; Pastor, S. J.; Chinn, R.; Sclimenti, M. J.; Onstad, G. D.; Thruston Jr, A. D., Occurrence of a new generation of disinfection byproducts. Environ. Sci. Technol. 2006, 40, (23), 7175-7185. 5. Krasner, S. W., The formation and control of emerging disinfection by-products of health concern. Philos. Trans. A. Math. Phys. Eng. Sci. 2009, 367, (1904), 4077-95. 6. U. S. Environmental Protection Agency, National Primary Drinking Water Regulations; Stage 2 Disinfectants and Disinfection Byproducts Rule. Fed Reg 2006, 71, (2), 387-493. 7. Plewa, M. J.; Kargalioglu, Y.; Vankerk, D.; Minear, R. A.; Wagner, E. D., Mammalian cell cytotoxicity and genotoxicity analysis of drinking water disinfection byproducts. Environ. Mol. Mutagen. 2002, 40, (2), 134-142. 8. Narotsky, M. G.; Best, D. S.; McDonald, A.; Godin, E. A.; Hunter, E. S., 3rd; Simmons, J. E., Pregnancy loss and eye malformations in offspring of F344 rats following gestational exposure to mixtures of regulated trihalomethanes and haloacetic acids. Reprod. Toxicol. 2011, 31, (1), 59-65. 9. Villanueva, C. M.; Cantor, K. P.; Grimalt, J. O.; Malats, N.; Silverman, D.; Tardon, A.; Garcia-Closas, R.; Serra, C.; Carrato, A.; Castaño-Vinyals, G.; Marcos, R.; Rothman, N.; Real, F. X.; Dosemeci, M.; Kogevinas, M., Bladder cancer and exposure to water disinfection by-products through ingestion, bathing, showering, and swimming in pools. Am. J. Epidemiol. 2007, 165, (2), 148-156. 10. Waller, K.; Swan, S. H.; DeLorenze, G.; Hopkins, B., Trihalomethanes in drinking water and spontaneous abortion. Epidemiology 1998, 9, (2), 134-140. 11. National Toxicology Program Toxicity Effects. https://tools.niehs.nih.gov/cebs3/ntpviews/index.cfm?action=testarticle.toxicity&cas_nu mber=79-43-6 12. Plewa, M. J.; Simmons, J. E.; Richardson, S. D.; Wagner, E. D., Mammalian Cell Cytotoxicity and Genotoxicity of the Haloacetic Acids, A Major Class of Drinking Water Disinfection By-Products. Environ. Mol. Mutagen. 2010, 51, (8-9), 871-878. 13. Kargalioglu, Y.; McMillan, B. J.; Minear, R. A.; Plewa, M. J., Analysis of the cytotoxicity and mutagenicity of drinking water disinfection by-products in Salmonella typhimurium. Teratog. Carcinog. Mutagen. 2002, 22, (2), 113-28. 14. Zhang, S. H.; Miao, D. Y.; Liu, A. L.; Zhang, L.; Wei, W.; Xie, H.; Lu, W. Q., Assessment of the cytotoxicity and genotoxicity of haloacetic acids using microplatebased cytotoxicity test and CHO/HGPRT gene mutation assay. Mutat. Res. 2010, 703, (2), 174-9.


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29. Patel, M. S.; Roche, T. E., Molecular biology and biochemistry of pyruvate dehydrogenase complexes. Fed. Am. Soc. Exp. Biol. 1990, 4, (14), 3224-33. 30. Frey, P. A.; Flournoy, D. S.; Gruys, K.; Yang, Y. S., Intermediates in reductive transacetylation catalyzed by pyruvate dehydrogenase complex. Ann. N. Y. Acad. Sci. 1989, 573, 21-35. 31. Patel, M. S.; Korotchkina, L. G., The biochemistry of the pyruvate dehydrogenase complex*. Biochem. Mol. Biol. Educ 2003, 31, (1), 5-15. 32. Kato, M.; Chuang, J. L.; Tso, S. C.; Wynn, R. M.; Chuang, D. T., Crystal structure of pyruvate dehydrogenase kinase 3 bound to lipoyl domain 2 of human pyruvate dehydrogenase complex. The EMBO journal 2005, 24, (10), 1763-74. 33. Kato, M.; Li, J.; Chuang, J. L.; Chuang, D. T., Distinct Structural Mechanisms for Inhibition of Pyruvate Dehydrogenase Kinase Isoforms by AZD7545, Dichloroacetate, and Radicicol. Structure 2007, 15, (8), 992-1004. 34. Bonnet, S.; Archer, S. L.; Allalunis-Turner, J.; Haromy, A.; Beaulieu, C.; Thompson, R.; Lee, C. T.; Lopaschuk, G. D.; Puttagunta, L.; Bonnet, S.; Harry, G.; Hashimoto, K.; Porter, C. J.; Andrade, M. A.; Thebaud, B.; Michelakis, E. D., A Mitochondria-K+ Channel Axis Is Suppressed in Cancer and Its Normalization Promotes Apoptosis and Inhibits Cancer Growth. Cancer Cell 2007, 11, (1), 37-51. 35. Berg, J. M.; Tymoczko, J. L.; Stryer, L., Entry to the Citric Acid Cycle and Metabolism Through It Are Controlled. . In Biochemistry, 5th edition ed.; W H Freeman: New York, 2002. 36. Wagner, E. D.; Plewa, M. J., CHO cell cytotoxicity and genotoxicity analyses of disinfection by-products: an updated review. . J. Environ. Sci. 2017, Submitted. 37. Tindall, K. R.; Stankowski Jr, L. F., Molecular analysis of spontaneous mutations at the gpt locus in Chinese hamster ovary (AS52) cells. Mutat. Res. 1989, 220, (2-3), 241253. 38. Wagner, E. D.; Rayburn, A. L.; Anderson, D.; Plewa, M. J., Analysis of mutagens with single cell gel electrophoresis, flow cytometry, and forward mutation assays in an isolated clone of Chinese hamster ovary cells. Environ. Mol. Mutagen. 1998, 32, (4), 360368. 39. Wagner, E. D.; Rayburn, A. L.; Anderson, D.; Plewa, M. J., Calibration of the single cell gel electrophoresis assay, flow cytometry analysis and forward mutation in Chinese hamster ovary cells. Mutagenesis 1998, 13, (1), 81-84. 40. Lissens, W.; De Meirleir, L.; Seneca, S.; Liebaers, I.; Brown, G. K.; Brown, R. M.; Ito, M.; Naito, E.; Kuroda, Y.; Kerr, D. S.; Wexler, I. D.; Patel, M. S.; Robinson, B. H.; Seyda, A., Mutations in the X-linked pyruvate dehydrogenase (E1) alpha subunit gene (PDHA1) in patients with a pyruvate dehydrogenase complex deficiency. Hum. Mutat. 2000, 15, (3), 209-19. 41. Patel, K. P.; O'Brien, T. W.; Subramony, S. H.; Shuster, J.; Stacpoole, P. W., The spectrum of pyruvate dehydrogenase complex deficiency: clinical, biochemical and genetic features in 371 patients. Mol. Genet. Metab. 2012, 106, (3), 385-94. 42. Kaplon, J.; Zheng, L.; Meissl, K.; Chaneton, B.; Selivanov, V. A.; Mackay, G.; van der Burg, S. H.; Verdegaal, E. M.; Cascante, M.; Shlomi, T.; Gottlieb, E.; Peeper, D. S., A key role for mitochondrial gatekeeper pyruvate dehydrogenase in oncogene-induced senescence. Nature 2013, 498, (7452), 109-12.


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43. Stacpoole , P. W.; Moore , G. W.; Kornhauser , D. M., Metabolic Effects of Dichloroacetate in Patients with Diabetes Mellitus and Hyperlipoproteinemia. N. Engl. J. Med. 1978, 298, (10), 526-530. 44. Li, C.; Meng, G.; Su, L.; Chen, A.; Xia, M.; Xu, C.; Yu, D.; Jiang, A.; Wei, J., Dichloroacetate blocks aerobic glycolytic adaptation to attenuated measles virus and promotes viral replication leading to enhanced oncolysis in glioblastoma. Oncotarget 2015, 6, (3), 1544-1555. 45. James, M. O.; Cornett, R.; Yan, Z.; Henderson, G. N.; Stacpoole, P. W., Glutathione-dependent conversion to glyoxylate, a major pathway of dichloroacetate biotransformation in hepatic cytosol from humans and rats, is reduced in dichloroacetatetreated rats. Drug Metab Dispos 1997, 25, (11), 1223-7. 46. Tong, Z.; Board, P. G.; Anders, M. W., Glutathione transferase zeta-catalyzed biotransformation of dichloroacetic acid and other alpha-haloacids. Chem Res Toxicol 1998, 11, (11), 1332-8. 47. Anderson, W. B.; Board, P. G.; Anders, M. W., Glutathione transferase zetacatalyzed bioactivation of dichloroacetic acid: reaction of glyoxylate with amino acid nucleophiles. Chem Res Toxicol 2004, 17, (5), 650-62. 48. Anderson, W. B.; Board, P. G.; Gargano, B.; Anders, M. W., Inactivation of glutathione transferase zeta by dichloroacetic acid and other fluorine-lacking alphahaloalkanoic acids. Chem Res Toxicol 1999, 12, (12), 1144-9. 49. Ammini, C. V.; Fernandez-Canon, J.; Shroads, A. L.; Cornett, R.; Cheung, J.; James, M. O.; Henderson, G. N.; Grompe, M.; Stacpoole, P. W., Pharmacologic or genetic ablation of maleylacetoacetate isomerase increases levels of toxic tyrosine catabolites in rodents. Biochem Pharmacol 2003, 66, (10), 2029-38. 50. Blackburn, A. C.; Matthaei, K. I.; Lim, C.; Taylor, M. C.; Cappello, J. Y.; Hayes, J. D.; Anders, M. W.; Board, P. G., Deficiency of glutathione transferase zeta causes oxidative stress and activation of antioxidant response pathways. Mol Pharmacol 2006, 69, (2), 650-7. 51. Kahlert, S.; Reiser, G., Requirement of glycolytic and mitochondrial energy supply for loading of Ca2+ stores and InsP(3)-mediated Ca2+ signaling in rat hippocampus astrocytes. J. Neurosci. Res. 2000, 61, (4), 409-420. 52. Cemeli, E.; Wagner, E. D.; Anderson, D.; Richardson, S. D.; Plewa, M. J., Modulation of the cytotoxicity and genotoxicity of the drinking water disinfection byproduct lodoacetic acid by suppressors of oxidative stress. Environ. Sci. Technol. 2006, 40, (6), 1878-83. 53. Stalter, D.; O'Malley, E.; von Gunten, U.; Escher, B. I., Fingerprinting the reactive toxicity pathways of 50 drinking water disinfection by-products. Water Res 2016, 91, 19-30. 54. Saunier, E.; Benelli, C.; Bortoli, S., The pyruvate dehydrogenase complex in cancer: An old metabolic gatekeeper regulated by new pathways and pharmacological agents. Int. J. Cancer 2016, 138, (4), 809-17.


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FIGURES

GAPDH kinetics inhibition induced by per µM HAAs (Index values (±SE))

100

80

60

40

20

0 IAA

BAA

CAA

BCA

DBA

DCA

TBA

TCA CDBA BDCA

Treatment Group [HAAs]

Figure 1. Percent reduction in GAPDH activity per µM of different HAAs as measured by NADH production. Among all the HAAs, the monoHAAs (red, specifically, IAA and BAA) were the strongest inhibitors of GAPDH activity, whereas CAA, di- (green), and tri-HAAs (blue) had more modest or no effects. Abbreviations: CAA, chloroacetic acid; BAA, bromoacetic acid; IAA, iodoacetic acid; DCA, dichloroacetic acid; DBA, dibromoacetic acid; BCA, bromochloroacetic acid; TCA, trichloroacetic acid; TBA, tribromoacetic acid; BDCA, bromodichloroacetic acid; CDBA, dibromochloroacetic acid


26


500

*

400

* *

300

*

*

*

200

* * 0

TC A

50

50 0 C µM D BA 30 0 BD µM C A 30 0 µM

µM

µM 90

0 TB A

D

A

BA

90 0

µM

µM 50 BC

D C

A

15

µM C AA

15

3 IA A

iv e eg at N

BA A

nt

µM

ro l

0

µM

100

co

PDC Activity as nmol of NADH/min/mg protein as Percent of the Concurrent Negative Control (±SE)

Page 27 of 29

Treatment Groups [HAA]

Figure 2. Effect of HAAs on CHO PDC activity (nmol NADH/min/mg protein). PDC activity for each HAA treatment group was normalized against each concurrent negative control and the negative control shown here is the average of all the negative control groups. The * indicates a statistically significant difference from the concurrent negative control, SE is standard error.

Abbreviations: CAA, chloroacetic acid; BAA, bromoacetic acid; IAA, iodoacetic acid; DCA, dichloroacetic acid; DBA, dibromoacetic acid; BCA, bromochloroacetic acid; TCA, trichloroacetic acid; TBA, tribromoacetic acid; BDCA, bromodichloroacetic acid; CDBA, dibromochloroacetic acid


27


Page 28 of 29

pmol ATP/mg Protein Normalized with Concurrent Negative Control (±SE)

250

* 200

150

100

*

*

* *

50

50 0 µM TC A 50 0 C µM D BA 30 BD 0 µM C A 30 0 µM

µM

TB A

90

0

D BA

A

90 0

µM

µM 50 BC

D

C A

µM

µM

30 0

3

15

C AA

BA A

IA A

N eg at iv e

C on

tro

µM

l

0

Treatment Groups [HAA]

Figure 3. Effect of HAAs on cellular ATP levels (pmol ATP/mg protein). ATP levels for each HAA treatment group are expressed as a percent of the concurrent negative control; the negative control shown here is the average of all the negative control groups. * indicates a statistically significant difference from the negative control, SE is standard error. Abbreviations: CAA, chloroacetic acid; BAA, bromoacetic acid; IAA, iodoacetic acid; DCA, dichloroacetic acid; DBA, dibromoacetic acid; BCA, bromochloroacetic acid; TCA, trichloroacetic acid; TBA, tribromoacetic acid; BDCA, bromodichloroacetic acid; CDBA, dibromochloroacetic acid


28

Page 29 of 29


187x98mm (300 x 300 DPI)


Haloacetic Acid Water Disinfection Byproducts Affect Pyruvate

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