Synergistic Treatment of Mixed 1,4-Dioxane and Chlorinated Solvent

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Synergistic Treatment of Mixed 1,4-Dioxane and Chlorinated Solvent Contaminations by Coupling Electrochemical Oxidation with Aerobic Biodegradation Jeramy R. Jasmann, Phillip B. Gedalanga, Thomas Borch, Shaily Mahendra, and Jens Blotevogel Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b03134 • Publication Date (Web): 12 Oct 2017 Downloaded from http://pubs.acs.org on October 13, 2017

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Environmental Science & Technology

Synergistic Treatment of Mixed 1,4-Dioxane and Chlorinated Solvent Contaminations by Coupling Electrochemical Oxidation with Aerobic Biodegradation Jeramy R. JasmannA,†, Phillip B. GedalangaB, Thomas BorchA,C,D, Shaily MahendraB, Jens BlotevogelC,*

A

Department of Chemistry, Colorado State University, Fort Collins, Colorado, CO 80523, USA

B

Department of Civil and Environmental Engineering, University of California, Los Angeles, CA 90095, USA

C

Department of Civil and Environmental Engineering, Colorado State University, Fort Collins, CO 80523, USA

D

Department of Soil and Crop Sciences, Colorado State University, Fort Collins, CO 80523, USA

†

Current address: U.S. Geological Survey, National Research Program, Boulder, CO 80303, USA

TOC Art

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ABSTRACT

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Biodegradation of the persistent groundwater contaminant 1,4-dioxane is often hindered

3

by the absence of dissolved oxygen and the co-occurrence of inhibiting chlorinated solvents.

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Using flow-through electrolytic reactors equipped with Ti/IrO2-Ta2O5 mesh electrodes, we show

5

that combining electrochemical oxidation with aerobic biodegradation produces an over-additive

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treatment effect for degrading 1,4-dioxane. In reactors bioaugmented by Pseudonocardia

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dioxanivorans CB1190 with 3.0 V applied, 1,4-dioxane was oxidized 2.5 times faster than in

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bioaugmented control reactors without an applied potential, and 12 times faster than by abiotic

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electrolysis only. Quantitative polymerase chain reaction analyses of CB1190 abundance,

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oxidation-reduction potential, and dissolved oxygen measurements indicated that microbial

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growth was promoted by anodic oxygen-generating reactions. At a higher potential of 8.0 V,

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however, the cell abundance near the anode was diminished, likely due to unfavorable pH and/or

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redox conditions. When coupled to electrolysis, biodegradation of 1,4-dioxane was sustained

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even in the presence of the common co-contaminant trichloroethene in the influent. Our findings

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demonstrate that combining electrolytic treatment with aerobic biodegradation may be a

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promising synergistic approach for the treatment of mixed contaminants.

17 18 19

INTRODUCTION

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1,4-Dioxane, a widely used solvent stabilizer, is a persistent organic pollutant frequently

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observed in groundwater impacted by chlorinated volatile organic compounds.1-3 Recent reports

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of contaminated site data from across the United States highlight the high probability of co-

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occurrence of 1,4-dioxane with vapor degreasers such as trichloroethene (TCE) and 1,1,1-

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trichloroethane (1,1,1-TCA).3-5 1,4-Dioxane’s miscibility in water and low sorption affinity to

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soil make it highly mobile in groundwater, frequently leading to large plume development.

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While often at sub-mg/L concentrations in dilute groundwater plumes, 1,4-dioxane

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concentrations can occur at tens or hundreds of mg/L in source zones and industrial

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wastewater.3,6-8 1,4-dioxane is highly recalcitrant, and commonly applied remediation

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technologies for chlorinated solvent co-contaminants, like sorption and air stripping, have been

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ineffective for its removal. Advanced oxidation processes (AOPs) involving UV light can

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produce highly reactive hydroxyl radicals (•OH) to mineralize 1,4-dioxane, 9-12 but the high cost 2 ACS Paragon Plus Environment

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typically limits their use to treatment of relatively small water volumes and ex situ treatment.

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Thus, the development of more cost-effective treatment options is critically needed.

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Recent laboratory studies have documented successful aerobic biodegradation of 1,4-

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dioxane either co-metabolically13-16 or metabolically,16-18 and one study demonstrated microbially

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driven Fenton-based degradation19 of 1,4-dioxane in a system alternating between aerobic and

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anaerobic conditions. These studies indicate that bioremediation of 1,4-dioxane is generally

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possible under suitable conditions. However, three factors may limit the site-specific

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biodegradation potential for 1,4-dioxane: (1) lack of dissolved oxygen for aerobic microbial

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respiration, (2) low activity of indigenous bacterial populations, and (3) inhibition of microbial

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metabolism by chlorinated solvents.20-22 Currently, all pure culture isolates capable of 1,4-

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dioxane biodegradation require O2 as the terminal electron acceptor.13-18 This constraint has been

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prohibitive to natural attenuation approaches since 1,4-dioxane contamination often occurs in

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anoxic water conditions. Recent investigations by Mahendra and coworkers20,22 revealed that

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TCE, 1,1,1-TCA, and the abiotic transformation products 1,1-dichloroethene (1,1-DCE) and cis-

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1,2-dichloroethene (cis-DCE) caused metabolic inhibition of the 1,4-dioxane-metabolizing

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bacteria Pseudonocardia dioxanivorans CB1190. The observed decrease in biodegradation rates

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were attributed to delayed ATP production and down-regulation of the 1,4-dioxane

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monooxygenase (dxmB) and aldehyde dehydrogenase (aldH) genes essential to the production of

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enzymes known to be significant in the 1,4-dioxane degradation pathway.23 However, these

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studies also indicated that the inhibition was non-competitive and reversible, opening up the

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opportunity for full metabolic recovery once the chlorinated compounds were removed. Hand

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and coworkers provided similar evidence for TCE-caused inhibition of 1,4-dioxane

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biodegradation in two co-metabolizing bacteria, Mycobacterium vaccae JOB5 and Rhodococcus

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jostii RHA1.21 Despite this inhibitory effect, groundwater concentration data and detection of in

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situ biomarkers for 1,4-dioxane-metabolizing bacteria from 2000 to 2016 provide encouraging

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evidence for natural 1,4-dioxane attenuation in some plumes co-contaminated with TCE.4,24,25

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Still, observable biodegradation was limited to aerobic regions of the aquifers and negatively

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correlated with concentrations of chlorinated volatile organics. Thus, in order for biodegradation

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of 1,4-dioxane to become a widely used remediation strategy, some form of biostimulation or

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augmentation is required, along with an economical treatment process to address inhibiting co-

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contaminants such as TCE.13,26 3 ACS Paragon Plus Environment


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Electrochemical degradation has previously been shown to effectively mineralize

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chlorinated solvents both by reductive dechlorination and •OH mediated oxidation.27-29 1,4-

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dioxane is also degraded by electrochemical treatment via strongly oxidizing •OH radicals

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generated by the oxidation of H2O at the anode.30-32 Our previous work using abiotic flow-

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through, electrochemical reactors with Ti/IrO2-Ta2O5 electrodes confirmed that 1,4-dioxane

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(influent concentrations 0.3 to 41.4 mg/L) was degraded into CO2 and small organic acid and

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aldehyde intermediates32 known to be readily biodegradable.27,33-35 It has also been shown that

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chloride, often present in natural waters, can potentially be transformed to highly oxidized

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chlorine species by electrolysis, assisting in the mineralization of 1,4-dioxane and other

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contaminants,36 although these species could also be destructive to microbial cells.37-40 By taking

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advantage of the concurrent electrolysis of bulk water to evolve molecular oxygen from the

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anode half reaction, 2 H2O(l) → O2(g) + 4 H+(aq) + 4e-,41-43 a dramatic shift in redox potential from

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hypoxic to highly oxic conditions can be achieved. We thus hypothesized that electrochemical

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treatment would degrade 1,4-dioxane and microbially inhibiting co-contaminants such as TCE

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while simultaneously generating O2 essential to aerobic respiration, thus leading to enhanced

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aerobic biodegradation rates of 1,4-dioxane.33,34

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To test our hypothesis, we built flow-through electrolytic column reactors with

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dimensionally stable, Ti/IrO2-Ta2O5 mesh electrodes, which are commercially available and have

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several beneficial properties. Metal oxide electrodes, like Ti/IrO2-Ta2O5, have a higher oxidation

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state available (i.e. Ir4+ → Ir5+) close to the thermodynamic potential for aqueous oxygen

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evolution, making them excellent candidates for the electrocatalytic generation of O2 at low

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power requirements.41,44,45 Published comparisons have shown the relatively low oxygen

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evolution potential of Ti/IrO2-Ta2O5 (1.5 - 1.8 V vs. SHE, standard hydrogen electrode) also

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coincides with less disinfection by-product (DBP) formation compared to other electrode

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materials used for organic contaminant degradation such as PbO2, RuO2-IrO2, Pt-IrO2, and SnO2-

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Sb.45,46 Detailed reaction schemes describing O2 production and organic pollutant oxidation can

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be found in the Supporting Information (SI, Reactions 1 - 6). Furthermore, previous studies have

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demonstrated this mixed metal oxide coating to have favorable surface chemistry interactions and

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degradation efficiencies with chlorinated contaminants, like TCE.29,47,48 Finally, at a cost of ~400

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$/m2, it is substantially cheaper than many other electrodes such as boron-doped diamond anodes

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(~15,000 $/m2),37 and previous field demonstrations, including use as an in situ permeable 4 ACS Paragon Plus Environment

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electrolytic barrier for chlorinated solvent removal, have revealed a long service life (i.e., no loss

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in electrolytic performance during greater than 2 years of continuous treatment).28,49

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We used Pseudonocardia dioxanivorans CB1190 (hereafter “CB1190”) as the model

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bacterium for 1,4-dioxane biodegradation due to its versatile capabilities to metabolize a wide

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range of organic molecules (although not TCE), while also being able to rapidly metabolize 1,4-

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dioxane at rates ranging from 1.1 to 19.8 mg·min-1 per mg of protein in the biomass.17,18,50 From

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among the chlorinated solvents that have been characterized with respect to inhibiting 1,4-

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dioxane degradation by CB1190,22 TCE was chosen because the dichloroethene isomers are

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substantially more volatile and may easily be stripped by electrolytically produced gasses, and

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1,1,1-TCA has shown very little inhibition to CB1190 metabolism.

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Our specific objectives were to (a) determine whether anodic generation of O2 improves

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biodegradation rates of 1,4-dioxane and (b) investigate the impacts of voltage potentials and

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presence of TCE as co-contaminant on degradation kinetics of 1,4-dioxane and spatial CB1190

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abundance relative to the electrodes. Consequently, this is the first study to provide a

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fundamental basis for combined electrochemical and biological treatment for 1,4-dioxane-

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contaminated waters, both when occurring on its own and in the presence of potentially

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inhibiting co-contaminants.

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MATERIALS AND METHODS

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Chemicals. All chemicals were used as received. 1,4-Dioxane (99.5%, Burdick &

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Jackson), TCE (99.5%, Alfa Aesar), 1,1-DCE (99%, Alfa Aesar), and vinyl chloride (99%, Alfa

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Aesar) were used to prepare calibration standards and/or influent contaminant mixtures while

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dichloromethane (99.96%, OmniSolv Millipore EMD) was used as an extraction solvent prior to

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quantitation of chemical analytes. Additionally, deuterated 1,4-dioxane-d8 (99% and 99 atom

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%D, Sigma Aldrich) was used for isotopic dilution quantitation.

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Sterilization and Disinfection Protocol. All glassware, metal laboratory equipment, and

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sand used for abiotic control experiments were dry heat-sterilized in a furnace at 232°C for 12-18

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hours or autoclaved at 121°C and 23 psig for 20 minutes (Tuttnauer 2870 EP Autoclave). Non-

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autoclavable materials, i.e. work bench surface and the PVC sand column reactors, were

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disinfected either with 70% ethanol or 10% bleach solutions. 5 ACS Paragon Plus Environment


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Bacterial Strain and Culture Conditions. The model 1,4-dioxane metabolizing

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bacterium, Pseudonocardia dioxanivorans strain CB1190, was harvested by a 1% (v/v) transfer

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from an actively growing pure culture into ammonium mineral salts (AMS) nutrient medium

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similar to previously reported procedures (Appendix A in Supporting Information).17,22 The AMS

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medium’s main constituents, 660 mg/L (NH4)2SO4 and 1000 mg/L MgSO4·7H2O, serve as

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important nitrogen and sulfur sources for CB1190. In addition, the AMS medium serves as a

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proxy for groundwater since it consists of a complex mixture of other inorganic ions commonly

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found in groundwater environments such as Ca2+, Fe2+, Zn2+, Mn2+, SO42- and Cl-. Aqueous

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CB1190 cultures were grown aerobically in AMS medium and sequentially harvested by 1 - 10

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% (v/v) transfers into larger and larger volumes of fresh AMS solution, ending in 4.0-L

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polycarbonate Nalgene containers, which were then used to inoculate silica sand (Figure S3).

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Spikes of 1,4-dioxane were added every 2 to 5 days, maintaining levels at 80 - 100 mg/L, to

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provide ongoing renewal of the sole carbon and energy source. Throughout the process, aqueous

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and sand cultures were incubated at 30°C with continuous agitation at 100 rpm to

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homogeneously mix the 1,4-dioxane carbon source and micronutrients, and infuse oxygen. Only

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“active” CB1190 cultures were used to inoculate sand, and only “active” batches of this

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homogeneously mixed inoculated sand were used to pack into bioaugmented column reactors.

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Initial CB1190 population densities (planktonic and sessile) were measured by qPCR analysis

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immediately following transfer into column reactors. Targeting “active” cultures was meant to

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provide sand containing CB1190 communities at peak population density and metabolic activity

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prior to initiating each flow-through column experiment. We characterized “active” CB1190

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cultures as those with (a) consistent, rapid 1,4-dioxane biodegradation (biodegradation rates > 10

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mg/L/day), (b) high CB1190 bacteria populations detected on aerobic count plates (1:1000

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dilution having > 5 colonies/cm2 of petrifilm plate), and (c) evidence of the culture experiencing

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mid- to late-exponential growth phase indicated by increased bacterial ATP production (Figure

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S3). To track ATP concentrations over time, luminescence analysis (BioTek Synergy HT) was

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completed within two hours of sampling using the Promega BacTiter-GloTM Microbial Cell

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Viability Assay following Promega Technical Bulletin #TB337 and Protocol for Measuring ATP

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from Bacteria Bulletin. Bacterial abundance was monitored by counting bacterial colonies on 3M

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Petrifilm Aerobic Count Plates following manufacturer’s protocol.


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Flow-Through Column Reactors. Bench-scale flow-through experiments were

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performed using 10-cm I.D. x 45-cm long clear PVC column reactors (Figure 1 and Figure S1),

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packed with 8-12 mesh quartz silica sand and two permeable, disc-shaped Ti/IrO2-Ta2O5 mesh

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electrodes installed perpendicular to flow (78.5 cm2 cross-sectional area, 1mm thick with 1.0 x

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2.8 mm diamond-shaped openings, Corrpro Companies). All column reactors were operated in

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the dark, under opaque plastic, to prevent potential photolysis of 1,4-dioxane. PVC column

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reactors were chemically disinfected (70% ethanol) between experiments and repacked with

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CB1190-inoculated silica sand for all bioaugmented experiments, and repacked with heat-

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sterilized silica sand (232°C for 18 hours) for all non-bioaugmented control experiments. One

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column reactor was used for all 3V experiments, a second column reactor for all 8V experiments,

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and a third was used for all 0V experiments.

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In addition to the influent (12.5 cm before anode) and effluent (32.5 cm downstream of

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anode) liquid sample ports, three additional liquid ports (L1, L2, and L3) and solid ports (S1, S2,

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and S3) were installed along the column flow path at distances of 2.5, 12.5, and 22.5 cm

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downstream of the leading anode (Figure 1). Thus, L1 (S1) is located midway between the

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leading anode (positive polarity) and trailing cathode (negative polarity) which have 5.0 cm of

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separation between them.

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-12.5

0

2.5

12.5

22.5

RE2

RE1

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32.5cm

PVC PVC tube endcap

gas vents

Direction of Flow S1

S2

S3

Leffluent

Linfluent

+

L1

-

L2

L3

Contaminant Feedstock 1,4-Dioxane in AMS medium (TCE as co-contaminant)

Effluent Reservoir

+ - anode & cathode working electrodes S L RE

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solid sample ports liquid sample ports Ag/AgCl reference electrode (redox) silica sand (8/12 mesh)

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Figure 1. Schematic of flow-through electrolytic reactors used for non-bioaugmented electrolytic degradation, bioaugmented biodegradation and combined electro-biodegradation experiments. Solid sample ports have removable threaded PVC plugs (1” diameter) to allow access to sand within the reactor. Liquid sample ports have 3-way valves. Gas vents are exhausted into large tedlar bags.

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Influent feedstock (in 20-L glass carboys) was composed of degassed, hypoxic AMS

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medium spiked with 100 mg/L 1,4-dioxane (and additional 5 mg/L TCE for co-contaminant

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experiments). A high 1,4-dioxane concentration was chosen so that CB1190 biodegradation

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rates would not be limited by electron donor availability and to assess degradation performance

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under challenging conditions, such as in source zones and industrial wastewaters.3,5 The specific

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conductivity of the medium was 3.4 - 3.6 mS/cm and initial dissolved oxygen concentration

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(DO) after degassing was 2.9 - 3.1 mg/L. The pH was adjusted to 6.9 ± 0.1 using NaOH before

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adding phosphate buffer (KH2PO4 / K2HPO4). Completely anoxic feedstock conditions could not

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be maintained without the addition of unwanted oxygen scavenger compounds that may have

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interfered with electrochemical reactions or biological functions. All flow-through sand column

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experiments were operated at 22 ± 2°C (ambient temperature) and a flow rate of 1.07 mL/min. 8 ACS Paragon Plus Environment

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Constant feedstock flow rates (Q) of 1.07 mL·min-1 were confirmed each day by volumetric

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analysis. To account for the sand column reactor porosity (ϕ, average of 0.43) and the flux

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through the reactor’s cross-sectional area (A, 78.5 cm2), flow rate is expressed as its seepage

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velocity (vs) of 46 cm/d, since vs = Q / (A · ϕ) = (1.07cm3·min-1 x 1440min·day-1) / (78.5cm2 x

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0.43). Ten different experimental regimes were tested for the treatment of 100 mg/L 1,4-dioxane

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feedstock under the following conditions: 3V ± CB1190 ± TCE, 8V ± CB1190 ± TCE, and 0V +

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CB1190 ± TCE (Figure S1). To minimize sorption effects, feedstock solution was flowed

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through each column reactor prior to each new treatment regime until influent and effluent

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contaminant concentrations were stable (± 3% coefficient of variation). The target voltage was

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then initiated (0, 3.0 or 8.0 V), and effluent concentrations were monitored until achieving

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steady-state conditions (i.e. stable effluent concentrations over time). Steady-state conditions

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were achieved within 3 - 5 pore volumes (3 - 5 days), and replicate aqueous samples (n ≥ 3) of

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the influent and effluent were taken at 1 - 2 day intervals. Aqueous samples were filtered with

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Acrodisc 0.45-µm nylon membrane syringe filters (Pall Corporation) and extracted into

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dichloromethane solvent prior to analysis for 1,4-dioxane and chlorinated ethenes using an

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Agilent 6890N gas chromatograph (GC) coupled to an Agilent 5973N mass spectrometer (MS)

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in full scan mode (m/z 40-250 Da) and selected ion monitoring (SIM) mode. 1-µL injections

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were made using a 4:1 split flow ratio and an inlet temperature of 250°C. The GC was equipped

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with a Restek Rxi-624Sil MS mid-polarity column (30 m x 0.25 mm ID x 1.4 µm df). The

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Helium carrier gas was to set to constant flow at 2.0 mL/min. The initial oven temperature was

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held at 40°C for 2 minutes, then ramped at 8°C/min to 100°C, followed by an additional ramp of

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40°C/min to 160°C and held for 1 minute.

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The MS was programmed to scan for m/z 62 and 64 (vinyl chloride) in segment 1 from

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1.0 to 3.52 minutes after injection. Segment 2 was set to scan for m/z 61 and 96 (DCE isomers)

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until 5.0 minutes when segment 3 began scanning for m/z 95 and 130 (TCE). Segment 4 started

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at 6.25 minutes scanning for m/z 64 and 96 (1,4-dioxane-d8) as well as m/z 58 and 88 (1,4-

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dioxane). Each ion was assigned a dwell time of 100 µs. Quantitation with GC/MS(SIM) for all

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chlorinated compounds was performed using external calibration standards. External standard

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calibration was also used for 1,4-dioxane in experiments with 1,4-dioxane as the only

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contaminant in the feedstock. Isotopic dilution calibration was use to quantify 1,4-dioxane in all

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co-contaminant experiments, using 1,4-dioxane-d8 as the internal standard to correct for analyte 9 ACS Paragon Plus Environment


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losses due to sample preparation or variations in instrument response. Calibration was obtained

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by plotting the ratio of the analyte signal to the 1,4-dioxane-d8 signal as a function of the analyte

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standard concentration. Further analytical method descriptions are provided in the Supporting

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Information.

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Physicochemical Measurements. Daily measurements were made of voltage and current

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between the anode and cathode (Fluke Multimeter). Solution oxidation-reduction potential (ORP)

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was measured against Ag/AgCl reference micro-electrodes (RE-5, World Precision Instruments)

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at two locations within the column, RE1 and RE2 in Figure 1. Solution pH was measured for all

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experimental regimes in the influent, the effluent, and from aqueous ports L1, L2, and L3 along

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the column flow path (Figure 1). The pH was measured using a pH electrode (±0.1 pH unit

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precision) and confirmed by pH indicator strips at beginning and end of experiments. Specific

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conductivity and dissolved oxygen levels were measured comprehensively only for the

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bioaugmented/mixed contaminant experiments using Hach HQ40d meter with graphite

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conductivity and luminescent DO probes. Calibration of the DO probe with auto salinity

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corrections was performed according to the water-saturated air (100%) procedure in

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manufacturer’s operation manual.

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Quantitative Polymerase Chain Reaction (qPCR). Nucleic acid primers for the dxmB

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gene, which codes for the dioxane monooxygenase β subunit, were used to quantify this target by

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qPCR analysis as described in Gedalanga, et al.23 Additionally, the dxmB gene target was used as

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a proxy for CB1190 population abundance due to its known requirement for 1,4-dioxane

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biodegradation.23 At the start and completion of each ~2-week flow-through experiment, 0.5-mL

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liquid samples and 0.5-g solid sand samples were collected into 2.0-mL microcentrifuge tubes,

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sealed, and frozen at -20°C. Liquid aliquots were sampled via passive flow from valves at the

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influent, inter-column locations L1, L2, L3, and effluent (duplicates collected from each

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location). Removable, threaded PVC plugs at ports S1, S2, and S3 provided access to collect

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duplicate sand samples from center and outer portions of each reactor with heat-sterilized, 1.0-cm

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diameter metal tubes (detailed description in SI). Replicate qPCR analyses were performed for

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each duplicate sample collected, and the average of the four values obtained was used to estimate

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CB1190 abundance at each location along the column flow path.

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In preparation for qPCR analysis, DNA was extracted from sand and liquid phase samples

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using a bead beating method followed by phenol/chloroform purification.23 Briefly, liquid 10 ACS Paragon Plus Environment

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samples were centrifuged to pellet the biomass for 3 minutes at 13,200 x g. The supernatant was

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discarded and DNA was extracted from the pelleted biomass by addition of 0.25 mL extraction

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buffer, 0.1 mL 10% sodium dodecyl sulfate, and 1 mL saturated phenol to all tubes. Samples

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were heated at 65˚C for 2 minutes followed by bead beating for 2 minutes using a minibead

256

beater-16 (Biospec Products, Bartlesville, OK). Lysed samples were incubated at 65˚C for 8 min

257

followed by another round of bead beating for 2 min. Samples were centrifuged and the lysate

258

was transferred to a sterile 1.7-mL microcentrifuge tube for phenol/chloroform purification.

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Nucleic acid extracts were resuspended in 100 µL of nuclease-free H2O and stored at -80˚C until

260

further analyses.

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All qPCR reactions were performed in a total volume of 20 µL containing 1X Kapa Sybr

262

Fast qPCR Master Mix (Kapa Biosystems, Wilmington, MA), 0.25 µM of each primer

263

(IDTDNA, San Diego, CA), and 2 µL of template DNA. All reactions were performed on an

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Applied Biosystems StepOnePlus real-time PCR system (Life Technologies, Carlsbad, CA) as

265

previously described23 and were accompanied with a melt curve analysis to confirm the

266

specificity of qPCR products.

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(Minimum Information for Publication of Quantitative Real-Time PCR Experiments) can be

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found in Table S3.

Additional details following MIQE reporting requirements

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RESULTS AND DISCUSSION

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1,4-Dioxane Degradation in the Absence of Co-contaminant. Investigations on

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coupled electrochemical and biological treatment of 1,4-dioxane were performed at flow rates of

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1.07 mL/min, or seepage velocity of 46 cm/d, in sand-packed flow-through column reactors

275

equipped with permeable Ti/IrO2-Ta2O5 mesh electrodes (Figure 1). In non-bioaugmented

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(“abiotic”) reactors at 3.0 V and 8.0 V in the absence of TCE, only 14.6 and 11.7 mg 1,4-dioxane

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were oxidized per hour per m2 of electrode surface area, respectively (uncertainty of ± 13.9 mg·h-

278

1

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with electrochemical oxidation, additional electrode pairs would need to be added sequentially

280

down the flow path to increase hydraulic retention time and improve overall degradation

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capacity. Although higher concentrations of reactive •OH radicals would be generated at 8.0 V

282

(2.34 mA/cm2) than at 3.0 V (0.20 mA/cm2), these two voltage applications for non-

·m-2, Figure 2). Thus, in a full-scale application treating high contaminant concentrations only



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bioaugmented columns exhibited an insignificant difference in degradation rate (p = 1.0). The

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reason for 8.0 V producing equal (or slightly lower than 3.0 V) removal rates can be justified by

285

the overall reaction rate being controlled by mass transfer setting the upper constraint on

286

degradation rate. The limitation on oxidation of 1,4-dioxane by •OH radicals on, or very near, the

287

electrode surface is compounded in the 8.0 V reactor by more pronounced visible coverage of the

288

mesh electrodes with gas bubbles, thereby reducing hydraulic conductivity and creating

289

preferential flow paths.32,51,52

290

291 292 293 294 295 296 297 298

Figure 2. 1,4-Dioxane degradation rates for non-bioaugmented and P. dioxanivorans CB1190 bioaugmented reactors in the absence (striped) and presence (solid) of TCE co-contaminant (n ≥ 3). The percentage of TCE removed during the mixed co-contaminant experiments is represented by the shaded region of the pie charts above the corresponding bar plot. Error bars of ±13.9 mg·h1 ·m-2 were generated by transforming a precision calculation of the mean absolute deviation for instrumental measurements of influent 1,4-dioxane concentrations (calculations shown in Table S1). 12 ACS Paragon Plus Environment

Page 13 of 28


299

For bioaugmented experiments, the sand was inoculated with “active” CB1190

300

transferred at, or near, peak population density and activity, as described in the Materials and

301

Methods section. In the CB1190 bioaugmented control column (no voltage) with degassed

302

feedstock (initial DO of 3.0 ± 0.1 mg/L, ORP of 0.98 V vs. SHE), a 1,4-dioxane degradation rate

303

of 68.7 mg·h-1·m-2 was observed, about 5 times faster than in either non-bioaugmented

304

electrolytic column, and demonstrating that the microaerophilic CB1190 has the capacity to

305

metabolize 1,4-dioxane even at hypoxic DO levels (~2-3 mg/L). However, previous

306

investigations have shown that the 1,4-dioxane biodegradation capacity of CB1190 would cease

307

in a purely anoxic environment. When electrolysis and biodegradation processes were combined,

308

degradation rates substantially increased in an over-additive manner to 169 mg·h-1·m-2 at 3.0 V

309

and 101 mg·h-1·m-2 at 8.0 V. This degradation rate of 169 mg·h-1·m-2 in the bioaugmented reactor,

310

the highest achieved in this study, was 12 times higher than non-bioaugmented 3.0 V electrolysis,

311

15 times higher than non-bioaugmented 8.0 V electrolysis, and 2.5 times higher than the

312

bioaugmented control reactors without applied potential. The improvement of electro-

313

biodegradation rates over biotic treatment alone would be expected to be even greater had we

314

been able to achieve maintain completely anoxic conditions in preparing the feedstock solutions.

315

The dramatic increase in 1,4-dioxane removal rate has to be attributed to anodic O2 generation

316

resulting in an advantageous environment for aerobic biodegradation processes42 since the

317

coupled treatment regime at 3.0 V performed significantly better than either treatment on its own.

318

The manifestation of a highly oxidative environment was evidenced by the considerable

319

generation of O2 gas bubbles at the anode and by elevated solution ORP measurements at the

320

anode of 1.8 V and 4.4 V vs. SHE for the electro-bioreactors at 3.0 and 8.0 V, respectively (Table

321

1). We were unable to perform DO measurements in these reactors; however, DO concentrations

322

measured in analogous bioaugmented reactors with TCE present exhibited extraordinarily high

323

concentrations of 11.0 and 15.7 mg/L at 3.0 and 8.0 V, respectively (Table S2). Since ORP

324

measurements in Table 1 show comparable mean values near 1.7 ± 0.3 V for all experiments with

325

3.0 V applied, it is reasonable to use ORP as a proxy for revealing dissolved oxygen shifts and

326

assume that all 3.0 V reactors generated DO concentrations in the range of 5 to 11 mg/L. Hence,

327

by providing O2 as a terminal electron acceptor for aerobic metabolism, concurrent electrolysis

328

and biodegradation processes outperformed the degradation efficiencies of all other regimes

329

tested. 13 ACS Paragon Plus Environment


330 331 332

Page 14 of 28

Table 1. Physicochemical parameters measured along the column flow path. Data is divided into 1,4-dioxane degradation experiments conducted without TCE (a) and with 5 mg/L of TCE cocontaminant (b).

333 (a)

Non-bioaugmented, 1,4-dioxane (-) TCE Sample port

2

3V and 0.2 mA/cm *

Bioaugmented with CB1190, 1,4-dioxane (-) TCE

2

2

8V and 2.3 mA/cm

0V and 0 mA/cm

2

3V and 0.3 mA/cm

2

8V and 1.6 mA/cm

distance downgradient

pH

ORP (V)

pH

ORP (V)

pH

ORP (V)

pH

ORP (V)

pH

ORP (V)

influent

7.0

0.6

7.0

0.6

6.8

0.7

6.8

0.98 ± 0.5

6.8

0.98 ± 0.5 4.4

S1 = 2.5 cm

8.4 ± 3.1

1.6

9.1 ± 2.9

4.2

6.8

0.3

4.7

1.8

3.1 ± 0.8

S2 = 12.5 cm

6.9

-1.0

6.8

-1.1

6.8

0.1

6.5

0.1

5.8

0.0

S3 = 22.5 cm

7.0

0.1

6.9

-0.1

6.8

-0.2

6.7

0.1

6.3

-0.2

effluent =32.5 cm

6.9

0.3

6.8

0.1

-0.2

6.8

0.2

6.3

0.2

standard deviation**

± 0.3

± 0.05

± 0.3

± 0.05

6.8 ± 0.3

± 0.2

± 0.3

± 0.2

± 0.3

± 0.3

(b)

Non-bioaugmented, 1,4-dioxane (+) TCE Sample port

distance downgradient

2

3V and 0.2 mA/cm * pH

ORP (V)

Bioaugmented with CB1190, 1,4-dioxane (+) TCE 2

0V and 0 mA/cm2

8V and 1.2 mA/cm pH

ORP (V)

pH

ORP (V)

3V and 0.2 mA/cm2 pH

ORP (V)

8V and 1.1 mA/cm2 pH

ORP (V)

influent

7.0

0.6

7.0

0.6

7.0

0.5

7.0

0.5

7.0

0.5

S1 = 2.5 cm

6.5 ± 1.7

1.7

2.3 ± 0.5

3.7

7.0

0.2

7.3 ± 0.6

1.6

5.3

S2 = 12.5 cm

7.5 ± 1.7

0.0

9.0

-0.2

7.0

-0.1

7.0

0.1

12.0 11.3 ± 1.2

S3 = 22.5 cm

7.0

0.2

6.8 ± 0.5

0.3

7.0

-0.1

7.0

0.3

9.5 ± 0.9

0.4

effluent =32.5 cm standard deviation**

6.6

0.3

±0.3

± 0.03

5.5 ± 0.3

0.0

0.4

7.0

-0.1

7.0

0.7

4.2

0.8

± 0.2

±0.3

± 0.1

± 0.3

± 0.2

± 0.3

± 0.4

ORP, oxidation reduction potential vs. SHE (standard hydrogen electrode) *all current densities are averages of n =3, with uncertainties less than ± 0.01 mA/cm2 **the standard deviation (n=3) is explicitly written for measurements when exceeding the common one shown below each column

334

335

Although electrolysis provides O2 necessary for aerobic biodegradation, electrochemical

336

processes may also cause cellular death associated with the production of biologically destructive

337

hydroxyl radicals (SI, Reactions 1 to 6)53 or highly oxidized chlorine species (e.g., hypochlorous

338

acid) and other DBPs.37,54 In addition, considerable pH fluctuations near the electrodes may be

339

harmful to microbial populations as well. Electrolysis commonly produces an acidic boundary

340

layer at the anode surface due to the oxidation of water molecules to form H+ ions, and more

341

alkaline conditions surround the cathode due to H+ reduction to H2 gas and production of OH-

342

ions in solution.55 Hence, it was important that we investigated the impacts of these potentially

343

adverse conditions on the viability of CB1190 by analyzing both liquid (planktonic) and

344

solid/sand (biofilm) aliquots at various positions along the column reactor to locate zones of

345

microbial inhibition or proliferation adjacent to, or downstream of, the anode. Quantitative PCR 14 ACS Paragon Plus Environment

Page 15 of 28


346

data on the dxmB gene biomarker was used as a proxy for CB1190 cell abundance to elucidate

347

preferential location(s) of microbial proliferation, either in biofilm or planktonic forms, leading

348

to a better understanding of optimum spacing between multiple electrode pairs or reactor

349

dimensions.

350

Upon CB1190 inoculation of the sand used in columns for the 0 V, 3.0 V and 8.0 V

351

treatments of 1,4-dioxane in the absence of TCE, initial measurements of mean abundance values

352

after packing the reactors were 4.1×108 cells/mL for planktonic form and 1.3×107 cells/g for

353

CB1190 present as biofilm on sand particles (Figure 3 a,b). These levels are thought to represent

354

maximum population density achievable on this sand media since the cultures were growing

355

under ideal conditions up until this point. Considering the total volume of water and total mass of

356

sand in the reactors, we note that similar to previous reports,56 the planktonic biomass was equal

357

to or slightly greater than biofilm-associated biomass.

358

After two weeks of flow-through operation in the biological control (0 V) reactor, the pH

359

remained stable at 6.8, the planktonic CB1190 abundances dropped from 3.7×108 cells/mL at

360

port L1 to 1.0×107 at the effluent site down gradient. The biofilm levels also dropped by two

361

orders of magnitude down to 1.5×105and 2.1×105 cells/g at solid sample ports S3 and S5,

362

respectively. Thus, CB1190 growth rates were at least sufficient enough to overcome the steady-

363

state biomass losses of 1.0×107 cells/mL measured in the effluent caused by hydraulic shear

364

forces.57 Although these losses in microbial populations were observed, the remaining CB1190

365

was in quantities high enough to produce the 1,4-dioxane biodegradation rate observed here of

366

68.7 mg·h-1·m-2.18,22 Table 1 shows that ORP values along the column were steadily decreasing

367

from 0.7 to -0.2 V vs. SHE, likely due to biochemical oxygen demand for CB1190’s cellular

368

respiration. This decrease in CB1190 communities correlated to more negative ORP levels would

369

add credence to the need for adding charged electrodes to generate beneficial oxygen

370

downgradient.

371



Page 16 of 28

372 373 374 375 376 377 378 379 380

Figure 3. P. dioxanivorans CB1190 bioaugmented plots showing planktonic abundance (a,c) and biofilm abundance (b,d) of CB1190 cells along the horizontal flow path in the absence (a,b) and presence (c,d) of TCE co-contaminant, plotted as a function of distance from the anode. Sampling ports 1, 2, 3 and effluent correspond to 2.5, 12.5, 22.5 and 32.5 cm downstream from the anode. Any samples that were below the qPCR method limits of detection (LOD), 10 cells/mL for liquid samples and 40 cells/g for solid samples, were plotted just below LOD line. The error bars represent the average range of qPCR duplicates (2 analytical replicate analyses of each duplicate sample).

381 382

At the low stimulation voltage of 3.0 V, ORP levels reached 1.8 V vs SHE and pH levels

383

dropped to 4.7 between the electrodes, which is near the growth range previously reported for

384

CB1190 from pH 5.0 - 8.0.17 After the cathode, the pH rapidly returned to the starting influent

385

pH of 6.8 (Table 1).

386

experimental and modeling studies have demonstrated that their high reactivity limits their

387

existence to a narrow zone of less than 1 µm from the anode surface.55,58-60 Consequently, any

Although free radicals like •OH can be damaging to cellular life,53


Page 17 of 28


388

adverse conditions caused by the presence of •OH are only expected in the immediate vicinity of

389

the anode. Despite these risks, high mean planktonic abundances near 1.5×108 cells/mL along the

390

reactor flow path revealed that the overall conditions remained conducive to cellular growth

391

(Figure 3 a,b) and reported greater overall 1,4-dioxane degradation rates than biological control

392

rates. Our measurements revealed that CB1190 populations were not homogenous at different

393

distances from the anode. At sample port 1, between the electrodes, biofilm counts were below

394

the qPCR detection limit (BDL, < 40 cells/g); and planktonic cell counts dropped slightly from

395

4.1×108 initially to 6.8×107 cells/mL after equilibrating to the 3.0 V system for two weeks. The

396

subsequent CB1190 population rebound to 3.0×108 planktonic cells/mL and 1.3×107 sessile

397

cells/g at the location 12.5 cm downstream of the anode reveals a possible “sweet spot” for

398

bacterial proliferation along the flow path where dissolved oxygen is still abundant (likely in the

399

4 - 7 mg/L range) and the distance away from the electrodes was sufficient to avoid deleterious

400

effects. Physicochemical measurements in Table 1 show both locations 12.5 and 22.5 cm

401

downstream of the anode support this hypothesis in that solution ORP was 0.1 V vs SHE and the

402

pH returned to circumneutral (6.5-6.7). Biofilms are dynamic microbial communities capable of

403

interchangeable transitions between sessile and planktonic modes of growth as a response to

404

different environmental cues, including carbon source, oxygen saturation and pH of the

405

media.61,62 The existence of localized maximum cell counts of both biofilm and planktonic forms

406

could indicate that environmentally-tolerant biofilms may be the foundation of a stable CB1190

407

microbial community, with dispersed planktonic cells primarily responsible for higher rates of

408

carbon (1,4-dioxane and transformation products) metabolism. This type of survival strategy has

409

been documented in other bacteria and could warrant further study into the predominance of the

410

role played by these planktonic CB1190 cells dispersed from densely packed and specialized

411

biofilm subpopulations.63,64

412

When a higher potential of 8.0 V was applied, an elevated solution ORP of 4.4 V vs.

413

SHE was measured after the anode, which can be explained by the expected increase in anodic

414

generation of O2 and other reactive oxygen species (ROS) due to the higher voltage and current

415

density.55 We hypothesized that this occurrence, in combination with a drastic drop in pH from

416

6.8 to 3.1, could create conditions less hospitable for microbial growth and/or survival. This was

417

confirmed with measurements between the electrodes showing biofilm levels dropped to below

418

the qPCR detection limit and a substantial 4 order of magnitude decrease in planktonic 17 ACS Paragon Plus Environment


Page 18 of 28

419

abundance was observed as well (Figure 3 a,b). And although planktonic cells did rebound to

420

2.3×108 cells/mL 12.5 cm downstream from the anode, biofilm counts remained below LOD for

421

the remainder of the column. Thus, the decline in both life cycle modes of CB1190 communities

422

in the 8.0 V column is understood to be the principal reason for why lower overall 1,4-dioxane

423

degradation rates were obtained from the 8.0 V column compared with the 3.0 V column. In

424

addition, the lower degradation rates provide further support for the probable necessity of a

425

critical mass of biofilm being present, acting as a base for planktonic dispersal, in order for

426

optimal degradation rates to be achieved.

427

Finally, we note that ~104 counts (3.0 V) and ~102 counts (8.0 V) of dxmB genes were

428

detected in the non-bioaugmented reactor experiments (Figure S4). However, these data points

429

may be due to the fact that non-bioaugmented experiments were performed after augmented ones

430

for logistical reasons. Thus, we suspect that these detections are the result of extracellular DNA

431

released from lysed CB1190 cells during the column disinfection and sand heat-sterilization

432

process.

433

TCE Co-contaminant Impact on 1,4-Dioxane Degradation. Non-bioaugmented

434

electrolytic experiments performed in the presence of TCE co-contaminant (Figure 2) exhibited

435

1,4-dioxane degradation rates of 0.06 ± 13.9 mg·h-1·m-2 at 3.0 V (maximum ORP 1.7 V vs. SHE)

436

and 6.4 ± 13.9 mg·h-1·m-2 at 8.0 V (maximum ORP 3.7 V vs. SHE). These outcomes were not

437

significantly different from each other (p = 1.0), nor were they significantly different from

438

equivalent experiments spiked solely with 1,4-dioxane (p = 0.79 at 3.0 V, p = 0.92 at 8.0 V)

439

because rates of abiotic (electrolytic) 1,4-dioxane removal were controlled by mass transfer.58,65

440

In addition, 1.4 mg/L TCE (27% of the initial 5 mg/L) was removed at 3.0 V and 3.0 mg/L TCE

441

(60%) at 8.0 V (Figure 2 and Figure S5), showing higher electrolytic removal than for 1,4-

442

dioxane. This is expected because 1,4-dioxane degradation rates are limited to anode-generated

443

•

444

in addition to reductive dechlorination by direct electron transfer at the cathode. Additionally,

445

overall degradation kinetics for TCE can be enhanced by favorable surface chemistry interactions

446

with IrO2 electrode surfaces.47 GC/MS(SIM) analysis of effluent did not detect any DCE isomer

447

or vinyl chloride intermediates. Furthermore, no other organic transformation products appeared

448

as new chromatographic peaks in GC/MS full scan analysis, signifying that TCE was

449

mineralized, or volatilized/stripped by the effervescent oxygen and hydrogen gas bubbles.

OH radical oxidation pathways, whereas oxidation of TCE can occur via •OH radical pathways


Page 19 of 28


450

Previous studies on the electrochemical treatment of TCE have reported carbon dioxide, carbon

451

monoxide, chloride, formate, acetate, and chloroxyanions as stable products, but failed to detect

452

organochlorine intermediates, indicating that these may be too short-lived to accumulate in the

453

bulk solution.27,35

454

In the 0 V biological control column at 46 cm/d flow velocity, the addition of 5 mg/L

455

TCE to the 100 mg/L 1,4-dioxane feedstock lowered 1,4-dioxane biodegradation rates from 68.7

456

mg·h-1·m-2 (in the absence of TCE) to 37.5 mg·h-1·m-2 (Figure 2). This outcome is supported by

457

previously mentioned studies that show inhibitory effects of TCE on the biodegradation rates of

458

CB1190.22 Steady-state analysis revealed that no TCE removal had occurred, confirming

459

previous reports that CB1190 is not capable of biodegrading TCE20 and serving as an

460

experimental control that sorption did not contribute to TCE removal from the aqueous phase

461

after steady-state flow-through conditions had been achieved. The pH of 6.8 remained constant

462

throughout the entire biological control column and the solution ORP decreased from 0.5 in the

463

influent down to -0.1 V vs. SHE, analogous to our earlier experiments performed in the absence

464

of TCE (Table 1).

465

In the mixed contaminant experiments, 1,4-dioxane oxidation rates were again highest

466

when both abiotic and biodegradation processes were combined, achieving rates of 98.4 mg·h-

467

1

468

was minor in non-bioaugmented experiments; thus, the vast majority of 1,4-dioxane oxidation

469

observed here can be attributed to electrolytic enhancement of aerobic biodegradation rates. The

470

improved treatment is understood to be the result of increased biodegradation kinetics stimulated

471

by anodic production of molecular oxygen and the simultaneous benefit from electrochemical

472

removal of the inhibitive TCE co-contaminant. This claim is supported by DO measurements. As

473

stated above, O2 concentrations between the electrodes at port L1 increased to 11.0 mg/L at 3.0 V

474

and 15.6 mg/L at 8.0 V compared to 2.4 mg/L in the hypoxic bioaugmented control column, and

475

remained above 6 mg/L throughout the entire column reactors, while the biological control

476

experiment showed steadily decreasing DO values down to 1.9 mg/L along the flow path (Table

477

S2). Although DO measurements were not made in the experimental treatments of 1,4-dioxane in

478

the absence of TCE, the similarly decreasing trend in ORP values correlated to distances farther

479

downstream of the electrodes (Table S2) would signify that DO concentrations would also

480

decrease downstream. However, it may be critically important to note that when TCE is present

·m-2 at 3.0 V and 94.5 mg·h-1·m-2 at 8.0 V. Abiotic electrochemical oxidation of 1,4-dioxane



Page 20 of 28

481

as an inhibitor to CB1190 metabolism, it appears that the decreased DO levels along the flow

482

path could cause the precipitous drop in planktonic populations to below 2.0×108 cells/mL

483

measured at the effluent ports in 0V, 3V and 8.0 V reactors (Figure 3 c). Therefore, in situ

484

treatment of hypoxic waters (e.g. groundwater and wastewater), with much longer hydraulic

485

retention times would benefit from repeating intervals of electrodes to maintain high levels of

486

dissolved oxygen and high biodegradation rates.

487

At the higher stimulation voltage (8.0 V), 3.1 mg/L (62%) of TCE was removed (Figure

488

2 and Figure S5), minimizing adverse effects on biodegradation by CB1190 such that 1,4-

489

dioxane removal rates were nearly equivalent to rates when TCE was not present (94.5 compared

490

to 101 mg·h-1·m-2, p = 0.90). The recovery in metabolic biodegradation rates observed here is

491

consistent with conclusions by Zhang and co-workers (2016) that inhibition was dose-dependent

492

and reversible once chlorinated compounds were removed.22 Despite the removal of 2.05 mg/L

493

TCE (41%) with 3.0 V applied, the remaining 2.9 mg/L TCE appeared to decrease 1,4-dioxane

494

biodegradation efficiency by 40% under these conditions. Overall, our data demonstrate that

495

aerobic biodegradation rates can be enhanced by upstream electrolysis, and that complete

496

removal of TCE is not required to obtain effective electro-biodegradation of 1,4-dioxane;

497

although more investigation is needed to better understand whether a certain low threshold

498

concentration for TCE relative to CB1190 abundance may be needed to eliminate any noticeable

499

inhibition effects.

500

Adverse microbial impacts are observed with the presence of TCE since qPCR analyses

501

for the bioaugmented experiments revealed that mean planktonic cell counts under all voltage

502

conditions were generally orders of magnitude lower than comparable experiments in the absence

503

of TCE (Figure 3 a-d, Table S4).20-22 In contrast, the mean biofilm counts in the presence of TCE

504

tended to be higher than mean values when TCE was not present, possibly indicating a stress

505

response to toxicity posed by the presence of chlorinated solvents.66 Physicochemical

506

measurements also reinforced our expectation for less-viable conditions near the 8.0 V electrode.

507

For mixed contaminant experiments, the 8.0 V bioaugmented reactor recorded a highly oxidative

508

ORP of 5.3 V vs. SHE, along with highly alkaline pH levels of 12.0, 11.3 and 9.5 being

509

measured at 2.5, 12.5, and 22.5 cm downstream of the anode; whereas, in the 0 and 3.0 V

510

bioaugmented reactors the ORP values did not exceed 1.6 V vs. SHE and the pH remained

511

between 7.0 - 7.3 (Table 1). Much lower CB1190 planktonic abundance (5.2 x 103 cells/mL) was 20 ACS Paragon Plus Environment

Page 21 of 28


512

observed between the electrodes when 8.0 V was applied, when compared to the 0 V (1.7 x 105

513

cells/mL) and 3.0 V (1.6 x 105 cells/mL) treatments (Figure 3 c,d). Thus, it is plausible that

514

sparse CB1190 populations in the 8.0 V experiment are caused by TCE inhibition of

515

growth/metabolism compounded by the rapid increase in pH and ORP levels, and potential for

516

greater abundance of reactive oxygen species.18 The high pH level does not appear to be caused

517

by reaction pathways isolated to having TCE present, since both of the non-bioaugmented

518

experiments without TCE had alkaline pH values of 8.4 and 11.0 for 3.0 and 8.0 V, respectively.

519

Our experimental results were not able to produce a definitive trend or explanation for why some

520

experiments were acidic at the 2.5 cm sampling location while others were basic. Since the pH

521

continuum from the acidic anode to the alkaline cathode just 5.0 cm away is very steep

522

(especially with higher voltages), it is conceivable that variability in flow and/or mixing in the

523

inter-electrode space could contribute to the range of pH values observed. It is also feasible that

524

a high inter-electrode pH may arise when CO2 gas, present in the influent or generated from

525

organic carbon mineralization, is effervescently released through the gas vents at the acidic

526

anodes, and thus no longer has the buffering capacity provided by carbonic acid.

527

The sessile CB1190 counts for bioaugmented columns at 0 V and 3.0 V were able to

528

maintain dense biofilms between the electrodes, measuring 3.4 x 107 and 2.1 x 106 cells/g,

529

respectively. The higher biofilm counts relative to the planktonic counts in this region between

530

the electrodes can be explained by biofilms generally having a greater ability to withstand pH

531

changes and short-lived reactive oxygen species.67 The aqueous conditions within the 8.0 V

532

experiment appear to have been too harsh to establish stable biofilm communities, such that

533

sessile counts were below LOD between the electrodes (and at 22.5 cm downstream) with

534

biofilm counts only being observed at the 12.5 cm “sweet spot” reaching 2.3 x 106 cells/g (Figure

535

3 d). This becomes a disadvantage if biodegradation activity is dependent on newly dispersed

536

planktonic cells from biofilms as the primary source of active cells. Interestingly, there was a

537

substantial spike in sessile cell abundance at 12.5 cm downstream experienced in both the 3.0

538

and 8.0 V electro-bioreactors, reaching 107-108 cells/g. This would lend further support to our

539

“sweet spot” hypothesis for optimum bacterial growth. Based upon the consistency of the

540

location of peak sessile cell counts just downstream of the electrodes, it appears that biofilm

541

formation is being used as a strategy of colonization in a nutrient-dense (1,4-dioxane carbon

542

source), oxygen-enriched, and circumneutral pH region (usually pH 6.4 ± 0.6), where dispersion 21 ACS Paragon Plus Environment


Page 22 of 28

543

of actively metabolizing planktonic cells can then occur, providing the majority of observable

544

biodegradation.67

545

Technological

Implications.

Our

results

demonstrate

that

the

coupling

of

546

electrochemical oxidation with aerobic biodegradation is an effective, synergistic approach for

547

the treatment of waters contaminated with 1,4-dioxane, and may provide an effective solution to

548

the problems associated with co-occurring inhibitors. We did not experience clogging in any of

549

the experiments, indicating that bioaugmentation of aquifers with CB1190 is a viable strategy.

550

Successful establishment of CB1190 biofilm is expected to be possible in a range of flow

551

regimes. Although tests at different flow rates were outside the scope of this study, other research

552

results have shown that bacterial biofilm thickness and survivability did not change significantly

553

with altered flow rates.57,68 In the absence of a co-contaminant, where the enhancement is mainly

554

based on oxygen generation, low voltages (i.e., 3.0 V) slightly above the oxygen evolution

555

potential of the electrode are preferable to avoid extreme pH and redox conditions unfavorable

556

for microbial growth, and may be a viable strategy to limit DBP formation. Furthermore, lower

557

voltages are preferable due to longer electrode service life, reduced power cost, and more

558

uniform flow with less gas bubbles.37,54,55

559

When microbially-inhibiting co-contaminants such as TCE are present, more electrodes

560

may be required in order to mineralize or transform the additional contaminants into less toxic or

561

more readily biodegradable intermediates.33,69 If the chlorinated co-contaminants are not fully

562

removed during electrolytic treatment, a subsequent reductive treatment step may be effective,

563

which could also aid in the removal of any generated DBPs.70

564

This study provides convincing evidence that 1,4-dioxane metabolism by CB1190 can be

565

electrochemically enhanced despite the production of strong oxidants at the anode. Our data

566

suggest that microbial abundance is lowest between the electrodes, likely due to destructive

567

effects of short-lived chemical oxidants, and highest just downstream of the trailing cathode (near

568

12.5 cm downstream of the anode) where dissolved oxygen concentrations stimulate microbial

569

metabolism. Thus, microbial populations are able to thrive downstream of the electrodes, and the

570

generation of strong oxidants (e.g., •OH) can provide significant concomitant benefits of

571

oxidizing recalcitrant and/or toxic co-pollutants in addition to 1,4-dioxane.

572

For technology implementation, specific aquifer conditions (e.g., contaminant

573

concentrations, water quality parameters, soil composition, and indigenous microbial 22 ACS Paragon Plus Environment

Page 23 of 28


574

communities) or wastewater composition need to be taken into consideration and tested to

575

validate contaminant degradation efficiencies. These preliminary test results and specific

576

treatment targets would guide decisions regarding the number of sequential electrodes required

577

(more electrode surface area for more contaminant mass removal), but our results suggest that

578

sufficient spacing between electrode pairs should be arranged to promote maximum microbial

579

activity in the favorable downstream conditions. It is conceivable that additional synergistic

580

mechanisms could also be promoted, such as stimulation of other intrinsic microbes capable of

581

metabolizing transformation products and co-contaminant parent compounds71 or electrolytic

582

transformation

of

a

27,33,72-74

persistent

parent

compound

into

more

readily

biodegradable

583

intermediates.

Although other mixed contaminants waters were not explored in this

584

study, the capability of this technology being effective in a flow-through system could lend itself

585

to feasible treatment applications for landfill leachate, domestic wastewater, and industrial

586

effluent as well.69,75 The joint benefits of having tunable electrolysis to degrade recalcitrant

587

pollutants while simultaneously supporting intrinsic or augmented contaminant-degraders may be

588

one innovative approach needed to overcome the technological and cost limitations faced when

589

treating waters with mixed, persistent contaminants.

590 591 592 593

AUTHOR INFORMATION

594

Corresponding Author* E-mail: [email protected]. Phone: +1-970-491-8880. Fax:

595

+1-970-491-8224.

596

Notes: The authors declare no competing financial interest.

597 598 599

ACKNOWLEDGMENTS

600

Funding for this work was provided by The Chemours Company (to J.B.), E. I. du Pont de

601

Nemours and Company (to J.B. and contract no. MA-03653-13 to S.M.) and The Dow Chemical

602

Company (contract no. 244633 to S.M.). We thank Maria Irianni-Renno for assistance with ATP

603

analysis, and Michelle Myers and Shu Zhang for assisting with DNA extractions.

604 23 ACS Paragon Plus Environment


Page 24 of 28

605

ASSOCIATED CONTENT

606

Supporting Information. The Supporting Information is available free of charge on the ACS

607

Publications website at DOI: …

608

It includes schematics and photos of the experimental design, details on analytical procedures

609

and CB1190 inoculation procedures, and calculations used to determine flow-through

610

degradation rates and instrumental uncertainty. Additional plots and tables show TCE removal,

611

DO measurements, and pH values along the column reactor path.

612 613 614

REFERENCES

615 616 617 618 619 620 621 622 623 624 625 626 627 628 629 630 631 632 633 634 635 636 637 638 639 640 641 642 643 644 645 646 647

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Synergistic Treatment of Mixed 1,4-Dioxane and Chlorinated Solvent

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