Natural attenuation and anaerobic benzene detoxification processes

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Natural attenuation and anaerobic benzene detoxification processes at a chlorobenzene-contaminated industrial site inferred from field investigations and microcosm studies Wenjing Qiao, Fei Luo, Line Lomheim, Elizabeth Erin Mack, Shujun Ye, Jichun Wu, and Elizabeth A. Edwards Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b04145 • Publication Date (Web): 27 Nov 2017 Downloaded from http://pubs.acs.org on November 27, 2017

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Environmental Science & Technology

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Natural attenuation and anaerobic benzene

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detoxification processes at a chlorobenzene-

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contaminated industrial site inferred from field

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investigations and microcosm studies

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Wenjing Qiao1,2, Fei Luo2, Line Lomheim2, Elizabeth Erin Mack3, Shujun Ye1*, Jichun Wu1,

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Elizabeth A. Edwards2*

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1

Key Laboratory of Surficial Geochemistry, Ministry of Education; School of Earth Sciences and

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Engineering, Nanjing University, Nanjing 210046, China; 2

Department of Chemical Engineering and Applied Chemistry, University of Toronto, Toronto

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M5S 3E5, Canada 3

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DuPont Corporate Remediation Group, Wilmington, Delaware 19805, USA

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*Corresponding author: Shujun Ye: Tel. (+86) 2589684150; Fax (+86) 2583686016; email:

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[email protected]; Elizabeth A. Edwards: Tel. (+1) 4169463506; Fax (+1) 4169788605; email:

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[email protected];

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ACS Paragon Plus Environment


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Abstract: A five-year site investigation was conducted at a former chemical plant in Nanjing,

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China. The main contaminants were 1,2,4-trichlorobenzene (TCB) reaching concentrations up to

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7300 µg/L, dichlorobenzene (DCB) isomers, monochlorobenzene (MCB) and benzene. Over

19

time, these contaminants naturally attenuated to below regulatory levels under anaerobic

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conditions. To confirm the transformation processes and to explore the mechanisms, a

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corresponding laboratory microcosm study was completed demonstrating that 1,2,4-TCB was

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dechlorinated to 1,2-DCB, 1,3-DCB and 1,4-DCB in approximately 2%/10%/88% molar

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proportions. The DCB isomers were dechlorinated via MCB to benzene, and finally benzene was

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degraded under prevailing sulfate-reducing conditions. Dechlorination could not be attributed to

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known dechlorinators Dehalobacter or Dehalococcoides, while anaerobic benzene degradation

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was mediated by microbes affiliated to a Deltaproteobacterium ORM2, previously associated

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with this activity. Unidentified organic compounds, possibly aromatic compounds related to past

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on-site production processes, were fueling the dechlorination reactions in situ. The microcosm

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study confirmed transformation processes inferred from field data and provided needed

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assurance for natural attenuation. Activity-based microcosms studies are often omitted from site

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characterization in favor of rapid and less expensive molecular surveys. However, the value of

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microcosm studies for confirming transformation processes, establishing electron balances,

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assessing co-contaminant inhibition and validating appropriate monitoring tools is clear. At

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complex sites impacted by multiple compounds with poorly characterized transformation

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mechanisms, activity assays provide valuable data to incorporate into the conceptual site model

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to most effectively inform remediation alternatives.

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1. Introduction

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Chlorinated benzenes (CBs) have been widely released into the environment as a result of their

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intensive use in the manufacture of pesticides, herbicides, dyes and other chemicals, as solvents

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or reactants.1 CBs persist in sediment, accumulate in the food chain and are potential

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carcinogens.2, 3 Air sparging4, 5 combined with soil vapor extraction,6 groundwater pump-and-

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treat,7 granular iron permeable reactive barriers8 and bioremediation9 have all been used to

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remediate sites contaminated with volatile organic compounds (VOCs), including CBs. Of these

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alternatives, bioremediation is a cost-effective and long term solution in many cases,10,

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although uncertainties in the rate and extent of transformation often limit application.

11

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CBs can be biodegraded aerobically and anaerobically. Aerobic CB biodegradation has been

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studied extensively. For example, microbes belonging to Burkholderia12, Enterobacter13,

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Pseudomonas13-16 can grow on CBs as the sole carbon and energy source17. In situ, the

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availability of dissolved oxygen is often limiting due to low water solubility18 and rapid

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consumption by aerobic microorganisms,19 and the difficulty and expense associated with

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injecting sufficient oxygen underground.20 Moreover, CBs may form dense non-aqueous phase

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liquids (DNAPLs) and migrate to deep anaerobic regions such as confined aquifers,21-24 making

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them inaccessible to aerobic degradation. Thus, anaerobic transformation processes are important

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determinants of the fate of these compounds in situ. Anaerobically, CBs serve as terminal

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electron acceptor for respiration and undergo sequential reductive dehalogenation to less

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chlorinated products. Anaerobic reductive dechlorination of CBs has been studied in a variety of

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mixed and some pure cultures derived from sewage, sediment, soil or sludge. Dehalococcoides

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mccartyi

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tetrachlorobenzenes (except 1,2,3,5-TeCB) to 1,3,5-trichlorobenzene (TCB).25,

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dechlorinates

hexachlorobenzene

(HCB),


pentachlorobenzene 26

(QCB),

Dehalobium


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1,3,5-TCB.27

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chlorocoercia

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Dehalococcoides mccartyi CBDB1 dechlorinates CBs with three or more chlorines to 1,3,5-

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TCB, and then to 1,3- and 1,4-dichlorobenzene (DCB).28-30 D. mccartyi DCMB5 dechlorinates

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HCB to 1,2,4-TCB, 1,3,5-TCB and 1,4-DCB.31 Comparatively few studies report microbes

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responsible for dechlorination of lesser chlorinated DCB isomers and monochlorobenzene

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(MCB). Enrichment cultures containing three different Dehalobacter spp. dechlorinated all

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chlorobenzene isomers to MCB and benzene.21-23 An anaerobic consortium capable of

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dechlorinating all dichlorobenzene isomers except 1,4-DCB to benzene was recently described,

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possibly involving a Geobacter sp.24 Complete dechlorination of CBs results in the accumulation

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of benzene, which is a known human carcinogen32 and much more toxic than any of its

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chlorinated parent compounds; thus dechlorination does not always lead to detoxification.

DF-1

dechlorinates

HCB,

QCB,

and

1,2,3,5-TeCB

to

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The transformation of benzene to CO2 completes the detoxification process for chlorinated

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aromatics. While benzene is readily degraded aerobically, anaerobic benzene biodegradation is

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much less well understood, and the metabolic pathway is still unknown.33 Benzene can be

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anaerobically metabolized under iron-reducing,34 nitrate-reducing,35, 36 sulfate-reducing,37, 38 and

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methanogenic conditions, and the microbes responsible have been identified in some cases.39, 40

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For example, a Deltaproteobacterium referred to as ORM2 was shown to be the benzene-

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metabolizing population under sulfate and methanogenic conditions40. An enrichment culture

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capable of reductive dechlorination of MCB to benzene was mixed with a benzene degrading

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methanogenic culture (containing ORM2) demonstrating transformation of MCB beyond

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benzene to nontoxic CO2 and CH4 in lab cultures. 41

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Natural attenuation at sites contaminated with chlorinated ethenes and ethanes has been studied

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extensively.42-46 However, at chlorobenzene-contaminated sites, most of previous research has

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focused on aerobic oxidation15, 47-49 whereas comparably less information is available on anoxic

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processes 19, 50, especially documenting complete detoxification of chlorobenzenes. In this study,

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a five-year site investigation was conducted at a former chemical plant contaminated with 1,2,4-

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TCB and many other compounds. In parallel, a two-year anaerobic laboratory microcosm study

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was completed to provide a solid basis on which to base a remediation strategy for 1,2,4-TCB

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and its daughter products at this site.

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2. Materials and Methods

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2.1 Field Site Description and Investigations

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The site in Nanjing, China, was the location of a former chemical plant that manufactured 2-

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nitro-4-methylphenol, 2-amino-4-methylphenol and the optical brightener 2,2'-[(E)-1,2-

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Ethenediyl]bis(5-methyl-1,3-benzoxazole) (Figure S1) from 1999 to 2010. The ~9,500 m2 site

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contained the former production facilities and waste and chemical storage areas in the North, and

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a former office room to the South (Figure S2).

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The site investigation consisted of a series of sampling campaigns performed between 2012 to

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2016 (Table S1, Figure S3). Detailed soil boreholes and monitoring well information, and

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sampling collection methods are provided in Table S1 and additional details are provided in the

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Supporting Information (SI). A preliminary investigation in June 2012 delineated contaminants

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and their distribution. Seven shallow groundwater samples were taken and analyzed for VOCs,

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heavy metals, and total cyanide. An additional site investigation was conducted in May 2013. At

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this time, a membrane interface probe (MIP) test was conducted to provide real-time direct

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measurements of geological conditions and volatile organic compound distributions at depth

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(details provided in the SI). Groundwater parameters, including pH, dissolved oxygen and ORP,

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were measured. Seven supplementary groundwater samples were taken and analyzed for semi-

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VOCs and VOCs. Slug Tests were performed to determine the hydraulic conductivity.

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Subsequent third and fourth sampling rounds were conducted in September 2014 and October

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2016 where only two wells could be sampled because the other monitoring wells had been

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backfilled by the local government. Semi-VOCs and VOCs were analyzed in these samples.

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Analyses of field samples were carried out by commercial laboratories (Table S1).

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2.2 Laboratory Microcosm Study

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Samples and Reagents. Sediment and groundwater samples were taken in July 2014 to set up

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the laboratory microcosms. Sediment samples were taken at depths 1.5-2 m below ground

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surface close to borehole S1, which was thought to have highest dechlorinating activity, and

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were stored in aluminum core liners, sealed with parafilm immediately to keep anoxic.

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Groundwater samples were collected at the same time from well W4 in dark brown glass bottles

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filled to the top. All samples were kept in a cooler with ice packs on site, and then were stored

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refrigerated and sent to the laboratory in Canada by express delivery.

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CBs, benzene, methanol, ethanol, acetone and sodium lactate were purchased from Sigma-

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Aldrich at the highest purity available (98% for 1,3-DCB and sodium lactate, ≥99% purities for

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all other chemicals). Sodium lactate was dissolved in sterile anaerobic Milli Q water to a 6%

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concentration by mass. ACS grade ferrous sulfate was purchased from British Drug House (now

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part of Merk). Ferrous sulfate was dissolved in 0.1 M HCl solution (Aldrich) to make a 500 mM

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solution. Potassium hydrogen phthalate was purchase from Fisher Scientific with ≥98.5 % purity.

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Anaerobic Microcosm Study. Sediment and groundwater were used to set up microcosms in

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250 mL bottles, with details of the setup provided in the SI. Four treatments were prepared: 1)

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sterile/poisoned controls (Bottles 1-4); 2) in-situ conditions (Bottles 5-8); 3) electron donor-

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amended treatment (Bottles 9-12); and 4) a benzene only treatment (Bottles 13-16) (Table S2).

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Microcosms were amended and sampled as described below.

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The first three treatments (sterile, in-situ conditions, and donor-amended) initially received a

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“CB mixture” (consisting of 1,2,4-TCB, 1,2-, 1,3-, 1,4-DCBs and MCB) at approximate in situ

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concentrations of 29, 3.6, 3.8, 15 and 18 µM, respectively. Subsequently, these CB-amended

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active treatments were spiked with benzene on Day 80 to a concentration of 7 µM to test if other

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compounds interfere with benzene degradation. The “donor-amended” set received a mixture of

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Methanol, Ethanol and 6% sodium Lactate (“MEL”) at a ratio of 7/5/122 by volume to provide

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equal electron equivalents of each donor. The volume of MEL added was calculated to provide

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an electron donor to chlorinated electron acceptor (CBs) ratio of 10:1 initially, and 5:1 where

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noted. Once degradation was observed, selected compounds were re-amended as necessary or in

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order to test various hypotheses (See Figure S4). For example, after the first dose of 1,2,4-TCB

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was depleted on Day 126, Bottle 5 and 10 were re-amended with “CB mixture”, while the other

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replicates in these sets were re-amended with only 1,2,4-TCB on Day 166, to limit accumulation

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of DCB and MCB daughter products. To clearly identify the dechlorination products of 1,2,4-

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TCB, bottles 7 (in situ conditions) and 9 (donor-amended) were purged with CO2/N2 on Day 370

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to remove accumulating daughter products prior to re-amending with 1,2,4-TCB. Bottle 9 was

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also re-amended with donor. To confirm dechlorination of 1,2-DCB and 1,3-DCB, samples from

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Bottles 5 and 10 were used to inoculate (using a 10% transfer) two bottles containing sterile,

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defined iron sulfide-reduced mineral medium.51 These bottles (5-T1 and 10-T1) received mixture

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of 1,2- and 1,3-DCB and then each compound separately in subsequent transfers (10-T2a and 10-

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T2b; see Figure S4). Methanol and Ethanol (ME) were amended to these transfer cultures at 5

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times electron equivalents required for complete dechlorination. A third 10% transfer culture

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from a 1,3-DCB-amended microcosm (Bottle 10-T3) was amended with MCB dissolved in a

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methanol/ethanol mixture. To assess 1,4-DCB dechlorination, Bottle 11 was purged to remove

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all remaining VOCs prior to re-amending with 1,4-DCB dissolved in a methanol/ethanol mixture.

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A 10% transfer culture from Bottle 11 (Bottle 11-T1) was amended with 1,4-DCB in acetone to

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try to reduce methane production (Figure S4).

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A set of microcosms (the fourth treatment) was designed to investigate anaerobic benzene

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biodegradation, and these microcosms received repeated doses of benzene at concentrations of 9

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µM (Bottles 13-15) and 100 µM (Bottle 16). Sulfate was naturally present in the groundwater

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and was replenished on Day 148 at a concentration of 300 µM. To enrich benzene-degrading

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microbes, a 10% culture from Bottle 16 (Bottle 16-T1) was inoculated into mineral medium, and

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was amended with 80 µM benzene, which was gradually increased to 330 µM per feeding.

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For comparative purposes, rates of dechlorination or degradation were simply calculated ஼೅భ ି஼೅మ

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assuming zero-order kinetics as V =

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CT1 and CT2 are the concentrations at times T1 and T2.

்మ ି்భ

, where v is the dechlorination rate (µM/day) and

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Analytical Methods. 1,2,4-TCB was analyzed using an Agilent 5890 GC-FID equipped with

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an Agilent J&W DB-WAX column (30 m × 0.53 mm). DCBs, MCB, benzene and methane were

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quantified by an Agilent 7890 GC-FID equipped with an Agilent J&W GS-Q column (30 m ×

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0.53 mm). Anions, including Cl-, NO3-, NO2-, SO42-, were analyzed by a Dionex DX-100 series

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ion chromatography equipped with an IonPac AS18 anion exchange column and an AG18 guard

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column. TOC measurements were carried out using a TOC-VCPN (SHIMADZU, Japan) analyzer

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combined with an OCT1 8-port sampler using the non-purgeable organic carbon (NPOC)

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method.52 Supernatants from all parent bottles were sampled to measure total organic carbon

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(TOC) concentration after ~580 days’ incubation to attempt to close the mass balance.

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Additional method details are provided in the SI.

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DNA Extraction. Samples (slurry) from three arbitrary bottles on Day 0 and all the treatment

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bottles on Day 372 or 379 were collected to extract DNA using the PowerSoil DNA isolation kit

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(MoBio Laboratories, Solana Beach, CA) according to the manufacturer’s protocol except that

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DNA was eluted in 50 µL sterile UltraPure distilled water (Invirogen, Carlsbad, CA). A

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NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE) was used to

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assess DNA concentrations and quality.

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Quantitative PCR (qPCR) Analyses. Dehalococcoides (Dhc), Dehalobacter (Dhb),

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Deltaproteobacterium ORM2 and total Bacteria (TotBac) 16S rRNA genes were enumerated

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using qPCR with published primers as listed in Table S3. Serial dilutions of plasmid preparations

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containing corresponding 16S rRNA gene fragments were used to generate standard curves.

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Each qPCR reaction was performed in duplicate. All manipulations were conducted in a PCR

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cabinet (ESCO Technologies, Hatboro, PA). Analyses were conducted with a Bio-Rad CF96

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Touch real-time modular thermal cycler platform and CFX Manager software

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additional details of qPCR reactions).

53

(see SI for

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3. Results

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3.1 Field Site Investigation

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The general hydrogeology at the site to 22m below ground surface consisted of 3 layers: a 2-4

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m thick unconfined aquifer composed of silty clay, situated above a 3-5m thick aquitard

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composed of clay and a deeper confined aquifer composed of silty clay/clayey silt (Figure S3).

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The vadose zone and the unconfined aquifer consisted of continuous clay with extremely low

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hydraulic conductivities of 0.002-0.07 m/day determined by slug tests, and the well screens were

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set at least one meter below the water table at 3 meter intervals. The unconfined groundwater

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velocities were from 10-5 to 2.5×10-4 m/day, making penetration of oxygen virtually impossible.

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Groundwater concentrations measured during the preliminary site investigation in June 2012

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are shown in Figure 1. The primary contaminant identified was 1,2-4-TCB and concentrations in

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samples from four locations (W2, S1, S3, S4) exceeded the Maximum Contaminant Levels

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(MCL) of 70 µg/L.54 Specifically, 1,2,4-TCB at boreholes S1 and S4 were as high as 4800 µg/L

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and 7300 µg/L (>1% of 1,2,4-TCB solubility; possibly indicating DNAPL), and the

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concentrations of benzene, MCB, 1,4-DCB in the two sampling wells were all higher than

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respective MCLs (5, 100, 75 µg/L).54 The most contaminated wells were below the former waste

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water basin (S1) and chemical production facilities (W2, S3, S4) in the northeastern portion of

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the site. The samples from wells located below the former office area and slag pool showed very

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low 1,2,4-TCB concentrations (W3, S2) or none detected (W1).

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During the site investigation of May 2013, a different set of wells were sampled. Certain

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additional groundwater parameters were measured, and these are shown in Table S4. Most

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groundwater samples were anoxic, except from wells W4 and W8. Methane was detected at

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depths of 2-4 m below ground surface during the MIP tests, indicating methanogenic conditions.

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In May 2013, only 1,2,4-TCB concentrations in two samples (W2, DW5) were above the MCL

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(Figure 1); these wells were located in the center of the former production facility and waste

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water basin, consistent with the results obtained in 2012. However, the concentration of 1,2,4-

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TCB in shallow well SW5 was much lower than in deep well DW5, possibly because the 1,2,4-

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TCB had migrated down as a DNAPL over time. Concentrations in the four samples taken in

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2013 from locations outside of the former production facility and waste water basin (W4, W6-8)

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were low or below detection.

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The last two sampling campaigns conducted in September 2014 and October 2016 were

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limited to 2 locations (S1 and W4). Results in 2014 revealed that the only contaminants close to

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or above MCLs were dechlorination daughter products 1,4-DCB and MCB (Figure 1).

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Concentrations of other compounds, including 1,2,4-TCB, 1,2-DCB, 1,3-DCB and benzene had

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substantially decreased (Figure 1). By 2016, all contaminants analyzed were below respective

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MCLs (Figure 1).

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Three locations were sampled over multiple years to allow direct comparisons (Figure 2). At

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borehole S1, concentrations of all compounds measured decreased to lower than respective

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MCLs from 2012 to 2016 (Figure 2). Considering the low permeability of the clay at the site and

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the extremely low groundwater velocities, dilution from precipitation and groundwater flow only

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played a small role in the observed decreases. Penetration of oxygen is also unlikely excluding

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aerobic biodegradation. Rather, anaerobic transformation reactions are the most likely

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explanation. A closer look at wells W2 and W4 sampled at one-year intervals reveals a pattern

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corresponding to reductive dechlorination, where parent compound 1,2,4-TCB concentrations

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decreased, while concentrations of daughter products (DCBs, MCB and benzene) increased

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(Figure 2). Concentrations of dechlorination intermediates 1,4-DCB and MCB increased the

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most, suggesting that these compounds were relatively more slowly degraded.

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In summary, the multi-year site investigation results indicated that the contamination was

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primarily in the vicinity of the former production facilities and waste water basin in the

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northeastern portion of the site. The main VOC released into the groundwater was 1,2,4 TCB

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used as a solvent in production, and it likely existed as a DNAPL (See supplemental method

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244

details). Benzene and lesser chlorinated benzenes were not used or manufactured at the site. TCB

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was evidently dechlorinated by indigenous microbes to the suite of chlorobenzenes and benzene

246

detected at the site. The site data also suggest that the products 1,4-DCB and MCB were most

247

slowly dechlorinated.

248 249

3.2 Laboratory Microcosms

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Overview of Microcosm Results. No significant change in chlorobenzene concentration

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occurred over more than one year in poisoned control bottles spiked with mercuric chloride and

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sodium azide (Figure S5), while substantial transformation was observed in all active bottles.

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The addition of electron donor did not change dechlorination patterns. Donor-amended bottles

254

differed only by higher methane production. Although overall trends were similar among the

255

replicates in each set, each bottle displayed slightly different transformation rates. Therefore,

256

degradation profiles are presented for one representative bottle of a series in the main text (e.g.,

257

Figure 3), while the data for all remaining bottles are provided in SI (Figure S6-S9).

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Corresponding TOC and anions measurements for all treatment bottles are provide in Table S5-

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S6. Some sorption of CBs onto the sediments in microcosms was observed, ranging from 12 to

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28% of added substrate for MCB and DCB isomers, up to ~55% for 1,2,4-TCB (Table S2D).

261

Dechlorination of 1,2,4-TCB. In chlorobenzene treatments (Bottles 5-12), dechlorination of

262

1,2,4-TCB (~16 µM) began after a lag of approximately 60 days, and 1,2,4-TCB was depleted by

263

Day 150 (Figure 3A and Figure S6-S9). Following subsequent re-spikes, the dechlorination rate

264

of 1,2,4-TCB increased from ~0.1 to 0.5-1.0 µM/day over a period of 355 days. A maximum

265

dechlorination rate of 2.5 µM/day was observed in bottle 9 after ~600 days’ incubation. To

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clearly quantify dechlorination products, bottles were purged and re-amended with 1,2,4-TCB

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after one year of incubation. The products 1,2-DCB, 1,3-DCB and 1,4-DCB were produced at a

268

molar ratio of approximately 2%/10%/88% (Figure 3A and Figure S8).

269

Dechlorination of DCB Isomers. 1,2-DCB and 1,3-DCB initially present in the microcosms

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(Bottles 5-12) were dechlorinated to MCB after a lag of approximately 200 days (Figure 3A and

271

Figure S6-S9) at rates 0.03-0.1 µM/day, while 1,4-DCB seemed to simply accumulate from TCB

272

dechlorination. Transfer cultures were set up to further investigate dechlorination of DCB

273

isomers. In Bottles 5-T1 and 10-T1, fed a mix of 1,2- and 1,3-DCBs, MCB was produced after a

274

lag of 40 to 120 days (Figure S10). Dechlorination rates of 1,2-DCB and 1,3-DCB increased

275

substantially from 0.6 µM/day to over 20 µM/day in transfers (Figure S11). Dechlorination of

276

recalcitrant 1,4-DCB was finally observed on day 706 in Bottle 11 after supplementation with

277

mineral medium (Figure S9), suggesting that 1,4-DCB dechlorination was nutrient- or vitamin-

278

limited. Subsequent dechlorination of 1,4-DCB was also observed in transfer Bottle 11-T1

279

(Figure 3B).

280

Dechlorination of MCB to Benzene. Hints of benzene production were detected in Bottle 10-

281

T2b amended with 1,3-DCB only, where clear dechlorination of MCB to benzene was first

282

observed (Figure S11B). In transfer culture Bottle 10-T3, with MCB alone, dechlorination of

283

MCB to benzene was also observed (Figure 3C), indicating that MCB dechlorination was not co-

284

metabolically dependent on 1,3-DCB dechlorination.

285

Anaerobic Degradation of Benzene. Benzene was observed to degrade after an ~150 day lag

286

in most of the original chlorobenzene- and benzene-amended microcosms, regardless of electron

287

donor amendment (Figure 3A and Figure S7-S9). Only in the two Bottles 5 and 10 was benzene

288

persistent; these two bottles were the only two re-amended with a mixture of 1,2,4-TCB and

289

dichlorobenzenes, rather than just 1,2,4-TCB (Figure S6), suggesting benzene inhibition from the

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dichlorobenzenes. We also established microcosms from the start (Bottles 13-16, Figure S12)

291

with only benzene to eliminate possible competition from other donors. In these microcosms,

292

benzene degradation began after a much shorter lag of ~ 45 days, with corresponding depletion

293

of sulfate. After transfer of Bottle 16 into mineral medium (Bottle 16-T1) and progressive

294

increase in benzene concentration on successive feedings, a remarkably rapid benzene-

295

degrading, sulfate-reducing enrichment culture was obtained (Figure 3D) with a degradation rate

296

of 19 µM/day more than 100 times faster than the initial rate of 0.14 µM/day.

297

Microbial analysis by qPCR. Copies of 16S rRNA genes for Dhc, Dhb, ORM2-like

298

Deltaproteobacteria, and total bacteria were analyzed in samples from Day 0 and from samples

299

taken more than 370 days later (Table S7, Figure 4). For both Dhc and Dhb, these increases are

300

not substantial (Figure 4A), nor do they correlate well with extent of dechlorination (Table S7B).

301

The corresponding Dhc and Dhb yields were in general quite low (Table S8) compared to

302

reported growth yields of Dhc (4.1×107 cells per µmol Cl- released)31, Dhb (1.62×106 cells per

303

µmol Cl- released)23, and these same populations never represented more than 0.2% (Dhc),

304

0.02% (Dhb) of the total bacteria in these bottles (Table S7B), suggesting that Dhc and Dhb were

305

not responsible for the observed dechlorination.

306

In these same CB-amended bottles, copies of putative benzene-degrading ORM2-like 16S

307

rRNA sequences were sustained in the benzene-degrading chlorobenzenes-amended bottles, but

308

decreased in Bottles 5 and 10 where benzene was not observed to degrade (Figure 4B). In the

309

microcosms and transfers where only benzene was amended (Bottles 13-16, 16-T1), ORM2-like

310

gene copies increased by more than 2-3 orders of magnitude (Figure 4B). The ability to measure

311

cell yields improved with increasing mass of benzene degraded, and yields eventually exceeded

312

published yields (2.41×107 cells per µmol benzene consumed)40, and the ORM2-like population

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313

grew to represent up to 60% of the total bacterial population (Figure 4B), indicating that the

314

observed benzene transformation was mediated by ORM2-like microbes.

315 316

4. Data Analysis and Discussion

317

4.1 Mole and electron balances

318

A mole balance comparing initial to final chlorinated benzene parent and daughter products

319

was calculated for each microcosm (Table S9) to verify analytical procedures and transformation

320

products. 1,2,4-TCB was primarily transformed to 1,4-DCB (~80%) with the remainder going to

321

1,2- and 1,3-DCBs. This is further confirmed in Bottle 7 and 9 when 1,2,4-TCB was re-amended

322

after all VOCs were purged (Figure 3A and S7). Moreover, the subsequent increase in MCB (1.2

323

±0.01µmol) approximately equals to the decrease of 1,2-DCB and 1,3-DCB (0.96±0.04µmol).

324

The two recalcitrant compounds, 1,4-DCB and MCB, could be further transformed (Figures

325

3B/3C, S9, S11B) in re-amended microcosms and transfers. The overall chlorobenzene

326

recoveries were 87%-105% (Table S9) suggesting that little conversion of MCB to benzene

327

occurred in the original microcosms, but only later in targeted microcosms that were re-amended

328

with DCBs or MCB.

329

An overall electron balance was performed to understand the relationships between electron

330

donors (MEL, benzene, TOC) and acceptors (SO42-, CO2, CBs,) in all microcosms (Table 1). The

331

high concentration of TOC (Table S5) and sulfate (Table S6) in the site groundwater (from

332

sulfuric acid) are the result of leaks from the chemical production processes at the site (Figure

333

S1). The compounds responsible for the TOC serving as electron donors are unknown but may

334

be related to the production process reactions described in Figure S1. In the electron balance, we

335

ignored electrons going to biomass synthesis which is expected to be < 8% of total donor

15 / 30



336

consumed in these anaerobic systems55. Given that exogenous electron donor addition did not

337

stimulate dechlorination, we wanted to better understand the contribution of the TOC in site

338

groundwater to transformation processes. Reasonable electron balances (Acceptor/Donor) of 46-

339

84% in the original microcosms (Bottles 5-16) and 110% in Bottle 16-T1 were obtained (Table

340

1). The consumption of TOC found in the groundwater (Table S5) accounted for a large fraction

341

of electron donor consumption, driving sulfate reduction primarily, as well as dechlorination.

342

Dechlorination accounted for ~1% of the electrons consumed. The exogenous donor added

343

(MEL) did not affect dechlorination, but appears to have led simply to methanogenesis (Table 1).

344

The transformation of chlorobenzenes and benzene was accompanied by a decrease in TOC and

345

sulfate. Of the possible electron acceptors for benzene degradation, only sulfate depletion could

346

account for the benzene consumed (Table 1). Sulfate concentrations in microcosms was initially

347

4 mM, and decreased substantially during benzene degradation; nitrate concentrations were

348

below 0.2 mM in all microcosms (Table S6), and methane concentrations did not increase

349

sufficiently (Table 1). In all cases except for the transfer Bottle 16-T1, sulfate consumption

350

exceeded measured donor consumption, indicating that additional sources of donor beyond

351

measured TOC were present but not accounted for, perhaps present sorbed to the soil in the

352

microcosms. In the benzene-amended 10% transfer bottle with little soil and defined mineral

353

medium, the electron balance was essentially closed. Interestingly, in microcosm bottle 16, that

354

was amended with a higher concentration of benzene compared to the other bottles in this series

355

(bottles 13-15), the TOC was not consumed; perhaps the presence of 100 µM benzene inhibited

356

the degradation of the compounds measured as TOC.

357 358

4.2 Microbial Characterization

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359

Quantitative PCR is well-accepted for tracking microbial phylotype abundances in

360

groundwater and microcosm samples. Dehalococcoides strain CBDB129 and strain DCMB531

361

have been reported to dechlorinate 1,2,4-TCB to 1,3- and 1,4-DCB or 1,4-DCB only, with no

362

further dechlorination of DCB isomers were reported. Certain strains of Dehalobacter can

363

dechlorinate TCB and all DCB isomers, but production of 1,2-DCB from 1,2,4-TCB has not

364

been seen.23 The primers (Table S3) used in this study specifically bind the known

365

chlorobenzenes dechlorinating Dhb and Dhc but we did not detect their growth, suggesting that

366

other dehalogenating microbes were present, particularly since all three DCB isomers were

367

produced from 1,2,4-TCB – a novel dechlorinating pattern (Figure 5). A complete 16S rRNA

368

amplicon survey is in progress and will be reported in a subsequent manuscript.

369

The qPCR data clearly establish a major role for ORM2-like Deltaproteobacteria in benzene

370

degradation under sulfate-reducing conditions at this site. Compared to other benzene-degrading

371

enrichment cultures in our own and other’s laboratories, the rates achieved in the transfers are 3-

372

4 times greater, and the abundance of a similar and consistent phylotype is exciting as it may be

373

a valuable biomarker for field investigations. It is particular important for remediation to be able

374

to observe benzene degradation in the presence of other compounds, like MCB, and in the

375

presence of other electron donors, in order to detoxify the parent compounds. In this study,

376

benzene degradation remarkably continued in the presence of high concentrations of unknown

377

donors in TOC (Bottles 6-8) and even in the MEL donor-amended bottles (Bottles 9, 11,12).

378

Benzene degradation was however inhibited by the addition of DCB isomers (but not TCB) as

379

seen in Bottles 5 and 10. In these two bottles, benzene was not degraded, and the corresponding

380

copies of ORM2-like sequences did not increase (Figure 4B).

381

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382

Field Implications. The characterization of the site, though far from comprehensive, strongly

383

suggested that natural attenuation of chlorobenzenes and benzene was occurring in situ. The

384

thorough laboratory microcosm study results confirmed that the indigenous microbes were

385

capable of completely transforming chlorobenzenes to non-toxic CO2 via benzene, and provided

386

needed assurance for affirming natural attenuation processes in situ. 1,4-DCB and MCB were

387

more slowly transformed both at the site and in laboratory microcosm. However, the latest two

388

samples (S1, W4) analyzed in October 2016 revealed that benzene was completely degraded, and

389

1,4-DCB and MCB concentrations had decreased to below MCLs. The degradative activities

390

recovered from both field and lab investigations are illustrated in Figure 5. The microcosm

391

activity data was essential to confirming assumptions based on field data, provided information

392

on relative rates, provided a means to assess potential molecular monitoring tools and brought to

393

light the presence and important unexpected beneficial role of unknown electron donors and

394

sulfate in the groundwater at the site. The surprisingly high anaerobic benzene degrading

395

activity, perhaps enriched because of the aromatic nature of the compounds manufactured at the

396

site, promoted the full detoxification of 1,2,4-TCB, DCB isomers and MCB at this site.

397

Given the site hydrogeological conditions comprising silty clay and clayey silt with extremely

398

low permeability, commonly used remediation technologies like pump-and-treat, soil vapor

399

extraction and many others requiring injection are simply impossible to implement. Rather,

400

future work should continue to monitor and quantify the rate of natural attenuation at the site,

401

monitoring key rate-limiting electron donors. Further research is on-going to determine if the

402

indigenous microbes at the Nanjing site resemble those previously described to degrade DCB,

403

MCB and benzene and to identify the unknown electron donors in the site groundwater that led

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404

to a microbial community capable of complete dechlorination and mineralization of site

405

contaminants.

406 407

ASSOCIATED CONTENT

408

Supporting Information. The Supplemental files contain Figures S1-S12, Tables S1-S9, and

409

method details. Included is information on site layout and wells, products manufactured at the

410

site, site hydrogeology, groundwater analyses and field data measurements. Additional

411

information on the microcosm study includes, experimental design, additional degradation

412

profiles of chlorobenzenes and benzene not shown in main text, donor and acceptor

413

concentration data, TOC, IC data, mole balances and qPCR raw and processed data.

414

Notes: The authors declare no competing financial interest.

415

Acknowledgement: The authors thank Jian Guo and Dong Wang for the field investigations and

416

gratefully acknowledge funding from E.I. du Pont de Nemours and Company, the National

417

Natural Science Foundation of China (NSFC) grant No. 41472212, the Ontario China Research

418

Innovation Fund No. 2016YFE0101900, and the Government of Canada through Genome

419

Canada and the Ontario Genomics Institute (OGI-102). WQ was supported by the China

420

Scholarship Council.

421

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422

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423 424 425 426 427 428 429 430 431 432 433 434 435 436 437 438 439 440 441 442 443 444 445 446 447 448 449 450 451 452 453 454 455 456 457 458 459 460 461 462 463 464 465 466

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Biodegradation 2008, 19, (4), 463-480. 18. Boopathy, R., Factors limiting bioremediation technologies. Bioresource Technol. 2000, 74, (1), 63-67. 19. Kaschl, A.; Vogt, C.; Uhlig, S.; Nijenhuis, I.; Weiss, H.; Kastner, M.; Richnow, H. H., Isotopic fractionation indicates anaerobic monochlorobenzene biodegradation. Environ. Toxicol. Chem. 2005, 24, (6), 1315-1324. 20. Farhadian, M.; Vachelard, C.; Duchez, D.; Larroche, C., In situ bioremediation of monoaromatic pollutants in groundwater: a review. Bioresource Technol. 2008, 99, (13), 52965308. 21. Fung, J. M.; Weisenstein, B. P.; Mack, E. E.; Vidumsky, J. E.; Ei, T. A.; Zinder, S. H., Reductive Dehalogenation of Dichlorobenzenes and Monochlorobenzene to Benzene in Microcosms. Environ. Sci. Technol. 2009, 43, (7), 2302-2307. 22. Nelson, J. L.; Fung, J. M.; Cadillo-Quiroz, H.; Cheng, X.; Zinder, S. H., A role for Dehalobacter spp. in the reductive dehalogenation of dichlorobenzenes and monochlorobenzene. Environ. Sci. Technol. 2011, 45, (16), 6806-6813. 23. Nelson, J. L.; Jiang, J.; Zinder, S. H., Dehalogenation of chlorobenzenes, dichlorotoluenes, and tetrachloroethene by three Dehalobacter spp. Environ. Sci. Technol. 2014, 48, (7), 3776-3782. 24. Zhou, X.; Zhang, C. F.; Zhang, D. D.; Awata, T.; Xiao, Z. X.; Yang, Q.; Katayama, A., Polyphasic characterization of an anaerobic hexachlorobenzene-dechlorinating microbial consortium with a wide dechlorination spectrum for chlorobenzenes. J. Biosci. Bioeng. 2015, 120, (1), 62-68. 25. Fennell, D. E.; Nijenhuis, I.; Wilson, S. F.; Zinder, S. H.; Haggblom, M. M., Dehalococcoides ethenogenes strain 195 reductively dechlorinates diverse chlorinated aromatic pollutants. Environ. Sci. Technol. 2004, 38, (7), 2075-2081. 26. Löffler, F. E.; Yan, J.; Ritalahti, K. M.; Adrian, L.; Edwards, E. A.; Konstantinidis, K. T.; Müller, J. A.; Fullerton, H.; Zinder, S. H.; Spormann, A. M., Dehalococcoides mccartyi gen. nov., sp. nov., obligately organohalide-respiring anaerobic bacteria relevant to halogen cycling and bioremediation, belong to a novel bacterial class, Dehalococcoidia classis nov., order Dehalococcoidales ord. nov. and family Dehalococcoidaceae fam. nov., within the phylum Chloroflexi. Int. J. Syst. Evol. Micr. 2013, 63, (2), 625-635. 27. Wu, Q. Z.; Milliken, C. E.; Meier, G. P.; Watts, J. E. M.; Sowers, K. R.; May, H. D., Dechlorination of chlorobenzenes by a culture containing bacterium DF-1, a PCB dechlorinating microorganism. Environ. Sci. Technol. 2002, 36, (15), 3290-3294. 28. Adrian, L.; Szewzyk, U.; Wecke, J.; Gorisch, H., Bacterial dehalorespiration with chlorinated benzenes. Nature 2000, 408, (6812), 580-583. 29. Jayachandran, G.; Gorisch, H.; Adrian, L., Dehalorespiration with hexachlorobenzene and pentachlorobenzene by Dehalococcoides sp. strain CBDB1. Arch. Microbiol. 2003, 180, (6), 411-416. 30. Holscher, T.; Gorisch, H.; Adrian, L., Reductive dehalogenation of chlorobenzene congeners in cell extracts of Dehalococcoides sp strain CBDB1. Appl. Environ. Microbiol. 2003, 69, (5), 2999-3001. 31. Poritz, M.; Schiffmann, C. L.; Hause, G.; Heinemann, U.; Seifert, J.; Jehmlich, N.; von Bergen, M.; Nijenhuis, I.; Lechner, U., Dehalococcoides mccartyi Strain DCMB5 Respires a Broad Spectrum of Chlorinated Aromatic Compounds. Appl. Environ. Microbiol. 2015, 81, (2), 587-596.

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32. Dean, B. J., Recent findings on the genetic toxicology of benzene, toluene, xylenes and phenols. Mutat. Res. 1985, 154, (3), 153-181. 33. Abu Laban, N.; Selesi, D.; Rattei, T.; Tischler, P.; Meckenstock, R. U., Identification of enzymes involved in anaerobic benzene degradation by a strictly anaerobic iron ‐reducing enrichment culture. Environ. Microbiol. 2010, 12, (10), 2783-2796. 34. Zhang, T.; Bain, T. S.; Nevin, K. P.; Barlett, M. A.; Lovley, D. R., Anaerobic benzene oxidation by Geobacter species. Appl. Environ. Microb. 2012, 78, (23), 8304-8310. 35. Ulrich, A. C.; Edwards, E. A., Physiological and molecular characterization of anaerobic benzene-degrading mixed cultures. Environ. Microbiol. 2003, 5, (2), 92-102. 36. Luo, F.; Gitiafroz, R.; Devine, C. E.; Gong, Y.; Hug, L. A.; Raskin, L.; Edwards, E. A., Metatranscriptome of an anaerobic benzene-degrading, nitrate-reducing enrichment culture reveals involvement of carboxylation in benzene ring activation. Appl. Environ. Microbiol. 2014, 80, (14), 4095-4107. 37. Kleinsteuber, S.; Schleinitz, K. M.; Breitfeld, J.; Harms, H.; Richnow, H. H.; Vogt, C., Molecular characterization of bacterial communities mineralizing benzene under sulfatereducing conditions. FEMS Microbiol. Ecol. 2008, 66, (1), 143-157. 38. Abu Laban, N.; Selesi, D. e.; Jobelius, C.; Meckenstock, R. U., Anaerobic benzene degradation by Gram-positive sulfate-reducing bacteria. FEMS Microbiol. Ecol. 2009, 68, (3), 300-311. 39. Noguchi, M.; Kurisu, F.; Kasuga, I.; Furumai, H., Time-Resolved DNA stable isotope probing links Desulfobacterales-and Coriobacteriaceae-related bacteria to anaerobic degradation of benzene under methanogenic conditions. Microbes Environ. 2014, 29, (2), 191-199. 40. Luo, F.; Devine, C. E.; Edwards, E. A., Cultivating microbial dark matter in benzene‐ degrading methanogenic consortia. Environ. Microbiol. 2016, 18, (9), 2923-2936. 41. Liang, X. M.; Devine, C. E.; Nelson, J.; Lollar, B. S.; Zinder, S.; Edwards, E. A., Anaerobic Conversion of Chlorobenzene and Benzene to CH4 and CO2 in Bioaugmented Microcosms. Environ. Sci. Technol. 2013, 47, (5), 2378-2385. 42. Sturchio, N. C.; Clausen, J. L.; Heraty, L. J.; Huang, L.; Holt, B. D.; Abrajano, T. A., Chlorine isotope investigation of natural attenuation of trichloroethene in an aerobic aquifer. Environ. Sci. Technol. 1998, 32, (20), 3037-3042. 43. Lollar, B. S.; Slater, G. F.; Sleep, B.; Witt, M.; Klecka, G. M.; Harkness, M.; Spivack, J., Stable carbon isotope evidence for intrinsic bioremediation of tetrachloroethene and trichloroethene at area 6, Dover Air Force Base. Environ. Sci. Technol. 2001, 35, (2), 261-269. 44. Davis, J. W.; Odom, J. M.; DeWeerd, K. A.; Stahl, D. A.; Fishbain, S. S.; West, R. J.; Klecka, G. M.; DeCarolis, J. G., Natural attenuation of chlorinated solvents at Area 6, Dover Air Force Base: characterization of microbial community structure. J. Contam. Hydrol. 2002, 57, (1), 41-59. 45. Clement, T. P.; Truex, M. J.; Lee, P., A case study for demonstrating the application of U.S. EPA's monitored natural attenuation screening protocol at a hazardous waste site. J. Contam. Hydrol. 2002, 59, (1), 133-162. 46. Imfeld, G.; Nijenhuis, I.; Nikolausz, M.; Zeiger, S.; Paschke, H.; Drangmeister, J.; Grossmann, J.; Richnow, H. H.; Weber, S., Assessment of in situ degradation of chlorinated ethenes and bacterial community structure in a complex contaminated groundwater system. Water Res. 2008, 42, (4-5), 871-882. 47. Thullner, M.; Schäfer, W., Modeling of a Field Experiment on Bioremediation of Chlorobenzenes in Groundwater. Bioremediation Journal 1999, 3, (3), 247-267.

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48. Vogt, C.; Alfreider, A.; Lorbeer, H.; Hoffmann, D.; Wuensche, L.; Babel, W., Bioremediation of chlorobenzene-contaminated ground water in an in situ reactor mediated by hydrogen peroxide. J. Contam. Hydrol. 2004, 68, (1-2), 121-141. 49. Dominguez, R. F.; da Silva, M. L. B.; McGuire, T. M.; Adamson, D.; Newell, C. J.; Alvarez, P. J. J., Aerobic bioremediation of chlorobenzene source-zone soil in flow-through columns: performance assessment using quantitative PCR. Biodegradation 2008, 19, (4), 545553. 50. Stelzer, N.; Imfeld, G.; Thullner, M.; Lehmann, J.; Poser, A.; Richnow, H.-H.; Nijenhuis, I., Integrative approach to delineate natural attenuation of chlorinated benzenes in anoxic aquifers. Environ. Pollut. 2009, 157, (6), 1800-1806. 51. Edwards, E. A.; Grbić-Galić, D., Anaerobic degradation of toluene and o-xylene by a methanogenic consortium. Appl. Environ. Microbiol. 1994, 60, (1), 313-322. 52. Rice, E. W.; Baird, R. B.; Eaton, A. D.; Clesceri, L. S., Standard methods for the examination of water and wastewater (22th ed). American Public Health Association.: Washington, DC, 2012; Vol. 2. 53. Molenda, O.; Quaile, A. T.; Edwards, E. A., Dehalogenimonas sp. Strain WBC-2 Genome and Identification of Its trans-Dichloroethene Reductive Dehalogenase, TdrA. Appl. Environ. Microbiol. 2016, 82, (1), 40-50. 54. https://www.epa.gov/your-drinking-water/table-regulated-drinking-water-contaminants organic . (Accessed October 31, 2016). 55. Rittmann, B. E.; McCarty, P. L., Environmental biotechnology: principles and applications. Tata McGraw-Hill Education: 2012.

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582 583

Figure 1. Contaminant concentrations measured in site samples by year in wells (W) or soil

584

boreholes (S): 2012 (Top Left); 2013 (Bottom Left); 2014 (Top Right); 2016 (Bottom Right).

585

N.D.: not detected.

586

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587 588

Figure 2. Comparison of site contaminant concentrations by location over time from samples

589

from within the former production facility area. Soil borehole S1 (Left panel); well W2 (Middle

590

panel); well W4 (Right panel). N.D.: Not detected.

591 592 593 594 595 596

25 / 30



597 598 599

Figure 3. Concentration profiles observed in microcosm and transfer bottles. (A) Representative

600

bottle 7 containing sediment and groundwater under in situ conditions. These microcosms were

601

initially amended with a chlorobenzene (CB) mixture containing 1,2,4-TCB, DCBs and MCB, at

602

concentrations of 12 µM (1,2,4-TCB/1,4-DCB), 2.8 µM (1,2-/1,3-DCB) and 17 µM (MCB).

603

Dechlorination profiles were similar in all 8 replicate microcosms (regardless of added electron

604

donor) and graphs and data for other bottles are provided in the SI (Figures S6-S8 and Table S2).

605

Concentrations measured before Day 370 are shown on the left axis while concentrations

606

measured after purging on Day 370 are shown on the right axis. The arrows represent different

607

feeding events and with substrates indicated. (B) Dechlorination of 1,4-DCB (initial

608

concentration 280 µM) in Bottle 11-T1 amended with acetone at 5X the electron equivalents

609

(eeq) required for dechlorination. (C) Dechlorination of MCB (initial concentration 115 µM) in

610

Bottle 10-T3 amended with methanol/ethanol at 5X eeq. (D) Benzene and sulfate degradation

611

profiles Bottle 16-T1 amended with 80 µM benzene initially that was gradually increased to 330

612

µM. Degradation profiles for all original benzene only-amended microcosms are provided in

613

Figure S12. See Figure S4 for a summary of microcosm transfer history.

26 / 30


Page 26 of 30

Page 27 of 30


614 615

Figure 4. Summary of 16S rRNA gene abundance (in copies per ml) of Dehalococcoides (Dhc),

616

Dehalobacter (Dhb), ORM2-like Deltaproteobacteria and Total Bacteria (TotBac) as determined

617

by qPCR in all original microcosms and in transfer cultures. “N.M.”: not measured. Error bars

618

represent positive standard deviation of the mean in triplicate or quadruplicate microcosms,

619

except for measurements of total bacteria in CB treatments where data are average and range of

620

duplicates. Panel A presents data for Dhc, Dhb and total bacteria in poisoned control (Bottles 1-

621

4) and CB-amended treatments (Bottles 5-8, and Bottles 9-12 with electron donor added). The

622

sampling time is indicated under each treatment. The initial samples on Day 0 were collected

623

from three random microcosms as they were all set up identically prior to amendments. Panel B

624

provides qPCR data for ORM2-like organisms and total bacterial copies in all CB-amended

625

bottles as well as benzene-only bottles. The latter include bottles amended with 9 µM of

626

benzene (Low, Bottles 13-15), 100 µM of benzene only added treatment (High, Bottle 16) and

627

80-330 µM of benzene only amended in transferred culture (Transfer, Bottle 16-T1). Copies of

628

ORM2-like sequences in Bottle 5 and 10 are not combined with other bottles in these series and

629

displayed individually because benzene was recalcitrant in these two microcosms.

27 / 30



630 631

Figure 5. Dechlorination pathways for chlorobenzene dechlorination mediated by Dehalobium

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DF-127 (red arrows), Dehalococcoides mccartyi strains 19525, CBDB129, and DCMB531 (blue

633

arrows), Dehalobacter23 (green arrows), and in this study (pink arrows).

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28 / 30


Page 28 of 30

Page 29 of 30

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TOC/ Abstract Art

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Page 30 of 30

Table 1. Electron balances in active microcosm bottles and transfer Bottle 16-T1

Microcosm Description

Under in situ conditions With electron donor added Amended with Benzene only High Benzene 10% Transfer

Electron equivalents of Acceptors Consumed (µeeq/Bottle)

Electron Equivalents of Electron Donors Consumed (µeeq/Bottle)

Sulfate Reduction

Methane Production

Dechlori nation

Sum

MEL3

Benzene

TOC4

Sum

Donor/Ac ceptor Recovery (%)

5

2610

203

13

2820

0

5

1310

1310

46%

6

2170

45

14

2230

0

101

1290

1390

62%

7 8 9

2250 2170 2670

55 34 754

13 19 13

2320 2220 3430

0 0 988

113 92 32

1260 1250 1280

1380 1340 2300

59% 60% 67%

10

2660

1450

17

4130

988

6

1340

2330

56%

11

2540

775

17

3340

988

99

1250

2340

70%

12

2400

689

17

3110

988

116

1340

2440

78%

13

2500

51

2550

0

824

1320

2150

84%

14

2800

60

2860

0

822

1310

2130

74%

15

3150

197

3350

0

820

1350

2170

65%

161

5160

84

5240

0

2500

35

2530

48%

16-T12

2590

46

2640

0

2930

n.m.

2930

111%

Bottle no.

N/A

638

Note:

639

1. Bottle 16 was amended with ~10 times more benzene initially (100µM vs 9µM); note little TOC consumed in this bottle

640

2. 10% inoculated from Bottle 16 into anaerobic medium; n.m.: TOC not measured.

641

3. MEL is an abbreviation for electron donor mix: Methanol, Ethanol and Lactate.

642

4. Assuming 5 eeq/mole carbon.


30

Natural attenuation and anaerobic benzene detoxification processes

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