Natural Attenuation in Streambed Sediment Receiving Chlorinated

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Natural Attenuation in Streambed Sediment Receiving Chlorinated Solvents from Underlying Fracture Networks Burcu Simsir, Jun Yan, Jeongdae Im, Duane Graves, and Frank E Löffler Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.6b05554 • Publication Date (Web): 22 Mar 2017 Downloaded from http://pubs.acs.org on March 29, 2017

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Environmental Science & Technology

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Natural Attenuation in Streambed Sediment Receiving Chlorinated Solvents from

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Underlying Fracture Networks

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Burcu Şimşir1,2,3,4, Jun Yan2,3,4,5,6, Jeongdae Im7, Duane Graves8 and

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Frank E. Löffler1,2,3,4,6*

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TN 37996; 2Center for Environmental Biotechnology, University of Tennessee, Knoxville,

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TN 37996, USA; 3Biosciences Division, Oak Ridge National Laboratory, Oak Ridge, TN

Department of Civil and Environmental Engineering, University of Tennessee, Knoxville,

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37831; 4Joint Institute for Biological Sciences (JIBS), Oak Ridge National Laboratory,

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Oak Ridge, TN 37831; 5Key Laboratory of Pollution Ecology and Environmental

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Engineering, Institute of Applied Ecology, Chinese Academy of Sciences, Shenyang,

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Liaoning 110016, P.R.China; 6Department of Microbiology, University of Tennessee,

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Knoxville, TN 37996; 7Department of Microbiology, University of Massachusetts,

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Amherst, MA 01002; 8Geosyntec Consultants, Knoxville, TN 37919.

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*

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University of Tennessee

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Department of Microbiology

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M409 Walters Life Science Bldg.

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Knoxville, TN 37996

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Phone: (865) 974-4933

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Fax:

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E-mail: [email protected]

Corresponding author

(865) 974-4007

1 ACS Paragon Plus Environment

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Abstract

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Contaminant discharge from fractured bedrock formations remains a remediation

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challenge. We applied an integrated approach to assess the natural attenuation potential of

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sediment that forms the transition zone between upwelling groundwater from a chlorinated

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solvent-contaminated fractured bedrock aquifer and the receiving surface water. In situ

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measurements demonstrated that reductive dechlorination in the sediment attenuated

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chlorinated compounds before reaching the water column. Microcosms established with

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creek sediment or in situ incubated Bio-Sep beads degraded C1-C3 chlorinated solvents to

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less chlorinated or innocuous products. Quantitative PCR and 16S rRNA gene amplicon

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sequencing revealed the abundance and spatial distribution of known dechlorinator

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biomarker genes within the creek sediment, and demonstrated that multiple dechlorinator

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populations degrading chlorinated C1-C3 alkanes and alkenes coinhabit the sediment.

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Phylogenetic classification of bacterial and archaeal sequences indicated a relatively

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uniform distribution over spatial (300 meters horizontally) scale, but Dehalococcoides and

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Dehalobacter were more abundant in deeper sediment, where 5.7 ± 0.4 × 105 and 5.4 ± 0.9

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× 106 16S rRNA gene copies per gram of sediment, respectively, were measured. The

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microbiological and hydrogeological characterization demonstrated that microbial

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processes at the fractured bedrock-sediment interface were crucial for preventing

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contaminants reaching the water column, emphasizing the relevance of this critical zone

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environment for contaminant attenuation.

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Key-words: Chlorinated solvents, natural attenuation, reductive dechlorination,

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organohalide respiration, Dehalococcoides, Dehalobacter, fractured rock contaminant

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discharge, critical zone 2 ACS Paragon Plus Environment


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Introduction

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Chlorinated solvents have been widely used in a variety of industrial, military, and

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household applications since the 1940s.1, 2 Their extensive use, improper handling and

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disposal practices as well as accidental spills resulted in widespread subsurface and

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groundwater contamination.1, 3 Common chlorinated solvents including tetrachloroethene

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(PCE), trichloroethene (TCE), carbon tetrachloride (CT), and 1,1,1-trichloroethane

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(TCA)3 tend to form dense non-aqueous phase liquids (DNAPLs), which move

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gravitationally along interconnected fractures and form pools in low points in fractured

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bedrock formations. A significant mass of chlorinated solvents in the fractured rock site

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diffuse into low-permeability zones,4-6 and back diffusion into water-bearing fractures

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serves as long-term source of groundwater contamination.5, 7, 8

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In the last two decades, various in situ remediation technologies, including

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bioremediation, thermal and chemical treatments have been successfully applied to treat

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chlorinated solvent contamination in porous medium aquifers;3, 9 however, the remediation

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of fractured bedrock formations remains challenging due to difficulties in characterizing

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complicated fracture networks, back diffusion of contaminant from low-permeability

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zones, and the challenge of targeted delivery of remedial fluids.2, 4, 10-12 An alternate

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remedial approach focuses on treatment at the fractured bedrock-sediment interface, where

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contaminated groundwater discharges to surface waters. Recent studies have shown that

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such hyporheic zones are “hotspots” of microbial activities playing relevant roles for

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contaminant attenuation.13-16

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Organohalide-respiring bacteria (OHRB) use chlorinated hydrocarbons as terminal

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electron acceptors, and a number of species belonging to different genera (e.g., Geobacter,

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Dehalobacter (Dhb), Desulfitobacterium, and Sulfurosprillum) have been demonstrated to 3 ACS Paragon Plus Environment

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couple reductive dechlorination of PCE to TCE or cis-1,2-dichloroethene (cis-DCE) with

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energy conservation.9 In contrast, complete reductive dechlorination to environmentally

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benign ethene appears to be restricted to some strains of the species Dehalococcoides

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mccartyi (Dhc).9 Dhb are involved in dechlorination of a range of chlorinated compounds

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including chlorinated aromatics,17 chlorinated ethanes,18 chlorinated methanes,19, 20 as well

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as PCE.21 Dehalogenimonas spp. have been implicated in dehalogenation of chlorinated

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alkanes,22 and recently in reductive dechlorination of trans-1,2-dichloroethene (trans-

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DCE) to VC.23 Studies demonstrated significant correlations between the abundance of

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OHRB and observed in situ dehalogenation activities.9 Therefore, 16S rRNA genes and

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reductive dehalogenase (RDase) genes from known OHRB serve as biomarkers to assess

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in situ bioremediation activity and potential.9, 24

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At a former metal manufacturing facility located adjacent to Third Creek, a

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Tennessee River tributary in Knoxville, TN (Figure 1), chlorinated solvents, primarily

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PCE, TCE, TCA, and CT were released and penetrated the underlying fractured bedrock

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formation. Although no DNAPL source zones could be identified, dissolved phase

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concentrations of total chlorinated volatile organic compounds (cVOCs) exceeded 10 mg

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L-1 in bedrock monitoring wells indicative of free-phase chlorinated solvents (Table S1).

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Spent solvents are the primary source of groundwater contamination and, as such, the

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solvents would be co-contaminated with the oils and grease from the cleaning operation.

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Additionally, machining and lubricating oils and mineral spirits were used throughout the

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history of the facility but in much smaller quantities than the chlorinated solvents. These

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limited, and often localized, sources of hydrocarbons were not quantified during site

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assessments but are thought to have supported the modest and incomplete microbial

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transformation of contaminants observed in the fractured bedrock (Table S1), indicating 4 ACS Paragon Plus Environment


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the need for alternative remedies at the Third Creek site. This study evaluated the role of

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the streambed sediment as a natural barrier preventing contaminant discharge into Third

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Creek surface water. Integrated efforts characterizing the hydrogeological (e.g., flow

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paths) and microbiological site conditions at the Third Creek site demonstrated efficient

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natural attenuation in the sediment.

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Material and Methods

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Chemicals. Chlorinated compounds were of >99% purity. PCE and CT were purchased

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from ACROS Organics (Morris Plains, NJ), TCE was obtained from Fisher Scientific

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(Pittsburgh, PA), and cis-DCE, vinyl chloride (VC), ethene, TCA, dichloromethane

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(DCM), chloroform (CF), chloromethane (CM), 1,2-dichloropropane, 1,1-dichloroethane

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(DCA), and chloroethane (CA) were obtained from Sigma-Aldrich-Fluka (St. Louis, MO).

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Site characterization. The manufacturing site is bounded by Third Creek on its west side

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(Figure 1). The facility used chlorinated solvents as degreasers from the mid-1930s ending

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in the late 1990s. During the course of manufacturing activities, chlorinated solvents,

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primarily PCE, TCE, and, to a smaller extent, TCA and CT, were released resulting in

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contamination of the underlying groundwater-bearing fractured bedrock (Figure S1-2).

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Direct observation of soil and bedrock cores failed to locate DNAPL although

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contaminant concentrations as high as 120 mg/L were measured in non-flowing fractures

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(Figure S3). The presence of cis-DCE, VC, DCA and CF in several bedrock monitoring

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wells indicated that some contaminant transformation occurred; however, high

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concentrations of parent compounds and no ethene and ethane formation indicated limited

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dechlorination capacity within the fracture network (Table S1). Groundwater seepage rates

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were measured with leak-tested seepage meters.25 Horizontal and vertical bedrock 5 ACS Paragon Plus Environment

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groundwater flow direction was evaluated using seasonal groundwater elevations from

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monitoring wells screened at multiple depths and staff gauges in the creek. The SI

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provides additional details about the site characterization efforts.

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Sediment pore water diffusion sampling. To measure concentrations of cVOCs,

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methane, ethene, and ethane, as well as geochemical parameters, sediment pore water

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diffusion samples were collected with depth-discrete diffusion samplers loaded with 40-

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mL glass vials.26 Before placement of the samplers into the sediment, the 40-mL glass

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vials were filled with deionized water and covered with either polyethylene film for cVOC

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sampling, or a porous, nonwoven fabric for measurement of geochemical parameters. The

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samplers were installed in the same locations as the seepage meters 0-0.55 m (0-1.8 ft)

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below the sediment surface, and left in the creek for 2 weeks to achieve equilibrium with

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the pore water. After recovery of the samplers, the vials were immediately sealed and

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shipped to analytical laboratories (SiREM, Guelph, Canada and Microbac, Maryville, TN)

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for volatile fatty acids (VFAs), cVOC and anions measurements.

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Sediment collection. Grab sediment samples and cores were collected from locations #1,

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#2 and #3 (Figure 1). These locations were chosen based on the observed sediment cVOC

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concentrations and the site’s hydrogeological characteristics. Top layer sediment samples

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were collected using autoclaved, sealable glass containers (Mason jars). Deeper sediment

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layers were obtained using direct push tools (AMS, Inc., American Falls, ID). The plastic

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liners and caps were wiped with 70% ethanol before use. All other materials (spatulas,

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containers, etc.) were autoclaved and aseptic techniques were applied to the extent feasible.

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All core samples were immediately transferred to sterile Mason jars, filled completely

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with creek water to exclude air, capped and placed in a cooler with ice packs.



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Depth-resolved sediment collection. Depth-discrete diffusion samplers were employed to

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collect sediment at multiple depths at Location #3. The diffusion sampler was loaded with

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40-mL glass vials evenly spaced over a length of 99 cm (3.2 ft). The vial openings were

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covered with plastic mosquito netting (1-mm mesh size) held in place with rubber bands.

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The loaded sampler was pushed into the sediment to a depth of about 55 cm (1.8 ft). A

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second sampler loaded with customized Bio-Trap samplers (about 200 Bio-Sep beads per

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sampler, Microbial Insights, www.microbe.com) was placed in the sediment in the same

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location. After a 1-month incubation period, the samplers were removed and the glass

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vials with sediment material were immediately closed with sterile Teflon-lined rubber

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septa. The vials and the Bio-Sep bead cartridges were placed individually in Ziploc bags,

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immediately transferred to a cooler with ice and transported to the laboratory for

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microcosm setup and DNA extraction.

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Sample handling. Grab samples and sediment core samples from the same locations were

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combined and mixed inside an anoxic glove box filled with H2/N2 (3/97%, vol/vol).

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Approximately 100 g of sediment material from each of the three sampling locations was

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transferred to sterile plastic tubes and stored at -80°C. The remaining sediment materials

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were kept at 4°C and microcosms were established within 1 week of sample collection.

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Sediment materials collected in the 40-mL glass vials from eight different depths were

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individually transferred to sterile plastic tubes, homogenized and stored at -80°C for

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molecular analyses. The Bio-Trap samplers collected from eight depths were opened

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inside the glove box and about 200 Bio-Sep beads per depth sample were transferred to a

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sterile plastic tube for DNA extraction and microcosm experiments. All procedures used

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strictly aseptic techniques. To prevent cross contamination, sediment samples from

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different locations were not handled simultaneously. 7 ACS Paragon Plus Environment

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Microcosm setup. For sediment microcosm setup inside the anoxic glove box,

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approximately 4 g (wet weight) of the sediment from locations #1, #2, or #3 were

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transferred to sterile 60-mL glass serum bottles. Twenty-six mL of anoxic, bicarbonate-

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buffered mineral salts medium27 amended with 5 mM lactate was added to each bottle

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before the vessels were sealed with autoclaved butyl rubber stoppers (Geo-Microbial

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Technologies, Inc., Ochelata, OK, USA). Neat PCE, TCE, cis-DCE, VC, TCA, DCA, 1,2-

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dichloropropane, CT, CF, DCM, and CM were added to triplicate microcosms with 5 or

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10 µL Hamilton glass syringes (Hamilton 85925 and 80370) to achieve initial aqueous

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phase concentrations of approximately 0.2 mM (12.5-33.1 mg/L). One microcosm for

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each treatment was autoclaved for 60 min at 121°C. An additional set of live control

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microcosms received all amendments except the cVOC additions.

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To obtain depth-resolved information about microbial reductive dechlorination

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activity, additional microcosms were established with in situ incubated Bio-Sep beads

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collected at Location #3. Inside the glove box, five beads were transferred to sterile 60-mL

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glass serum bottles containing 30 mL of reduced mineral salts medium amended with 5

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mM lactate. The bottles were closed with butyl rubber stoppers and PCE, TCE, cis-DCE,

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VC, or TCA were added to reach aqueous phase concentrations of approximately 0.2 mM.

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Initially, cVOCs could not be measured due to sorption to the Bio-Sep beads but

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quantitative analysis was possible in transfer cultures. After a 2-month incubation period,

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3% (vol/vol) culture suspension without beads was transferred to fresh medium amended

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with 5 mM lactate and 0.2 mM of the respective cVOC. Enrichment cultures that showed

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no reductive dechlorination received 6 mL of H2 to ensure that electron donor availability

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was not a limitation. All microcosms and enrichment cultures were incubated statically at

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room temperature in the dark and monitored over a 20-month incubation period. 8 ACS Paragon Plus Environment


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DNA isolation, qPCR, and 16S rRNA gene amplicon sequencing. To assess the spatial

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distribution of known dechlorinators in relation to the three different sampling locations

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and sediment depth at Location #3, PCR, quantitative PCR (qPCR), and 16S rRNA gene

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fragment amplicon sequencing were performed. DNA was extracted from 0.25 g of wet

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sediment using the MoBio PowerSoil DNA Isolation Kit (MO BIO, Carlsbad, CA). DNA

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extraction from Bio-Sep beads (~160 beads/depth) was performed by Microbial Insights

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using established procedures.28 Published primer sets and PCR conditions were used to

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amplify total bacterial,29 Dhc,30 Dhb,31 Dehalogenimonas22 and Geobacter lovleyi strain

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SZ32 16S rRNA genes and dcpA encoding a 1,2-dichloropropane-to-propene RDase.33 For

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increased sensitivity of detection, a nested PCR approach was applied 34 (see SI for

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details). qPCR to enumerate total bacterial, Dhc and Dhb 16S rRNA genes, as well as the

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bvcA, vcrA, tceA, and cfrA RDase genes used published primers and probes and followed

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established protocols (Table S2).

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To compare the microbial communities at locations #1, #2 and #3, amplification of

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the hypervariable V1-V3 and V3-V5 regions of the 16S rRNA gene and subsequent

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pyrosequencing of the PCR amplicons was performed with barcoded-primers35-38 (Table

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S2). Library preparation was performed as described38 with minor modifications (SI), and

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pyrosequencing was performed on a 454 FLX Life Sciences Genome Sequencer (Roche

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Diagnostics) according to manufacturer’s instructions. Sequence data analyses followed

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established procedures (see SI).

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Analytical Methods. cVOCs, ethene, ethane, propene and methane were monitored using

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an Agilent 7890 gas chromatograph (GC) equipped with a flame ionization detector and a

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DB-624 capillary column (60 m by 0.32 mm with a film thickness of 1.18 µm) as


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described39. Details of the method for cVOCs measurements by SIREM Laboratory, are

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given in SI.

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Results.

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Third Creek site hydrological features. Available hydrogeological data sets from 46

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sampling locations on both sides of Third Creek consistently indicated that groundwater

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has an upward gradient, a vertical gradient, and horizontal gradients sloping toward the

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creek from both sides. The groundwater seepage meters placed in creek sediment

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confirmed, as predicted by vertical (Figure 1) and horizontal gradients (not shown), that

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the creek received groundwater with maximum, mean and median seepage rates of 3,380,

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721 and 205 mL/m2 day, respectively, at Location #3. The broad range of measured

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seepage rates were likely influenced by the creek stage, sampling location, and

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groundwater elevation. The lowest mean and median seepage rates of 47 and 5.4 mL/m2

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day, respectively, were measured near Location #1, and occasionally negative values (i.e.,

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losing water) were observed. Assuming the highest rate of seepage (3,380 mL/m2day), a

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sediment depth of 0.52 meters, a sediment porosity of 20 percent, and no retardation of

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cVOC migration in the sediments, the shortest retention time for groundwater seeping

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through the sediment into the creek was calculated to be approximately 30 days.

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The seepage measurements and vertical and horizontal groundwater gradients

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suggest that Third Creek receives groundwater from underlying fractures downstream of

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Location #1 (Figure 1). The elevation data suggest that the creek transitions from a losing

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to a gaining creek along the perimeter of the contaminated area near Location #1 (see SI

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for details). Based on the average creek widths of 7.3 meters between locations #1 and #2

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and 10.4 meters between locations #2 and #3, segment lengths of 226.5 meters between 10 ACS Paragon Plus Environment


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locations #1 and #2 and 77.1 meters between locations #2 and #3, the yearly volume of

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groundwater seeping into Third Creek between locations #1 and #3 was estimated at

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42,000 to 170,000 L/year using median and mean seepage rates, respectively.

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Third Creek site geochemical characteristics. At Location #3, sediment pore water

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measurements detected cis-DCE, VC, ethene, ethane and methane in the deep sediment,

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whereas significantly lower cVOC concentrations were measured near the sediment-

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surface water interface. Concentrations varied with depth (Figure 2) and maximum ethene,

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ethane and methane concentrations of 0.25, 0.08 and 5.2 mg/L, respectively, were

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measured near the sediment-surface water interface. This observation suggests the

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formation of ethene and ethane as reductive dechlorination products with their further

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degradation occurring in shallower sediment layers. Using Location #3 ethene, ethane, and

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cVOC data, the change in ethene and ethane concentration from 0.52 meters to 0.22

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meters depth accounts for approximately 92 percent of the loss in cVOCs on a mole basis.

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cVOC concentrations were lower near the sediment-surface water interface and the cis-

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DCE and VC concentrations were less than the method quantification limit of < 0.01 mg/L.

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Maximum cis-DCE and VC concentrations of 0.78 mg/L and 0.33 mg/L, respectively,

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were observed in the deeper sediment at Location #3. The lack of cVOC detections at the

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sediment/surface water interface could be the result of dilution with the creek water;

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however, the upward flow of water into the creek, the lack of cVOC detections below the

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sediment/water interface, and the presence of methane in the shallow sediment indicate

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that the influence of surface water on pore water cVOC concentrations is inconsequential.

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With the exception of cis-DCE (0.03 mg/L) in the deep sediment at Location #1, no

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cVOCs were detected in sediment pore waters collected at locations #1 and #2 (Table S3).

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TCE concentrations were generally below 0.01 mg/L in the deep sediment, but TCE was 11 ACS Paragon Plus Environment

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occasionally detected near the sediment-surface water interface (Figure 2, Table S3). In

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addition, concentration gradients of VFAs, sulfate, and chloride were observed in the

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sediment pore water at Location #3. VFA concentrations of 374 mg/L in the deep

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sediment pore water decreased to 4 mg/L in pore water collected near the sediment-

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surface water interface, and sulfate concentrations increased from

Natural Attenuation in Streambed Sediment Receiving Chlorinated

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