Microbial Mats as a Biological Treatment Approach for Saline

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Microbial Mats as a Biological Treatment Approach for Saline Wastewaters: The Case of Produced Water from Hydraulic Fracturing Benay Akyon, Elyse Stachler, Na Wei, and Kyle Bibby Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/es505142t • Publication Date (Web): 13 Apr 2015 Downloaded from http://pubs.acs.org on April 20, 2015

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Environmental Science & Technology

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Microbial Mats as a Biological Treatment Approach for

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Saline Wastewaters: The Case of Produced Water from

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Hydraulic Fracturing

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Benay Akyon1, Elyse Stachler1, Na Wei1, Kyle Bibby1,2*

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Systems Biology, University of Pittsburgh, Pittsburgh, PA 15261, USA

Department of Civil and Environmental Engineering and 2Department of Computational and

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*

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[email protected], 412-624-9207

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Keywords: Microbial mats, biodegradation, biological treatment, saline wastewater, hydraulic

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fracturing, wastewater, produced water, flowback water

Corresponding Author: Kyle Bibby, 709 Benedum Hall, Pittsburgh, PA 15261

ACS Paragon Plus Environment


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ABSTRACT

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Treatment of produced water, i.e. wastewater from hydraulic fracturing, for reuse or final

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disposal is challenged by both high salinity and the presence of organic compounds. Organic

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compounds in produced water may foul physical-chemical treatment processes, or support

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microbial corrosion, fouling, and sulfide release. Biological approaches have potential

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applications in produced water treatment, including reducing fouling of physical-chemical

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treatment processes and decreasing biological activity during produced water holding; however,

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conventional activated sludge treatments are intolerant of high salinity. In this study, a biofilm

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treatment approach using constructed microbial mats was evaluated for biodegradation

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performance, microbial community structure, and metabolic potential in both simulated and real

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produced water. Results demonstrated that engineered microbial mats are active at total

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dissolved solids (TDS) concentrations up to at least 100,000 mg/L, and experiments in real

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produced water showed a biodegradation capacity of 1.45 mg COD/gramwet-day at a TDS

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concentration of 91,351 mg/L. Additionally, microbial community and metagenomic analyses

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revealed an adaptive microbial community that shifted based upon the sample being treated and

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has the metabolic potential to degrade a wide array of contaminants, suggesting the potential of

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this approach to treat produced waters with varying composition.

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INTRODUCTION

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Advances in high-volume hydraulic fracturing and horizontal drilling techniques have enabled

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oil and gas production from unconventional reservoirs and have altered the current and future

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energy landscape. In 2012, more than 34% of U.S. natural gas was produced from

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unconventional resources,1 and that percentage is expected to increase.2 Additionally, shale gas

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resources are globally distributed,3 with worldwide exploration expected to begin in the coming

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decades. High-volume hydraulic fracturing involves the injection of 10-20 million liters of

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fracture fluid at high pressure to fracture the target formation and stimulate reservoir

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permeability.4 Following well completion, a portion (5% to greater than 100%) of the fracture

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fluid mixed with subsurface brine returns to the surface as produced water.5, 6 Fracture fluid is

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typically 99% water and sand, with the remaining 1% comprised of chemicals to regulate pH,

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viscosity, and reduce friction, precipitation, scaling and biological fouling.4,

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typically has an elevated total dissolved solids (TDS) concentration that ranges from 5,000 to

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300,000 mg/L,7-9 including high concentrations of sodium, calcium, barium, strontium, chloride,

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bromide and naturally occurring radioactive material (NORM).9,

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typically contain a large suite of poorly defined organic compounds.11 A large volume of this

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water is produced upon well completion and is known as ‘flowback water’; however, wells

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continue to produce water during their entire operation. Here, the term ‘produced water’ is used

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to refer to all wastewater generated during unconventional well operation. Produced water

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characteristics are unique to each geological formation.10, 12

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Water management challenges associated with hydraulic fracturing, including produced water

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disposal and water sourcing for fracturing, have emerged at the forefront of the public and

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regulatory discussion regarding hydraulic fracturing. Due to large volumes and high salt


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Produced water

Produced waters also


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concentrations, disposal and treatment options for produced water are limited. Treatment of

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produced water in municipal wastewater treatment plants for surface disposal is no longer a

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viable alternative with new regulations in effect.4 Deep well injection is one of the most common

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methods for produced water disposal; however, some regions (e.g. Pennsylvania) have limited

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deep well disposal capacity.4 Additionally, induced seismic activity has been associated with

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deep well injection,13 suggesting the potential for future regulatory limitations to this disposal

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approach. Finally, in some regions with suitable disposal capacity, there are concerns about the

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environmental impacts of fresh water sourcing.6

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Biological treatment is a promising and underexplored treatment technology to remove organic

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compounds in high-salinity produced water. Biological treatment approaches may be used in

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conjunction with physical-chemical treatment to limit energy costs and membrane fouling for

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both produced water reuse in future hydraulic fracturing operations and final disposal.

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Additionally, reduced concentrations of organic compounds due to biological treatment would

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limit heterotrophic microbial growth during produced water holding, decreasing the need for

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biocide use. Recently, the effect of dissolved solids on chemical oxygen demand (COD)

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biodegradation in sequencing batch reactors was examined.14 While the biological removal of

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COD decreased the membrane fouling potential, a significant decrease in COD degradation with

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increasing salt concentration was observed,14 prompting further investigation into the suitability

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and potential of biological treatment approaches for produced water. These results suggest the

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need for more halo-tolerant biological treatment approaches.

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Microbial mats, clustered biofilms of mixed microbial communities,15,

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hypersaline habitats, with abundant microbial diversity.16, 20 Microbial mats have been used as a

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bioremediation technique for more than twenty years, and are tolerant of high salinity as well as


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occur naturally in

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metal and metalloid toxicity.17 An early study demonstrated oil biodegradation by microbial mats

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developing in an oil-contaminated area.18, 19 Hypersaline microbial mats are capable of removing

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25-85% of model petroleum compounds, with performance decreasing as salinity increases.19-21

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Various approaches have been attempted to cultivate microbial mats,16, 17, 22-24 including using

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glass wool,17 coconut mesh,22 polyester fiber,23 silica particles,24 and grass silage16 as a growth

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scaffold. Among these, grass silage has been found to perform well due to stimulation of rapid

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microbial growth by providing a scaffold surface to support microbial growth and an initial

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supply of nutrients including lactic acid, amino acids and various minerals.16

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In the current study, microbial mats were constructed using grass silage, the degradation of

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model compounds at various TDS concentrations was tested in both simulated and real produced

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water, and biodegradation rates were evaluated. In addition, 16S rRNA and shotgun

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metagenomic sequencing was performed to evaluate the microbial community structure and

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metabolic potential of microbial mats treating produced water.

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MATERIALS AND METHODS

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Construction of engineered microbial mats

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To prepare microbial mats, window screen was cut in approximately 2.5 centimeter diameter

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circles, filled with grass silage (1 gram), and sewn together (Fig S1). Growth medium for the

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microbial mats was comprised of 25 g/L Luria Bertani (LB) broth in deionized water (Synergy-R

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purification system with 18.2 MΩ resistance) amended with 50,000 mg/L TDS (35 g/L NaCl, 15

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g/L CaCl2). The growth media was seeded with 10% (v/v) of a mixed stock of produced water

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and activated sludge from municipal wastewater. Prepared mats were then placed in the growth



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medium in a 2 L plastic beaker and mixed continuously for 21 days to maintain aerobic

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conditions. The same growth batch of microbial mats was used for each set of experiments.

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Preparation of the test media

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Microbial mats guar gum degradation capacity was tested in both synthetic and real hydraulic

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fracturing produced water; acetate degradation was tested only in synthetic produced water.

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Acetate was utilized as a model simple organic molecule, as a fermentation and breakdown

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product from other complex organic molecules in fracturing fluid, such as guar gum, and has

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previously been identified in produced water.25 In experiments conducted with synthetic

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produced water media, NaCl and CaCl2 were added at a Na/Ca mass ratio of 3.510 to supply TDS

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concentrations of 0, 50,000, 100,000, or 200,000 mg/L. For the synthetic produced water acetate

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degradation experiment, a 5,000 mg/L acetate stock solution was prepared with 6.94 g/L sodium

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acetate in deionized water. The stock solution was used to provide a 2,500 mg/L acetate

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concentration to all test conditions. In the synthetic produced water guar gum degradation

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experiment, guar gum solution was prepared using a modified approach of Lester et al.14 Guar

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gum is a commonly used chemical in fracturing fluid to increase viscosity14 and was used here as

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a representative complex organic COD source. Typical guar gum dosage in fracturing fluid

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ranges from 600-4800 mg/L.26 Briefly, 3,000 mg/L guar gum was prepared with deionized water,

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the supernatant was withdrawn after an 18-hour settling period and filtered through glass fiber

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filters (Fisher Scientific, Pittsburgh, PA). The resulting filtrate had a COD value of

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approximately 2,500 mg/L. Real produced water experiments were conducted with two different

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produced water samples, Sample A and Sample B, together with each sample diluted one half,

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Sample A1/2 and Sample B1/2. Sample characteristics are shown in Table 1. Sample A (182,702

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mg/L TDS) was produced water from a well in southwest PA, and the biocide used in the


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fracturing fluid was glutaraldehyde. Sample B (18,400 mg/L TDS) was from an open produced

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water holding pond containing water from multiple wells in southwest PA that was maintained

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with chlorine dioxide. All test conditions were supplemented with 2,500 mg COD/L guar gum as

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described above. As the produced water samples used had low biodegradable COD

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concentrations, guar gum addition was utilized to more accurately provide a degradation rate

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estimate. Chemical details of all test media are included in Table 1.

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Experimental Procedure

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Prior to use, microbial mats were rinsed for 1.5 hours at 70 rpm in 50,000 mg/L TDS synthetic

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produced water to limit the carryover of the cultivation media to the test medium. Following

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rinsing, the wet weight of the mats was recorded and they were placed into 10 mL of test media

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in 6-well plates (Corning Costar, Tewksbury, MA). For an initial homogenization period, 6-well

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plates were placed on a shaker table at 110 rpm for 15 minutes and time zero sampling was

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performed. Microbial mats were continuously shaken at 110 rpm throughout each experiment.

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All conditions were sampled at 0, 6, 24, 48, and 72 hours. Successive loadings of test media were

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performed, with each loading lasting 72 hours. The 72 hour loading period was chosen based

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upon preliminary tests demonstrating limited substrate removal following this time period. Three

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biological replicates were conducted for each test condition. Surface samples of the mats were

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taken at the beginning and at the end of each cycle and stored at -20°C for later microbial

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analysis.

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High performance liquid chromatography (HPLC, Agilent Technologies 1200 Series) and

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HACH (Loveland, CO) COD kits were used to quantify acetate and guar gum, respectively. In

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acetate experiments with synthetic produced water, 0.5 mL of liquid was withdrawn from each



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well and centrifuged (Fisher Scientific, Accuspin Micro 17) at 4000xg for 5 minutes.

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Supernatants were transferred to slip syringes (Fisher Scientific, Luer-Slip Syringes, 3 mL

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capacity) and filtered through syringe filters (Fisher Scientific, Cellulose Syringe Filter, 0.2µm)

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into microcentrifuge tubes (Fisher Scientific, 1.5 mL) and 0.2 mL from each filtered sample was

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analyzed by HPLC. The HPLC was equipped with a refractive index detector and a Rezex ROA-

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Organic Acid H+ (8%) column (Phenomenex Inc., Torrance, CA). The column was eluted with

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0.005 N H2SO4 as a mobile phase at a flow rate of 0.6 mL/min at 50°C with a quaternary pump.

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In guar gum experiments, 0.5 mL of liquid was withdrawn from each well, diluted 20 times with

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deionized water, and filtered through 0.45 µm filter paper (Millipore MF-EMD, Billerica MA).

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Filtered samples were stored at -20°C prior to analysis. The COD of the filtered samples were

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measured with HACH LR (low range) COD vials and a DR850 HACH Colorimeter. Due to the

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high salt content of samples, 0.5 g of mercuric sulfate (Acros Organics, Geel, Belgium) was

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added to COD vials to decrease chloride interference.27 Standard errors were calculated with

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biological triplicates of each test condition. Technical replicates of COD measurements were

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performed for each time point.

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Rate Analysis and Kinetics

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Rate analysis was performed on all experiments to analyze COD removal. The assumed

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theoretical COD value from acetate was 0.92 g acetate/g COD (64 g COD/mole acetate). The

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second and third loadings of each experiment were utilized to determine the removal rates.

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Reaction rate constants are shown in Table S1. The highest and lowest kinetic constants were

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selected (denoted as highest and lowest performance, respectively) for each TDS condition.

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Microbial Ecology


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DNA was extracted from each sample using the PowerSoil DNA Isolation Kit (MO BIO

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Laboratories, Carlsbad, CA) and quantified using a Qubit (Life Technologies, Carlsbad, CA).

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The 16S rRNA region was PCR amplified using the 515F primer and 806R barcoded sequencing

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primers as previously described.28 Additional details are included in the Supplemental

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Information. Sequence reads were analyzed using QIIME 1.7.0.29 The sequences were

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demultiplexed and filtered to remove sequences that did not contain a correct sample barcode

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and did not have a quality score of Q20 or higher. Demultiplexed and quality filtered sequences

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are available on MG-RAST30 (accession 4576856.3). Closed reference OTUs were predicted

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from the quality filtered sequences (pick_closed_reference_otus.py) and assigned taxonomy

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based on the Greengenes 13_8 database.31 For alpha diversity estimation, a single rarefaction

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was run using 4500 random sequences from each sample and alpha diversity was calculated

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using the observed_species metric.

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DNA for metagenomic sequencing was prepared using Illumina’s Nextera XT DNA Sample

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Preparation Kit (San Diego, CA) without kit normalization. Samples were denatured and pooled

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to a final concentration of 12 pM. Sequencing was performed with a MiSeq reagent Kit V2 (500

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cycles) (Illumina, San Diego, CA). Sequences were quality trimmed in CLC Genomics

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Workbench 7.0.3 (CLC Bio, Aarhus, Denmark) by the following parameters: a Q score of Q30

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or higher, a length greater than 100 bp, and the maximum number of ambiguities allowed was 2.

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Quality filtered sequences were then assembled using a kmer of length 20 and a minimum contig

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length of 500 bps. Assembled contigs were submitted to the MG-RAST server for annotation and

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are available under accession numbers 4570162.3, 4570163.3, 4570164.3, and 4570165.3. In

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MG-RAST, contigs were functionally annotated by the Subsystems pathways and KEGG

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Orthology, extracted, and taxonomically assigned using Galaxy/MGTAXA.32



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Statistical analysis was performed in Minitab statistical software (Versions 16 and 17, State

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College, PA). One-way ANOVA analysis was conducted on biological removal data and a

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pairwise comparison of each data set was performed. Confidence intervals (95%) and unequal

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variances were assumed. Statistical analysis of removal rates was conducted by ANOVA

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General Linear Model by selecting TDS as covariate with 95% confidence interval. For

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microbial ecology statistical analysis, an ANOVA General Linear Model was run with 2 factors

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(sample and cycle) along with interaction from the two factors with an individual family as the

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response for taxonomic diversity or the number of observed species for alpha diversity. Grouping

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information was obtained using Tukey comparisons at a 95% confidence level.

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RESULTS

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Synthetic Produced Water Experiments

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Results from synthetic produced water experiments with acetate and guar gum are provided in

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Figure 1A and B, respectively. The first loading cycle demonstrated significant performance

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variability for all conditions, potentially due to growth media carryover or microbial community

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adaptation. For acetate, the removal performance of the 0 and 50,000 mg/L total dissolved solids

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(TDS) conditions were statistically indistinguishable (p=0.735), while the 100,000 mg/L TDS

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condition demonstrated statistically significant decreased performance (p0.99 in all conditions) and lowest

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removals (R2>0.96 for 0 and 50,000 mg/L and R2 =0.53 for 100,000 mg/L TDS) within a 72 hour

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period. The second and third produced water loading cycles of each experiment were evaluated



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to determine removal rates (Figure 2). Comparable removal rates were observed in 0 and 50,000

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mg/L TDS conditions for degradation of each substrate, with reduced rates at 100,000 mg/L

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TDS. No degradation was observed at 200,000 mg/L TDS. Microbial mats demonstrated higher

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guar gum degradation rates in real produced water than in synthetic produced water (p=0.008).

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Microbial Ecology

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The microbial ecology of mats treating Sample A1/2, Sample B, and Sample B1/2 was analyzed

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using high-throughput 16S rRNA sequencing. The ecology of mats treating Sample A was not

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analyzed, as no degradation was observed. Samples were taken and analyzed prior to the first

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loading and following the first and third loadings. After quality filtering, the number of

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sequences ranged from 5,144–36,602 per sample (Table S2). Figure 3 shows the average class

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level abundances for each condition. The taxonomic composition of the microbial mats differed

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by the salinity of the sample being treated, while biological replicates tended to be more similar

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in composition. ANOVA was performed on family level diversity to compare the microbial

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community of the initial mats with the ecology of mats following operational cycles (Table S3).

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Sample A1/2 exhibited a significant increase in Idiomarinaceae, specifically the genus

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Idiomarina. All samples except Sample A1/2 at the end of the first loading showed a significant

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decrease in the Campylobacteraceae, specifically the genus Arcobacter. Sample B showed a

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significant increase in the Rhodospirillaceae family while Sample B1/2 showed a slight increase,

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corresponding to the genus Novispirillum, which is not generally considered to be halotolerant.

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This is reflected in Figure 3 by the large increase in Alphaproteobacteria in the Sample B mats at

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the end of the third loading.


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Alpha diversity was calculated to estimate the number of species observed in each sample (Table

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S2). An ANOVA General Linear Model was run on the data to observe statistical differences

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between sample types and cycles. The mats at time zero, along with the Sample A1/2 mats were

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found to have a significantly lower number of species than microbial communities from Sample

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B from the later cycles.

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Four metagenomes were generated in this study: two mats treating Sample A1/2 (91,351 mg/L

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TDS) and two mats treating Sample B (18,400 mg/L TDS), all from the end of the third loading.

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These conditions were chosen to provide replication and a range of salinity concentrations, while

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maximizing sequencing coverage. Results from sequencing are included in Tables S4-S6. While

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differing slightly from 16S rRNA sequencing data, metagenomic taxonomic data (Figure 3)

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exhibited general trends from 16S rRNA sequencing, such as an increase in the

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Alphaproteobacteria population in the Sample B mats. At the genus level, the Sample A1/2

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metagenomes also captured an increase in the Idiomarina population similar to the 16S rRNA

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data (Table S5). The metagenomes were analyzed for carbon metabolism pathways and stress

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response subsystems. Guar gum is composed of a mannan backbone with galactose residues

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attached.33 In addition to the enzymes necessary for guar gum degradation, the mat samples

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included necessary genes for diverse carbohydrate degradation, including xylose, ribose,

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fructose, sucrose, and glycogen (Figures S2 and S3). Additionally, the metagenomes evidenced a

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wide range of stress response mechanisms. Mechanisms of osmotic stress response in the mat

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communities include choline and betaine uptake and sodium antiporters.

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To relate functional annotations with observed shifts in the microbial community, functionally

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annotated metagenomic sequences were extracted and taxonomically classified. Specific

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functional annotations investigated included carbohydrate, galactose, and mannose/fructose



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metabolism, and oxidative and osmotic stress response (Figure 4). Results demonstrated that

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functional microbial community changes mirrored taxonomic shifts. Specifically, analyses

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showed an increased abundance of Alphaproteobacteria, Flavobacteriia and Sphingobacteriia in

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Sample B compared to Sample A1/2, and Sample A1/2 demonstrated an increased abundance of

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Gammaproteobacteria and Bacteroidia. Moreover, analyses at the genus level (Table S4)

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indicated that the most abundant genera associated with carbohydrate metabolism were

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Halomonas, Marinobacterium, and Marinobacter in Sample A1/2, and Vibrio and

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Flavobacterium in Sample B.

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DISCUSSION

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This study demonstrates the ability of engineered microbial mats to treat saline hydraulic

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fracturing produced water. Several concerns (e.g. fouling, souring, and corrosion) regarding

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produced water holding and reuse have emerged, generally requiring some level of treatment

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before reuse, and biocide application during holding. Currently, nearly all produced water

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treatment is physical-chemical and generally involves transportation to a centralized facility.

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Biological produced water treatment approaches may be combined with physical-chemical

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treatment to reduce process fouling, or applied prior to produced water holding to reduce

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biological activity. The development of on-site and low-cost treatment options, including biofilm

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processes such as microbial mats, will encourage produced water reuse in future hydraulic

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fracturing operations, reducing the environmental impacts of fresh water sourcing, produced

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water transportation and disposal, and excess chemical usage. For example, biocide application

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is commonly used to control biological activity during produced water holding, often with

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limited efficacy.34, 35 Biological treatment to remove available electron donors has the potential

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to decrease heterotrophic microbial growth and the necessity of biocide use. It is envisioned that


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microbial mats may be used as either an on-site technology during produced water holding or

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coupled with physical-chemical treatment to reduce process fouling, although additional

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evaluation is necessary to determine the best application of this technology.

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Microbial mat treatment experiments conducted for two different substrates in synthetic and real

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produced water demonstrated similar trends. For all experiments, the first sample loading

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showed significant performance variability. This is likely due to the acclimation period of the

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microorganisms36 or growth medium leaching from the mats to the test medium. Our approach

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was to treat the first cycle as a wash and adaption step and exclude it from further rate

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evaluation. After several successive loadings, the biodegradation performance decreased with a

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steeper decrease at higher TDS concentrations. This behavior could be a result of a reduction in

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mat activity due to nutrient limitation, salinity stresses, or starvation between cycles. The highest

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biodegradation rates were observed at 0 and 50,000 mg/L TDS conditions for both acetate and

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guar gum. The 100,000 mg/L TDS condition exhibited a decreased rate, and no biological

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substrate removal was observed in the 200,000 mg/L TDS condition. Tests in real produced

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water reflected trends observed in synthetic produced water. A similar trend was seen in real

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produced water experiments, where the performance of mats decreased in Sample A1/2 (91,351

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mg/L TDS) and no biodegradation occurred in Sample A (182,702 mg/L TDS), similar to the test

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conditions with salt concentrations of 100,000 and 200,000 mg/L TDS, respectively. Improved

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COD removal in real produced water is likely due to constituents in the produced water, such as

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trace minerals, organics, nutrients, or salts. Additionally, it has been shown that trace minerals

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(e.g. Mg, Fe) have a significant effect on cell viability during starvation conditions.37 As

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demonstrated by microbial mats treating synthetic produced water, salinity had a strong role in

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driving the performance of the mats. In real produced water, uncharacterized compounds,



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including biocides, may have influenced the performance; however, these results demonstrate the

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ability of microbial mats to perform in actual produced water samples.

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The influence of salinity on microbiological treatment performance has long been recognized.38

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Previous studies have demonstrated that activated sludge treatment systems experience a sharp

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decrease in COD removal efficiency above TDS concentrations of 10,000 mg/L,39,

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decrease in the COD removal efficiency from 85% to 59% when the TDS concentration

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increased from zero to 50,000 mg/L.40 A recent study evaluating the efficiency of activated

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sludge to treat synthetic produced water demonstrated a 60% removal efficiency in 31 hours at a

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salt concentration of 45,000 mg/L.14 The current study demonstrates guar gum COD removals of

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66% and 45% COD at 50,000 and 100,000 mg/L TDS concentrations, respectively. A rate

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analysis (Figure 2) demonstrated that the acetate degradation rate by microbial mats was higher

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than the guar gum degradation rate, likely due to the simpler chemical structure of acetate. Both

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acetate and guar gum showed similar removal rate trends with increasing salt concentration.

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Microbial ecology analyses were performed to further the understanding of degradation and

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survival mechanisms of microbial communities in engineered microbial mats. The microbial mat

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community composition was strongly driven by the sample being treated. Mats treating higher

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TDS samples exhibited a lower alpha-diversity, while the microbial community of mats treating

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lower TDS samples diverged throughout the study, suggesting salinity constrained microbial

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diversity. Idiomarina increased with each treatment cycle in the higher salinity Sample A1/2,

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suggesting this may be an important population in mats treating higher salinity produced water.

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This genus is comprised of moderate halophiles that are generally strict aerobes41, 42 and have

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been isolated from various marine environments, including a hypersaline solar saltern.43 They

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have a high content of iso-branched fatty acids and certain species produce large amounts of


40

and a

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extracellular polymeric substances (EPS) to facilitate biofilm formation, both of which would

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assist growth in the harsh conditions of high salinity produced water,44,

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salinity protection for the biofilm. Idiomarina cannot use carbohydrates as their sole carbon and

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energy sources, instead using amino acids to obtain carbon and energy,44 and cannot utilize

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mannose for energy. This suggests that they cannot grow on guar gum alone,41,

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require amino acids synthesized by the microbial community for survival.

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Microbial community analyses of microbial mats in this study demonstrate similar trends as

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previous studies of the microbial communities in oil and gas wastewater treatment and hydraulic

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fracturing produced

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Gammaproteobacteria classes have previously been found to degrade 80-100% of polycyclic

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aromatic hydrocarbons in crude-oil contaminated marine water.46 Additionally, microbial

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community analysis of an acetate-fed denitrification reactor showed increased abundance of

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Gammaproteobacteria with increasing salinity,47 consistent with trends observed here. Finally,

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previous studies of the microbial ecology of early stage produced water have identified the

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predominance of Alphaproteobacteria, Gammaproteobacteria, Bacteroidia, and Clostridia,48, 49

369

consistent with the dominant microbial groups identified in microbial mats following three

370

cycles of treatment.

371

Functional metagenomic analysis demonstrated the presence of metabolic pathways necessary to

372

degrade the substrate (guar gum) and respond to the saline conditions. To gain a more

373

fundamental understanding of how microbial community shifts affected the desired functional

374

potential of the microbial community, we extracted functionally annotated genes and assigned

375

them to microbial taxa. Taxonomic assignment of functional metagenomic subsets largely

376

mirrored the shifts observed via 16S rRNA microbial community profiling, suggesting that

water.

Moderate halophiles

from


45

and may provide

42

and likely

the Alphaproteobacteria

and


Page 18 of 29

377

observed shifts were to maximize functional potential. Specifically, in Sample A1/2 the taxa

378

associated with carbon metabolism demonstrated an increased abundance of Halomonas,

379

Marinilabilia and Marinobacterium (Table S6). Strains of Halomonas are able to grow at salt

380

concentrations of 3-15% (w/v) and utilize D-galactose and D-mannose.50 Moreover, certain

381

strains of Marinilabilia are known to have α- and β-glucosidase and α-galactosidase,51,

382

some species of Marinobacterium can use D-galactose as a sole carbon source.53 In Sample B, a

383

higher abundance of the genera Vibrio and Flavobacterium was observed in subsystems

384

associated with carbohydrate metabolism. Certain strains of Vibrio are known to produce

385

extracellular or capsular polysaccharide consisting several types of sugars including D-galactose

386

and D-mannose, suggesting related metabolisms.54-57 Several recently isolated species of

387

Flavobacterium can tolerate salt concentrations up to 5% (w/v NaCl) and produce acid from D-

388

mannose,

389

Rhodobacteraceae and Hyphomonadaceae, were observed to be significantly more abundant in

390

the osmotic and oxidative stress response categories in the less saline Sample B, suggesting that

391

the observed increase in abundance of Alphaproteobacteria was related to the decreased salinity.

392

Taxonomic assignment of functionally annotated metagenomes demonstrated that observed

393

microbial community shifts were in response to the composition of the samples being treated.

394

Mats treating both Sample B and Sample B1/2 became more diverse in the number of operational

395

taxonomic units (OTUs) that could be detected with each successive treatment cycle. Most of

396

these newly detected taxa were present at low relative abundance and are not known halophiles.

397

These emergent species were likely present at low relative abundance from initial cultivation,

398

and were able to become a larger percentage of the mat microbial community with the reduction

399

in salinity; however, as the produced water was not sterilized prior to use, the potential for

D-galactose

and

D-fructose.58

Classes

of

Alphaproteobacteria,


52

and

including

Page 19 of 29


400

growth of native microorganisms cannot be excluded. Mat microbial communities demonstrated

401

compositional differences by the end of the first loading, suggesting rapid adaptation to new

402

environments, such as produced waters with changing chemical and biological characteristics.

403

Additionally, metagenome data evidenced the potential of mat microbial communities to adapt to

404

different substrates, as well as different salt and metal concentrations through various stress

405

response mechanisms. Produced water characteristics vary both between and within

406

unconventional oil and gas plays.10,

407

microbial mats for the treatment of produced water of various composition and TDS

408

concentrations by maintaining a population of diverse microbes that can grow to suit a particular

409

environment. Additional work is necessary to identify the role of cultivation conditions on mat

410

performance, and optimal cultivation conditions.

411

In the current study, the biodegradation performance of engineered microbial mats was

412

investigated in both synthetic and real produced water, with acetate and guar gum utilized to

413

simulate simple and complex substrates. Our experiments demonstrate that engineered microbial

414

mats are capable of degrading COD in a broad range of salt concentrations. Furthermore,

415

microbial ecology analyses demonstrate that engineered microbial mats are capable of adapting

416

to different produced waters to effectively degrade COD. Results suggest the potential

417

applicability of microbial mats for produced water treatment within a wide range of salinity

418

concentrations, and rates were found to be zero-order. While further work is necessary to

419

understand the scale-up requirements of microbial mats for produced water treatment, microbial

420

mats represent an emerging biological treatment technology to encourage produced water reuse,

421

improve the performance of physical-chemical treatment approaches, remove organic

422

constituents, and reduce biocide application. Biological produced water treatment approaches,

12

These results demonstrate the potential for utilizing



423

including microbial mats, have the potential to decrease the operational costs and improve the

424

efficiency of treating produced water from hydraulic fracturing. Ultimately, improved produced

425

water treatment will serve to address a significant source of public and regulatory concern

426

surrounding the environmental impacts of the hydraulic fracturing process.

427

428

ACKKNOWLEDGEMENTS:

429

This work was funded by a United States National Science Foundation EAGER award

430

(1353858) to KB.

431

432

Supporting Information Available:

433

This information is available free of charge via the Internet at http://pubs.acs.org/.

434

435

REFERENCES:

436 437 438 439 440 441 442 443 444 445 446 447

1.

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38. McCarty, P. L., Anaerobic waste treatment fundamentals. Public works 1964, 95, (9), 107-112. 39. Uygur, A.; Kargı, F., Salt inhibition on biological nutrient removal from saline wastewater in a sequencing batch reactor. Enzyme and Microbial Technology 2004, 34, (3), 313318. 40. Kargi, F.; Dincer, A. R., Biological treatment of saline wastewater by fed batch operation. Journal of Chemical Technology and Biotechnology 1997, 69, (2), 167-172. 41. Ivanova, E. P.; Romanenko, L. A.; Chun, J.; Matte, M. H.; Matte, G. R.; Mikhailov, V. V.; Svetashev, V. I.; Huq, A.; Maugel, T.; Colwell, R. R., Idiomarina gen. nov., comprising novel indigenous deep-sea bacteria from the Pacific Ocean, including descriptions of two species, Idiomarina abyssalis sp. nov. and Idiomarina zobellii sp. nov. International journal of systematic and evolutionary microbiology 2000, 50, (2), 901-907. 42. Martínez-Cánovas, M. J.; Béjar, V.; Martínez-Checa, F.; Páez, R.; Quesada, E., Idiomarina fontislapidosi sp. nov. and Idiomarina ramblicola sp. nov., isolated from inland hypersaline habitats in Spain. International journal of systematic and evolutionary microbiology 2004, 54, (5), 1793-1797. 43. Choi, D. H.; Cho, B. C., Idiomarina seosinensis sp. nov., isolated from hypersaline water of a solar saltern in Korea. International journal of systematic and evolutionary microbiology 2005, 55, (1), 379-383. 44. Hou, S.; Saw, J. H.; Lee, K. S.; Freitas, T. A.; Belisle, C.; Kawarabayasi, Y.; Donachie, S. P.; Pikina, A.; Galperin, M. Y.; Koonin, E. V., Genome sequence of the deep-sea γproteobacterium Idiomarina loihiensis reveals amino acid fermentation as a source of carbon and energy. Proceedings of the National Academy of Sciences 2004, 101, (52), 18036-18041. 45. Donachie, S. P.; Hou, S.; Gregory, T. S.; Malahoff, A.; Alam, M., Idiomarina loihiensis sp. nov., a halophilic γ-Proteobacterium from the Lō ‘ihi submarine volcano, Hawai ‘i. International journal of systematic and evolutionary microbiology 2003, 53, (6), 1873-1879. 46. Arulazhagan, P.; Vasudevan, N., Role of a moderately halophilic bacterial consortium in the biodegradation of polyaromatic hydrocarbons. Marine pollution bulletin 2009, 58, (2), 256262. 47. Osaka, T.; Shirotani, K.; Yoshie, S.; Tsuneda, S., Effects of carbon source on denitrification efficiency and microbial community structure in a saline wastewater treatment process. Water research 2008, 42, (14), 3709-3718. 48. Murali Mohan, A.; Hartsock, A.; Bibby, K. J.; Hammack, R. W.; Vidic, R. D.; Gregory, K. B., Microbial community changes in hydraulic fracturing fluids and produced water from shale gas extraction. Environmental science & technology 2013, 47, (22), 13141-13150. 49. Cluff, M. A.; Hartsock, A.; MacRae, J. D.; Carter, K.; Mouser, P. J., Temporal changes in microbial ecology and geochemistry in produced water from hydraulically fractured Marcellus Shale gas wells. Environmental science & technology 2014, 48, (11), 6508-6517. 50. Mata, J. A.; Martínez-Cánovas, J.; Quesada, E.; Béjar, V., A detailed phenotypic characterisation of the type strains of Halomonas species. Systematic and Applied Microbiology 2002, 25, (3), 360-375. 51. Kumar, S.; Subramanian, S.; Raghava, G. P. S.; Pinnaka, A. K., Genome sequence of the marine bacterium Marinilabilia salmonicolor JCM 21150T. Journal of bacteriology 2012, 194, (14), 3746-3746.



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52. Shalley, S.; Kumar, S. P.; Srinivas, T. N. R.; Suresh, K.; Kumar, P. A., Marinilabilia nitratireducens sp. nov., a lipolytic bacterium of the family Marinilabiliaceae isolated from marine solar saltern. Antonie van Leeuwenhoek 2013, 103, (3), 519-525. 53. Kim, Y.-G.; Jin, Y.-A.; Hwang, C. Y.; Cho, B. C., Marinobacterium rhizophilum sp. nov., isolated from the rhizosphere of the coastal tidal-flat plant Suaeda japonica. International journal of systematic and evolutionary microbiology 2008, 58, (1), 164-167. 54. Bramhachari, P.; Dubey, S., Isolation and characterization of exopolysaccharide produced by Vibrio harveyi strain VB23. Letters in applied microbiology 2006, 43, (5), 571-577. 55. Wai, S. N.; Mizunoe, Y.; Takade, A.; Kawabata, S.-I.; Yoshida, S.-I., Vibrio cholerae O1 strain TSI-4 produces the exopolysaccharide materials that determine colony morphology, stress resistance, and biofilm formation. Applied and environmental microbiology 1998, 64, (10), 36483655. 56. Enos-Berlage, J. L.; McCarter, L. L., Relation of capsular polysaccharide production and colonial cell organization to colony morphology in Vibrio parahaemolyticus. Journal of bacteriology 2000, 182, (19), 5513-5520. 57. Thompson, F. L.; Iida, T.; Swings, J., Biodiversity of vibrios. Microbiology and molecular biology reviews 2004, 68, (3), 403-431. 58. McCammon, S. A.; Bowman, J. P., Taxonomy of Antarctic Flavobacterium species: description of Flavobacterium gillisiae sp. nov., Flavobacterium tegetincola sp. nov., and Flavobacterium xanthum sp. nov., nom. rev. and reclassification of [Flavobacterium] salegens as Salegentibacter salegens gen. nov., comb. nov. International journal of systematic and evolutionary microbiology 2000, 50, (3), 1055-1063.

605

606

607 608 609 610 611 612 613


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614


Table 1. Chemical characteristics of Synthetic Produced Water, Sample A, and Sample B Constituent

615

Sample A

Sample B

Na (mg/L) 47,107.2 5,272 Ca (mg/L) 16,509.14 1,691 Mg (mg/L) 1,820.2 193 Ba (mg/L) 328.26 14.61 Sr (mg/L) 1,888.39 1,051.3 Fe (mg/L) 19 4.22 Cl (mg/L) 115,277.6 13,867 COD (mg/L) 1,865 440 TDS (mg/L) 182,702 18,400 pH 5.89 7.35 Days after Fracture 20 N/A a : Example synthetic media given for 50,000 mg/L TDS condition.

616 617

618 619

TOC Art


Synthetic Produced Watera 17,500 5,000 0 0 0 0 27,500 2,500 50,000 6.66 N/A


620


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621 622 623 624 625


Figure 1. Microbial mat treatment of various substrates. A. Acetate removed in synthetic produced water as a function of time; B. COD (guar gum) removed in synthetic produced water as a function of time; C. COD (guar gum) removed in real produced water as a function of time. Each 72-hour loading is denoted by a dark vertical line. Data shown is the average of biological triplicates for each conditions. Error bars represent +/- 1 standard deviation.

626 627 628 629 630 631 632 633 634

Figure 2. Microbial mat substrate removal rates in synthetic and real produced water. Data shown is the average of biological triplicates for each conditions. Error bars represent +/- 1 standard deviation.



Page 28 of 29

635 636 637 638 639

Figure 3. Normalized plot of average class abundance for triplicate (microbial community) or duplicate (metagenome) microbial mats treating guar gum amended produced water. ‘Other’ indicates sequences that could not be annotated to the class level or of classes that comprised less than 2% of relative abundance.

640


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641 642 643 644

Figure 4. Taxonomic assignment to metagenomic data from microbial mats associated with carbon metabolism and stress response pathways. ‘Other’ indicates classes below 2% abundance in all metabolisms in averaged samples.

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Microbial Mats as a Biological Treatment Approach for Saline

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