The effects of biocide use on the microbiology and geochemistry of produced water in the Eagle Ford formation, Texas, U.S.A.

Journal of Petroleum Science and Engineering 135 (2015) 1–9

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The effects of biocide use on the microbiology and geochemistry of produced water in the Eagle Ford formation, Texas, U.S.A. Eugenio-Felipe U. Santillan a,n, Wanjoo Choi a, Philip C. Bennett b, Juliette Diouma Leyris a a Statoil—Production and Water Management, Research Development and Innovation, 10111 Richmond Avenue, Westchase Corporate Center, Houston, TX 77042, USA b The University of Texas at Austin, Department of Geological Sciences, Jackson School of Geosciences, 1 University Station C1100, Austin, TX 78712, USA

art ic l e i nf o

a b s t r a c t

Article history: Received 17 April 2015 Received in revised form 29 July 2015 Accepted 30 July 2015 Available online 11 August 2015

A side effect of hydraulic fracturing (HF) is the inoculation of the subsurface with surficial microorganisms, many of which could affect production and the chemistry of produced water (PW). There is little knowledge of what organisms are selected for once brought to the subsurface. The current practice to control microbial growth is to include a biocide in the HF fluids. However, biocides add a significant cost to these fluids and has potential consequences to health, safety, and the environment. In this study, we evaluate the microbial community composition of PW and freshwater in the Eagle Ford Shale (EF) to determine what organisms survive in production wells during the HF process. We also assess the effectiveness and necessity of the tri-n-butyl tetradecyl phosphonium chloride (TTPC) biocide for controlling microbial growth in these wells. At greater than 4 km depth, temperatures in EF are 4160 °C conditions that do not support viable microbial communities, but at shallower depths, microbial communities may be viable. Freshwater and PW samples were collected and cultured in representative subsurface conditions at 128 °C and 60 °C, in the presence or absence of biocide and measured for their composition and viability using molecular methods and microscopy. Results indicate the presence of a diverse microbial community and these communities contain populations of microorganisms that may have the potential to affect production such as through hydrocarbon degradation and bioclogging. Furthermore, laboratory experiments simulating EF downhole conditions show that cells lose viability due to high temperatures, regardless of biocide. At lower temperatures, 4 50% of viable cells remain in cultures even with biocide present. Results imply that at depth, introduced cells are sterilized due to temperature while at shallower depths, TTPC may not be completely effective in biomass control, prompting the need to further explore options for microbial control. & 2015 Elsevier B.V. All rights reserved.

Keywords: Hydraulic fracturing Eagle Ford formation Produced water Tri-n-butyl tetradecyl phosphonium chloride

1. Introduction During hydraulic fracturing (HF), surficial water is pumped into a nearly impermeable shale reservoir under pressure in order to create fractures. The newly created permeability allows for easier extraction of oil and gas. The water used for HF is composed largely of freshwater from impoundment ponds at the site of HF, as well as a complex mixture of other compounds including proppant, corrosion inhibitors, acids, and biocides all designed to optimize the process. Because of HF, microbial communities living in shale reservoirs, which will affect reservoir permeability and geochemistry, will n Corresponding author. Present address: 647 Contees Wharf Rd., Edgewater, MD 21037, USA. E-mail addresses: [email protected] (E.-F. Santillan), [email protected] (W. Choi), [email protected] (P.C. Bennett), [email protected] (J. Diouma Leyris).

http://dx.doi.org/10.1016/j.petrol.2015.07.028 0920-4105/& 2015 Elsevier B.V. All rights reserved.

undoubtedly change. The composition of the altered community will be a combination of microorganisms native to the subsurface as well as surficial microorganisms that are introduced when the freshwater from impoundment ponds is pumped down wells (Mohan et al., 2013; Struchtemeyer and Elshahed, 2012; Struchtemeyer et al., 2012; Wuchter et al., 2013). These surficial organisms are likely dominated by aerobic, heterotrophic, or photosynthetic microorganisms due to their exposure to sunlight as well as O2 from the atmosphere. Once introduced to the subsurface, their presence and distribution change to reflect the anaerobic, saline, and thermophilic conditions of the reservoir (Mohan et al., 2013; Struchtemeyer et al., 2011). Prior to HF, microbial growth is limited due to the low-permeability of shales, preventing the movement of nutrients, carbon sources, and electron acceptors to cells and providing little space for cells to grow and divide (Fredrickson et al., 1997). However, with the creation of fractures also comes the release of carbon sources and nutrients and the exposure of redox gradients, which can favor microbial growth,

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especially those among organisms capable of surviving the stresses of the subsurface environment. Enrichment of organisms in wells can result in both negative and positive consequences to production and produced water (PW) chemistry as microbial metabolic activities often catalyze many kinetically inhibited geochemical reactions. Sulfate reducers, produce sulfide, a component that accelerates the corrosion of pipes (Mohan et al., 2013; Struchtemeyer et al., 2011, 2012; Struchtemeyer and Elshahed, 2012; Wuchter et al., 2013). Furthermore, sulfate reducers can produce significant amounts of sulfide, resulting in reservoir souring as well as safety concerns. Microbial activities in production wells may also have positive effects by enhancing production. Organisms, for example, can produce various surfactants, helping with the emulsification and extraction of petroleum in reservoirs (e.g., Almeida et al., 2004; Banat, 1995; Belcher et al., 2012; Darvishi et al., 2011; Ghojavand et al., 2008; Khire and Khan, 1994; Merchant and Banat, 2012). Furthermore, through their various metabolic activities, microorganisms may also produce CO2 that causes oil swelling. They can also form biofilms to alter subsurface wettability and pressure and release acids to dissolve rocks and increase permeability (Sen, 2008). In addition to effects on production, microbial activities may impact the composition of produced water (PW), which can be an important consideration during its disposal. PW is a combination of the original formation water, HF fluids, natural gas, and hydrocarbons. PW, which is more saline than the original fluids used to perform HF, presents a challenge for disposal because of its salinity, radioactivity, and presence of toxic components (Gregory et al., 2011). Enhanced microbial activities within wells can further influence the composition of PW by the addition of metals through enhanced rock dissolution, through the addition of gases like sulfide, carbon dioxide, and methane due to metabolism, through the degradation of components in HF fluids, and through the addition of further hydrocarbons due to enhanced oil degradation at depth. The current practice in HF is to control microbial growth by adding biocides such as tri-n-butyl tetradecyl phosphonium chloride (TTPC) in the HF fluid mixture to prevent both corrosion and souring (Moore and Cripps, 2012). The composition of these biocides can vary, and are often proprietary, though they are generally glutaraldehyde-based, cocodiamine-based, or phosphonium-based (Kramer et al., 2008). Kill studies have shown that many of these biocides are effective at eliminating 4 99% of bacteria in both planktonic and biofilm cultures (Kramer et al., 2008). However, bacterial enumeration tests are culture-based, relying heavily on the response of culturable microbes in known growth media. Given that 99% of microorganisms are unculturable especially in natural environments, these kill tests may not be completely representative (e.g., Madigan, 2012). TTPC is one particular biocide in use, chosen for multiple reasons including its solubility in water, its low volatility, and its photo-stability. However, the biocide is highly toxic to higher organisms, and does present safety concerns to humans as well as aquatic organisms. While biocide use is well documented in conventional operations (e.g., Struchtemeyer et al., 2012) little is known about its effect on organisms living in the shale reservoirs. Furthermore, biocide necessity may vary by well and by location. Within the Eagle Ford Shale (EF), biocide may not be necessary as downhole temperatures are in excess of 160 °C, hotter than the known thermal limits of life (e.g., Clarke, 2014). However, within the Marcellus Shale, downhole temperatures are near 60 °C suggesting microorganisms could have a significant effect on production at depth. In addition to cost, biocide use has important implications towards health, safety, and the environment, prompting interest in

alternative methods for microbial control due to its toxicity. The objective of this project was twofold: (1) to assess what organisms are present in wells at a representative HF site and their potential effects on production and PW chemistry and (2) to determine how effective biocide use is on controlling biomass using microscopy, a method that is not typically used by service companies to validate biocide success.

2. Field site Impoundment pond or PW samples were obtained from locations from Runge, Texas at the Eagle Ford Shale. Water from these impoundment ponds were sourced from the local Carrizo-Wilcox aquifer as well as surface runoff from precipitation. Two impoundment ponds (IP1 and IP2, respectively) were sampled. IP1 was located approximately 4 km from the well while IP2 was located approximately 100 m away. The sampled well (PW1) is a well that is drilled to a depth of approximately 4.1 km. Downhole temperatures are at minimum, 160 °C with pressures at approximately 9000 to 11,500 psi.

3. Materials and methods Impoundment and PW samples were collected to characterize the organisms present during HF operations. Culture work was used to determine the factors that influence microbial community composition in the subsurface. 3.1. Field sampling Equipment used for microbiological analysis was autoclaved at 122 °C and 15 psi for 30 min prior to fieldwork. When steam sterilization was unavailable, equipment was sterilized using a 10% sodium hypochlorite solution followed by a rinse with a 10% sterile sodium thiosulfate as a neutralizer. Bottles for cation analysis were acid washed prior to use. Two sets of filters were collected from field sites: one set of filters was used exclusively for DNA extraction to determine subsurface microbial community composition. A second set of filters was of impoundment water, and was used for culture experiments and to determine freshwater microbial community composition. Between 7 to 8 L of water was collected from the impoundment ponds into a sterile collection vessel. Water from this vessel was immediately filtered in the field using a peristaltic pump through a 0.2 mm filter. Unfiltered water from the ponds was used to measure temperature, pH, and conductivity in the field using an Ultrameter (Myron L Company, Carlsbad, CA). Measurements of ferrous and total iron and sulfide were also performed on filtrate using the phenanthroline and methylene blue methods, respectively (CHEMetrics, Midland, VA). Samples for cation, anion, dissolved inorganic carbon (DIC) and dissolved organic carbon (DOC), and alkalinity were analyzed in the laboratory. Samples for cation analysis were acidified to 5% with trace metal grade HNO3 upon return to the laboratory 24 h later. Filters from freshwater ims pounds were stored in 50 ml tubes (Falcon ) at 4 °C in impoundment unfiltered water for cultivation and DNA extraction. Dust samples collected from the surrounding area were also added to these samples as a portion of the microbial community present at IP1 and IP2 are likely windblown and sourced from dust surrounding the area. Filters of produced water, used exclusively to determine subsurface microbial community composition, were asceptically stored in WhirlPak™ bags and preserved at
70 °C using a liquid nitrogen dewar prior to DNA extraction. Because there was far less

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produced water available for sampling, filtrate was used for geochemical analyses in both the field and in the laboratory. 3.2. Water analysis Cation and trace metal analyses were performed using an Agilent 7500ce Quadrupole inductively coupled plasma mass spectrometer (ICP-MS) at the University of Texas at Austin Jackson School of Geosciences. Anions were analyzed by ion chromatography. Alkalinity was measured through titration using 0.1 N H2SO4 to the total alkalinity endpoint of pH 4.5. Dissolved inorganic carbon and dissolved organic carbon were measured using a carbon analyzer. 3.3. Culture experiments A microbial suspension used for inoculation was created by sonicating impound filters in a water bath sonicator (Fisher Scientific, Pittsburg, PA) along with vigorous vortexing to remove cells attached to filters. Approximately 500 mg of dust samples were added to each suspension as well, in order to simulate the contribution of the surrounding environment (e.g., windblown microorganisms and soil microorganisms) to the IP microbial community. Initial cell counts were performed using bac live/dead stain (Life Technologies, Inc., Carlsbad, CA) to determine total cell counts as well as dead cells. A synthetic PW was created using a water analysis of PW from January 2014 and autoclaved using per liter of distilled water: 0.34 g NaHCO3, 0.048 g MgSO4, 0.0035 g K2HPO4, 3.13 g CaCl2, 0.20 g NH4Cl, and 12.86 g NaCl. Approximately 30 ml of media was dispensed into serum bottles containing 500 mg of sterile drill cuttings, and 200 mL of filter sterilized condensate as a carbon source. For experiments with biocide, media was supplemented with 5 ml TTPC as this biocide is the typical biocide used in the EF (www.fracfocus.org). Reactors were then inoculated using 3 ml of the various inocula. Shale samples for culture experiments were obtained from drill cuttings from the nearby Wessendorf well, from 3 different depths: between 13,210 to 13,220 ft., 13,220 to 13,230 ft., and 13,230 to 13,240 ft. Equal amounts of drill cuttings were ground using a ceramic mortar and pestle. Drill cuttings were dispensed in media bottles and autoclaved prior to use in culture experiments. In addition, abiotic controls were set up containing media and drill cuttings that were not inoculated. These were used as comparisons to determine the effects of both microorganisms and biocide on PW chemistry and to subtract out the interference from suspended particles resulting from the drill cuttings. Cultures were kept in serum bottles capped by butyl stoppers to prevent gas exchange with the atmosphere. Experiments were organized following the setup in Table 1. High temperature Table 1 Experimental conditions for laboratory cultures. Inoculum

Culture condition

Incubation

None

           

24 h

IP1

IP2

128 °C 128 °C þ biocide 60 °C 60 °C þ biocide 128 °C 128 °C þ biocide 60 °C 60 °C þ biocide 128 °C 128 °C þ biocide 60 °C 60 °C þ biocide

2 weeks 24 h 2 weeks 24 h 2 weeks

3

incubations were performed at 128 °C at 15 psi using an autoclave for 24 h. Low temperature cultures (60 °C) were incubated 2 weeks at atmospheric pressure. Live and dead cells were enumerated using previously stated methods. Samples were also analyzed for cations, anions, and alkalinity, as well as community composition. 3.4. Community sequence analysis Prior to DNA extraction, PW filters were soaked in filter-sterilized 1% sodium dodecyl sulfate for approximately 1 h (Parkar et al., 2001; Simoes et al., 2005). Filters were then sonicated as previously described and centrifuged to concentrate biomass. For impoundment pond inocula and cultures, approximately 10 to 25 ml of sample was concentrated through centrifugation. For both PW and impoundment samples, DNA was then extracted using the Mo Bio Power Biofilm DNA extraction kit (Mo Bio Laboratories, Inc., Carlsbad, CA). Sequencing was performed at Molecular Research LP (Shallowater, TX) using 454 sequencing of the 27F primer to detect bacteria and 349F primer to detect archaea. Sequence data were processed using QIIME (Caporaso et al., 2010). Sequences were demultiplexed followed by denovo chimera removal through Uchime. Operational taxonomic units (OTUs) were defined at the 97% level using BLASTn against a curated GreenGenes database. Values for diversity were also calculated using reciprocal Simpson’s index. Briefly, the reciprocal Simpson’s index summarizes the probability that two randomly picked taxa within a community will be the same. Low values in this calculation represent communities consisting of only a few species while higher values indicate diverse communities.

4. Results Table 2 summarizes the water composition of the three different field sites along with field measurements. The table also contains data from previous sampling periods at PW1. Bicarbonate concentrations were estimated at the PW1 September 2013 sampling through charge balance adjustment as these values were not measured. Sampled waters are generally near-neutral pH, Na
Cl type waters. Total dissolved solids is an order of magnitude higher in PW compared to the freshwaters from the IP sites and contains more ferrous iron and sulfide than the impoundment samples suggesting the water is more reducing than the impoundment ponds. PW, especially the July sampling, contains more ammonia, boron, phosphorous, and barium than impoundment ponds, likely a signature of HF fluids. All samples are at or close to saturation with carbonates. The environmental samples also revealed the presence of multiple classes of bacteria. At IP2, most classes were Synechococcophycideae along with Alphaproteobacteria and Betaproteobacteria. Classes at IP1 consisted of Actinobacteria, Alphaproteobacteria, and Betaproteobacteria. The major classes present at PW1 include the Actinobacteria, Gammaproteobacteria, Alphaproteobacteria, Bacilli, and Phycisphaerae, with other classes at 5% relative abundance and less. In particular, the genuses Microbacterium, Mycobacterium, Crenothrix, and Gordonia composed 44% of the organisms present at PW1. Archaea detected at both IP1 and IP2 consisted of the classes Methanomicrobia, Crenarchaeota, and Thaumarchaeota with Methanomicrobia dominating sequences. No Archaea were detected in PW1 (Fig. 1 and Table 3). Based on rarefaction curves showing observed taxa (Fig. 2), all three sites still had unsampled diversity. Values for observed taxa show the number of different OTUs present, with IP1 having the highest number followed by IP2 and then PW1. Values for the reciprocal Simpson’s index show that both IP1 and PW1 contain

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Table 2 Water chemistry for environmental samples and laboratory cultures. Values are in ppm unless otherwise stated. T (°C) pH

PW1 (Sept) PW1 (Jan) PW1 (July) IP1 IP2 Negative Hi T Negative Hi T þBiocide IP1 Inoculum Hi T IP1 Inoculum þ Biocide IP2 Inoculum Hi T IP2 Inoculum Hi T þBiocide Negative Lo T Negative Lo Tþ Biocide IP1 Inoculum Lo T IP1 Inoculum Lo T þBiocide IP2 Inoculum Lo T IP2 Inoculum Lo T þBiocide

21.1 21.6 34.3 30.2 31.4 128 128 128 128 128 128

7.28 7.62 5.79 6.8 7.02 6.74 6.61 6.65 6.51 6.7 6.47

60 60 60 60

6.88 6.49 6.67 6.42

60 60

6.65 6.5

Eh (mV) SC (mS) Fe(II) Fe(tot) Na þ


0.13 0.26

0.34

30,700 28,300 26,630 1060 2046

8.4

9.1

0.12

0.29

Kþ

Ca þ þ

Mg þ þ

NH4 þ

HCO3- Cl-

SO4



29 30 38 20 7 5 7 6 7 5 6

100 0 0 8 10 9 9 8 13

1971 199 226 198 384 100 295 133 196 117 327

9040 70 10,600 42 11,098 17 151 37 298 132 9775 56 13,253 77 8947 91 12,538 84 8770 82 12,084 92

63 0.0 15
2.3 19 0.1
2.7 24 0.0 4.6 23 0.02
0.4 26 0.1
2.0 40 57.0
17.4 4 0.9
1.8 9 48.6
18.5 25 0.7
1.5 25 49.5
17.9

Si

P

% Charge balance error

SICalcite

5420 5160 5194 141 365 5137 4932 4741 4484 4671 4467

124 897 83 1130 55 1293 8 23 16 10 34 853 37 1023 33 791 32 992 33 771 36 924

5106 4965 4710 4320

34 34 34 30

862 989 815 887

5 6 8 7

9 10 8 9

303 331 278 338

9733 12,619 8911 11,463

42 73 99 67

9 0.2 10 37.2 63 0.8 15 38.4

-2.5
15.2
2.1
17.1

0.74 0.43 0.48 0.34

4724 33 4563 33

799 939

6 6

8 10

260 320

8856 12,401

90 85

8 0.6 10 37.5


1.7
18.1

0.43 0.41

more rare OTUs in comparison to IP2, which is composed largely of the Synechococcophycideae (Table 4). Initial cell counts for the IP2 inoculum were on the order of 106 cells/ml while cells from the IP1 inoculum were on the order of 108 cells/ml. Cell counts for IP2 cultures increased approximately 2 orders of magnitude during cultivation, though there was a lot of variability in cell counts due to interference from shale particles during imaging. This was corrected for by subtracting particles counted from negative control cultures from counts in inoculated cultures. Cell counts for IP1 cultures stayed within the magnitude of 108 cells/ml during cultivation. After incubation at 128 °C, total cell counts and dead cell counts appear the same for both inocula regardless of biocide presence (Fig. 3A).

1.46 0.92
0.64
0.85
0.70 0.72 1.16 0.78 0.89 0.76 1.05

Cell counts for 60 °C cultures ranged between 107 and 10 cells/ml, with variability in counts owing to the interference of shale particles. However, in most cultures, there are more viable cells present than dead cells, with the exception of IP1 cultures without biocide. The percentage of dead cells present for IP2 cultures with biocide is 52% and 53% for IP1 culture with biocide (Fig. 3B). Fig. 1 also lists the organisms present in the 60 °C cultures on the class level. The composition of bacteria in non-biocide cultures differed from those of biocide-containing cultures. Non-biocide cultures showed more diversity and evenness in organismal distribution than biocide cultures, with multiple classes represented. Laboratory cultures and PW1 showed a few taxa in common, 8

Fig. 1. Microbial community composition of field locations sampled on the class level.

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5

Table 3 Bacteria present at PW1 on the class level, and species levels and descriptions of related organisms. Class

Relative abundance (%)

Genus and relative abundance (%)

Putative description

Actinobacteria

35.68

Microbacterium (14%) – Microbacterium oleivorans (4%) – Microbacterium hydrocarbonoxydans ( 40.1%) Mycobacterium (12%) – Mycobacterium petroleophilum (6%) Gordonia (6%) – Gordonia alkanivorans ( 41%) Thermobifida ( 40.1%) – Thermobfida fusca ( 4 1%) Crenothrix (12%) Nitrosococcus ( 4 0.1%) Methylobacterium (1%) Bradyrhizobium ( 41%) Thiobacillus ( 41%) Methylocystis ( 40.1%) Xanthobacter ( 40.1%) Bacillus (5%) – Bacillus subtilis ( 40.1%)

Hydrocarbon degradation (Schippers et al., 2005a)

Gammaproteobacteria

13.73

Alphaproteobacteria

11.94

Bacilli

7.04

Phycisphaerae Synechococcophycideae Betaproteobacteria Clostridia

5.07 3.98 3.80 3.48

Actinobacteria

3.20

Sphingobacteria

3.00

Planctomycetacia Nitrospira Other

2.29 0.11 6.71

Leptothrix Clostridium (2%) Desulfosporosinus ( 40.1%) Nocardioides (2%) – Nocardioides oleivorans (0.2%) Lewinella (1%) Flexibacter ( 41%) Acidimicrobiales ( 4 1%) Planctomyces (1%) Thermodesulfovibrio (4 1%)

Hydrocarbon degradation (Iizuka et al., 1975)

Hydrocarbon degradation (Arenskotter et al., 2004) Bio-polymer degrader, thermophile (McGrath and Wilson, 2006) Iron oxidizer (Walter, 1997) Ammonia oxidizer (Klotz et al., 2006) C1 compound consumer, methanotroph (Peyraud et al., 2012) Nitrogen fixer (Kulakov et al., 2002) Sulfur oxidizer, acid producer, acidophile (Robertson and Kuenen, 2006) Methanotroph (Krentz et al., 2010) Alcohol consumer (Rong et al., 2009) Surfactant production (Gudina et al., 2013)

Phototrophs (Waterbury et al., 1986) Iron oxidizer (Corstjens et al., 1992) Fermenter, gas producer, degrades complex carbon compounds (Mainguet and Liao, 2010) Sulfate reducer (Mayeux et al., 2013) Hydrocarbon degrader (Schippers et al., 2005b) Degrades complex carbon compounds (Khan et al., 2007) Denitrifying bacteria (Wu and Knowles, 1995) Acidophile (Clum et al., 2009) Halophiles (Fuerst et al., 1997) Sulfate reducer, thermophile (Sekiguchi et al., 2008) Cytophagia, Thermoleophilia, Deltaproteobacteria, Acidimicrobia, Planctomycea, Nostocophycideae, Armatimonadia, Flavobacteria, tm7 (candidate division), Epsilonproteobacteria, Verrucomicrobiae, Caldilineae, Oscillatoriophycideae, Erysipelotrichi, Rubrobacteria, Gloeobacterophycideae, Chloroflexi, Spartobacteria

Table 4 Alpha diversity values for microbial communities from EF field samples.

IP1 IP2 PW1

Fig. 2. Rarefaction curve of observed phylotypes present at the three sampling locations.

though in different relative abundances. However, the samples also differed markedly in the presence and absence of specific organisms as well as their relative abundances. Non-biocide

Observed taxa

Reciprocal simpsons index

% Similarity to PW1

1926 1187 1063

82.283 5.152 39.726

13 17 100

cultures were dominated by Prochlorococcus, and Massilia. Biocide containing cultures showed high abundances of Serratia, Acinetobacter, and Agrobacterium and each biocide culture had far fewer organisms present in comparison to non-biocide cultures. Most of the organisms detected in laboratory cultures have been identified with the biodegradation of petroleum products and are listed in Table 5. Water analyses of samples containing TTPC show higher Clconcentrations than non-biocide cultures because TTPC is composed of a phosphonium cation and a chloride counterion. As a result, the analysis is unbalanced for biocide containing cultures as the analysis does not account for the phosphonium cation (Table 2). There is no clear difference in major ion concentrations between biotic and abiotic cultures at both 128 and 60 °C. However, cultures containing biocide had higher concentrations of metals such as Mn, Cu, Co, Zn, and As relative to cultures that contained no biocide regardless of the presence of cells. Phosphate concentrations in cultures containing biocide were two orders of magnitude higher than those without biocide. For 60 °C cultures, these metals also show elevated concentrations in cultures

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Fig. 4. Metal concentrations in PW1 as well as laboratory cultures with and without biocide.

containing biocide in comparison to those that do not, though not as pronounced as with 128 °C cultures (Fig. 4).

5. Discussion

Fig. 3. Total and dead cell counts for cultures at (A) 128 °C and (B) 60 °C.

Approximately 17% of phylotypes present at IP2 and 12% from IP1 are present at PW1 (Table 4). These include the classes Actinobacteria, Betaproteobacteria, and Gammaproteobacteria (Fig. 1). Organisms not shared between PW1 and the impoundment ponds could be a mixture of organisms from locations that were not sampled, e.g., drilling muds or pipelines, or organisms native to the subsurface. While shared organisms are present between the impoundment ponds and PW1, their distribution at PW1 differs, showing that the environment at PW1, which is likely hot, saline, anaerobic, and rich in hydrocarbons, will change organismal distribution. For example, organisms at IP2 are dominated by cyanobacteria belonging to the Synechococcophydiceae. At PW1, these organisms are also present but at a much lower abundance, likely because population size drastically decreased following the removal of light from the environment as well as the introduction of warm, saline conditions reaching 160 °C. A surprisingly diverse microbial community exists at PW1 despite the stresses presented by the subsurface (Table 4 and Fig. 1). We had initially hypothesized that PW1 would be a low diversity environment due to the high downhole temperatures at EF as well as the toxicity of HF fluids. However, the diversity at PW1 suggests

Table 5 Petroleum degrading bacteria on the genus level found in laboratory cultures and their relative abundances. Petroleum degrading bacteria found in laboratory cultures IP1 Lab Cultures

IP2 Cultures

IP1 Lab Culturesþ Biocide

IP2 Lab Culturesþ Biocide

Arthrobacter (15%) Porphyrobacter (14%) Roseomonas (5%) Hyphomonas (2%) Rubrobacter (2%) Novosphingobium (2%) Porphyrobacter (14%)

Brachymonas (10%) Porphyrobacter (9%) Nocardioides (6%) Serratia (4%) Pseudomonas (3%) Brevundimonas (1%) Roseomonas ( 41%)

Acinetobacter (40%) Nocardioides (13%) Pseudomonas (10%) Serratia (7%) Sphingobium (5%)

Serratia (52%) Sphingomonas (18%) Novosphingobium (12%) Sphingobium (5%)

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an environment that can accommodate multiple microbial niches all contributing to subsurface geochemistry. Many of these potential niches are presented in Table 3. Many of the organisms found in PW1 are related to bacteria isolated from oil wells or oil spills and have been shown to degrade crude oil and other hydrocarbons. Because downhole conditions in the EF are too hot to support viable bacteria at 4.1 km, the organisms detected are likely residing along the well bore. These organisms, which are putatively thriving on hydrocarbons, may have an effect by decreasing the amount of extracted product. In contrast, some organisms identified in this study have previously been shown to benefit production. Bacillus subtilis, for example, is used in microbially enhanced oil recovery (MEOR) because it is capable of producing surfactants to emulsify oil and make it more recoverable (Gudina et al., 2013; Lazar et al., 2007). While surfactant production at 4.1 km depth may not be possible within the EF, it may be possible in other reservoirs where temperatures at depth are cooler than in the EF (e.g., 4160 °C). In this case, including additives to HF fluids to select for these organisms may aid increasing production. Other organisms identified may potentially be taking advantage of components in HF fluids and breaking them down (Table 3). These have implications towards the processes that will eventually remove these compounds from the environment. Organisms such as Thermobifida, and Xanthobacter, for example, catalyze the breakdown of complex carbon compounds such as gels, friction reducers, and biocide within the HF fluids, which would otherwise remain in the subsurface for some time (McGrath and Wilson, 2006; Rong et al., 2009). Other organisms, such as Methylobacterium can consume C1 compounds, and may be degrading alcohols added (Peyraud et al., 2012). The high ammonium present in the HF fluids may be selecting for organisms that can cycle these compounds such as Flexibacter and Nitrosococcus (Wu and Knowles, 1995). Over time, stimulation of many of these organisms could aid in well clean up after the HF process has been completed. There is also evidence that iron cycling is occurring, as with the detection of Crenothrix and Leptothrix, organisms that can oxidize ferric iron to ferrous iron. This makes sense as reduced iron was measured at PW1 (Table 2). These organisms have been previously documented in water quality studies and their presence at PW1 could result in bioclogging, especially if the well is located in an iron-rich area (Walter, 1997). Even though PW1 has been treated with TTPC, these organisms are still present. Some sulfur cycling may also be implied at PW1. Organisms like Desulfosporosinus and Thermodesulfovibrio may be taking advantage of the abundant sulfate in PW (Mayeux et al., 2013). Many wells within the EF are known to have the corrosive H2S, and PW1 is one well where there was some measured (Table 2). In addition, TTPC has been shown to be effective against sulfate reducers and the lack of sulfate reducers present at PW1 may also be a reflection of the efficacy of TTPC against these organisms (Struchtemeyer et al., 2012). Other sulfur metabolizing organisms like Thiobacillus are probably taking advantage of the H2S, oxidizing it, and producing sulfuric acid in the process (Robertson and Kuenen, 2006) further contributing to corrosion (Table 3). While other shale plays have shown that methanogenesis can be important for gas production (e.g., Mohan et al., 2013; Wuchter et al., 2013), this is likely not the case within the EF. Archaea, especially methanogens, were detected in both impoundment ponds. Interestingly, none of these archaea are present at PW1 suggesting that the archaea introduced into PW1 do not survive, are outcompeted, or that archaea are in such low abundance that they are not detectable. Our image-based assessments, while showing a great deal of uncertainty due to interference from shale particles, provide

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implications towards the effectiveness of TTPC. The thermal limits of life are generally considered to be at 122 °C (e.g., Clarke, 2014) and the near or complete cell death that occurs in high temperature cultures reflects this (Fig. 3). This also implies that organisms introduced into EF are likely killed downhole due to heat sterilization, suggesting biocide may not be necessary below 4.1 km. Furthermore, these results suggest that the organisms detected at PW1 are likely to come from shallower in the subsurface. Our data from the live/dead stains on 60 °C incubations also suggests a significant number of surviving cells for cultures exposed to biocide, a contradiction to previous culture-based assessments (Kramer et al.). Previous biocide studies have deemed biocides to be effective after only 1 to 2 days of bacterial growth monitoring or have been used with representative monocultures rather than environmental samples (Kramer et al., 2008). While the initial stress of TTPC may be enough to eliminate the majority of cells within cultures over short time scales, the two week incubation period in our experiments would have allowed for surviving cells to reproduce. Those cells that are enriched will be biocide resistant and will have fewer organisms to compete for resources. Abiotic controls were set up as a means to determine the differences inoculated cultures have to PW chemistry. In both temperature incubations, we observed few differences between the biotic and abiotic cultures suggesting surviving cells did not have a major effect on rock dissolution for the time scales of the experiment. Furthermore, no sulfur smell was observed in the cultures indicating sulfate reduction may not be a major metabolism among cells in biotic cultures. However, the amount of ferrous iron and dissolved organic carbon were not measured and organisms present in biotic cultures may be metabolizing either through dissimilatory iron reduction or through fermentation. Furthermore, the incubation times used may not be enough to allow for measurable changes to be observed. Laboratory cultures and PW1 show common organisms, though in different relative abundances. Some of the dominant genuses, such as Serratia, were not detected in the PW. Furthermore, all laboratory cultures had fewer organisms present than PW1 and were dominated by specific bacteria. For non-biocide cultures, these genuses included Prochlorococcus, and Massilia. Biocide containing cultures showed high abundances of Serratia, Acinetobacter, and Agrobacterium and each biocide culture had far fewer organisms present in comparison to non-biocide cultures (Table 5 and Fig. 3). Most of the organisms found in both sets of cultures have been associated with the biodegradation of petroleum products, likely because the growth media contained both condensate and organic-rich drill cuttings, selecting for these organisms (e.g., Fida et al., 2013; Gu et al., 2013; Kappell et al., 2014; Kim and Crowley, 2007; Lal and Khanna, 1996; Mansur et al., 2014; Thomas et al., 2013). The differences in diversity between biocide and non-biocide cultures shows that biocide is a stress to microbial communities. The few organisms present in biocide-containing cultures are represented by ubiquitous genuses that are highly adaptable to multiple environments, such as Serratia and Acinetobacter. These organisms are not present at PW1, showing that our cultures only reflect biocide-resistant bacteria in surficial conditions and not at 4.1 km depth. This also further highlights the fact that the microbial community present at PW1 is reflected by other aspects beyond just the surficial microbial community. The organismal distribution in laboratory cultures differs from that of PW and appears more similar to those of the original inocula, especially for the non-biocide cultures (Fig. 3). This shows that while we tried to capture many of the conditions of the subsurface through the water chemistry, mineralogy, carbon sources, and HF components, there were other components in

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PW1 that were not considered such as temperatures and environments, other carbon sources, pressure, or electron acceptors. The community measured at PW1 is also an aggregate of organisms along the entire wellbore, spanning multiple temperatures and environments. Furthermore, the organisms present in laboratory cultures are representative of surficial and soil organisms, although PW1 contains other organisms not present in soils that are likely native to the subsurface and introduced into the subsurface through other methods. The amount of biocide used for culturing was approximately 5%, well above the amounts used in HF fluid, which is at 0.0037% (www.fracfocus.org). The commercial concentration of TTPC is at approximately 5–10% solution, corresponding to a minimum concentration of 30 ppm. In solution, this suggests an active ingredient corresponding between 1.5 to 3 ppm. Our cultures utilize several orders of magnitude more biocide, at approximately 5000 ppm. Even at this high concentration, over half of the total biomass survives, showing that high doses of TTPC do not necessarily correspond to increased sterilization. While many of the organisms present in TTPC-amended cultures may be resistant to biocide, items present within the growth media, such as the shale particles, may also be interacting with TTPC, decreasing its efficacy. While biotic cultures did not show major differences in comparison to abiotic cultures, all samples containing biocide showed changes in water composition. Cultures containing biocide were especially cloudy, and biocide appears to have allowed for the shale particles within the cultures to disperse much more easily than cultures that did not. This could be due to the fact that TTPC is partly hydrophobic, allowing for easier extraction of organic matter within the organic rich EF drill cuttings (Kramer et al., 2008). Previous work has also shown that high organic loads present in the media can decrease the effectiveness of TTPC (Struchtemeyer et al., 2012). A decrease in TTPC’s effectiveness may also be due to its adsorption onto the shale particles in the growth media. The organicrich, crushed drill cuttings present in the growth media appears to have allowed for the sorption and removal of much of the TTPC, resulting in lowered biocide activity and the continued presence of organisms within the media. This interaction with shale particles in the growth media may also explain the increased metal concentrations observed in TTPC containing media (Fig. 4). The adsorption of the biocide onto the organic rich shale may have allowed for the release of other metals attached on the same particles. Furthermore, many organic molecules are able to chelate metals present in rocks, further allowing for metal release (Appelo and Postma, 2005). Many of the metals detected in the cultures do have toxic effects at high concentrations due to their abilities to oxidize organic molecules (Gadd, 2010). TTPC not only has important implications to public safety due to its toxicity, it may also have the unintended consequences of adding further toxicity to PW and this should be a consideration when determining biocide use and PW disposal.

6. Conclusions and implications Our data suggests that at 4.1 km depth at the Eagle Ford, temperatures are too high to support viable microorganisms further providing evidence that sulfide present within EF is likely abiogenic. At decreased temperatures (e.g., 4128 °C), such as along the wellbore at depths shallower than 4.1 km, active microbial communities exist and these organisms are mixture of organisms present from the surface, organisms native to the subsurface, and organisms introduced into the environment through other means we did not account for. Furthermore, the diversity of the environment within the wellbore is rich, suggesting an

environment high in nutrient availability, electron acceptors, and carbon sources which can support multiple different microbial niches. This community may have many different effects on production ranging from biofouling, to corrosion, to oil degradation, though it is difficult to make a complete conclusion based solely on sequence data. Through sequencing, we have determined the community composition of this environment which in turn allows us to hypothesize the potential contribution to production. While the benefits of these organisms are mixed, aiming to completely sterilize wells may ignore the potential positive roles these organisms could play, especially in terms of enhanced oil recovery. Furthermore, the conventional biocide used to sterilize wells during HF appears ineffective at completely sterilizing the microbial community, even extremely high dosages. While biocide does drastically decrease diversity and biomass, it also appears to have the unintended side effect of selecting for the most resilient organisms. These organisms, having little competition due to the sterilization of the rest of the microbial community, have also been implicated in petroleum degradation and could potentially be decreasing yields. While the effect of these organisms was not measurable over the two week experimental incubation, their effects may be more drastic over long time scales and should be further investigated. The fact that organisms are present and viable despite the addition of biocide suggests the need to consider alternate methods for microbial control. Doing so can result in a significant reduction in HF costs and can reduce impacts on both health and the environment. The organisms examined in this study constitute only one well. However, given the heterogeneity in the subsurface, and the different geochemical conditions of different shale plays, microbial communities will be different in each HF well. In order to maximize the efficacy of HF fluids, fluid composition could potentially be tailored to reflect different microbial communities in different wells. Doing so could allow for the right biocide uses or for innovative methods to enhance production in various wells. Based on its mixed success at sterilizing microbes, its cost as part of the HF fluid mix, as well as its potential environmental impacts, we suggest that further tests on TTPC efficacy should be performed in comparison to other biocides and in other HF sites in order to truly maximize the effectiveness of HF fluids.

Acknowledgments We would acknowledge the help of the Statoil Eagle Ford asset team. In particular, we would like to thank Zethais Mujica, Marianne Gudmundseth Nilsen, and Amy Paddock. Additional thanks go to Lichun Wang, Anita Skarstad, Nate Miller, and Scot Dowd.

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