Inhibition of Biodegradation of Hydraulic Fracturing Compounds by

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Inhibition of biodegradation of hydraulic fracturing compounds by glutaraldehyde: Groundwater column and microcosm experiments Jessica D. Rogers, Imma Ferrer, Shantal S. Tummings, Angela Bielefeldt, and Joseph N. Ryan Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b02316 • Publication Date (Web): 07 Aug 2017 Downloaded from http://pubs.acs.org on August 8, 2017

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Environmental Science & Technology

Inhibition of biodegradation of hydraulic fracturing compounds by glutaraldehyde: Groundwater column and microcosm experiments Jessica D. Rogers1, Imma Ferrer1, Shantal S. Tummings1,2, Angela R. Bielefeldt 1, and Joseph N. Ryan1*

1

Department of Civil, Environmental and Architectural Engineering University of Colorado Boulder, Boulder, CO 80309 2

Department of Civil, Environmental, and Geodetic Engineering The Ohio State University; Columbus, OH 43210 * Corresponding author, phone: (303)492-0772 e-mail: [email protected] address: 607 UCB, Boulder, CO 80303

ACS Paragon Plus Environment


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1 1 2

ABSTRACT The rapid expansion of unconventional oil and gas development has raised concerns

3

about the potential contamination of aquifers; however, the groundwater fate and transport of

4

hydraulic fracturing fluid compounds and mixtures remains a significant data gap. Degradation

5

kinetics of five hydraulic fracturing compounds (2-propanol, ethylene glycol, propargyl alcohol,

6

2-butoxyethanol, and 2-ethylhexanol) in the absence and presence of the biocide glutaraldehyde

7

were investigated under a range of redox conditions using sediment-groundwater microcosms

8

and flow-through columns. Microcosms were used to elucidate biodegradation inhibition at

9

varying glutaraldehyde concentrations. In the absence of glutaraldehyde, half-lives ranged from

10

13 d to >93 d. Accurate mass spectrometry indicated that a trimer was the dominant aqueous-

11

phase glutaraldehyde species. Microbial inhibition was observed at glutaraldehyde trimer

12

concentrations as low as 5 mgL-1, which demonstrated that the trimer retained some biocidal

13

activity. For most of the compounds, biodegradation rates slowed with increasing

14

glutaraldehyde concentrations. For many of the compounds, degradation was faster in the

15

columns than the microcosms. Four compounds (2-propanol, ethylene glycol, propargyl alcohol,

16

and 2-butoxyethanol) were found to be both mobile and persistent in groundwater under a range

17

of redox conditions. The glutaraldehyde trimer and 2-ethylhexanol were more rapidly degraded,

18

particularly under oxic conditions.


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2 19

INTRODUCTION

20

The rapid expansion of unconventional oil and gas (UOG) development has raised

21

concerns about the potential contamination of aquifers.1-3 Possible pathways for groundwater

22

contamination include surface spills of hydraulic fracturing fluids (HFF) or flowback and

23

produced waters (referenced as “UOG wastewaters”) and subsurface releases due to wellbore

24

integrity failure.2, 4-7 Several recent studies have reported the occurrence of organic

25

contaminants in groundwater potentially linked to UOG extraction.4, 8-10 These findings

26

underscore the need for exposure assessments to understand potential risks to aquifers; however,

27

significant knowledge gaps include frequency of releases, toxicity data, and the mobility and

28

persistence of HFF compounds and mixtures in groundwater.1, 11-13 Many HFF compounds are

29

individually biodegradable.14 Several studies have observed biodegradation of HFF mixtures;

30

however, most measured the removal of bulk parameters such as dissolved organic carbon,15-17

31

and data on specific compounds within the HFF mixture remain limited.18 These studies

32

identified several co-contaminant interactions that affected the fate and transport of HFF

33

compounds, including inhibition of biodegradation by high salinity16, 19 or the presence of

34

biocides.18

35

Five HFF compounds were selected for this study: 2-propanol, ethylene glycol,

36

propargyl alcohol, 2-butoxyethanol, and 2-ethylhexanol (Table 1). The combined frequency of

37

use and/or expected mobility and persistence in groundwater makes these compounds more

38

likely to be transported in the event of a release.12 Four of the compounds (2-propanol, ethylene

39

glycol, propargyl alcohol, and 2-butoxyethanol) have been identified in groundwater samples

40

potentially impacted by UOG extraction.9, 10, 20, 21



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3 41

Glutaraldehyde was chosen to investigate biodegradation inhibition of the five HFF

42

compounds in the presence of a biocide. Biocides are frequently used in HFFs to prevent

43

biological growth leading to clogging, corrosion, and generation of hydrogen sulfide gas.22-25

44

Glutaraldehyde’s biocidal activity is attributed to electrophilic aldehyde groups, which cross-link

45

amines within bacterial membrane proteins.22, 26-28 Glutaraldehyde is relatively mobile in

46

groundwater and is considered readily biodegradable under aerobic and anaerobic conditions at

47

sub-inhibitory concentrations.29 Reported glutaraldehyde inhibitory concentrations range from 5

48

up to 200 mgL-1.23, 24, 29 At high concentrations typically used in HFFs, glutaraldehyde is

49

inhibitory to the biodegradation of other compounds.18 Glutaraldehyde polymerizes at alkaline

50

pH30, 31 and multiple hydrated and polymerized species can exist in equilibrium.26, 32

51

Glutaraldehyde is rapidly polymerized under downhole conditions (high temperatures and

52

alkaline pH) and larger polymers are precipitated; thus, in the wastewaters returning to the

53

surface glutaraldehyde may be entirely depleted or present only as water-soluble oligomers.31

54

The biocidal efficacy and environmental behavior of glutaraldehyde oligomers are not well

55

characterized.31

56

The objectives of this study were to characterize the fate and transport of five HFF

57

compounds with increased groundwater exposure potential in aquifer sediments and to examine

58

inhibition effects of glutaraldehyde on the biodegradation of the selected HFF compounds.

59

Removal kinetics of five HFF compounds in the absence and presence of glutaraldehyde were

60

measured under a range of reduction-oxidation (redox) conditions using sediment-groundwater

61

microcosms and flow-through columns. Rates were measured under both oxic and anoxic

62

environments to capture a range of conditions relevant for groundwater transport. Microcosms

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were used to elucidate biodegradation inhibition at varying glutaraldehyde concentrations.


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4 64

Flow-through columns packed with aquifer sediment were used to measure fate and transport

65

parameters under conditions more representative the in situ groundwater environment.

66 67 68

METHODS Aquifer material and groundwater composition

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Sediments were collected from the Arapahoe Formation between the depths of 70 and

70

120 m during the drilling of a domestic well at a location in the Denver-Julesburg Basin in

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Colorado. The Arapahoe Formation is heavily utilized as a domestic and agricultural aquifer.33,

72

34

73

The composition and hydrologic properties were typical for a sandstone formation, including low

74

organic carbon content (0.10% w/w; Table S1).

75

Collected sediments were homogenized and stored saturated with native groundwater at 4°C.

A synthetic groundwater representative of the Arapahoe Formation with respect to major

76

ions and pH was used in all experiments (details in Supporting Information). The synthetic

77

groundwater was dominated by calcium and bicarbonate with a pH of 7.9 (Table S1). In the

78

columns, the bromide concentration was increased as a conservative tracer.

79 80 81

Hydraulic fracturing fluid compounds HFF compound concentrations were determined from the average concentration for

82

hydraulic fracturing performed in the Denver-Julesburg Basin for 2010–2015 as reported on the

83

FracFocus Chemical Disclosure Registry35 (Table 1). Glutaraldehyde was obtained as a 70%

84

solution in water, and 2-propanol, ethylene glycol, propargyl alcohol, 2-butoxyethanol and

85

2-ethylhexanol were obtained as 99% reagents (Sigma-Aldrich).

86

The HFF compounds were quantified using a gas chromatograph equipped with a

87

flame-ionization detector with direct aqueous injection (GC-FID; 7890a, Agilent Technologies;



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method details and detection limits in SI). For treatments with initial glutaraldehyde

89

concentrations of 5 mgL-1, glutaraldehyde was measured using colorimetric reagents (Hach),

90

quantified by absorbance at 600 nm using a spectrophotometer (Cary 100, Agilent Technologies;

91

method details in SI). Glutaraldehyde polymerization was evaluated using liquid

92

chromatography coupled with quadrupole time-of-flight mass spectrometry (LC/Q-TOF-MS)

93

using the Ferrer and Thurman method.32

94 95 96

Microcosms Sediment-groundwater microcosm experiments were conducted using sacrificial

97

sampling. For each treatment, 18-30 individual microcosms were constructed, and duplicates

98

were sacrificed at each time point. In each microcosm, HFF reagents were dissolved in synthetic

99

groundwater (100 mL) and added to 125 mL pre-baked borosilicate glass serum bottles with 25 g

100

of saturated sediments. The dry-weight equivalent of the saturated sediments was 20±0.2 g

101

(oven-dried at 150C for 2 h), or a 5:1 ratio of water to sediment masses. Bottles were mixed

102

continuously on an orbital shaker (150 rpm) in the dark at 20±2°C.

103

Four treatments were prepared under both oxic and anoxic conditions: the HFF mixture

104

with no glutaraldehyde (“GA-0”) and with initial glutaraldehyde concentrations of 5 mgL-1

105

(“GA-5”), 50 mgL-1 (“GA-50”), and 100 mgL-1 (“GA-100”). An abiotic control with the HFF

106

mixture, 50 mgL-1 glutaraldehyde, and 1.0 gL-1 sodium azide (NaN3, high purity grade, Amresco)

107

was used to distinguish abiotic and biotic removal. Oxic serum bottles were covered loosely

108

with aluminum foil. For anoxic treatments, the groundwater was de-aerated by purging with

109

>99% N2 for 1 h prior to the addition of the HFF mixture. Anoxic microcosms were prepared

110

under a N2 atmosphere and sealed with Teflon-lined septa.


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Microcosm samples were collected using sterile needles and glass syringes. Dissolved

112

oxygen (DO) and pH were measured immediately using a luminescent DO probe (Hach

113

LDO101) and a pH electrode (Hach PHC301). Samples collected for HFF compound analysis

114

were syringe-filtered (0.2 μm, polyethersulfone membrane, Pall Corporation) and stored in 2 mL

115

amber glass vials capped with a Teflon-lined silicon septa without headspace. Samples were

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stored at 4C for less than 14 days before analysis. First-order removal rate coefficients were fit

117

using OriginPro 201636 and included any acclimation periods. The significance of rate

118

coefficients was analyzed using an F-test executed in OriginPro (α = 0.01 confidence level). To

119

avoid substantial extrapolation, half-lives (t1/2) were only reported when at least 40% removal

120

was observed by the final sample. Suspended adenosine triphosphate (ATP) was monitored

121

using a luminescence assay and luminometer (PhotonMaster, LuminUltra; method details in SI).

122

Major anions and cations were analyzed using inductively coupled plasma-optical emission

123

spectroscopy (3410, ARL) and ion chromatography (4500I, Dionex), respectively.

124 125 126

Flow-through columns Two stainless steel columns (1 m length, 0.1 m internal diameter) were constructed

127

following the design of Pitoi et al.37 with ten sample ports located along the length (Fig. S1;

128

additional details in SI). Sample ports consisted of stainless steel needles with Teflon Luer-lock

129

hubs that extended 0.05 m to the center of the column. Columns were wet-packed with sediment

130

and operated under saturated up-flow conditions. After an initial equilibration period with the

131

synthetic groundwater, the combined groundwater and HFF mixture was injected continuously at

132

the base of each column at a rate of 2.4±0.2 mLh-1 using a peristaltic pump drive



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(Masterflex 07522-30). The average linear flow velocity was 0.018 md-1, which resulted in a

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water residence time of 56 d.

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Two injection solutions were applied: a biotic column with the HFF mixture and an initial

136

glutaraldehyde concentration of 50 mgL-1, and an abiotic control column with the HFF mixture,

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50 mgL-1 glutaraldehyde, and 1.0 gL-1 NaN3. The injection solutions were initially oxic and the

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columns were allowed to develop gradients to simulate redox zones characteristic of

139

groundwater contaminant plumes. Fresh injection solutions were prepared every 14 d and were

140

stored in 3 L sample bags with stainless steel fittings (FlexFoil, SKC).

141

Samples were collected by attaching a glass syringe with a Teflon Luer lock fitting to the

142

needle hub at each port. Prior to collecting a sample, 2 mL of pore water was purged from each

143

port. The first sample was collected 15 d after starting the injection solution, and subsequently at

144

30-day intervals. The injection solution was applied continuously for 232 d.

145

For each sample, DO and pH and were measured immediately. Samples collected for

146

HFF compounds and inorganic ions were syringe-filtered and quantified as described above.

147

After the d 232 sample, column sediments were analyzed for attached ATP (method details in

148

SI). Retardation coefficients (R) and first-order removal rate coefficients were determined using

149

curve-fitting functions in OriginPro, and sediment partition coefficients (Kd) were calculated

150

from fitted R values (details in SI). The significance of rate coefficients was analyzed using an

151

F-test executed in OriginPro. Half-lives were reported when at least 40% removal was observed

152

by the final sample port.

153 154


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RESULTS Glutaraldehyde speciation Accurate mass spectrometry indicated that a trimer was the dominant aqueous-phase

158

glutaraldehyde species in all samples, including d 0 (on average 72% of the total response of all

159

glutaraldehyde species; Table S5). The monomer and dimer were not present. The tetramer was

160

detected (on average 28% of total response) but no larger polymers were identified. The relative

161

abundance of the trimer and tetramer did not change with time, and was consistent across

162

varying concentrations (5-100 mgL-1). The relative abundance of the tetramer was slightly

163

greater at higher pH (25% at pH 7.7; 32% at pH 8.0). A time point comparison showed that the

164

relative concentrations (C/C0) of the trimer measured by LC/Q-TOF-MS corresponded closely

165

with C/C0 measured by GC-FID (Table S6), confirming that the GC-FID method measured the

166

trimer. Thus, our results report the groundwater fate and transport of the glutaraldehyde trimer.

167

Two trimer ions were identified in the mass spectra: a triply-hydrated trimer (m/z = 341)

168

and a doubly-hydrated fragment (m/z = 323) indicating the loss of one water molecule.32 An ion

169

with all aldehyde groups hydrated was not present. The retention times for the two ions were

170

identical, which demonstrates that dehydration occurred in the mass spectrometer source;

171

therefore, one aldehyde group in the trimer remained unbound by water in solution.

172 173

Microcosms

174

Microbial activity. Microbial activity was monitored using suspended ATP as an

175

indicator.38 Under oxic conditions, lags in ATP production increased and overall magnitude of

176

ATP concentrations decreased as glutaraldehyde concentrations increased (Fig. 1a). There was

177

no production of ATP in the GA-100 treatment or the abiotic control. Under anoxic conditions



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there was a lag in the GA-5 compared to the GA-0 treatment, and no production of ATP in the

179

GA-50 and GA-100 treatments or the abiotic control (Fig. 1b).

180

Redox conditions. In all oxic treatments (open to the atmosphere), DO remained

181

saturated throughout the experiment and nitrate (NO3-) depletion was observed in the GA-0,

182

GA-5, and GA-50 treatments (Fig. S3a). No significant changes were observed in other redox-

183

active species monitored in any treatment, including the abiotic control (data not shown). In all

184

anoxic treatments (deaerated and capped with a nitrogen headspace), DO remained below

185

detection. NO3- was depleted in the GA-0 and GA-5 treatments (Fig. S3b). No NO3- depletion

186

was observed in the GA-50 and GA-100 treatments, and no changes were observed in the other

187

redox-active species monitored for any treatment, including the abiotic control (data not shown).

188

The pH was stable for all treatments (7.4–8.2; Fig. S4).

189

HFF compound removal. The glutaraldehyde trimer was quickly removed in the oxic

190

GA-5 and GA-50 treatments (t1/2 = 5 and 15 d, respectively; Table 2; Fig. 2a), and there was not

191

a significant difference between the GA-100 treatment and the abiotic control, where 15%

192

removal was observed (Table S7). High variability was observed between GA-50 experimental

193

duplicates over d 50-77. Because glutaraldehyde was completely removed in both duplicates by

194

d 93, duplicates with limited glutaraldehyde removal during d 50-77 were considered outliers

195

and excluded from kinetic models for all HFF compounds. Under anoxic conditions,

196

glutaraldehyde was removed in the GA-5 treatment (t1/2 = 12 d; Fig. S5a). There was not a

197

significant difference between the GA-50 and GA-100 treatments and the abiotic control, where

198

37% removal occurred.

199 200

Under oxic conditions, 2-ethylhexanol was removed in the GA-0 and GA-5 treatments (t1/2 = 13 and 24 d, respectively; Table 2; Fig. 2b). Removal in the GA-50 and GA-100


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treatments was not significantly different than the abiotic control (t1/2 values of 68, 68, and 71 d).

202

Similarly, 2-propanol was removed in oxic GA-0 and GA-5 treatments (t1/2 = 71 and 66 d;

203

Fig. 2c), and removal in the GA-50 and GA-100 treatments was not significantly different than

204

the abiotic control (t1/2 = 94, 103, and 101 d). For both 2-ethylhexanol and 2-propanol, removal

205

occurred in the abiotic control (64% and 54%, respectively). Under anoxic conditions, there was

206

limited removal (93 d. Enhanced removal attributable to biodegradation was observed in the

275

GA-0 treatment for 2-ethylhexanol and 2-propanol. There was evidence of slow 2-butoxyethanol

276

degradation in the oxic GA-0 treatment, and no enhanced removal of propargyl alcohol

277

attributable to biodegradation. There was no evidence of ethylene glycol degradation in the

278

GA-0 treatment.

279

Anoxic conditions. Under anoxic conditions, degradation of the HFF compounds was

280

limited in the absence of glutaraldehyde. In the GA-0 treatment, the only compound that

281

degraded, albeit slowly, was 2-butoxethanol (Table S7).

282 283

Degradation of glutaraldehyde trimer

284

Abiotic removal. Abiotic transformation of the glutaraldehyde trimer was apparent in the

285

microcosm controls. Removal in the oxic abiotic control was not statistically different from the

286

oxic GA-100 or anoxic GA-50 and GA-100 treatments (15-25%), and may be due to sorption.

287

Further polymerization was likely not a significant removal mechanism because the relative

288

response of the glutaraldehyde trimer and tetramer were steady with time. More removal was

289

observed in the anoxic than the oxic abiotic control. The azide added to the abiotic controls to

290

inhibit microbial activity can be reduced to ammonium (NH4+).41 Ammonium was not

291

quantified, but if azide was reduced to NH4+ under anoxic conditions, ammonia could have


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cross-linked with the trimer’s available aldehyde group18, 26, 27 and caused the enhanced removal

293

of the glutaraldehyde trimer observed in the anoxic but not the oxic control.

294

Oxic conditions. Under oxic conditions, degradation of the glutaraldehyde trimer slowed

295

as concentrations increased. Degradation was faster in the GA-5 than the GA-50 treatment

296

(t1/2 = 5 and 15 d, respectively), and there was limited evidence of biodegradation in the GA-100

297

treatment relative to the abiotic control. Following a lag of approximately 30 d, the extent of

298

degradation was highly variable between duplicates in the GA-50 treatment (Fig. 2a). When fit

299

for only the period of rapid decay in some duplicates (36-64 d), the GA-50 half-life increased to

300

5 d, which suggests that the glutaraldehyde trimer was rapidly degraded once its concentration

301

was reduced to a sub-inhibitory level. Large variability between time points and duplicates was

302

frequently observed during the microcosm experiments, which may be a consequence of our

303

sacrificial sampling design where an individual microcosm bottle was used for each sample. For

304

instance, small differences between individual microcosms (e.g., the initial microbial consortia,

305

fraction organic carbon) could result in larger differences over time. This is particularly true for

306

low biomass populations.42 The slower degradation rate of the glutaraldehyde trimer with

307

increasing concentrations provides additional evidence of biodegradation as opposed to further

308

polymerization: because glutaraldehyde polymerization follows second-order kinetics,26, 31 we

309

would likely observe faster rates at higher concentrations if removal was due to further

310

polymerization. This is in agreement with the consistency of the relative abundance of the trimer

311

and tetramer at varying glutaraldehyde concentrations. Finally, the pH of the biotic treatments

312

was overall slightly lower than that of the abiotic controls, so increased polymerization due to

313

more alkaline pH would not be expected.



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15 314

Anoxic conditions. Biodegradation of the glutaraldehyde trimer was slower under anoxic

315

conditions than under oxic conditions but followed the trend of slower rates with increasing

316

concentrations. The trimer was degraded in the GA-5 treatment (t1/2 = 12 d). There was no

317

evidence of biodegradation in the GA-50 or GA-100 treatments.

318 319 320

Glutaraldehyde inhibition of HFF compound biodegradation The glutaraldehyde trimer was found to be inhibitory to the indigenous microorganisms.

321

The ATP concentrations did not increase until glutaraldehyde trimer concentrations decreased

322

(Figs. S12 and S13), which demonstrates toxicity of the biocide.43, 44 Under oxic conditions,

323

there was a lag in ATP production relative to the GA-0 treatment that grew longer with

324

increasing initial glutaraldehyde concentrations (Fig. 1a). In the GA-100 treatment, ATP did not

325

increase relative to the abiotic control, which indicated that microbial activity was completely

326

inhibited. Under anoxic conditions, a lag in ATP production occurred in the GA-5 treatment,

327

and microbial activity was completely inhibited in the GA-50 and GA-100 treatments (Fig. 1b).

328

Glutaraldehyde’s biocidal mode of action is non-specific;28 thus, diminished ATP production

329

under anoxic conditions likely reflected overall low metabolic activity.38, 45 These results

330

indicate that microbial inhibition occurred at glutaraldehyde trimer concentrations as low as

331

5 mgL-1 and that complete inhibition occurred above 50 mgL-1, consistent with the lower range

332

of previously reported inhibitory concentrations.29 The observed inhibition demonstrates that the

333

glutaraldehyde trimer retained some biocidal activity, in agreement with LC/Q-TOF-MS analysis

334

which indicated the presence of an unbounded aldehyde group. This finding is consistent with

335

that of McLaughlin et al., 18 who reported biocidal effects when one of the glutaraldehyde

336

monomer’s two aldehyde groups was bound.


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16 337

Microbial inhibition was reflected in the biodegradation kinetics of several of the HFF

338

compounds (Fig. S14, Table S9). The glutaraldehyde trimer was inhibitory to

339

glutaraldehyde-degrading microorganisms – under oxic conditions, biodegradation of the trimer

340

slowed as concentrations increased. Biodegradation rates of 2-ethylhexanol and 2-propanol were

341

two to five times slower in the presence of the glutaraldehyde trimer. Under anoxic conditions,

342

the only biodegradation inhibition observed was for the glutaraldehyde trimer itself: the other

343

HFF compounds did not degrade sufficiently to evaluate inhibition effects.

344 345 346

Fate and transport in columns The HFF compounds all had low Kd values (Table 2). For these neutral organic

347

compounds, sorption to sediment organic matter was likely the most significant mechanism

348

contributing to the Kd.46 The relative order of the calculated Kd values agreed well with the

349

predicted sequence based on estimated organic carbon partition coefficients (Table S8), and the

350

low Kd values are consistent with the low organic carbon content of the sediments (0.1% w/w).

351

The low Kd values indicate that the six HFF compounds would be highly mobile in aquifers with

352

low organic carbon and would be only minimally separated during transport (not accounting for

353

removal by other mechanisms).

354

The only compound with significant abiotic removal in the columns was the

355

glutaraldehyde trimer, for which transformation was faster in the abiotic column than in the

356

microcosm abiotic controls. While the relative abundance of both oligomers was steady with

357

time, an increase in the relative abundance of the tetramer from 25% at pH 7.7 to 32% at pH 8.0

358

suggests that further polymerization could have occurred at pH >8.0. However, the dependence

359

of glutaraldehyde polymerization on pH likely does not fully account for the differences between



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17 360

the microcosm and column abiotic controls because all three control experiments had a similar

361

pH range (7.8-8.2). One possible explanation is nucleophilic substitution of the trimer’s

362

available aldehyde group27 by increased concentrations of bromide46 added to the columns as a

363

conservative tracer. Mass spectrometric analysis was not conducted for abiotic control samples

364

due to high azide concentrations; thus, the formation of potential abiotic transformation products

365

was not confirmed. Our results suggest that abiotic transformations of glutaraldehyde may be

366

significant, but more work is needed to better characterize potential mechanisms.

367

ATP concentrations were higher in biotic compared to the abiotic control column, which

368

were at or near the detection limit (Fig. S7), consistent with the inference of biodegradation as a

369

primary removal mechanism in the biotic column. ATP concentrations in the biotic column were

370

generally lower than the clean aquifer sediments. The high concentrations of the HFF mixture

371

could have inhibited the indigenous microbial consortia and contributed to the overall slow

372

degradation rates observed for most compounds.16, 47 Nevertheless, for many of the HFF

373

compounds, biodegradation was greater in the biotic column than the microcosms.

374

The initially oxic biotic column developed redox gradients characteristic of groundwater

375

contaminant plumes. Oxygen and nitrate were depleted over a transport distance of 0.04 m

376

(Fig. S8a). Thus, the biotic column had a short distance of mixed oxygen- and nitrate-reducing

377

conditions and was anoxic throughout the remaining length. Oxygen depletion in the abiotic

378

control column (Fig. S8b) was attributed to reduction by iron(II)-containing minerals in the

379

sediments (Table S1) because the ATP results indicated no biological activity.

380

In the biotic column, the glutaraldehyde trimer was rapidly degraded in the oxygen- and

381

nitrate-reducing zone (t1/2 = 0.5 d) at a rate much faster than the corresponding oxic GA-50

382

microcosm. The faster biodegradation in the column could be due to the formation of biofilms


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18 383

on the sediments,48 which are known to have decreased susceptibility to glutaraldehyde.24, 25, 49, 50

384

Additionally, more abiotic transformation occurred in the columns than in the microcosms,

385

which could have helped reduced the biocide to sub-inhibitory concentrations.18 The

386

approximately 30 d acclimation period in the biotic column was similar to that observed in the

387

GA-50 microcosm treatment.

388

Ethylene glycol degraded slowly in the biotic column under anoxic conditions prevalent

389

throughout most of the length (t1/2 = 82 d), while there was no evidence of degradation in the

390

anoxic microcosms over 93 d. Up to a 60 d acclimation period was observed in the biotic

391

column; thus, slow acclimation of the indigenous microbial consortia could have contributed to

392

limited biodegradation in the microcosms. There was evidence of slow degradation of

393

2-ethylhexanol and 2-propanol in the biotic column under anoxic conditions (Table S7), while no

394

biodegradation was observed in the anoxic microcosms. Similarly to glutaraldehyde, greater

395

degradation in the columns than in the microcosms may be due to increased resistance of the

396

attached biomass to environmental stressors.25 Breakthrough curves suggested a long

397

acclimation period prior to 2-propanol degradation in the biotic column; thus, limited removal in

398

the anoxic microcosms may be a result of the indigenous microbial community not having

399

sufficient exposure time to acclimate. There was no evidence of degradation of

400

propargyl alcohol or 2-butoxyethanol in the biotic column, consistent with limited removal in the

401

anoxic microcosm.

402 403 404 405

Environmental implications Under both oxic and anoxic conditions, our measured removal rates of the HFF compounds were generally slower than those reported by other studies at comparable



Page 20 of 34

19 406

concentrations, even in the absence of glutaraldehyde (Table 2).16, 29, 39, 40, 51-62 Site-specific

407

variability is a general limitation of biodegradation studies. Differences in sediment properties,

408

nutrient and electron acceptor availability, groundwater pH, and indigenous microbial diversity

409

and abundance can all have large effects on the fate and transport behavior of contaminants.42, 47

410

The overall limited removal may be due to the pristine nature of the Arapahoe Formation

411

sediments, which were collected from a depth where prior exposure to organic contaminants

412

would not be expected and indigenous biomass density and population diversity are typically

413

low.45, 48, 60 Slow proliferation of microorganisms capable of degrading the HFF compounds due

414

to initially low populations42, 63 and limited or slow adaptation of the indigenous microbial

415

community may have contributed to long acclimation periods in the biotic column and the

416

variability between microcosm duplicates. Given the long acclimation periods observed for our

417

continuous injection scenario, the HFF compounds would be removed to a lesser extent during a

418

shorter release under otherwise similar conditions (Fig. S10). The high concentration of our HFF

419

mixture also may have contributed to the overall limited degradation observed. The

420

concentrations used in all experiments were representative of a release of HFF prior to injection.

421

Other release scenarios would likely involve lower concentrations of these compounds,31, 32, 64 or

422

concentrations may be reduced by dilution or hydrodynamic processes. At lower concentrations,

423

faster biodegradation may be expected.47 Greater removal may also be expected for other HFF

424

release scenarios, such as transport through surficial aquifer sediments with higher organic

425

carbon content or greater microbial density and diversity following a surface spill. While

426

environmental fate and transport can be highly variable, our results demonstrate that some of the

427

HFF compounds, including 2-propanol, ethylene glycol, propargyl alcohol and 2-butoxyethanol,

428

can be both mobile and persistent in groundwater under a range of redox conditions, which is


Page 21 of 34


20 429

consistent with their recent detections in groundwater field samples.9, 10, 20, 21 Glutaraldehyde and

430

2-ethylhexanol, both found to be more rapidly degraded particularly under oxic conditions, have

431

not been reported in any environmental samples.

432

Our results demonstrate that biodegradation inhibition in the presence of the biocide

433

glutaraldehyde may be relevant for the fate and transport of HFF compounds. However, the

434

timing of a spill is significant because glutaraldehyde may be rapidly removed under downhole

435

conditions,31 and so far glutaraldehyde has not been reported in any UOG wastewater samples in

436

any form (monomer or polymer).32 Thus, biodegradation inhibition may be of most concern for

437

accidental releases of pre-injection HFF containing glutaraldehyde. Biocides are sometimes

438

applied to UOG wastewater surface impoundments,25, 65 representing an additional release

439

scenario in which inhibition may be pertinent.

440

The management of UOG wastewaters is emerging as a significant issue with respect to

441

potential environmental contamination.5, 66 While our study demonstrates inhibition of a

442

microbial consortia indigenous to pristine aquifer sediments, evidence suggests that microbes

443

present in UOG wastewaters are resistant to both biocides and high salinity.15, 23, 67 During a spill

444

of UOG wastewater, any microbes present could also be released to the environment.48

445

Additionally, reactions downhole may alter the composition of UOG wastewaters compared to

446

the pre-injection HFF.28, 31, 68, 69 Thus, the degradation potential of compounds within the

447

complex matrix of actual UOG wastewaters under environmentally-relevant conditions is an

448

important research need. Overall, evidence shows that some HFF compounds can be naturally

449

attenuated in groundwater under a range of redox conditions; however, much work remains to

450

better characterize fate and transport of HFF mixtures and UOG wastewaters to improve our

451

understanding of exposure potential for groundwater resources.



Page 22 of 34

21 452

AUTHOR INFORMATION

453

Corresponding Author: *E-mail: [email protected]. Phone: (303)492-0772.

454

Notes: The authors declare no competing financial interest.

455 456

ACKNOWLEDGEMENTS

457

This research is supported by the AirWaterGas Sustainability Research Network funded by the

458

National Science Foundation (CBET-1240584) and the U.S. Environmental Protection Agency

459

STAR Fellowship (FP 91745101). We thank Dr. E. Michael Thurman for assistance with

460

LC/Q-TOF-MS analysis, Troy Burke for assistance with FracFocus data, and Dr. Fred Luiszer

461

for assistance with ICP-OES and IC analysis.

462 463

ASSOCIATED CONTENT

464

Supporting Information Available: Additional detail on methods; nine tables and 14 figures

465

showing rate coefficients, column breakthrough curves, geochemical results, and inhibition

466

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

467


Page 23 of 34


22 468 469 470 471 472 473 474 475 476 477 478 479 480 481 482 483 484 485 486 487 488 489 490 491 492 493 494 495 496 497 498 499 500 501 502 503 504 505 506 507 508 509 510 511 512

REFERENCES (1) Adgate, J.L.; Goldstein, B.D.; McKenzie, L.M. Potential public health hazards, exposures and health effects from unconventional natural gas development. Environmental Science & Technology 2014, 48, (15), 8307-8320; DOI: 10.1021/es404621d. (2) Vengosh, A.; Jackson, R.B.; Warner, N.; Darrah, T.H.; Kondash, A. A critical review of the risks to water resources from unconventional shale gas development and hydraulic fracturing in the United States. Environmental Science & Technology 2014, 48, (15), 8334-8348; DOI: 10.1021/es405118y. (3) Sherwood, O.A.; Rogers, J.D.; Lackey, G.; Burke, T.L.; Osborn, S.G.; Ryan, J.N. Groundwater methane in relation to oil and gas development and shallow coal seams in the Denver-Julesburg Basin of Colorado. Proceedings of the National Academy of Sciences of the United States of America 2016, 113, (30), 8391-8396; DOI: 10.1073/pnas.1523267113. (4) Gross, S.A.; Avens, H.J.; Banducci, A.M.; Sahmel, J.; Panko, J.M.; Tvermoes, B.E. Analysis of BTEX groundwater concentrations from surface spills associated with hydraulic fracturing operations. Journal of the Air & Waste Management Association 2013, 63, (4), 424-32; DOI: 10.1080/10962247.2012.759166. (5) Vidic, R.D.; Brantley, S.L.; Vandenbossche, J.M.; Yoxtheimer, D.; Abad, J.D. Impact of shale gas development on regional water quality. Science 2013, 340, (6134); DOI: 10.1126/science.1235009. (6) Jackson, R.B. The integrity of oil and gas wells. Proceedings of the National Academy of Sciences of the United States of America 2014, 111, (30), 10902-10903; DOI: 10.1073/pnas.1410786111. (7) Li, H.S.; Son, J.H.; Carlson, K.H. Concurrence of aqueous and gas phase contamination of groundwater in the Wattenberg oil and gas field of northern Colorado. Water Research 2016, 88, 458-466; DOI: 10.1016/j.watres.2015.10.031. (8) Drollette, B.D.; Hoelzer, K.; Warner, N.R.; Darrah, T.H.; Karatum, O.; O'Connor, M.P.; Nelson, R.K.; Fernandez, L.A.; Reddy, C.M.; Vengosh, A.; Jackson, R.B.; Elsner, M.; Plata, D.L. Elevated levels of diesel range organic compounds in groundwater near Marcellus gas operations are derived from surface activities. Proceedings of the National Academy of Sciences of the United States of America 2015, 112, (43), 13184-13189; DOI: 10.1073/pnas.1511474112. (9) DiGiulio, D.C.; Jackson, R.B. Impact to underground sources of drinking water and domestic wells from production well stimulation and completion practices in the Pavillion, Wyoming, Field. Environmental Science & Technology 2016, 50, (8), 4524-36; DOI: 10.1021/acs.est.5b04970. (10) Llewellyn, G.T.; Dorman, F.; Westland, J.L.; Yoxtheimer, D.; Grieve, P.; Sowers, T.; Humston-Fulmer, E.; Brantley, S.L. Evaluating a groundwater supply contamination incident attributed to Marcellus Shale gas development. Proceedings of the National Academy of Sciences of the United States of America 2015, 112, (20), 6325-6330; DOI: 10.1073/pnas.1420279112. (11) Yost, E.E.; Stanek, J.; DeWoskin, R.S.; Burgoon, L.D. Overview of chronic oral toxicity values for chemicals present in hydraulic fracturing fluids, flowback, and produced waters. Environmental Science & Technology 2016, 50, (9), 4788-4797; DOI: 10.1021/acs.est.5b04645.



Page 24 of 34

23 513 514 515 516 517 518 519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556 557

(12) Rogers, J.D.; Burke, T.L.; Osborn, S.G.; Ryan, J.N. A framework for identifying organic compounds of concern in hydraulic fracturing fluids based on their mobility and persistence in groundwater. Environmental Science & Technology Letters 2015, 2, (6), 158-164; DOI: 10.1021/acs.estlett.5b00090. (13) Yost, E.E.; Stanek, J.; Burgoon, L.D. A decision analysis framework for estimating the potential hazards for drinking water resources of chemicals used in hydraulic fracturing fluids. Science of the Total Environment 2017, 574, 1544-1558; DOI: 10.1016/j.scitotenv.2016.08.167. (14) Stringfellow, W.T.; Domen, J.K.; Camarillo, M.K.; Sandelin, W.L.; Borglin, S. Physical, chemical, and biological characteristics of compounds used in hydraulic fracturing. Journal of Hazardous Materials 2014, 275, 37-54; DOI: 10.1016/j.jhazmat.2014.04.040. (15) Strong, L.C.; Gould, T.; Kasinkas, L.; Sadowsky, M.J.; Aksan, A.; Wackett, L.P. Biodegradation in waters from hydraulic fracturing: Chemistry, microbiology, and engineering. Journal of Environmental Engineering 2014, 140, (5); DOI: 10.1061/(ASCE)EE.1943-7870.0000792. (16) Kekacs, D.; Drollette, B.D.; Brooker, M.; Plata, D.L.; Mouser, P.J. Aerobic biodegradation of organic compounds in hydraulic fracturing fluids. Biodegradation 2015, 26, (4), 271-287; DOI: 10.1007/s10532-015-9733-6. (17) Mouser, P.J.; Liu, S.; Cluff, M.A.; McHugh, M.; Lenhart, J.J.; MacRae, J.D. Redox conditions alter biodegradation rates and microbial community dynamics of hydraulic fracturing fluid organic additives in soil-groundwater microcosms. Environmental Engineering Science 2016, 33, (10); DOI: 10.1089/ees.2016.0031. (18) McLaughlin, M.C.; Borch, T.; Blotevogel, J. Spills of hydraulic fracturing chemicals on agricultural topsoil: Biodegradation, sorption, and co-contaminant interactions. Environmental Science & Technology 2016, 50, (11), 6071-6078; DOI: 10.1021/acs.est.6b00240. (19) Lester, Y.; Yacob, T.; Morrissey, I.; Linden, K.G. Can we treat hydraulic fracturing flowback with a conventional biological process? The case of guar gum. Environmental Science & Technology Letters 2013, 1, (1), 133-136; DOI: 10.1021/ez4000115. (20) Hildenbrand, Z.L.; Carlton, D.D., Jr.; Fontenot, B.E.; Meik, J.M.; Walton, J.L.; Taylor, J.T.; Thacker, J.B.; Korlie, S.; Shelor, C.P.; Henderson, D.; Kadjo, A.F.; Roelke, C.E.; Hudak, P.F.; Burton, T.; Rifai, H.S.; Schug, K.A. A comprehensive analysis of groundwater quality in the Barnett Shale region. Environmental Science & Technology 2015, 49, (13), 8254-62; DOI: 10.1021/acs.est.5b01526. (21) Hildenbrand, Z.L.; Carlton, D.D., Jr.; Meik, J.M.; Taylor, J.T.; Fontenot, B.E.; Walton, J.L.; Henderson, D.; Thacker, J.B.; Korlie, S.; Whyte, C.J.; Hudak, P.F.; Schug, K.A. A reconnaissance analysis of groundwater quality in the Eagle Ford shale region reveals two distinct bromide/chloride populations. Science of the Total Environment 2016. 575, 672-680. DOI: 10.1016/j.scitotenv.2016.09.070. (22) Kahrilas, G.A.; Blotevogel, J.; Stewart, P.S.; Borch, T. Biocides in hydraulic fracturing fluids: A critical review of their usage, mobility, degradation, and toxicity. Environmental Science & Technology 2015, 49, (1), 16-32. DOI: 10.1021/es503724k. (23) Vikram, A.; Lipus, D.; Bibby, K., Produced water exposure alters bacterial response to biocides. Environmental Science & Technology 2014, 48, (21), 13001-13009; DOI: 10.1021/es5036915.


Page 25 of 34


24 558 559 560 561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589 590 591 592 593 594 595 596 597 598 599 600 601 602 603

(24) Struchtemeyer, C.G.; Morrison, M.D.; Elshahed, M.S. A critical assessment of the efficacy of biocides used during the hydraulic fracturing process in shale natural gas wells. International Biodeterioration & Biodegradation 2012, 71, 15-21; DOI: 10.1016/j.ibiod.2012.01.013. (25) Gaspar, J.; Mathieu, J.; Yang, Y.; Tomson, R.; Leyris, J.D.; Gregory, K.B.; Alvarez, P.J.J. Microbial dynamics and control in shale gas production. Environmental Science & Technology Letters 2014, 1, (12), 465-473; DOI: 10.1021/ez5003242. (26) Migneault, I.; Dartiguenave, C.; Bertrand, M.J.; Waldron, K.C., Glutaraldehyde: Behavior in aqueous solution, reaction with proteins, and application to enzyme crosslinking. Biotechniques 2004, 37, (5), 790-802. (27) Williams, T.M.; McGinley, H.R. Deactivation of Industrial Water Treatment Biocides. In NACE International Corrosion Conference, National Association of Corrosion Engineers, San Antonio, TX, March 14-18, 2010. (28) Elsner, M.; Hoelzer, K. Quantitative survey and structural classification of hydraulic fracturing chemicals reported in unconventional gas production. Environmental Science & Technology 2016, 50, (7), 3290-3314; DOI: 10.1021/acs.est.5b02818. (29) Leung, H.W. Ecotoxicology of glutaraldehyde: Review of environmental fate and effects studies. Ecotoxicology and Environmental Safety 2001, 49, (1), 26-39; DOI: 10.1006/eesa.2000.2031. (30) Kawahara, J.; Ohmori, T.; Ohkubo, T.; Hattori, S.; Kawamura, M. The structure of glutaraldehyde in aqueous-solution determined by ultraviolet-absorption and light-scattering. Analytical Biochemistry 1992, 201, (1), 94-98. (31) Kahrilas, G.A.; Blotevogel, J.; Corrin, E.R.; Borch, T. Downhole transformation of the hydraulic fracturing fluid biocide glutaraldehyde: Implications for flowback and produced water quality. Environmental Science & Technology 2016, 50, (20), 11414-11423; DOI: 10.1021/acs.est.6b02881. (32) Ferrer, I.; Thurman, E.M. Analysis of hydraulic fracturing additives by LC/Q-TOF-MS. Analytical and Bioanalytical Chemistry 2015, 407, (21), 6417-28; DOI: 10.1007/s00216-015-8780-5. (33) Paschke, S.S. (ed.) Groundwater Availability of the Denver Basin Aquifer System, Colorado; Professional Paper 1770. U.S. Geological Survey: Reston, Va., 2011. (34) Leppert Associates, Inc. Arapahoe Aquifer Baseline Domestic Water Well Background Investigation Adams County, Colorado. Colorado Oil and Gas Conservation Commission: Denver, CO, 2007. (35) Groundwater Protection Council; Interstate Oil & Gas Conservation Commission. FracFocus Chemical Disclosure Registry. http://fracfocus.org/ (accessed June 2015). (36) OriginLab Corporation. OriginPro 2016. Northhampton, MA, 2016. (37) Pitoi, M.M.; Patterson, B.M.; Furness, A.J.; Bastow, T.P.; McKinley, A.J. Fate of N-nitrosomorpholine in an anaerobic aquifer used for managed aquifer recharge: A column study. Water Research 2011, 45, (8), 2550-60; DOI: 10.1016/j.watres.2011.02.018. (38) Patterson, J.W.; Brezonik, P.L.; Putnam, H.D., Measurement and Significance of Adenosine Triphosphate in Activated Sludge. Environmental Science & Technology 1970, 4, (7), 569-575; DOI: 10.1021/es60042a003. (39) U.S. National Library of Medicine. Toxicology Data Network (TOXNET) Hazardous Substances Data Bank (HSDB). http://toxnet.nlm.nih.gov/cgi-bin/sis/htmlgen?HSDB (accessed November 2016).



Page 26 of 34

25 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626 627 628 629 630 631 632 633 634 635 636 637 638 639 640 641 642 643 644 645 646 647

(40) Staples, C.A. A review of the environmental fate and aquatic effects of a series of C4 and C8 oxo-process chemicals. Chemosphere 2001, 45, (3), 339-346. (41) Skipper, H.D.; Westermann, D.T. Comparative effects of propylene oxide, sodium azide, and autoclaving on selected soil properties. Soil Biology and Biochemistry 1973, 5, 409-414. (42) Alexander, M., Biodegradation and Bioremediation; 2nd ed.; Academic Press: San Diego, CA, 1999. (43) Kennicutt, M.C. ATP as an indicator of toxicity. Water Research 1980, 14, (4), 325-328. (44) Dalzell, D.J.B.; Christofi, N. An ATP luminescence method for direct toxicity assessment of pollutants impacting on the activated sewage sludge process. Water Research 2002, 36, (6), 1493-1502. (45) Eydal, H.S.C.; Pedersen, K. Use of an ATP assay to determine viable microbial biomass in Fennoscandian Shield groundwater from depths of 3-1000 m. Journal of Microbiological Methods 2007, 70, (2), 363-373; DOI: 10.1016/j.mimet.2007.05.012. (46) Schwarzenbach, R.P.; Gschwend, P.M.; Imboden, D.M. Environmental Organic Chemistry; 2nd ed.; Wiley: Hoboken, NJ, 2003. (47) Alvarez, P.J.J.; Illman, W.A. Bioremediation and Natural Attenuation: Process Fundamentals and Mathematical Models. Wiley: Hoboken, N.J., 2006. (48) Griebler, C.; Lueders, T. Microbial biodiversity in groundwater ecosystems. Freshwater Biology 2009, 54, (4), 649-677; DOI: 10.1111/j.1365-2427.2008.02013. (49) Vikram, A.; Bomberger, J.M.; Bibby, K.J. Efflux as a glutaraldehyde resistance mechanism in Pseudomonas fluorescens and Pseudomonas aeruginosa biofilms. Antimicrobial Agents and Chemotherapy 2015, 59, (6), 3433-40; DOI: 10.1128/AAC.05152-14. (50) Simoes, L.C.; Lemos, M.; Araujo, P.; Pereira, A.M.; Simoes, M. The effects of glutaraldehyde on the control of single and dual biofilms of Bacillus cereus and Pseudomonas fluorescens. Biofouling 2011, 27, (3), 337-46; DOI: 10.1080/08927014.2011.575935. (51) Ejlertsson, J.; Alnervik, M.; Jonsson, S.; Svensson, B.H. Influence of water solubility, side-chain degradability, and side-chain structure on the degradation of phthalic acid esters under methanogenic conditions. Environmental Science & Technology 1997, 31, (10), 27612764; DOI: 10.1021/es961055x. (52) Nalli, S.; Cooper, D.G.; Nicell, J.A. Metabolites from the biodegradation of di-ester plasticizers by Rhodococcus rhodochrous. Science of the Total Environment 2006, 366, (1), 286-294; DOI: 10.1016/j.scitotenv.2005.06.020. (53) Bustard, M.T.; McEvoy, E.M.; Goodwin, J.A.S.; Burgess, J.G.; Wright, P.C. Biodegradation of propanol and isopropanol by a mixed microbial consortium. Applied Microbiology and Biotechnology 2000, 54, (3), 424-431. (54) Hollingsworth, J.; Sierra-Alvarez, R.; Zhou, M.; Ogden, K.L.; Field, J.A. Anaerobic biodegradability and methanogenic toxicity of key constituents in copper chemical mechanical planarization effluents of the semiconductor industry. Chemosphere 2005, 59, (9), 1219-1228; DOI: 10.1016/j.chemosphere.2004.11.067. (55) Suflita, J.M.; Mormile, M.R. Anaerobic biodegradation of known and potential gasoline oxygenates in the terrestrial subsurface. Environmental Science & Technology 1993, 27, (5), 976-978; DOI: 10.1021/es00042a022.


Page 27 of 34


26 648 649 650 651 652 653 654 655 656 657 658 659 660 661 662 663 664 665 666 667 668 669 670 671 672 673 674 675 676 677 678 679 680 681 682 683 684 685 686 687 688 689 690 691 692 693

(56) Loehr, R.C. Treatability Potential for EPA Listed Hazardous Wastes in Soil; EPA/600/2-89-011 (NTIS 89-166581); U.S. Environmental Protection Agency: Washington, D.C., 1989. (57) Agency for Toxic Substances and Disease Registry (ATSDR). Toxicological profile for 2-butoxyethanol and 2-butoxyethanol acetate; U.S. Department of Health and Human Services: Atlanta, GA, 1998. (58) Staples, C.A.; Boatman, R.J.; Cano, M.L. Ethylene glycol ethers: An environmental risk assessment. Chemosphere 1998, 36, (7), 1585-1613. (59) Staples, C.A.; Williams, J.B.; Craig, G.R.; Roberts, K.M. Fate, effects and potential environmental risks of ethylene glycol: A review. Chemosphere 2001, 43, (3), 377-383. (60) Klotzbucher, T.; Kappler, A.; Straub, K.L.; Haderlein, S.B. Biodegradability and groundwater pollutant potential of organic anti-freeze liquids used in borehole heat exchangers. Geothermics 2007, 36, (4), 348-361; DOI: 10.1016/j.geothermics.2007.03.005. (61) Mcgahey, C.; Bouwer, E.J. Biodegradation of ethylene-glycol in simulated subsurface environments. Water Science & Technology 1992, 26, (1-2), 41-49. (62) Mrklas, O.; Chu, A.; Lunn, S.; Bentley, L.R. Biodegradation of monoethanolamine, ethylene glycol and triethylene glycol in laboratory bioreactors. Water, Air, and Soil Pollution 2004, 159, (1), 249-263; DOI: 10.1023/B:WATE.0000049178.93865.d4. (63) Meckenstock, R.U.; Elsner, M.; Griebler, C.; Lueders, T.; Stumpp, C.; Aamand, J.; Agathos, S.N.; Albrechtsen, H.J.; Bastiaens, L.; Bjerg, P.L.; Boon, N.; Dejonghe, W.; Huang, W.E.; Schmidt, S.I.; Smolders, E.; Sorensen, S.R.; Springael, D.; van Breukelen, B.M. Biodegradation: Updating the concepts of control for microbial cleanup in contaminated aquifers. Environmental Science & Technology 2015, 49, (12), 7073-7081; DOI: 10.1021/acs.est.5b00715. (64) Orem, W.; Tatu, C.; Varonka, M.; Lerch, H.; Bates, A.; Engle, M.; Crosby, L.; McIntosh, J. Organic substances in produced and formation water from unconventional natural gas extraction in coal and shale. International Journal of Coal Geology 2014, 126, 20-31; DOI: 10.1016/j.coal.2014.01.003. (65) Mohan, A.M.; Hartsock, A.; Hammack, R.W.; Vidic, R.D.; Gregory, K.B. Microbial communities in flowback water impoundments from hydraulic fracturing for recovery of shale gas. FEMS Microbiology Ecology 2013, 86, (3), 567-580; DOI: 10.1111/15746941.12183. (66) Cozzarelli, I.M.; Skalak, K.J.; Kent, D.B.; Engle, M.A.; Benthem, A.; Mumford, A.C.; Haase, K.; Farag, A.; Harper, D.; Nagel, S.C.; Iwanowicz, L.R.; Orem, W.H.; Akob, D.M.; Jaeschke, J.B.; Galloway, J.; Kohler, M.; Stoliker, D.L.; Jolly, G.D. Environmental signatures and effects of an oil and gas wastewater spill in the Williston Basin, North Dakota. Science of the Total Environment 2016, 579, 1781-1793; DOI: 10.1016/j.scitotenv.2016.11.157. (67) Struchtemeyer, C.G.; Elshahed, M.S. Bacterial communities associated with hydraulic fracturing fluids in thermogenic natural gas wells in North Central Texas, USA. FEMS Microbiology Ecology 2012, 81, (1), 13-25; DOI: 10.1007/s00248-012-0073-3. (68) Hayes, T. Sampling and Analysis of Water Streams Associated with the Development of Marcellus Shale Gas; Final Report for the Marcellus Shale Coalition: Pittsburgh, PA, 2009. (69) Hoelzer, K.; Sumner, A.J.; Karatum, O.; Nelson, R.K.; Drollette, B.D.; O'Connor, M.P.; D'Arnbro, E.L.; Getzinger, G.J.; Ferguson, P.L.; Reddy, C.M.; Elsner, M.; Plata, D.L. Indications of transformation products from hydraulic fracturing additives in shale-gas



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wastewater. Environmental Science & Technology 2016, 50, (15), 8036-8048; DOI: 10.1021/acs.est.6b00430.


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28 697

Table 1. Fracturing fluid compounds studied in all experiments. compound

additive purpose a

FracFocus frequency (%) b

experimental concentration (mgL-1) c

groundwater occurrence d

glutaraldehyde

biocide

33.3

0 – 100 e

not reported

2-propanol

corrosion inhibitor, non-emulsifier, surfactant

50.1

240

Hildenbrand et al.20, DiGiulio and Jackson9

ethylene glycol

cross-linker, scale inhibitor, corrosion inhibitor, friction reducer

49.7

190

Llewellyn et al.10 f

propargyl alcohol

corrosion inhibitor

32.7

10 g

Hildenbrand et al.20, 21

2-butoxyethanol

surfactant, corrosion inhibitor, non-emulsifier

22.8

310

Llewellyn et al.10; DiGiulio and Jackson9

2-ethylhexanol

non-emulsifier, surfactant

7.2

30

not reported

structure

698

a

Function of additive in which each compound was identified as an ingredient.12

699

b

Percentage of FracFocus reports identifying use of compound.12

700

c

Concentration applied to all experiments, determined from average concentrations reported on

701

FracFocus for the Denver-Julesburg Basin.

702

d

Studies which have measured or cited the compound in groundwater samples.

703

e

Concentration varied for different treatments. Average concentration used in Denver-Julesburg

704

Basin of 100 mgL-1 was applied as maximum treatment concentration.

705

f

Reported in samples collected by the Pennsylvania Department of Environmental Protection.

706

g

Propargyl alcohol was applied at a concentration in the upper range determined from FracFocus

707

(as opposed to average) due to analytical limitations.

708



Page 30 of 34

29 709

Table 2. Observed sediment partition coefficients (Kd, columns only) and first-order half-lives

710

(t1/2) with  the standard error for flow-through columns and five microcosm treatments.

treatment

glutaraldehyde

literature b

anoxic microcosm

oxic microcosm

col

t1/2 (d)

Kd

2-propanol t1/2 (d)

Kd

ethylene glycol t1/2 (d)

2-butoxyethanol

2-ethylhexanol

t1/2 (d)

Kd

t1/2 (d)

Kd

> 63

0.068 0.020

> 66

0.105 0.012

biotic

0.5  0.06

abiotic

45  1

GA-0

n/a

71  7

> 64

> 64

> 64

13  1

GA-5

5  0.4

66  8

> 64

> 64

> 64

24  5

GA-50

15  1

94  11

> 93

99 30

79 19

68 13

GA-100

> 93

103 13

> 93

> 93

> 93

68 20

abiotic

> 93

101  9

> 93

> 93

> 93

71 12

GA-0

n/a

> 93

> 93

> 93

> 93

> 93

GA-5

12  1

> 93

> 93

> 93

> 93

> 93

GA-50

> 93

> 93

> 93

> 93

> 93

> 93

GA-100

> 93

> 93

> 93

> 93

> 93

> 93

abiotic

145  13

> 93

> 93

> 93

> 93

> 93

0.068 0.002

> 59

a

> 59

0.061 0.022

t1/2 (d)

ref.

t1/2 (d)

ref.

oxic

0.4-24; 10

29; 18

5-9

39

anoxic

0.32

29

10-21; 41-49

39; 54

82  9

Kd

propargyl alcohol t1/2 (d) Kd

> 59

t1/2 (d) 0.24-13.3; 4-24; 10-35 8-48

0.041 0.004

> 57 > 57

0.044 0.022

> 63

> 66

ref.

t1/2 (d)

ref.

t1/2 (d)

ref.

t1/2 (d)

ref.

61; 59; 62

13

56

6-13; 8-14

58; 57

4-15; 4-39

40; 39

59

na c

30; 30 - 284

51; 39

na

711

a

712

residence time (columns) or length of microcosm experiments. Due to small differences in

713

retardation, column residence time varied slightly between compounds.

714

b

Half-lives reported or estimated from published experimental data.

715

c

Kinetic data not available (na) in literature.

When

Inhibition of Biodegradation of Hydraulic Fracturing Compounds by

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