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Organic carbon amendments for enhanced biological attenuation of trace organic contaminants in biochar-amended stormwater biofilters Bridget A. Ulrich, Marta Vignola, Katelynn Edgehouse, David Werner, and Christopher P. Higgins Environ. Sci. Technol., Just Accepted Manuscript • Publication Date (Web): 19 Jun 2017 Downloaded from http://pubs.acs.org on June 19, 2017

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Organic carbon amendments for enhanced biological attenuation of trace organic contaminants in biochar-amended stormwater biofilters Bridget A. Ulrich,† Marta Vignola,§ Katelynn Edgehouse,‡ David Werner,§ and Christopher P. Higgins†,* †

Department of Civil and Environmental Engineering Colorado School of Mines, Golden, CO §

School of Civil Engineering and Geosciences Newcastle University, Newcastle upon Tyne, United Kingdom ‡

Department of Chemistry Cleveland State University, Cleveland, Ohio Submitted to Environmental Science & Technology *To whom correspondence should be addressed. E-mail: [email protected] Phone: (720)-984-2116 Fax: (303) 273-3413

Number of pages: Number of figures: Number of tables: Number of words:

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ABSTRACT

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This study sought to evaluate how dissolved organic carbon (DOC) affects attenuation of

38

trace organic contaminants (TOrCs) in biochar-amended stormwater biofilters. It was

39

hypothesized that (1) DOC-augmented runoff would demonstrate enhanced TOrC

40

biodegradation, and (2) biochar-amended sand bearing DOC-cultivated biofilms would

41

achieve enhanced TOrC attenuation due to sorptive retention and biodegradation.

42

Microcosm and column experiments were conducted; utilizing actual runoff, DOC from

43

straw and compost, and a suite of TOrCs. Biodegradation of TOrCs in runoff was more

44

enhanced by compost DOC than straw DOC (particularly for atrazine, prometon,

45

benzotriazole, and fipronil). 16S rRNA gene quantification and sequencing revealed that

46

growth-induced microbial community changes were, amongst replicates, most consistent

47

for compost-augmented microcosms, and least consistent for raw runoff microcosms.

48

Compost DOC most robustly enhanced utilization of TOrCs as carbon substrates,

49

possibly due to higher residual nutrient levels upon TOrC exposure. Sand columns

50

containing just 0.5 wt% biochar maintained sorptive TOrC retention in the presence of

51

compost-DOC-cultivated biofilms, and TOrC removal was further enhanced by

52

biological activity. Overall, these results suggest that co-amendment with biochar and

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compost may robustly enhance TOrC attenuation in stormwater biofilters; a finding of

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significance for efforts to mitigate the impacts of runoff on water quality.

55

INTRODUCTION

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Stormwater runoff has degraded urban water quality by transporting contaminants

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to receiving waters.1 Low Impact Development (LID) systems have gained popularity

58

because they remove many conventional contaminants during infiltration (e.g., metals


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and petroleum hydrocarbons).2,3 However, removal of trace organic contaminants

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(TOrCs) in LID systems is poorly understood, despite their broad presence in urban

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runoff.4 In particular, many toxic stormwater TOrCs, such as urban-use pesticides (e.g.,

62

phenylurea and triazine herbicides, phenylpyrazole insecticides),5–7 flame retardants (e.g.,

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organophosphates),8 and chemicals from vehicle fluids (e.g., benzotriazoles),9 are also

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relatively mobile; allowing them to pass through LID systems.10

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Amendment of LID systems with biochar has shown promise for improved

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removal of metals, bacteria, nutrients, and TOrCs.11–15 In particular, a recent study

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revealed remarkable TOrC removal in intermittently dosed, biochar-amended infiltration

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columns: >99% TOrC removal from spiked natural water was maintained throughout

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cumulative treatment of an annual treatment volume estimated for the Denver area.16

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However, it remains unclear if sorptive TOrC retention can be maintained throughout an

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entire system lifetime (i.e., more than 15 years, as limited by accumulation of non-

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degradable metals).17 It would therefore be desirable to stimulate biodegradation to

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reduce potential accumulation, as has been demonstrated previously for conventional

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stormwater biofilters.18 These systems contain an upper layer amended with organic

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carbon (e.g., compost or straw), stimulating biological activity by slowly releasing

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growth substrates (e.g., dissolved organic carbon, DOC; and nutrients such as nitrogen,

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N, and phosphorus, P). As this may also cause undesirable sorbent fouling and reduced

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TOrC sorptive retention,19 it is of interest to determine if DOC can allow enhanced

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biological TOrC attenuation despite this potential tradeoff.

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While several studies have evaluated the effects of DOC on microbial

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communities and TOrC biodegradation in wastewaters and surface waters,20–22 it remains


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unclear how the underlying phenomena will manifest in stormwater biofilters. TOrC

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biodegradation may be affected by increased overall microbial abundances, as well as

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growth-induced changes in microbial communities. For example, broader TOrC

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biodegradation potential has been demonstrated in wastewater communities with greater

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biodiversity (i.e., community richness and evenness),20 which has been linked to

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availability of ambient carbon and nitrogen.22 Further, DOC can affect TOrC-degrading

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activity by influencing biofilm formation and structure.23 DOC quality and composition

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may be drivers behind these phenomena. For example, easily utilizable carbon substrates

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may allow rapid microbial growth, while growth on more complex substrates may lead to

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broader substrate utilization potential.24 The carbon-to-nitrogen ratio (C:N) of leached

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DOC can also affect growth (and in turn TOrC utilization) by establishing either carbon-

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limited or nutrient-limited growth conditions.25,26 As DOC quality and composition can

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be controlled in biofilters through selection of organic carbon amendments, biofilters

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could potentially be designed to achieve optimal TOrC biodegradation.

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The objective of this study was to assess how DOC from organic carbon

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amendments affects TOrC attenuation in biochar-amended biofilters. Efforts were

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focused on biological processes, as relevant abiotic processes (i.e., sorptive TOrC

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retention in presence of biochar and DOC) were assessed in a previous study.14 It was

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hypothesized that (1) augmenting runoff with DOC would enhance TOrC biodegradation

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due to increased microbial growth, activity, and potentially growth-stimulated changes in

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community composition; and (2) biochar-amended media bearing DOC-cultivated

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biofilms would demonstrate enhanced TOrC attenuation due to a combination of TOrC

104

sorptive retention and biodegradation. The first hypothesis was assessed in microcosm


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experiments, and the second in column experiments. To maintain environmental

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relevance, these experiments utilized actual runoff to obtain representative microbial

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consortia, straw and woody plant-based compost to obtain DOC of varying quality, and a

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suite of TOrCs and transformation products (TPs) to represent compound classes that are

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likely to impact water quality (i.e., both toxic and mobile). The results from this study

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provided useful insight for the design of infiltration systems to mitigate impacts of runoff

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on receiving water quality.

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EXPERIMENTAL SECTION

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TOrC Analysis. The TOrC suite was selected in a previous study,4 and included atrazine,

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diuron, tris(3-chloro-2-propyl)phosphate (TCPP), tris(3-chloro-ethyl)phosphate (TCEP),

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prometon, benzotriazoles (1H-benzotriazole in microcosm experiments, 5-methyl-

116

benzotriazole in column experiments), fipronil, oryzalin, and 2,4-dichlorophenoxyacetic

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acid (2,4-D results are reported in the Supporting Information, because it behaved

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unusually relative to the other analyzed TOrCs for reasons that remain unclear).

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Transformation

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desethylatrazine, DEA; deisopropylatrazine, DIA), diuron (1-(3,4-dichlorophenyl)-3-

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methylurea, DCPMU; 3,4-dichloroaniline, DCA), and fipronil (fipronil sulfone and

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sulfide). TOrCs were measured by liquid chromatography tandem mass spectrometry

123

(LC-MS/MS), and ionized by electrospray ionization (ESI) in the positive or negative

124

mode (0.025 μg/L limit of quantitation, details in Method S1). Optima™-grade methanol

125

and water (Fischer Scientific) were used unless noted otherwise.

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DOC Extracts. Concentrated DOC extracts (used to augment DOC levels in

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microcosms) were prepared in runoff that was collected following three separate rain

products

(TPs)

were

analyzed

for


atrazine

(hydroxyatrazine;

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events, such that it was stored for less than 24 hours prior to use in microcosm, column,

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and DOC characterization experiments. Runoff was collected in sterilized glass jugs from

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a pipe discharging runoff from a residential neighborhood in Golden, CO. Approximately

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500 mL of sediment (leaves, sticks, and silt) was added to each jug to introduce bacteria

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and DOC. The runoff was equilibrated at room temperature on a shaker table overnight

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and centrifuged for 5 minutes at 800 RCF to remove large particles. To enable rapid

134

production of concentrated DOC extracts (to allow augmentation of runoff with DOC

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simulating leachate from rain events on the order of hours), runoff (2 L) containing either

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a woody plant-based compost (EcoGrow® compost, obtained from A1 Organics; Eaton,

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CO, 500 mL) or wheat straw (Longmont, CO; 2L, packed) was incubated at 30 °C on a

138

shaker table overnight in the dark (such that the leached DOC was minimally affected by

139

exposure to UV radiation).27 Extracts were filtered to 0.45 μm (such that minimal residual

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bacteria remained prior to subsequent sterilization by autoclave), analyzed with a

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Shimadzu Total Organic Carbon Analyzer to determine DOC and total nitrogen (TN),

142

then diluted with filtered runoff to adjust the concentration as necessary.

143

Fresh DOC extracts were analyzed by ultra-violet visible (UV-vis) spectroscopy

144

(Figure S1) from which specific UV absorption was determined (SUVA, absorption at

145

254 nm; listed in Table S4 with additionally calculated absorption parameters). SUVA is

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an indicator of DOC aromaticity,28 and negative associations between SUVA and DOC

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biodegradability have been previously reported for soil leachates.29 SUVA was higher for

148

the compost DOC (3.0 L·mgC-1·m-1 for compost DOC, versus 0.7 L·mgC-1·m-1 for straw

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DOC), likely because the compost was derived from lignin-rich, woody material.30 The

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C:N ratios for fresh compost and straw extracts were 8.3 and 15.8, suggesting that they


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provided carbon-limited and nutrient-limited growth conditions, respectively (considering

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an optimal C:N of 10).31 Higher residual (post-incubation) DOC levels in compost

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microcosms suggested that the compost DOC was less biodegradable (52% and 89%

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DOC biodegraded in compost and straw microcosms, respectively; Table S5). Lower

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residual levels of P (Table S6) and N (Table S5) in straw microcosms indicated nutrient-

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limited growth conditions.

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Microcosm Experiments. Three types of aqueous microcosm media (raw runoff, and

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runoff augmented with DOC from compost or straw) were used to prepare eight biotic

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microcosms (five monitored for TOrCs, three sacrificed for additional analyses) and five

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biologically inhibited controls. Erlenmeyer flasks (150 mL) were filled with 5 g sand

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(Sigma Aldrich) and 10 mL DOC extract or filtered raw runoff, then autoclaved. The

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microbial consortium was introduced with 40 mL of collected runoff, such that the DOC

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prior to incubation (and subsequent dilution) was 205 mg/L in straw and compost

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microcosms (6 mg/L DOC in raw runoff). Microcosms were aerobically incubated for 14

165

days in the dark at 30 °C on a shaker table to allow for microbial growth. Reduced

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dissolved oxygen (DO, Hach® DO probe) consumption in sacrificed microcosms

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indicated stabilized biological activity after this duration. Microcosms were then diluted

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to 125 mL with autoclaved synthetic stormwater (composition in Table S3), and a sodium

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azide (NaN3) solution (5 mL of 4 g/L in DI water, 154 mg/L final concentration) was

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added to control microcosms to inhibit biological activity (biotic microcosms diluted with

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5 mL autoclaved DI water). These dilutions allowed sufficient volume for numerous

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sampling events, and reduced the DOC to more representative levels (i.e., less than 50

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mg/L, Table S5).


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TOrCs were introduced by spiking microcosms with 52 μL of a methanol carrier

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solution (time 0), such that initial individual TOrC concentrations (40 μg/L) permitted

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small sample volumes without exceeding environmentally relevant levels.4 Spiked

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microcosms were shaken on a shaker table at room temperature and sampled over 76

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days (daily to weekly sampling). Prior to each sampling event, autoclaved DI water was

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added to correct for evaporation losses (individual volumes determined gravimetrically to

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avoid dilution). Aliquots (250 μL) were transferred with sterile pipette tips to 2 mL

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microcentrifuge tubes containing 200 μL internal standard solution (4 μg/L isotope

182

surrogates in methanol) and 1550 μL of Milli-Q™ water containing 500 mg/L sodium

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azide to prevent biodegradation during storage (4°C, less than 30 days). Solid phase

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extraction (SPE) was carried out immediately prior to LC-MS/MS analysis using

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Phenomenex Strata®-X 33u Polymeric Reversed Phase cartridges (30mg/1mL) following

186

the manufacturer’s recommended protocol. Nitrate (Nitrate TNT Plus Hach Kit), optical

187

density (OD, absorbance at 600 nm), and DO were also measured on 3 – 4 occasions

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(Figure S2).

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Column Experiments. Columns (Kimble Chase Economy FLEX-COLUMN® 6 cm

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length, 2.5 cm I.D.) were dry-packed with sand (Sigma Aldrich, 210 – 297 μm particles,

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column diameter: particle size greater than 50 to prevent boundary effects),32 or sand with

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0.5 wt% biochar (53 – 250 μM particle sizes, Milli-Q™-rinsed), and porosity was

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estimated gravimetrically after saturation with water (0.35 ± 0.1 for biochar columns,

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0.32 ± 0.1 for sand columns). Mountain Crest Gardens biochar (MCG-biochar, produced

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by gasification of pinewood by Gropro, Inc., in Etna, CA; material properties in Table

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S7) was selected for its high surface area (317 m2/g) and proven TOrC retention


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performance.14 While higher biochar doses may be desirable in full-scale systems (e.g.,

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greater than 5 wt%),16 a low dose was necessary to allow TOrC breakthrough within a

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reasonable experimental time frame. Batch sorption experiments with the DOC extract

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and biochar revealed that the biochar had similar sorption affinities for both types of

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DOC (Method S2, log Kd,DOC [L/kg] = 3.2).

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Biofilms were seeded on columns with concentrated solutions prepared similarly

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to the microcosm media. Over three cycles, undiluted, autoclaved straw and compost

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DOC extracts (>1000 mg/L DOC) were inoculated (either with runoff or the previously

205

incubated culture) and incubated until increases in OD indicated exponential growth

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(Method S3, growth curves in Figure S4). After flushing with CO2 to ensure saturation,

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columns were injected with one pore volume (PV, approximately 10 mL) of seeding

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solution at 1 mL/min in upflow, and then slowly cycled for 24 hours (0.05 mL/min, 0.6

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cm/hr linear velocity) to allow microbial attachment. Columns were then equilibrated

210

with synthetic stormwater containing 10 mg/L DOC at 0.21 mL/min (2.6 cm/hr linear

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velocity) for 10 days (influent reservoirs refreshed every 2-3 days) until effluent DO

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levels stabilized (Figure S5). Tracer tests using KBr (10 g/L) as a conservative tracer and

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a Hach® conductivity meter showed that hydraulic retention times were minimally

214

affected by biofilm formation (i.e., breakthrough less than 10 minutes before the

215

predicted 2.3 hour hydraulic retention time).

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Columns were then injected with synthetic stormwater containing 10 mg/L DOC

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and 10 μg/L TOrCs (methanol from TOrC carrier solution evaporated under N2) for 23

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days (675 PVs) and 18 days (528 PVs), respectively (injection times differed due to

219

slower microbial growth in the compost DOC seeding solution, Figure S3). Influents


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were refreshed every 24 hours to minimize TOrC biodegradation within the reservoir.

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NaN3 (400 mg/L) was added to influents for control sand-biochar columns (one control

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and three biotic replicates per DOC type). Following TOrC injection, columns were

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flushed with TOrC-free synthetic stormwater for the remainder of the experiment.

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Effluent samples (collected every 1 – 100 PVs over the total 1100 PVs) were preserved

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by diluting 10% with 4 g/L NaN3 (biotic columns only) and stored at 4°C in the dark.

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Isotope dilution was carried out immediately prior to LC-MS/MS analysis (ESI positive

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mode only due to constraints on sample analysis).

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Microbial Analyses. Microbial community composition and structure were investigated

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by high-throughput sequencing (pyrosequencing). Due to constraints on sample analysis,

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DNA extraction and sequencing was carried out for 3 representative biotic microcosms

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from each condition (9 total), and the column seeding solutions (Figure S4). The subset

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of microcosms chosen for microbial sequencing analysis included, from each condition,

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two TOrC-monitored microcosms (Figure S9; samples C1, C5, S1, S2, R1, and R5) and

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one unmonitored microcosm (samples C6, S6, and R6), such that community differences

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due to the TOrC sampling process could be identified if present. Immediately following

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the final sampling event, the sand and remaining aqueous media were centrifuged in

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sterile 50 mL Falcon tubes at 3220 RCF for 60 min. After decanting the liquid, the top

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500 mg of sand (wet weight) was collected for DNA extraction. DNA was extracted in

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duplicate using a PowerSoil DNA Isolation Kit (MO Bio Laboratories, Inc.), according to

240

the manufacturer’s instructions (stored at -80 °C until subsequent analysis).

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Polymerase Chain Reaction (PCR) amplification of the V4-V5 region of bacterial

242

16S rRNA genes for each DNA extract was conducted using fusion primers containing a


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PGM sequencing adaptor, a ‘GT’ spacer, and a unique 12 base pair Golay barcode to

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allow multiplex analyses (primers 515F: 5’-GTGNCAGCMGCCGCGGTAA-3’, and

245

926R: 5’-CCGYCAATTYMTTTRAGTTT-3’).33 PCR reactions were conducted using

246

the FastStart High Fidelity PCR System and the PCR Nucleotide Mix (Roche Diagnostics

247

GmbH), with the following thermocycle program: (i) 2 minutes denaturation at 95°C, (ii)

248

30 cycles of 30 minutes denaturation at 95°C, (iii) 30 second annealing at 56°C, (iv) 45

249

minutes elongation at 72°C, and (v) 7 minutes elongation at 72°C. PCR products were

250

purified using Agencourt AMPure XP reagent (Beckman Coulter) and quantified using a

251

Qubit dsDNA HS Assay kit (Invitrogen) on a Qubit® 2.0 Fluorometer. The single PCR

252

libraries were pooled in equimolar amounts, further purified using a Pippin Prep System

253

(Life Technologies) following the manufacturer’s protocol. Quantitative PCR (qPCR) of

254

overall bacterial 16S rRNA genes was also conducted to qualitatively compare gene copy

255

numbers among samples (Method S4).

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Sequencing was carried out with an Ion Torrent Personal Genome Machine®

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(PGM) System (Life Technology) at Newcastle University. A total of 1,863,582

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sequences were obtained and processed in QIIME (v 1.7.0) using the bioinformatics

259

pipeline. The sequences were trimmed to remove primers and barcodes, quality filtered

260

(considering a minimum quality score of 20) and chimera checked (ChimeraSlayer)

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resulting in 739,973 total sequences. Sequences were then clustered into Operational

262

Taxonomic Unit (OTU) at 97% sequence similarity level by the uclust algorithm, and a

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representative sequence from each OTU was selected and taxonomically identified using

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the Greengenes database34 and classified using the RDP naïve Bayesian rRNA

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classifier.35–37 Representative sequences and correspondent taxonomic assignment were


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used to build a table of OTU abundances at different levels of taxonomy. Microbial

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community diversity within and across the samples was determined using the QIIME

268

(v.1.7.0) pipeline and Primer v6 software.38,39 The Bray Curtis similarity metric was

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generated and represented by a 2-dimensional non-metric multidimensional scaling

270

(nMDS) plot.39 Additionally, an analysis of similarities (ANOSIM) was conducted on the

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Bray Curtis dissimilarity matrix (square root transformed) using PRIMER v6. Statistical

272

analysis was performed in R on the alpha diversity indices (Shannon index).

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RESULTS AND DISCUSSION

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TOrC biodegradation in microcosms. Biodegradation of most monitored TOrCs was

275

observed under at least one experimental condition, and generally occurred consistently

276

across replicates (Figures 1 and S6). Oryzalin and benzotriazole were biodegraded the

277

most quickly; while atrazine, prometon, fipronil, and diuron were biodegraded more

278

slowly. TCPP and TCEP were generally resistant to biodegradation, with the exception

279

that a single biotic raw runoff microcosm showed sudden and rapid degradation of both

280

compounds after 30 days (henceforth referenced as the ‘outlier’ microcosm).

281

Less than 20% removal was observed in the biologically inhibited control

282

microcosms for most TOrCs (Figure 1), suggesting that the effects of dilution and

283

reaction with NaN3 were minor relative to biodegradation. Removal of oryzalin and

284

benzotriazole in controls was greater than the other TOrCs, though still less than their

285

removal in biotic microcosms. This was potentially due to incomplete activation of

286

biological activity by NaN3; considering that these TOrCs were the most strongly

287

biodegraded overall, and reduced DO levels in DOC-augmented controls relative to raw

288

runoff controls suggested that microbial activity persisted to some degree despite the


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presence of NaN3 (Figure S2). The extent of biodegradation at the end of the experiment

290

was therefore assessed relative to the raw runoff controls (calculated extents of

291

biodegradation reported as fractions and μmols in Figures S8 and S9, respectively).

292

Interestingly, the outlier raw runoff microcosm demonstrated the greatest overall TOrC

293

biodegradation among the individual microcosms (i.e., highest sum of all monitored

294

TOrCs biodegraded, Figure S9), and is henceforth considered separately from the raw

295

runoff microcosms unless noted otherwise. Compost microcosms consistently

296

demonstrated the most overall TOrC biodegradation across experimental replicates, while

297

straw microcosms overall were not significantly different from raw runoff microcosms (p

298

values from the two-tailed student t test for comparison of raw runoff microcosms to

299

compost and straw microcosms were 0.0011 and 0.20, respectively; Table S8). The

300

compost microcosms demonstrated enhanced biodegradation of fipronil, benzotriazole,

301

atrazine, and prometon (p values = 0.0069, 0.0017, 0.0043, and 0.023, respectively, for

302

comparison to raw runoff); while straw microcosms demonstrated slightly enhanced

303

biodegradation of fipronil and benzotriazole (p = 0.0031 and 0.017, respectively, for

304

comparison to raw runoff).

305

In general, the most abundant monitored TPs for fipronil, diuron, and atrazine

306

were fipronil sulfone (Figure 1C), DCPMU (Figure 1D), and desethylatrazine (Figure

307

1E), respectively (additional TPs in Figure S7); except for the outlier microcosm, for

308

which the most abundant fipronil TP was fipronil sulfide. To determine if unmonitored

309

TPs were generated, mass balance errors were calculated according to differences in

310

concentration between (1) each parent TOrC in raw runoff controls, and (2) the sum of

311

each parent TOrC and their monitored TPs in biotic microcosms (Table S9). Atrazine and


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diuron showed low mass balance errors for the first 60 days, after which errors exceeded

313

10%. This coincided with the peak concentrations of the measured primary TPs,

314

suggesting biodegradation of primary TPs to unmonitored secondary TPs. In contrast,

315

fipronil mass balance errors exceeded 10% after just 13 days, suggesting that

316

unmonitored primary TPs may have been generated.

317

Trends in lag time (delays before biodegradation) across experimental replicates

318

provided insight into biodegradation mechanisms. For example, oryzalin, benzotriazole,

319

and fipronil were biodegraded in all DOC-augmented microcosms without a lag time;

320

suggesting biodegradation by co-metabolic processes (i.e., degradation by constitutively

321

expressed, non-specific enzymes). Atrazine and prometon, in contrast, were only

322

biodegraded in compost microcosms following a 30-day lag time. Depletion of

323

preferential carbon substrates may have caused this lag time, subsequently allowing

324

TOrC biodegradation due to adaptive expression of TOrC-degrading enzymes40,41 or

325

reduced substrate-substrate inhibitions.42 The greater aromatic character of the compost

326

DOC may have contributed to this behavior, as these more complex carbon substrates

327

may have been more slowly utilized and hence still present upon TOrC exposure. TCPP

328

and TCEP biodegradation in the outlier microcosm was particularly abrupt after the 30-

329

day lag time, potentially due to the sudden activity of specialist bacteria. Decreased DO

330

(Figure S2A), increased OD (Figure S2B), and reduced DOC (Table S5) relative to raw

331

runoff microcosms indicate accelerated growth in this microcosm following TOrC

332

addition, potentially due to methanol exposure (discussed further below).

333

Microcosm microbial communities. The 2D nMDS analysis showed that the microbial

334

communities in all DOC-augmented microcosms (i.e., both straw and compost


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microcosms) clustered apart from those in raw runoff microcosms (Figure 2A), indicating

336

DOC-stimulated, growth-induced changes in community structure. The communities of

337

the DOC-augmented microcosms were relatively similar to each other (58% Bray Curtis

338

similarity overall, see dendrogram in Figure S10), while compost DOC communities

339

showed the highest similarity among replicates (73%). As communities were more

340

similar among the DOC-augmented microcosms, the more significantly enhanced TOrC

341

biodegradation in compost microcosms relative to straw microcosms (as compared to the

342

raw runoff microcosms) was likely due to either subtle differences in communities (e.g.,

343

differences in relative abundances of specific OTUs), and/or other factors relating to

344

microbial abundance and activity. The outlier microcosm did not cluster with the other

345

microcosms, suggesting that microbial community differences caused its unique

346

biodegradation behavior. It is important to note that growth on methanol from the TOrC

347

carrier solution may have also affected communities. It is possible that this masked

348

community differences that had already developed during incubation, potentially

349

increasing the similarity among the DOC-augmented microcosm communities. For the

350

outlier microcosm, it is possible that growth of methanol-utilizing bacteria was stimulated

351

in the absence of other DOC sources, potentially contributing to its unique community

352

structure.

353

Gene copy numbers (GCNs) of overall bacterial 16S rRNA genes were

354

consistently high for compost microcosms, while they were lower and more variable for

355

raw runoff and straw microcosms (Figure 2B). Considering that low residual DOC levels

356

(Table S5) suggest that growth occurred in all DOC-augmented microcosms, the higher

357

GCNs in compost microcosms may have been due to greater abundances of active


15

Page 17 of 35


358

bacteria upon DNA extraction (at the end of the experiment). GCNs were higher in the

359

outlier microcosm than the raw runoff microcosms, further indicating accelerated growth

360

in this microcosm. Only small differences in the Shannon index were observed among the

361

microcosms (indicative of community richness and evenness, Table S10), therefore alpha

362

diversity likely had only minor effects on observed trends. This is unsurprising, as the

363

inoculum effectively fixed the species richness across microcosms (i.e., the same number

364

of species were introduced into all microcosms upon inoculation). While averaged

365

Shannon indices were higher for compost and straw microcosms than for raw runoff

366

microcosms (3.6 and 3.4, versus 3.2, respectively), differences were not statistically

367

significant (p = 0.072 and 0.66, respectively, for comparison to raw runoff microcosms).

368

Notably, an especially low Shannon index was observed for one straw microcosm (2.95,

369

microcosm S2, discussed below).

370

Figure 3 shows the most prevalent differences in OTU relative abundances that

371

were identified among the microcosms (family-level OTU bar plot in Figure S11).

372

Several OTUs that are known to facilitate nitrogen-cycling processes were enriched in

373

DOC-augmented microcosms relative to raw runoff; including those within the families

374

Rhodospirlliceae

375

Comamonadaceae (Comamonas; denitrification),45 and Rhodocyclaceae (Thauera;

376

denitrification).46 This reflects previous findings of coincidentally enhanced nitrogen-

377

cycling processes and TOrC biodegradation among wastewater communities.21,47 An

378

OTU from the genus Pseudomonas was enriched only in straw microcosms (particularly

379

microcosm S2, causing its low Shannon index; Figure S11). Though many Pseudomonas

380

species are known TOrC-degraders,48 the OTU's abundance was not positively associated

(N2-fixation),43

Nitrospirae

(Nitrospira;


nitrite-oxidation),44

16


Page 18 of 35

381

with individual or overall TOrC biodegradation (Spearman’s rank and Pearson

382

correlation tests described in Method S5, Tables S11 – S14). As Pseudomonas can

383

undergo metabolic regulation,49 this OTU may have initially out-competed other bacteria

384

for utilization of the straw-derived carbon substrates, potentially until residual levels of P

385

and N could no longer support growth (Tables S5 and S6). This potentially explains the

386

reduced TOrC biodegradation in straw microcosms relative to compost microcosms, as

387

microbial activity and hence TOrC biodegradation would presumably be diminished if

388

nutrients were depleted prior to TOrC introduction. Interestingly, the outlier microcosm

389

was enriched with OTUs within the family Sphingomonadaceae (markers in Figure 3),

390

which contains several genera of TOrC-degrading bacteria (e.g., Sphingomonas).50,51

391

While these OTUs could potentially be specialist TCEP and TCPP degrading bacteria,

392

they may also just covary with such bacteria.

393

Overall, while DOC-stimulated growth led to communities that were starkly

394

different from raw runoff communities, community differences among the DOC-

395

augmented microcosms were more subtle. However, the similarity in both growth and

396

community composition amongst replicates was the highest for the compost microcosms,

397

coinciding with the greatest overall TOrC biodegradation. One possible explanation for

398

this behavior is that the compost microcosms provided conditions that more robustly

399

promoted TOrC-degrading activity upon TOrC exposure. For example, higher residual N

400

and P levels may have facilitated the utilization of TOrCs as carbon sources, potentially

401

allowing sustained microbial activity throughout the experiment. These conditions may

402

have arisen due to several factors relating to DOC composition and quality, or a

403

combination thereof; including the initial C:N of the DOC (i.e., providing carbon-limited


17

Page 19 of 35


404

rather than nutrient-limited growth conditions), as well as effects of DOC aromaticity on

405

substrate utilization potential. In particular, the higher aromaticity of the compost DOC

406

may have led to adaptive expression of TOrC-degrading enzymes. This is further

407

supported by the biodegradation lag times observed for atrazine and prometon, which

408

also showed the most significantly enhanced biodegradation relative to straw microcosms

409

(p = 0.00077 and 0.00067, respectively, Table S8).

410

While the underlying phenomena behind the observed TOrC biodegradation

411

trends cannot be fully elucidated, these findings indicate that compost DOC may indeed

412

support TOrC-degrading microbial activity in stormwater biofilters more effectively than

413

straw DOC. However, the observed time-scale for TOrC biodegradation (weeks to

414

months), is much higher than hydraulic residence times for distributed infiltration

415

systems (hours to days). In this respect, biochar-amended biofilters may be especially

416

effective if sorptive retention is maintained, as increased TOrC retention times may allow

417

more time for biodegradation. The column experiments provided further insight into

418

these questions.

419

Column Experiments. Sand-only columns cultivated with compost DOC

420

demonstrated longer TOrC retention times than those cultivated with straw DOC (i.e.,

421

TOrC breakthrough was observed after approximately 100 PVs in sand-only compost

422

columns, versus less than 10 PVs for sand-only straw columns; Figures 4 and S12).

423

Compost columns also demonstrated greater microbial activity during TOrC injection and

424

flushing, as indicated by lower effluent DO levels (Figure S5). Notably, TOrC retention

425

times were enhanced by amendment of sand with just 0.5 wt% biochar (i.e., breakthrough

426

occurred approximately 100 PVs later in biochar-sand columns than in sand-only


18


Page 20 of 35

427

columns), despite the severe fouling expected to be caused by the concentrated seeding

428

solutions (with DOC concentrations potentially orders of magnitude above representative

429

levels). It is therefore likely that TOrC retention can be maintained in actual biofilters,

430

which presumably contain media with significantly higher biochar compositions (i.e.,

431

greater than 5 wt%).16

432

In addition to higher TOrC retention times, compost columns also demonstrated

433

greater percent removal of most TOrCs (i.e., lower recovery at the end of the experiment

434

as indicated by a Method of Moments analysis; Figure S13). This appeared to be at least

435

partially due to biological processes, as inhibition of biological activity with NaN3

436

reduced removal of atrazine by 20%, as well as TCEP and TCPP to a lesser degree

437

(though slowly receding concentrations during flushing may have slightly affected

438

recovery calculations). This could potentially indicate TOrC biodegradation within

439

columns (i.e., 20% greater atrazine removal in the absence of NaN3 potentially indicates

440

up to 20% biodegradation). This is an interesting result, as conventional porous media

441

transport theory suggests that only the dissolved TOrC fraction is bioavailable, and that

442

TOrCs are therefore biodegraded on the time-scale of the hydraulic retention time.52 Thus

443

biodegradation would be unlikely to amount to observable levels within the columns

444

unless biodegradation were particularly “rapid” (as the hydraulic retention time in

445

columns was approximately 2 hours).

446

Interestingly,

atrazine

retention

times

were

comparable

to

expected

447

biodegradation time-scales (i.e., 10-20 days) and appeared to be associated with atrazine

448

removal (i.e., biochar-sand columns demonstrated both atrazine retention times and

449

atrazine removal percentages that were nearly double those for sand-only columns).


19

Page 21 of 35


450

While a possible explanation for this behavior is the biodegradation of TOrCs sorbed to

451

biochar (which has been observed for some cases),53 this apparently contradicts

452

conventional transport theories. A more feasible explanation may be biodegradation of

453

TOrCs loosely associated with the immobilized DOC and the biofilm itself. Localized

454

biodegradation of these more readily desorbed TOrCs would continuously replenish

455

sorption sites, making it appear that sorbed TOrCs were biodegraded. Moreover, the

456

biofilm structure (potentially affected by DOC quality) could also affect TOrC

457

biodegradation (e.g., due to beneficial co-settlement of TOrC-degrading bacteria).23

458

Overall, the column experiments revealed that TOrC sorptive retention was

459

maintained by biochar-amended media bearing DOC-cultivated biofilms, and that overall

460

TOrC attenuation was improved in the presence of biological activity stimulated by

461

compost DOC. As it remains unclear if TOrCs were actually biodegraded within

462

columns, studies evaluating systems with longer TOrC retention times may provide

463

insight into whether TOrCs can be biodegraded significantly within biochar-amended

464

biofilters.

465

Environmental Implications. The findings of this study are of particular

466

significance for efforts to mitigate impacts of runoff on water quality through

467

optimization of contaminant removal in infiltration systems. In particular, in addition to

468

improving TOrC attenuation, co-amendment of biofilters with compost and biochar may

469

comprehensively enhance performance by mitigating undesirable leaching of metals and

470

nutrients from compost;54–56 making vegetated systems (which require compost for plant

471

growth) more attractive. Vegetated systems may be further advantageous for improving

472

TOrC attenuation: biological activity stimulated in the root zone may further improve


20


Page 22 of 35

473

TOrC biodegradation,57 while enhanced desorption in the presence of root exudates may

474

prevent TOrC accumulation on biochar.58,59 Remarkable TOrC removal has indeed been

475

observed in vegetated infiltration columns representing biochar-amended biofilters;16 yet

476

as no TOrC breakthrough was observed, it remains unclear whether TOrC removal is due

477

to sorption alone, or to a combination of sorption and biodegradation. Therefore,

478

additional studies seeking to identify and quantify TOrC biodegradation in vegetated,

479

biochar-amended biofilters are of interest to determine if TOrC accumulation can be

480

prevented throughout the system entire lifetime.

481

Supporting information. This material is available free of charge via the Internet at

482

http://pubs.acs.org, and contains additional figures and tables and methodological details,

483

as referenced throughout the text. The sequencing data has been made available to the

484

public through the EMBL European Bioinformatics Institute (Project PRJEB19822,

485

http://www.ebi.ac.uk/).

486

Author information

487

Corresponding author. Christopher P. Higgins.

488

E-mail: [email protected]

489

Phone: (720)-984-2116

490

Fax: (303) 273-3413

491

Acknowledgements.

492

Science Foundation Graduate Research Fellowship under Grant No. DGE-1057607 and

493

the National Science Foundation Engineering Research Center Program under

494

Cooperative Agreements EEC-1028968 (ReNUWIt) and EEC-1262655. Collaboration

This material is based upon work supported by the National


21

Page 23 of 35


495

with Newcastle University was facilitated by EPSRC grant EP/K021737/1 and Marie

496

Skłodowska-Curie Initial Training Network (MERMAID) grant 607492.

497

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organic compounds with varying polarity and planarity. Environ. Sci. Technol.

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FIGURES


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Figure 1. Concentrations of (A) oryzalin, (B) benzotriazole, (C) fipronil, (D) diuron, (E) atrazine, and (F) TCEP in microcosms (prometon and TCPP are shown in Figure S6). Error bars indicate standard deviations for replicate microcosms, and significant differences between inhibited controls and biotic microcosms are assumed to be due to biodegradation. Select transformation products (TPs) are also shown for atrazine (desethylatrazine), fipronil (fipronil sulfone and fipronil sulfide), and diuron (3,4dichloromethylphenyl urea, DCPMU). Additional TPs are shown in Figure S7. ACS Paragon Plus Environment

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Figure 2. (A) Two-dimensional non-metric multidimensional scaling (2D nMDS) plot of sequenced microcosm microbial communities (markers) and Bray Curtis similarities among samples (lines). Similarities are also shown in the dendrogram in Figure S11 (similarity analysis conducted using PRIMER v6). (B) Gene copy numbers per mass of DNA-extracted sand for each sequenced sample, determined by qPCR of total bacterial 16S rRNA genes (flocculation was observed in sample S1, likely causing its elevated gene copy numbers). Error bars indicate the standard deviation of triplicate measurements.


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Figure 3. Relative abundances of OTUs that were (i) identified at the genus level, and (ii) observed to be enriched by at least one order of magnitude relative to raw runoff (note the log-scale of the y axis). Bars show averages for replicate microcosms, with error bars showing the standard deviations. Markers show relative abundances for the outlier microcosm (not included in the averages for the raw runoff microcosms). Comparisons were made between averaged relative abundances, with the exception of the outlier microcosm, for which its individual values were compared to the averaged values for the other raw runoff microcosms.


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Figure 4. Breakthrough of (A, B) atrazine, (C, D) methylbenzotriazole, and (E, F) TCPP in sand-only and sand-biochar columns (0.5 wt% biochar), with biofilms cultivated from compost DOC (green) or straw DOC (yellow). Results for biologically inhibited control sand-biochar columns are shown by dotted lines. Columns were injected with 10 μg/L TOrCs, then flushed with TOrC-free synthetic stormwater. Diuron was exceptionally well removed overall, as has been observed previously.14,60


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Organic carbon amendments for enhanced ... - ACS Publications

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