UC Riverside Electronic Theses and Dissertations

UC Riverside UC Riverside Electronic Theses and Dissertations

Title The Fate, Metabolism and Toxicity of Select Contaminants of Emerging Concern in Terrestrial Environments

Permalink https://escholarship.org/uc/item/5nc806xq

Author Dudley, Stacia Lee

Publication Date 2019

Peer reviewed|Thesis/dissertation

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UNIVERSITY OF CALIFORNIA RIVERSIDE

The Fate, Metabolism and Toxicity of Select Contaminants of Emerging Concern in Terrestrial Environments

A Dissertation submitted in partial satisfaction of the requirements for the degree of

Doctor of Philosophy

Environmental Toxicology

Stacia L. Dudley

March 2019

Dissertation Committee: Dr. Jay Gan, Chairperson Dr. John Trumble Dr. David Volz

The Dissertation of Stacia L. Dudley is approved:

Committee Chairperson

University of California, Riverside

Acknowledgements

To my major professor, Dr. Jay Gan, thank you for your constant support, guidance and the countless hours spent helping to change me from a student into a scientist. I would also like to thank Dr. John Trumble, Dr. David Volz and Dr. Daniel Schlenk for welcoming me into their labs and for always being available to answer questions, both scientific and life-wise. A special thanks to the many members of the Gan Lab, particularly Dr. Qiuguo Fu, Dr. Chenliang Sun, Dr. Jei Wang, and Dr. Jaben Richards, for their time spent teaching me lab techniques, reviewing manuscripts, and hours spent helping me trouble- shoot. Thank you to my lab mates Douglas Wolf, Zachary Cryer and Allison Taylor for your friendship and constant willingness to lend me a hand, strong arm or second look. Thank you to my collaborators and friends Dr. Marcus Pennington, Dr. Luisa Becker-Bertotto, Sara Vliet, Dr. Scott Coffin, Marissa Giroux, and Michelle McGinnis for trusting me with your research and constantly inspiring me in my own work. Further, a big thank you to Jason Rothman for your aid with DNA extractions, bioinformatics and for not judging my microbial naiveté. Further, a big thank you to all my friends ranging from coast to coast, your texts, calls and visits have kept me sane. A special thanks to Sara Santos, Dr. Holly Mayton, Savannah Richards and Michelle McGinnis for nights/weekends of podcasts and fascinating conversation. Your brilliance constantly challenged me to think outside my comfort zone and your friendship has been invaluable. Finally, thank you to my brother Michael Dudley for his love and support throughout my entire life, no one could ask for a better big brother. To my mother Dean Caryn Beck-Dudley, you have been my role model and inspiration in my career and in my life. I would not be where I am today without your constant support, encouragement

and never-ending willingness to educate. Last, but certainly not least, I want to thank my husband Jonathan Pacha, you have always put my dreams and ambitions before your own. This is for you. These studies were supported in part by the following grants: U.S. Environmental Protection Agency STAR program (Award No: R835829) and the National Science Foundation- the Water SENSE Integrative Graduate Education Research Training program (No. DGE-1144635). Chapter 2 was previously published in the journal Environmental Pollution and Chapter 3 was previously published in the journal Science of the Total Environment.

Dedications

In Loving Memory of

Dr. Lynn Dudley

My Father and Mentor

ABSTRACT OF THE DISSERTATION

The Fate, Metabolism, and Toxicity of Select Contaminants of Emerging Concern in Terrestrial Environments

Stacia L. Dudley Doctor of Philosophy, Graduate Program in Environmental Toxicology University of California, Riverside, March 2019 Dr. Jay Gan, Chairperson

Increasing populations and shifting precipitation patterns put pressure on freshwater and food systems and incentivize governments and industries to exploit historically underutilized resources, such as recycled water and biosolids. Use of recycled water and biosolids in agricultural systems, however, comes with the potential risks of environmental and food contamination by trace organic contaminants including, contaminants of emerging concern (CECs). These compounds pose potential threats to humans and environmental health because they are designed to be biologically active at low concentrations and are considered “pseudo-persistent” due to their continuous release into the environment.

Using 14C tracing, mass spectrometry, stable-isotope labeling, and enzyme extractions to assess the fate, metabolism, and biological effects of four environmentally prevalent CECs (i.e. sulfamethoxazole, diazepam, naproxen and methyl paraben) in terrestrial organisms in hydroponic and artificial soil cultivations. These organisms included a model terrestrial invertebrate (Eisenia fetida), a model plant (Arabidopsis thaliana) and two crop plants (Cucumis sativus, Raphanus sativus,). Compounds were

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selected based on environmental prevalence and test organisms were selected due to their use in the literature, commercial availability, and global range.

Sulfamethoxazole was metabolized in A. thaliana cell cultures, and E. Fetida forming phase I and II metabolites. Diazepam was also metabolized in A. thaliana cell cultures and radish and cucumber seedlings, forming the phase I metabolites nordiazepam, temazepam and oxazepam, with the longevity corresponding to that of human metabolism. The major metabolites of naproxen and methyl paraben, O- desmethylnaproxen and p-hydroxybenzoic acid, respectively, were detected in treatment soils containing E. fetida and N4-acetylsulfamethoxazole was detected in E. fetida tissues, indicating CEC metabolism and excretion. Exposure to CECs resulted in changes to enzymes associated with detoxification and oxidative stress (i.e. glutathione-S- transerfase, glucuronosyl transferase, superoxide dismutase, and catalase) in crop plants and E. fetida. Our research indicates that terrestrial organisms can absorb and transform

CECs and that CECs can change biochemistry of the exposed organisms. Accordingly, it is crucial to consider CEC fate, transformation and effects on non-target organisms of

CECs when assessing risk in the agro-environment.

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Table of Contents

Chapter 1: Introduction ...... 1

1.1 Background ...... 1

1.2 Exposure and Fate of CECs in Plants ...... 3

1.2.1 Antibiotics ...... 4

1.2.2 Nonsteroidal Anti-Inflammatories ...... 6

1.2.3 Psychiatric Pharmaceuticals ...... 9

1.2.4 Antimicrobials and Preservatives ...... 11

1.2.5 Knowledge Gaps ...... 13

1.3 Phytotoxicity ...... 14

1.3.1 Antibiotics ...... 14

1.3.2 Nonsteroidal Anti-Inflammatories ...... 17

1.3.3 Psychiatric Pharmaceuticals ...... 18

1.3.4 Antimicrobials and Preservatives ...... 19

1.3.5 Mixtures ...... 20

1.3.5 Knowledge Gaps ...... 21

1.4 Exposure and Toxicity of CECs in Invertebrates ...... 21

1.4.1 Antibiotics ...... 22

1.4.2 Nonsteroidal Anti-Inflammatories ...... 23

1.4.3 Antimicrobials and Preservatives ...... 24

1.4.5 Mixtures ...... 25

1.4.5 Knowledge Gaps ...... 26

1.5 Problem Statement ...... 27

1.6 Research objectives ...... 27

Chapter 2: Metabolism of sulfamethoxazole in Arabidopsis thaliana cells and cucumber seedlings ...... 31

Abstract ...... 31

2.1 Introduction ...... 33

2.2 Materials and Methods ...... 36

2.2.1 Chemicals and solvents ...... 36

2.2.2 Arabidopsis thaliana Cell Incubation Experiment ...... 36

2.2.3 Whole Plant Hydroponic Cultivation Experiment ...... 39

2.2.4 Structural elucidation using mass spectrometry ...... 40

2.2.5 Data Analysis and Quality Control ...... 42

2.3 Results and Discussion ...... 43

2.3.1 Kinetics of parent, extractable and non-extractable residues ...... 43

2.3.2 Metabolism of Sulfamethoxazole in A. thaliana cells ...... 45

2.3.3 Uptake, Translocation and Transformation in Cucumber Seedlings ...... 48

2.4 Conclusion ...... 50

Supplementary Information ...... 61

Chapter 3: Formation of biologically active benzodiazepine metabolites in

Arabidopsis thaliana cell cultures and vegetable plants under hydroponic conditions ...... 69

Abstract ...... 69

3.1 Introduction ...... 71

3.2 Materials and Methods ...... 73

3.2.1 Chemicals and solvents ...... 73

3.2.3 Whole plant hydroponic cultivation ...... 76

3.2.4 Glycosyltransferase activity assay ...... 77

3.2.5 Structural elucidation using mass spectrometry ...... 78

3.2.6 Data analysis and quality control ...... 79

3.3 Results and discussion ...... 80

3.3.1 Kinetics of diazepam in A. thaliana cell cultures ...... 80

3.3.2 Metabolism of diazepam in A. thaliana cells ...... 80

3.3.3 Uptake and translocation of diazepam in cucumber and radish seedlings 82

3.3.4 Metabolism of diazepam in cucumber and radish seedlings ...... 84

3.3.5 Diazepam-induced changes to glycosyltransferase activity ...... 85

Supplementary Information ...... 94

Chapter 4: Bioaccumulation, metabolism and biochemical effects of contaminants of emerging concern in the earthworm species Eisenia fetida ...... 98

Abstract ...... 98

4.1 Introduction ...... 100

4.2 Materials and Methods ...... 103

4.2.1 Chemicals and solvents ...... 103

4.2.2 Test Soil ...... 103

4.2.3 Test Organism ...... 104

4.2.4 Bioaccumulation Experiment ...... 104

4.2.5 Assay of Antioxidant Enzyme Activities in E. fetida ...... 107

4.2.6 Assay of Lipid Peroxidation ...... 108

4.2.6 CEC Quantification using Mass Spectrometry ...... 108

4.2.7 Data analysis and quality control ...... 109

4.3 Results and Discussion ...... 110

4.3.1 Distributions of CECs among soil, porewater and earthworm ...... 110

4.3.2 Metabolism of CECs in E. fetida ...... 114

4.3.3 Effects of CECs on E. fetida antioxidant enzymes ...... 116

4.3.4 CEC-induced lipid peroxidation ...... 118

4.4 Conclusions ...... 118

Supplemental Information ...... 127

Chapter 5: Summary of Findings and Future Work ...... 134

5.1 Summary of Finding ...... 134

5.1.1 Metabolism of sulfamethoxazole in Arabidopsis thaliana cells and cucumber seedlings…………………………………………………………..133 5.1.2 Formation of biologically active benzodiazepine metabolites in Arabidopsis thaliana cell cultures and vegetable plants under hydroponic conditions…………………………………………………………………….134 5.1.3 Bioaccumulation, metabolism and biochemical effects of contaminants of emerging concern in earthworm Eisenia fetida …………………………..136 5.1.4 Overall conculsions……………………………………………………138

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5.1 Future Research ...... 139

References: ...... 139

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List of Tables

Table 1.1 Properties of contaminants of emerging concern and their major

metabolites ...... 31

Table 2.1 Summary of mass spectra information and proposed structures of

sulfamethoxazole and its metabolites ...... 52

Table 2.2 Detection of 14C-sulfamethoxazole in cucumber seedlings after 7 d

growth in nutrient solution ...... 54

Table 3.1 Extractable and non-extractable radioactivity in radish and cucumber

plants cultivated for 7d and 28d ...... 87

Table 4.1 Dissociation constants (Kd) for selected compounds in artificial soil

...... 119

Table 4.2 Bioconcentration factors and Bioaccumulation factors of selected

compounds in E. fetida………………………………………………………..120

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List of Figures

Figure 2.1 Sulfamethoxazole in the medium and Arabidopsis thaliana cells .... 55

Figure 2.2 Distribution of 14C activity in media and Arabidopsis thaliana cells ...

………………………………………………………………………………….56

Figure 2.3 Kinetics of proposed sulfamethoxazole metabolites in Arabidopsis

thaliana cells ...... 57

Figure 2.4 Proposed metabolism pathways of sulfamethoxazole in Arabidopsis

thaliana cells ...... 58

Figure 3.1 Diazepam in the medium and A. Thaliana cells ……………….…..88

Figure 3.2 Radioactivity in A. Thaliana cells ………………………..………..89

Figure 3.3 Diazepam in radish and cucumber plants cultivate for 7 d and

28d……………………………………………………………………………..90

Figure 3.4 Diazepam metabolites in radish and cucumber seedlings cultivated

for 7 d and 28 d …………………………………………………………….....91

Figure 3.5 Glycosyltransferase activity in radish and cucumber seedlings ...... 92

Figure 4.1 Kinetics of CECs in three matrices ...... 120

Figure 4.2 Kinetics of N4-acetylsulfamethoxazole in E. fetida...... 121

Figure 4.3 Kinetics of metabolites in earthworm and non-earthworm soils. ... 122

Figure 4.4 Activity of antioxidant enymes in earthworm exposed to CECs over

time…………………………………………………………………………....123

Figure 4.5 Malondialdehyde (MDA) content in earthworms...... 124

Chapter 1: Introduction

1.1 Background

Climate change and human population growth threaten sustainable development by placing stress on limited resources, including freshwater and agricultural productivity. As a result, the use of historically underutilized resources, such as treated wastewater (TWW) and biosolids (U.S. EPA, 1999; WHO, 1989), is now being harnessed to combat freshwater scarcity, improve soil fertility, and increase crop growth. Currently, ~54% of the biosolids produced in the United States are land applied, predominantly in agriculture, landscaping, and forestry (U.S. EPA, 1999).

Only ~8% of TWW in the United States is currently reclaimed and implemented for direct potable reuse or indirect potable reuse (U.S. EPA, 2017). The use of these underutilized resources is likely to increase in the future as governments and industries are also promoting their use to help communities adapt to a shifting climate, rising population, and increasing urbanization (Chen et al., 2013; Elgallal et al., 2016;

Marcus et al., 2017).

Population growth and increasing urbanization are giving rise to megacities

(≥10 million inhabitants) around the globe. In 2016 there were an estimated 31 megacities and the number is projected to reach 41 by 2030 (U.N., 2014). Such megacities face substantial challenges when it comes to waste disposal and water management, making the recycling of biosolids and TWW essential (Varis et al.,

2006). The need for TWW and biosolid recycling is also heightened in arid and semi-

arid areas where these resources are increasingly being used to improve agricultural output and conserve limited potable water resources (Tal, 2006).

The recycling of TWW and biosolids could be environmentally and economically beneficial, but significant concerns remain about their use in terrestrial environments (Boxall et al., 2012; Chen et al., 2013). TWW and biosolids have been shown to contain “contaminants of emerging concern” (CECs) (Focazio et al., 2008;

Kolpin et al., 2002). CECs are defined by the EPA as “chemicals and other substances that have no regulatory standard, have been recently discovered in natural water and may potentially cause deleterious effects in aquatic life at environmentally relevant concentration”. CECs include various compound classes such as antibiotics, nonsteroidal anti-inflammatory drugs, preservatives, and flame-retardants. The concentrations of various CECs generally range from ng L-1 to low µg L-1 in TWW and

µg kg-1 to mg kg-1 in biosolids (Clarke and Smith, 2011; Focazio et al., 2008; Kim et al., 2007; McClellan and Halden, 2010; Vanderford and Snyder, 2006). Literature reportedly documents the fate and toxicity of many CECs in various organisms in aquatic systems and raises concerns about the unintended consequences of the widespread use and release of CECs into the aquatic environment (Boxall et al., 2012;

Brooks, 2014; Daughton and Ternes, 1999; Harada et al., 2008; Jones et al., 2002;

Quinn et al., 2008; Sumpter and Johnson, 2005). For example, endocrine disruption and changes in male to female sex ratios have been reported in several fish species

(Ingerslev et al., 2003; Segner et al., 2003).

Data on environmental fate and toxicity of CECs in terrestrial environments are relatively scarce but crucial for understanding the risks to terrestrial organisms after

TWW or biosolids application. The effects of CECs in terrestrial organisms have garnered recent attention. There is increasing information on the effects of CECs in terrestrial plants, invertebrates, and vertebrates (Boxall et al., 2006; Laura J. Carter et al., 2014; Hillis et al., 2011; Malchi et al., 2014; Oaks et al., 2004; Pennington et al.,

2017, 2016). For example, studies have shown that pharmaceuticals can induce oxidative stress in plants (Chengliang Sun et al., 2018), exhibit phytotoxicity in crops

(Jin et al., 2009) and increase the time to adulthood in agricultural pests (Pennington et al., 2017).

1.2 Exposure and Fate of CECs in Plants

Several studies report the uptake of emerging contaminants from hydroponic solutions, spiked soils and soil irrigated with TWW or amended with biosolids or animal manure (Kumar et al., 2005; Malchi et al., 2014; Sabourin et al., 2012; Wu et al., 2010). However, these studies often report conflicting or contradictory results concerning the rate of uptake and the extent of translocation. Hydroponic studies often exhibit higher rates of uptake than those observed in spiked or amended soils studies.

For example, in a study conducted by Boonsaner and Hawker (2010) the maximum concentration of antibiotics in plant tissues was reached within 2 d in spiked water but took 6-8 d when plants were grown in contaminated soils.

In hydroponic systems, plant uptake and translocation are largely driven by the contaminants water solubility, log Kow (octanol-water partitioning coefficient) and/or the pH of the hydroponic solution and the potential for ionization, log Dow, (pH adjusted dependent octanol-water partitioning coefficient) (Tanoue et al., 2012; Wu et al., 2013). Whereas in soils, soil-specific processes such as soil-porewater partitioning, and transformations in soil, also contribute to contaminant uptake and accumulation in plants. Thus, the uptake rate and translocation of a contaminant in plants can vary widely depending upon soil and environmental conditions. For example, a soil with higher organic matter content can limit plant uptake of organic contaminants, due to stronger contaminant adsorption than a soil with lower organic matter content (Gao and Pedersen, 2005). Also, shifts in soil pH can result in ionization of ionizable organic contaminants, affecting the rate of plant uptake (Trapp, 2000).

1.2.1 Antibiotics

Antibiotics constitute one of the most extensively used pharmaceuticals classes for both human and livestock (Landers et al., 2012; U.S. CDC, 2016) and as such are nearly ubiquitously detected in wastewater effluent, biosolids and livestock manure

(Hu et al., 2010; Kolpin et al., 2002; McClellan and Halden, 2010). Relatively more studies have been reported on terrestrial plant uptake and translocation of antibiotics than other pharmaceuticals in the agro-environment, including studies conducted in hydroponic growth solutions, greenhouses, and under field conditions (e.g., Boxall et al., 2006; Kumar et al., 2005; Michelini et al., 2012; Zhang et al., 2017).

In hydroponic growth solution, the antibiotic sulfamethoxazole was taken up in the roots and translocated to leaves of four vegetable plants, including lettuce

(Lactuca sativa L.), spinach (Spinacia oleracea L.), cucumber (Cucumis sativus L.), and pepper plants (Capsicum annuum L.), with the concentration report to be significantly greater in the roots (translocation factor < 1.0) (Wu et al., 2013). In a 55 day hydroponic study, three antibiotics, i.e., tetracycline, cephalexin, and sulfamethoxazole, were found to be taken up and translocated into edible tissues of pakchoi (Brassica chinensis L.), with concentrations ranging from 6.9 - 11.8, 26.4 -

48.1, and 18.1 - 35.3 µg kg-1 for tetracycline, cephalexin and sulfamethoxazole, respectively (Zhang et al., 2017).

Several studies have also explored plant uptake of antibiotics from spiked soils

(e.g., Boxall et al., 2006; Michelini et al., 2012). For example, Boxall et al., (2006) exposed carrot (Daucus carota) and lettuce plants to soils spiked with 1 mg kg-1 of 7 antibiotics, i.e., sulfadiazine, trimethoprim, tylosin, amoxicillin, enrofloxacin, florfenicol, and oxytetracycline. After 103 d (lettuce) and 152 d (carrot) cultivation, antibiotics were quantified in both crops. However, the concentrations varied considerably among different antibiotics and between plant species. For example, amoxicillin was detected at < 1 µg kg-1 in lettuce tissues but was 24 µg kg-1 in carrot tissues (Boxall et al., 2006). Three sulfonamides, i.e., sulfadiazine, sulfamethazine, and sulfamethoxazole, were also reported to be taken up by pakchoi cultivated in spiked-soils, with sulfamethoxazole having the highest concentration among the three antibiotics throughout the 49 d cultivation (Li et al., 2013).

To better predict environmentally relevant risks from antibiotic uptake to human consumption, several studies have been carried out on crops grown in soils irrigated with spiked TWW and/or amended with livestock manure (Dolliver et al.,

2007; Jones-Lepp et al., 2010; Kumar et al., 2005). These studies showed that food crops were capable of taking up and accumulating antibiotics from wastewater and/or manure-amended soils; however, the levels were often very low. For example, chlortetracycline was taken up by corn (Zea mays L.), green onion (Allium cepa L.), and cabbage (Brassica oleracea L.) that were grown in soils amended with antibiotic- spiked manure (100 mg kg-1). However, the concentrations were low (2 – 17 µg kg-1)

(Kumar et al., 2005). Sulfamethoxazole and lincomycin were found to accumulate in lettuce tissues at concentrations up to 125 µg kg-1 and 822 µg kg-1, respectively, after irrigation with antibiotic-spiked synthetic wastewater at 1 mg L-1, (Sallach et al.,

2015). Similarly, in field studies, crops irrigated with TWW were found to take up antibiotics, including but not limited to, roxithromycin, clindamycin, ciprofloxacin, sulfamerazine, and sulfamethoxazole (Christou et al., 2017; Jones-Lepp et al., 2010;

Sabourin et al., 2012). However, in nearly every case the concentration of antibiotics in plant tissues was negligibly low.

1.2.2 Nonsteroidal Anti-Inflammatories

Nonsteroidal anti-inflammatories (NSAID) are the most commonly consumed class of pharmaceuticals in the world (Valcárcel et al., 2011). As such they are ubiquitously found in TWW, biosolids, and surfaces water (Daughton and Ternes,

1999; Focazio et al., 2008; McClellan and Halden, 2010). They have been reported to

accumulate in soils that receive TWW or biosolids (e.g., (Barron et al., 2010; Biel-

Maeso et al., 2018). Several studies have explored the potential for uptake and translocation of NSAIDs in plants, including in hydroponic systems, amended soils, and field studies (e.g., Carter et al., 2014; Christou et al., 2019, 2017; Jones-Lepp et al., 2010; Madikizela et al., 2018; Wu et al., 2015, 2014).

NSAIDs have a wide range of physicochemical range properties (e.g., Kow

0.46-4.51, pKa 4.15-9.38) and, as such, have displayed vastly different uptake and translocation rates (Madikizela et al., 2018). For example, in a hydroponic study the

NSAID diclofenac was observed to accumulate only in the roots of four vegetables

(spinach, lettuce, cucumbers, and peppers) while relatively high levels of acetaminophen were detected in the leaves (Wu et al., 2013). Similarly, a study exploring plant uptake of 14C labeled naproxen and diclofenac from hydroponic solutions showed that two vegetables, i.e., lettuce and collard greens (Brassica oleracea), were capable of accumulating both compounds, and both plants accumulated significantly more diclofenac than naproxen (Dodgen et al., 2013).

Radish and ryegrass were shown to absorb and accumulate diclofenac from soils spiked with the chemical at an initial concentration of 1 mg kg-1. However, after

40 d cultivation, the concentration of diclofenac in the plants was very low < 1 µg kg-1

(Carter et al., 2014). Greenhouse studies using soils amended with biosolids and field studies using TWW irrigation considered the uptake of NSAIDs under environmentally relevant conditions. For example, Cortés et al. (2013) conducted a greenhouse study in which soybeans (Glycine max) and wheat (Triticum aestivum)

were cultivated in biosolids-amended soils for 60 and 110 d. However, none of the four NSAIDs (naproxen, ketoprofen, diclofenac, and ibuprofen) was detected in the plant shoots. On the other hand, in a long-term field study (3 yr), diclofenac was found relatively high levels (up to11.63 µg kg-1) in the fruits of tomato plants after prolonged irrigation with TWW, as compared to sulfamethoxazole and trimethoprim (Christou et al., 2017). Further, in another field study, naproxen was detected in the edible tissues of various vegetables (< 1 µg kg-1) irrigated with TWW or TWW fortified with the chemical at 250 ng L-1 and grown until maturity (Wu et al., 2014).

Several NSAIDs have also been considered in the investigation of potential metabolism of pharmaceuticals in plant cell cultures and whole plants (Fu et al., 2017a,

2017b; Huber et al., 2012, 2009). The metabolism of diclofenac was investigated in four different plant systems, including a horseradish hairy root culture (Armoracia rusticana), barley (Hordeum vulgare), Arabidopsis thaliana cell culture, and

Arabidopsis thaliana whole plants (Fu et al., 2017a; Huber et al., 2012). However, the formation of diclofenac metabolites differed significantly by plant systems. For instance, while phase I hydroxylation was observed in all the systems, the horseradish hairy root cultures and barley formed a glucopyranoside as the major Phase II metabolite (Huber et al., 2012). Arabidopsis thaliana, on the other hand, produced acyl-glutamatyl-diclofenac as the major Phase II metabolite via direct conjugation (Fu et al., 2017a). Direct conjugation of naproxen and ibuprofen with glutamic acid and glutamine was also observed in Arabidopsis thaliana plants (Fu et al., 2017b). The metabolism of acetaminophen has also been studied in multiple plant systems,

including horseradish hairy root cultures and Indian mustard (Brassica juncea L.

Czern.). In these studies, direct glucuronisation, glucosidation, and sulfation were observed along with the formation of a reactive metabolite N-acetyl-p- benzoquinoneimine (Huber et al., 2009, 2010). Taken together these studies have highlighted the ability of plants to uptake and transform NSAIDs.

1.2.3 Psychiatric Pharmaceuticals

Several classes of psychiatric pharmaceuticals have been detected in TWW and biosolids including antidepressants, mood stabilizers, and antianxiety agents (Calisto and Esteves, 2009; Subedi et al., 2013; Yuan et al., 2013). Of these compounds, carbamazepine has been likely considered in probably the most in the agro- environment due to its stability during wastewater treatment and in the environment

(Clara et al., 2004). Carbamazepine has been oftenreported to be taken up by plants in both field and laboratory settings (e.g., Carter et al., 2014; Paltiel et al., 2016; Shenker et al., 2011; Winker et al., 2010; Wu et al., 2014, 2013). In hydroponic systems, carbamazepine has been shown to accumulate (BCF 10 - 100) in both roots and shoots of multiple plant species, including lettuce, spinach, cucumber, and peppers (Wu et al.,

2013). Cucumber was found to readily translocate carbamazepine when cultivated in hydroponic systems (Shenker et al., 2011). However, a high rate of translocation was not observed in cabbage plants cultivated in hydroponic systems (Herklotz et al.,

2010). In greenhouse studies, carbamazepine was reported to be taken up by cucumbers and ryegrass (Lolium perenne) grown in soils irrigated with TWW and urine (Shenker et al., 2011, Winker et al., 2010). In addition, Shenker et al., (2011)

reported that uptake into cucumbers was negatively correlated with soil organic matter content. In fields irrigated with TWW, trace levels of carbamazepine was found to accumulate in different parts of various vegetables (Wu et al., 2014). Carbamazepine was also reported to transfer to humans after consumption of contaminated vegetables

(Paltiel et al., 2016). The metabolism of carbamazepine in plants has also been investigated (Malchi et al., 2014; Wu et al., 2016). In carrot cell cultures five phase I metabolites of carbamazepine were observed to form over 22 d (Wu et al., 2016).

Further, 10,11-epoxycarbamazepine and 10,11-dihydroxycarbamazepine have been reported in carrots and sweet potatoes grown in fields irrigated with CEC-spiked

TWW (Goldstein et al., 2014).

Fluoxetine is an antidepressant prescribed for both human and animal consumption (Giorgi, 2012), resulting in fluoxetine being commonly detected in environmental samples (Brooks et al., 2003). In hydroponic cultivations fluoxetine was taken up by cauliflower (Brassica oleracea var. botrytis) and accumulated in the stems

(0.49 µg g-1) and leaves (0.26 µg g-1) ( Redshaw et al., 2008). In a greenhouse study exploring plant uptake of fluoxetine from soils irrigated with TWW and amended with biosolids fluoxetine accumulated in the roots (8.1 - 40.5 µg kg-1), but, it was not translocated to the leaves (Wu et al., 2010). In addition, fluoxetine displayed an opposite uptake pattern to that for carbamazepine, and showed a greater accumulation in plants grown in biosolid-amended soils as opposed to soil irrigated with TWW (Wu et al., 2010).

Benzodiazepines, are one of the most prescribed classes of pharmaceuticals

(Bachhuber et al., 2016). Of these, diazepam is among the most commonly detected pharmaceuticals in TWW, with concentration ranging from ng L-1 to low µg L-1

(Kosjek et al., 2012). Benzodiazepines have been shown to be taken up and accumulate in tissues of plants grown in treated hydroponic solutions or soils (Carter et al., 2018; Wu et al., 2013). In hydroponic solutions, diazepam has been observed to accumulate in both the leaves and roots of lettuce, spinach, cucumber, and pepper with

BCF of 10-100 (Wu et al., 2013)]. Further, in a greenhouse study exploring the uptake of seven benzodiazepines (chlordiazepoxide, clonazepam, diazepam, flurazepam, oxazepam, nordiazepam, triazolam, and temazepam), both silverbeets (Beta vulgaris) and radish (Raphanus sativus) crops took up and accumulated all seven benzodiazepines from the treated-soil (500 µg kg-1). Oxazepam was found to have the highest accumulation in both plants, with concentrations up to 14.2 µg g-1 in silverbeets and 5 µg g-1 in radishes (Carter et al., 2018). However, the fate of these pharmaceuticals in the agro-environment is still relatively unexplored, even though their physicochemical properties (moderate logKow, non-ionized) indicate a high potential for uptake by plants (e.g., Briggs et al., 1982; Carter et al., 2018; Laura J.

Carter et al., 2014; Trapp, 2009).

1.2.4 Antimicrobials and Preservatives

A multitude of antimicrobials and preservatives are used in health and grooming products, collectively known as personal care products (Pedrouzo et al.,

2011; Zulaikha Siti et al., 2015). Personal care products have garnered increased

scientific attention due to their presence in surface waters and concerns that some of these antimicrobials and preservatives (e.g., parabens) may be endocrine disruptors

(Gautam et al., 2014; Witorsch and Thomas, 2010). Of these, triclocarban and triclosan have been amongst the best studied compounds in the terrestrial environment due to their ubiquitous occurrence in biosolids and relative stability in soils after biosolid application (e.g., Cha and Cupples, 2009; Higgins et al., 2010; Wu et al., 2009; XIA et al., 2010).

Triclocarban and triclosan have been reported to be taken up by several crop species from hydroponic solutions. For example, after exposure to an aqueous solution mixture of triclocarban and triclosan (500 µg L-1) 11 different food crops, cucumber, tomato, cabbage, okra (Abelmoschus esculentus), pepper (Capsicum annuum), potato

(Solanum tuberosum), beet, onion (Allium cepa), broccoli, celery (Apium graveolens), and asparagus (Asparagus officianalis), were capable of taking up both compounds.

However, translocation from roots to the aerial tissue was ≤1.9% for triclocarban and ≤

3.7% for triclosan after 1 month of exposure (Mathews et al., 2014). Similarly, Wu et al. (2012) found triclocarban and triclosan to have a translocation factor (TF) < 0.01 in four vegetables (lettuce, spinach, cucumber, and peppers) cultivated in a hydroponic solution with two initial exposure concentrations (0.5 and 5 µg L-1). In a greenhouse study, triclocarban and triclosan were taken up in radish, carrot, and soybeans from biosolid-amended soils and, the greatest concentration was observed in the carrot root

(49.8 µg kg-1) after 45 d of treatment and decreased thereafter (Prosser et al., 2013).

However, in a three-year field study in which soils were amended with biosolids in

accordance with Ontario providence agricultural practices, the concentration of triclosan and triclocarban in the plant tissues was relatively steady and low (< 5.1 µg kg-1) (Prosser et al., 2013). Plants have also been shown to metabolize triclosan, forming 33 metabolites in horseradish cell cultures with the majority being phase II conjugates (Macherius et al., 2014). Further, one transformation product of the triclosan, methyl-triclosan, has been widely detected in environmental samples and is known to have greater toxicity than the parent compound (DeLorenzo et al., 2008;

Dhillon et al., 2015).

Parabens are common preservatives used in cosmetics, and among the most commonly detected CECs in TWW and biosolid. Parabens are of concern due to their endocrine disrupting potential (Błędzka et al., 2014). Parabens have been widely detected in surface waters and sediments (Ramaswamy et al., 2011; Venkatesan et al.,

2012). However, knowledge of their behavior, uptake, and transformation in terrestrial systems is comparatively limited. Methyl paraben was unstable in soil after application of biosolids, with the maximum concentration of 14.1 µg kg-1 reached after 5 h and decreasing to < 1 µg kg-1 after 35 d (Camino-Sánchez et al., 2016). In a biosolid- amended field, methyl paraben was the lone paraben detected in the biosolids but was not quantifiable in tomatoes, sweet corn, carrot and potatoes (Sabourin et al., 2012).

1.2.5 Knowledge Gaps

The above studies highlight the potential for CECs to enter the terrestrial environment, accumulate in plant tissues, and undergo transformations in plants.

However, the wide variations in plant uptake and translocation rates under different

soil and environmental conditions are currently not well understood and warrant further investigation. Further, it must be noted that the majority of currently published studies have focused on many of the same 20 or so CECs and explored their uptake in mostly the same plant species (Carvalho et al., 2014). There are over 1500 pharmaceutical compounds, alone, currently in circulation (Guo et al., 2016). Further, many of the current models have been shown to overestimate the concentration of

CECs in plant tissues (Prosser et al., 2014). In addition, no models have been able to take into account plant metabolism when determining the concentration and risk of

CECs in terrestrial plants. More research is needed on a wider swath of CECs with different physicochemical properties in a wider range of plants to improve risk assessment. Transformation of CECs in the environment, including through plant metabolism also needs to be further investigated to better understand their fate and risks in the terrestrial environment.

1.3 Phytotoxicity

CECs can potentially induce phytotoxicity. Literature pertinent to phytotoxicity of various classes of CECs is discussed below.

1.3.1 Antibiotics

Antibiotic exposure in plants has been widely studied due to previously observed phenotypic toxicity. Several studies showed decreases in root length and changes in shoot development of various plants exposed to several different classes of antibiotics including sulfamides, fluoroquinolones, and penicillins (Boxall et al., 2006;

Hillis et al., 2011; Jin et al., 2009). Most of these studies were conducted at antibiotic concentrations greater than those of environmental relevance and/or utilized artificial or hydroponic growth media. For instance, shoot and root growth of pinto beans

(Phaseolus vulgaris) grown in a nutrient solution spiked with two antibiotics, chlortetracycline and oxytetracycline, significantly decreased in a dose-dependent manner (Batchelder, 1981). Enrofloxacin, a fluoroquinolone, induced hormetic and toxic effects on post-germination growth in lettuce, cucumber, radish (Raphanus sativus) and barley (Phaseolus vulgaris) plants at concentrations ranging from 0.005 to

50 mg L-1 in laboratory conditions (Migliore et al., 2003).

Seed germination has also been studied as a potential biological end-point to assess toxicity to antibiotic exposure (e.g., Hillis et al., 2011; Jin et al., 2009; Pan and

Chu, 2016). The exposure effects on seed germination vary considerably by plant species and exposure chemical. In filter paper tests, sweet oat (Cichaorium endivia), rice (Oryza sativa L.) and cucumber (Cucumis sativus L.) seeds were negatively impacted when the seeds were exposed to aqueous solutions of increasing concentrations of six antibiotics, i.e., chlortetracycline, tetracycline, tylosin, sulfamethoxazole, sulfamethazine, and trimethoprim (Liu et al., 2009a). The EC10 and

EC50 for seed germination were, however, significantly different depending on the antibiotic and the plant species. Rice seeds exposed to sulfamethoxazole were the most

-1 -1 sensitive with an EC10 of 0.1 mg L but tylosin had an EC10 > 500 mg L. On the

-1 other hand, cucumber seeds exposed to sulfamethoxazole had an EC10 > 300 mg L

-1 but an EC10 of 0.17 mg L for chlortetracycline (Liu et al., 2009). Exposure to

antibiotics can also change plant nutrient and chemical compositions. For example, irrigation with water spiked with sulfamethoxazole and trimethoprim increased production in carbohydrate and soluble solid contents in tomatoes as compared to the plants irrigated with untreated water (Christou et al., 2019).

The mechanisms driving the phytotoxicity of antibiotics have also been explored. Antibiotics can be directly toxic to or indirectly affect plants. Indirect adverse effects can arise from antibiotic exposure that detrimentally affects mycorrhizal fungi, a vital plant-microbe interaction (Hillis et al., 2008). Direct toxicity can result when antibiotics interfere with plant hormones or chemical synthesis pathways, or damage chloroplasts, etc. For example, sulfamethoxazole was shown to directly disrupt the folate synthesis pathway in plants by blocking the action of dihydropteroate synthase (Brain et al., 2008; Schreiber et al., 2012). Tetracyclines was shown to interrupt mitochondrial proteostasis and damage plant chloroplasts (Wang et al., 2015). Interactions with plant hormones may also play a role in the observed phenotypic phytotoxicity. Erythromycin and tetracycline can promote the production of abscisic acid in plants (Pomati et al., 2004). Abscisic acid, a stress hormone, is crucial for plant responses to drought, salinity, heavy metals, among other stressors

(Pastori and Foyer, 2002), but antibiotic-induced production of this hormone can cause premature leaf and fruit detachment and inhibit seed germination.

Plants, depending upon species, can also detoxify antibiotics through reactions with phase II metabolic enzymes (Farkas et al., 2007). However, studies so far have shown significant variations among plant species. For example, the antibiotic

chlortetracycline was detoxified by glutathione conjugation via glutathione-S- transferase in maize (Zea mays), but glutathione-S-transferase did not efficiently catalyze the conjugation in pinto beans (Phaseolus vulgaris) (Farkas et al., 2007).

These detoxification reactions, likely produce a series of conjugated metabolites that have yet to be characterized. Understanding the extent of such conjugation is crucial for estimating the total antibiotic uptake, accumulation, and translocation of antibiotics in plants as the formation of conjugates may mask the total concentration, even though some of these conjugates may retain biological activity (Fu et al., 2017b).

1.3.2 Nonsteroidal Anti-Inflammatories

Several widely used NSAIDs, such as ibuprofen, acetaminophen, and diclofenac are amongst the most studied pharmaceuticals in the environment. Studies have shown that NSAIDs can induce toxicity to plants (e.g., An et al., 2009; González-

Naranjo and Boltes, 2014; Soares et al., 2016). Phytotoxicity, however, is often plant species and NSAID specific. For example, ibuprofen has been shown to inhibited root

-1 elongation in Sorghum bicolor at high concentrations, with EC50 of 232.64 mg L

(González-Naranjo and Boltes, 2014). However, in seed germination tests exposure to a hydroponic solution containing 1 mg L-1 ibuprofen, along with other fenamic acid class NSAIDs, increased the length of the primary root in lettuce but had no effect on radish (Schmidt and Redshaw, 2015). In the same study, diclofenac was observed to decrease the root-to-shoot ratio in radish seedlings cultivated in a sand/spiked-nutrient solution (10 µg L-1), but did not significantly affect the seed germination. However, protein content was not affected in maize (Zea mays L.) cultivated in soils irrigated

twice with different concentrations of acetaminophen (310 - 1240 mg L-1) but grain yields and seed germination were negatively impacted in a dose dependent-manner

(Hammad et al., 2018).

Plants can metabolize and detoxify NSAIDs. For example, plants were found to detoxify acetaminophen by conjugation with glutathione followed by conversion to cysteine and acetylcysteine conjugates (Huber et al., 2009; Sun et al., 2019). Similarly, diclofenac was found to be converted to glucose conjugates in barley and horseradish and glutamic acid conjugates in Arabidopsis thaliana (Fu et al., 2017a; Huber et al.,

2012). Arabidopsis thaliana cell cultures can detoxify ibuprofen via conjugation with sugars and amino acids (Marsik et al., 2017).

1.3.3 Psychiatric Pharmaceuticals

As mentioned above, pharmaceuticals used to treat psychiatric disorders are another group of frequently detected pharmaceuticals in environmental samples, particularly the anticonvulsant carbamazepine (Calisto and Esteves, 2009).

Carbamazepine exposure (1 mg L-1) has been seen to exhibit mycotoxicity to carrot mycorrhizal endpoints (a measure of hyphal growth and spore production) by decreasing the production of fungal spores (Hillis et al., 2008). Similarly, carbamazepine induced leaf necrosis, altered plant hormones and macronutrient concentrations, and reduced root growth at plant tissue concentrations of 1 to 4 mg kg-1 in zucchini (Cucurbita pepo) cultivated in soil spiked with chemical at 0.1 - 20 mg kg-1

(Knight et al., 2018). Information on the toxicity of benzodiazepines and fluoxetine in terrestrial plants is still limited; however, toxicity has been reported in aquatic plants

for these compounds, indicating that toxicity may also occur after exposure in terrestrial plants (Amy-Sagers et al., 2017).

1.3.4 Antimicrobials and Preservatives

Antimicrobials and preservatives are often added to personal care products to increase shelf life. They pass from the human body, largely unchanged, and ultimately end up in TWW, biosolids, and sewage sludge. (Carballa et al., 2004). Antimicrobials and preservatives have been detected in agricultural soils after irrigation with TWW and/or the application of biosolids, and can be taken up by plants (Cha and Cupples,

2009; Sabourin et al., 2012).

Two antimicrobials, triclosan and triclocarban, have attracted more attention because of their potential for endocrine disruption (Dodson et al., 2012) and phytotoxicity (e.g., Karnjanapiboonwong et al., 2011; Liu et al., 2009b; Ryan S.

Prosser et al., 2014). For example, triclosan significantly inhibited plant growth in

-1 -1 cucumber and rice seedlings with EC50 of 108 mg kg and 57 mg kg , respectively

(Liu et al., 2009b). Lettuce shoot mass also decreased in a dose-dependent manner after cultivation in soil amended with triclocarban-spiked biosolids (Prosser et al.,

2014). On the other hand, growth of radish, carrot, soybean, spring wheat, and corn plants grown in soils amended with biosolids containing environmentally relevant concentrations of triclosan and triclocarban, improved compared to un-amended soils; likely due to the positive impacts of biosolids addition (Prosser et al., 2014). Thus, plant species, concentrations, and growth media can significantly affect phytotoxicity of these CECs.

Similarly to antimicrobials, parabens have also garnered recent attention due to their potential for endocrine disruption and phytotoxicity (Dodson et al., 2012).

Polyparabens reportedly inhibited root growth in onion bulbs (Allium cepa) under hydroponic conditions (Herrero et al., 2012). Methyl paraben, the most commonly detected paraben, has been shown to inhibit seed germination in rice and mung bean

(Vigna radiata) in aqueous solutions at respective concentrations of ≥100 mg kg-1 and

≥ 200 mg kg-1. Methyl paraben decreased shoot growth and biomass in both rice and mung beans at a concentration of ≥ 200 mg kg-1 in soil (Kim et al., 2018).

1.3.5 Mixtures

Studies exploring the phytotoxicity of individual pharmaceuticals or classes of pharmaceuticals are useful to highlight high-risk compounds and/or the potential mechanism of toxicity. CECs are, however, often introduced into the environment in complex mixtures (Olofsson et al., 2012) and these mixtures can affect the uptake and translocation of individual compounds (Carter et al., 2014; Christou et al., 2016).

Some studies report positive effects on plants exposed to CEC mixtures under environmentally relevant conditions. For instance, TWW irrigation increased tomato and lettuce yield compared to freshwater irrigation (Aiello et al., 2007; Lu et al., 2016;

Margenat et al., 2017). Exposure of lettuce seedlings to a mixture of 11 CECs significantly altered plant metabolic pathways, including the citric acid cycle and pentose phosphate pathway, and decreased chlorophyll content in a dose-dependent manner (Hurtado et al., 2017). Also, exposure to 18 CECs at concentrations ranging

from 5 to 50 µg L,-1 induced oxidative stress in cucumber seedlings and caused upregulation of enzymes associated with detoxification reactions (Sun et al., 2018).

1.3.5 Knowledge Gaps

Literature on the toxicity of a number significant CECs (e.g., fluoxetine, benzodiazepines, parabens) to terrestrial plants is still very limited, and many of the studies have utilized concentrations that are orders of magnitude higher than those seen in the environment. Studies on the toxicity of mixtures in terrestrial plants are also limited, but warrant attention as several studies have indicated that mixtures can induce effects not observed from individual compounds (Carter et al., 2015; Christou et al.,

2016; Sun et al., 2018). The ability of plants to detoxify these compounds through metabolism also merit further research. Overall, more research is needed on the toxicity of a wider range of CECs in plants under environmentally relevant conditions to more accurately assess the impacts of CECs in the agro-environment.

1.4 Exposure and Toxicity of CECs in Invertebrates

The potential for exposure to, and toxicity of, CECs has been investigated in several aquatic invertebrate species. Toxicity end-points such as endocrine disruption, changes in growth, time to development, and mortality rates have been considered in these studies (Harada et al., 2008; Jones et al., 2002; Segner et al., 2003; Yamamoto et al., 2011). Studies addressing the effects of CECs on terrestrial invertebrates are, however, few. Of the published studies on terrestrial invertebrates, the earthworm

Eisenia fetida has been examined mainly due to their increased susceptibility and

ecological importance (OECD 207, 1984). Literature pertaining to toxicities of various classes of CECs to terrestrial invertebrates is discussed below.

1.4.1 Antibiotics

Like in terrestrial plants, antibiotics can also induce toxicity in terrestrial invertebrates. Exposure to environmentally relevant (and higher) concentrations of antibiotics caused mortality to earthworms and/or induced oxidative stress and genotoxicity in E. fetida. For instance, high concentrations (300 mg kg-1) of tetracycline and chlortetracycline inhibited antioxidant enzymes superoxide dismutase

(SOD) and catalase (CAT) while these enzymes were stimulated at lower doses (0.3-30 mg kg-1), and DNA damage was induced along a dose-dependent curve (Dong et al.,

2012; Lin et al., 2012). Also, chlortetracycline can reduced juvenile earthworm (EC50

= 96.1 mg kg-1) and cocoon counts (EC50 = 120.3 mg kg-1) in E. fetida (Lin et al.,

2012).

Environmentally relevant concentrations of three antibiotics, lincomycin, ciprofloxacin, and oxytetracycline increased mortality and development time in cabbage loopers (Trichoplusia ni) when reared on an artificial diet and treated tomato plants (Pennington et al., 2017). Further, the three antibiotics altered the microbiome inside cabbage loopers (T. ni) and mosquitos (Culex quinquefasciatus) but did not impact development time of mosquitoes (Pennington et al., 2017, 2016). However, antibiotic (lincomycin, ciprofloxacin, and oxytetracycline) exposure did not induce toxicity in aphids (Myzus persicae) reared on bell peppers (Capsicum annuum)

(Pennington et al., 2018). Antibiotic toxicity in terrestrial invertebrates, therefore,

appears to depends upon the specific antibiotics, concentrations, bioavailability, invertebrate species, and environmental conditions.

1.4.2 Nonsteroidal Anti-Inflammatories

Exposure to NSAIDs caused acute and sub-acute adverse effects in terrestrial invertebrates, including earthworms (Carter and Kinney, 2018; Chen et al., 2015). Pino et al. (2015) assessed lethality of E. fetida cultivated in artificial soil as a result of

-1 exposure to 18 pharmaceuticals. Ibuprofen had the lowest LC50 at 64.8 mg kg followed by diclofenac at 90.5 mg kg-1 (Pino et al., 2015). Exposure to diclofenac

-1 resulted in a dose-dependent decrease in survival (LC50 1099 mg kg ) and

-1 reproduction (EC50 170 mg kg ) of Folsomia candida in spiked soils (Chen et al.,

2015). However, it should be noted that these LC50 values were much higher than what may be expected in the real environment. At sub-acute concentrations (50 mg kg-1), diclofenac induced significant genotoxicity in Folsomia candida, including induction of the up-regulation of transcriptional processes and genes associated with the immune response (Chen et al., 2015).

Acetaminophen increased E. fetida mortality along both a dose-dependent curve (14 µg L-1- 14 mg L-1) and over time [7-28 d (Bulman, 2012)]. In the mosquito species Culex quinquefasciatus, exposure to water contaminated with an environmentally relevant concentration of acetaminophen resulted in increased susceptibility to Bacillus thuringien israelensis and increased larval development time

(Pennington et al., 2015). Acetaminophen at environmentally relevant concentrations also significantly increased days to adulthood in cabbage loopers (T. ni) reared on an

artificial diet. However, a similar effect was not observed when cabbage loopers (T. ni) were reared on acetaminophen-treated tomato plants (Pennington et al., 2017).

Similarly, the development time for aphids (M. persicae) reared on acetaminophen- treated bell pepper was not affected by acetaminophen (Pennington et al., 2018).

Therefore, like for other CECs, effeccts of NSAIDs on terrestrial invertebrates are species, compound, and environment specific.

1.4.3 Antimicrobials and Preservatives

Many antimicrobials and preservatives, including the common environmental contaminants triclocarban, triclosan, and parabens are amongst the most frequently detected in TWW and biosolids (Prosser and Sibley, 2015). Partitioning of these CECs into biosolids suggests that soil-dwelling organisms are at greater risks of exposure as they preferentially consume organic matter rich soils and biosolids (Jager et al., 2003).

Triclocarban, triclosan, and methyl-triclosan have been detected in the tissues of earthworms collected from field sites that were amended with biosolids 4 years prior to the worm collection (Macherius et al., 2014).

After 28-d exposure to triclosan at ≥50 mg kg-1 in soil E. fetida had significantly increased SOD and CAT activities and increased concentrations of malondialdehyde (MDA), a chemical indicative of lipid peroxidation and DNA damage in E. fetida (Lin et al., 2012). Lin et al. (2014) reported negative impacts of triclosan exposure (≥ 50 mg kg-1) on E. fetida reproduction including decreases in the number of cocoons and juveniles. Triclosan also decreased the biomass, shell diameter, and food intake in a terrestrial snail (Achatina fulica) at concentrations ≥ 40 mg kg-1.

Further, triclosan exposure increased CAT and SOD activities and MDA concentration in A. fulica in a dose-dependent manner (Wang et al., 2014). However, no adverse effects were observed in E. fetida cultivated in triclosan-amended biosolids at environmentally relevant concentrations (<1 mg kg-1) (Pannu et al., 2011).

Triclocarban is more persistent in the environment than triclosan and is known to bioaccumulate in earthworm tissues (Higgins et al., 2010b, 2010a; Macherius et al.,

2014). However, information on its toxicity to terrestrial invertebrates remains limited.

For example, in Synder et al. (2011) exposure to triclocarban at concentrations ≥ 77 mg kg-1 for 2-4 weeks resulted in a trend towards increased mortality; however, the variations in data were too high to discern any statistically significant trend.

Exposure ≥ 400 mg kg-1 to methyl paraben in soil resulted in increased abnormalities in earthworms (Eisenia andrei) where a normal survival-EC50 value of

397 mg kg-1 was estimated (Kim et al., 2018). An acute (≤14 d) exposure to methyl paraben in soil at ≥ 60 mg kg-1 increased F. candida mortality and chronic (≥28 d) exposure at concentrations ≥150 mg kg-1 decreased the reproductive rate (Kim et al.,

2018). However, methyl paraben is often detected at concentrations ranging from 15.9

- 203.0 µg kg-1 in sewage sludge, levels that are well below the concentrations where toxicity was observed (Chen et al., 2017).

1.4.5 Mixtures

Terrestrial invertebrates are more likely to be exposed to complex mixtures of

CECs in the environments, including agricultural soils (Yager et al., 2014). However, studies addressing the effects of CEC mixtures on terrestrial invertebrates are scarce.

E. fetida reared in biosolid-amended soils suffered a decrease in cocoon counts (1 - 4% biosolid) and a decrease in adult survival at 8 weeks with 4% biosolids amendment rates (Kinney et al., 2012). High, but environmentally relevant, rates of biosolids application (≥ 10 t ha-1) also caused mortality in E. fetida and F. candida; even lower application rates (~5 t ha-1) hindered reproduction in both species (Artuso et al., 2011).

However, biosolids are extremely complex in composition with numerous other constituents, including metals and salts, and it is difficult to attribute their effects solely to CECs. Environmentally relevant concentrations of mixtures of antibiotics, hormones, acetaminophen, and caffeine were found to extend development time, increase mortality, and alter the microbiome of mosquitos (C. quinquefasciatus) and cabbage loopers (T.ni) (Pennington et al., 2015, 2016, 2017a). Collectively, the aforementioned studies suggest that CECs may have increased toxicity when present in mixtures, and adverse effects at environmentally relevant concentrations are likely, but need to be further confirmed.

1.4.5 Knowledge Gaps

Currently, knowledge on the fate and toxicity of CECs in terrestrial invertebrates is still very preliminary. The limited studies conducted so far shown chemical-specific and species-specific effects of CECs in the environment. It is known that the bioavailability of these compounds and environmental conditions will play a large role in the overall accumulation and toxicity of the CECs in terrestrial invertebrates (van Straalen et al., 2005). However, information on bioavailability under differing environmental conditions is still lacking for most CECs. Similarly, the

potential for trophic transfer (e.g., Pennington et al., 2017) warrants further investigation. Overall, more research involving a wider range of compounds, especially in the context of environmental relevance, is needed to more accurately assess the fate and toxicity of CEC in terrestrial invertebrates.

1.5 Problem Statement

The studies highlighted above suggest that CECs are ubiquitous in the environment and that exposure, even at environmentally relevant concentrations, these contaminants may be hazardous for terrestrial organisms. However, studies also suggest that these organisms can metabolize, transform and detoxify these CECs. The interplay between the toxicological effects of CEC exposure and an organism’s ability to take up and metabolize these contaminant is poorly understood and serves as significant knowledge gaps in understanding the fate and risks of CECs in terrestrial environments. These gaps must be addressed to gain better risk assessments of CECs during the use of biosolids and treated wastewater in the agro-environment. To address these gaps, we carried out a series of experiments utilizing plant cell cultures, hydroponic cultivations, earthworm incubations, high-resolution mass spectrometry,

14C-tracing, and enzyme assays to systematically evaluate the fate, metabolism, and biological effects of sulfamethoxazole, diazepam, naproxen, and methyl paraben and their major metabolites in terrestrial organisms under laboratory conditions. The four

CECs were selected based on their detection in TWW and biosolids, their range of physicochemical properties and uses, and the paucity of information about their fate

and impacts in the literature (Table 1). The study systems included Arabidopsis thaliana cells cultures, radishes, cucumbers, and E. fetida. Organisms were selected due to their extensive use in the literature, commercial availability, and worldwide agricultural relevance.

1.6 Research Objectives

The ultimate goal of this project was to improve our understanding of the fate and potential adverse effects of CECs in terrestrial organisms to contribute to more accurate environmental risk assessments on the use of biosolids and treated wastewater in agriculture. Specifically, we sought to address some of these knowledge gaps by addressing the following specific research objectives:

1. To elucidate metabolic pathways of select CEC detoxification in a model plant

(A. thaliana) and vegetable (Cucumis sativas).

The metabolic pathways of a model CEC compound, sulfamethoxazole,

were determined in A. thaliana cells incubated for 96 h at 1 mg mL-1. Cell

extraction and high-resolution mass spectrometry were utilized to determine the

structure and kinetics of the metabolite formation. Hydroponic cultivations

were used to confirm metabolite presence in the whole plant.

2. To determine the differences in CEC metabolism caused by exposure conditions

and plant species.

Potential differences in CEC metabolism were comparatively

investigated using A. thaliana cells and cucumber (Cucumis sativus) and radish

(Raphanus sativus) seedlings grown in hydroponic solution following acute (7

d)/high concentration (1 mg L-1), and chronic (28 d)/low concentration (1 µg L-

1) exposures. Liquid chromatography paired with mass spectrometry, 14C

tracing, and enzyme extractions, were used to characterize metabolic phases.

3. To evaluate potential bioaccumulation and toxicological effects of select CECs

in the earthworm species Eisenia fetida.

Bioaccumulation and toxicological effects of four CECs, i.e., methyl

paraben, sulfamethoxazole, diazepam, and naproxen, in E. fetida exposed at

environmentally relevant concentrations were assessed after 21 d incubation in

an artificial soil. Extractions and chemical analysis of different compartments,

including earthworm tissue, soil, and soil-porewater, enzyme assays

(superoxide dismutase, catalase, glutathione-S-transferase) and lipid

peroxidation (malondialdehyde) analysis, were carried out to characterize

chemical distribution, bioaccumulation, and adverse biological effects.

Table 1: Properties of contaminants of emerging concern and their major metabolites

Name Activity Chemical CAS ID Molecular Pka1,2 Log 2 Formula Weight Kow

Diazepam Benzodiazepine C16H13ClN2O 439-14-5 284.7 3.4 2.82

Diazepam-d5 Benzodiazepine C16H8D5ClN2O 65854-76- 289.8 3.4 2.82 4

Nordiazepam Benzodiazepine and C15H11ClN2O 1088-11-5 270.7 2.85 2.79 Metabolite

Temazepam Benzodiazepine and C16H13ClN2O2 860-50-4 3 300.7 1.6 2.16 Metabolite

Oxazepam Benzodiazepine and C15H11ClN2O2 604-75-1 286.7 1.7 2.01 Metabolite

Oxazepam- Metabolite C21H19ClN2O8 6801-81-6 462.8 -0.87 1.37 Gluconoride

Methylparaben Preservative C8H8O3 99-76-3 152.2 8.5 1.96

Methylparaben-d4 Preservative C8H4D4O3 362049- 156.2 8.5 1.96 51-2

4-Hydroxybenzoic Metabolite C7H6O3 99-96-7 138.1 4.54 1.58 acid

Naproxen Anti- C14H14O3 52079-10- 230.3 4.15 3.18 inflammatory 4

Naproxen-d3 Anti- C14H13D3O3 958293- 233.3 4.15 3.18 inflammatory 77-1

O-desmethyl Metabolite C13H12O3 52079-10- 216.2 4.34 2.71 naproxen 4

Sulfamethoxazole Antibiotic C10H11N3O3S 723-46-6 253.3 6.1 0.89

Sulfamethoxazole- Antibiotic C10H7D4N3O3S 1020719- 257.3 6.1 0.89 d4 86-1

N4-acetyl Antibiotic C12H13N3O4S 21312-10- 295.3 5.68 0.86 sulfamethoxazole 7

1Wu et al 2015 2HSDB 2018 3Wishart DS, et al., 2018

Chapter 2: Metabolism of sulfamethoxazole in Arabidopsis thaliana

cells and cucumber seedlings

Abstract

Reclaimed water is a historically underutilized resource. However, with increased population growth and global climate change, reclaimed water is evolving into an economical and sustainable water resource to meet the needs of citizens, industries, and agriculture. The use of recycled water for agricultural irrigation comes with the potential risk of environmental and food contamination by pharmaceuticals and personal care products (PPCPs). The levels of PPCPs in plants will depend on translocation and metabolism in plant tissues. However, relatively little is known about the metabolism of PPCPs in plants. In this study, the metabolism of the antibiotic sulfamethoxazole was investigated in Arabidopsis thaliana cells as well as cucumber seedlings grown under hydroponic conditions. Using high-resolution mass spectrometry and 14C tracing allowed for sulfamethoxazole metabolism to be comprehensively characterized through all metabolic phases. Six phase I and II metabolites were identified in A. thaliana cell cultures and cucumber seedlings.

Sulfamethoxazole metabolism followed oxidation and then rapid conjugation with glutathione and leucine. Direct conjugation with the parent compound was also observed via acetylation and glucosylation. At the end of 96 and 168 h incubation, N4- acetylsulfamethoxazole was the major metabolite and >50% of the radiolabeled sulfamethoxazole became non-extractable in both A. thaliana cells and cucumber

seedlings suggesting extensive phase III metabolism and detoxification. The study findings provided information for a better understanding of the uptake and metabolism of sulfamethoxazole in higher plants, highlighting the need to consider metabolic intermediates and terminal fate when assessing the risk of PPCPs in the soil-plant continuum.

Key words: Pharmaceuticals, antibiotics, sulfamethoxazole, water reuse, plant uptake, metabolism

2.1 Introduction

Over the past two decades, pharmaceuticals and personal care products

(PPCPs) have emerged as contaminants of environmental concern due to their extensive use and continuous emission into the environment (Boxall et al., 2012;

Daughton and Ternes, 1999; Pedersen et al., 2005). PPCPs are released into the environment primarily through the disposal of treated wastewater and biosolids from wastewater treatment plants (WWTPs) (Carballa et al., 2004). As climate change and population growth places an increasing stress on freshwater resources, especially in arid and semi-arid regions, communities have turned to utilizing municipal treated water for agricultural irrigation, which may result in soil contamination by PPCPs

(Barnett et al., 2005; Marcus et al., 2017; Tal, 2006). Furthermore, the heavy use of some pharmaceuticals, particularly antibiotics, for disease control and growth promotion in intensive animal farming also contributes to contamination of agricultural fields when animal wastes are used for fertilization (Hu et al., 2010).

The presence of PPCPs in irrigation water and soil can lead to contamination of food crops if plants can substantially accumulate these compounds. Various studies over the last decade have sought to quantify plant uptake of PPCPs, and in general, only low levels of PPCPs have been found in edible tissues (ng/kg) (e.g., Wu et al.,

2014). The majority of studies to date have only targeted the parent form of PPCPs for analysis. However, plants have a cascade of enzymes that may extensively transform xenobiotics such as PPCPs after uptake (Celiz et al., 2009; Fu et al., 2017b). Recently several published studies have explored the metabolism of pharmaceuticals in plants

(e.g. Fu et al., 2017; Huber et al., 2012; LeFevre et al., 2015; Marsik et al., 2017).

Therefore, consideration of metabolism and biologically active metabolites is much needed for a better understanding of the fate and risks of PPCPs in the soil-plant system.

Higher plants have many detoxification enzymes similar to those in animals.

These enzymes function in plants as a ‘green liver’ (Sandermann, 1994). In general, metabolism of xenobiotics includes three phases. Phase I involves modification reactions such as oxidation, hydrolysis, and dealkylation reactions introducing reactive sites to the molecule. Phase II is characterized by conjugation with large polar biomolecules, such as sugars and amino acids, to further increase the polarity of the xenobiotic. Phase III is typified by sequestration, resulting in the formation of bound residues (Miller et al., 2016; Sandermann, 1992, 1994). As shown for many xenobiotics in mammals and plants metabolites from phases I and II often retain biological activity (Cribb et al., 1991; Osborne et al., 1990; Pichersky and Gang,

2000), and therefore should not be discounted.

In this study, sulfamethoxazole was selected as the compound of interest because of its prevalence in WWTP effluents and increasing concerns over the propagation of antibiotic resistance (Boucher et al., 2009; Ventola, 2015). Since its introduction in 1961 sulfamethoxazole has been widely prescribed due to its potency against both gram-positive and gram-negative bacteria (Hitchings, 1973). Currently, sulfamethoxazole has been detected from ng L-1 to µg L-1 in surface and effluent waters and µg kg-1 to mg kg-1 in soils and manure (Brausch et al., 2012; Hu et al., 2010).

Recent long-term studies of waste-water application under realistic field conditions have highlighted the potential for sulfamethoxazole to be taken up and translocated in crop plants, including to the fruit (Christou et al., 2017).

The structures of sulfamethoxazole metabolites, including conjugates from

Phase II metabolism, were identified using high-performance liquid chromatography coupled with time-of-flight high-resolution mass spectrometry (HPLC-TOF-HRMS) and further quantified using ultra-high performance liquid chromatography in tandem with a triple quadrupole mass spectrometry (UPLC-TQD-MS/MS). Furthermore,

Phase III terminal products in the form of bound residues were quantified using 14C labeling.

Arabidopsis thaliana cells were selected as the experimental organism due to their extensive use in the literature, commercial availability, and their membership in the commonly consumed Brassica family (e.g., cabbage, broccoli, kale). Further,

Arabidopsis thaliana plants are found worldwide under several common names (e.g.,

Wall cress, mouse-ear cress, shiro-inu-nazuna) and are consumed by a wide variety of animals as well as humans (Redei, 1992; van Poecke and Dicke, 2004). Cucumber

(Cucumis sativus) was selected in the hydroponic experiment due to the fact that it is often consumed raw, rapid growth, and amiability to soilless culture (Lineberge, 2018).

2.2 Materials and Methods

2.2.1 Chemicals and solvents

Non-labeled sulfamethoxazole was purchased from MP Biomedicals (Solon,

OH). Sulfamethoxazole-d4 was purchased from C/D/N Isotopes (Pointe-Claire,

Quebec, Canada) and 14C-labeled sulfamethoxazole was obtained from American

Radiolabeled Chemicals (Saint Louis, MO). Stock solutions of 14C-sulfamethoxazole and non-labeled sulfamethoxazole were prepared in methanol to reach a specific radioactivity of 1.2 × 103 dpm µL-1 and a chemical concentration of 1.0 mg mL-1, respectively. HPLC grade acetonitrile and methanol were used for extraction along with ultra pure water. Mobile phases were prepared using Optima™ LC/MS grade methanol and deionized water. Standards were prepared in HPLC grade methanol and stored in the dark at -20 °C. All solvents used in this study were purchased from

Fisher (Fair Lawn, NJ).

2.2.2 Arabidopsis thaliana Cell Incubation Experiment

PSB-D A. thaliana cell line (CL84840) was purchased from the Arabidopsis

Biological Resource Center (ARBC) at the Ohio State University (Columbus, OH).

The cells were maintained in liquid suspension culture at 25 °C and rotated at 130 rpm in the dark according to the ARBC protocol (Gehring et al., 2018). To explore metabolism of sulfamethoxazole in A. thaliana cells, 7 mL of cell culture was inoculated in 43 mL fresh culture and cultivated for 96 h at 25 °C and 130 rpm in the dark to produce the seed culture. A 30 µL aliquot of the non-labeled stock solution and

10 µL aliquot of 14C-sulfamethoxazole were spiked into 30 mL of A. thaliana cell culture, resulting in a nominal initial concentration of sulfamethoxazole of 1 µg mL-1 and a specific radioactivity of 1.2 × 103 dpm mL-1 (0.3% methanol). Simultaneously, control treatments were prepared by autoclaving cell suspensions before chemical spiking (non-viable cell control), flasks containing sulfamethoxazole without cells

(medium control), and flasks containing live cells but no sulfamethoxazole

(background control). These control treatments were used to determine adsorption, abiotic degradation, and potential toxicity to cells. The incubation lasted for 96 h, and triplicate containers were sacrificed at 0, 3, 6, 12, 24, 48 and 96 h.

At each sampling interval, the entire culture was transferred to a 50 mL polypropylene centrifuge tube and centrifuged at 10,000 rpm for 15 min. The supernatant was collected and stored at -20 °C until further analysis and the plant cells were placed at -80 °C before freeze-drying for 72 h. After drying, cells were fortified

-1 with 50 µL of 10 mg L sulfamethoxazole-d4 as a recovery surrogate. Cells were extracted using a modified method previously established in Wu et al. (2012). Briefly, cells are sonicated in a Fisher Scientific FS110H sonication bath (50/60 Hz, Pittsburgh,

PA) for 20 min with 30 mL acidified DI water (pH 4) followed by centrifugation at

10,000 rpm for 15 min. The supernatant was decanted into a new 50 mL centrifuge tubes. The cell matter was further extracted using 20 mL methyl tert-butyl ether

(MTBE), followed by 20 mL acetonitrile. The MTBE and acetonitrile supernatants were combined, dried under nitrogen at 35 °C, and re-constituted in 1.0 mL methanol.

The extract was then combined with the above water extract. The combined sample

extract was loaded onto a preconditioned 150-mg Oasis© HLB solid phase extraction

(SPE) cartridge and eluted with 20 mL methanol. The cleaned extract was dried under nitrogen and further recovered in 1.5 mL 50:50 MeOH:H2O (v/v). The growth media was acidified to pH 3 and similarly extracted and cleaned as described above.

Extraction recovery for the sample preparation protocol of the cell extract was 46% ±

13 and for the growth media was 57% ± 10.

Prior to instrument analysis, both cell and media extracts were transferred to micro-centrifuge tubes and centrifuged at 120,000 rpm in a bench-top SciLogex d2012 centrifuge (Rocky Kill, CT) and further filtered through a 0.22-µm polytetrafluoroethylene (PTFE) membrane (Millipore, Carrigtwohill, Cork, Ireland) into 2 mL glass vials. All final extracts in 2 mL glass vials were stored at -20 °C if not immediately analyzed.

At each time interval, 100 µL of the cell material extract or concentrated growth media was added to 6 mL Ultima Gold™ liquid scintillation cocktail (Waltham, MA) to measure the extractable 14C-radioactivity on a Beckman LS 5000TD Liquid

Scintillation Counter (LSC, Beckman, Fullerton, CA). Additionally, the extracted cell matter was air dried, and a 10 mg aliquot was combusted on an OX-500 Biological

14 Oxidizer (R. J. Harvey Instruments, Hillsdale, NJ). The evolved CO2 was captured in

15 mL Harvey Carbon-14 cocktail II and the 14C activity was measured to derive the fraction of bound residues from Phase III metabolism.

2.2.3 Whole Plant Hydroponic Cultivation Experiment

Uptake and metabolism of sulfamethoxazole were further evaluated using whole cucumber seedlings. Cucumber seeds were purchased from Fisher (Fairfield,

NJ). Seedlings were started in a commercially purchased organic soil (Parkview

Nursery Riverside, CA) in a growth chamber (22°C 16 h day/20°C 8 h night cycle; relative humidity of 75-80%). After the appearance of the first true leaves, plants of uniform size were selectively removed from pots, rinsed with DI water and placed in 1

L amber glass jars containing hydroponic solution (Oasis® 16-4-17 hydroponic fertilizer 3.16 g L-1). After acclimating for 4 d, each jar was treated with 100 µL of non-labeled sulfamethoxazole stock solution to arrive at a nominal concentration of 1

µg L-1. Simultaneously, 7.6 µL of 14C-sulfamethoxazole stock solution was added to reach an initial specific radioactivity of 8.6 × 103 dpm mL-1. Jars containing seedlings without sulfamethoxazole and jars containing sulfamethoxazole but no plant were similarly prepared as controls. After 7 d, seedlings were removed from the jars, and their roots were carefully rinsed with DI water and dried with paper towels. Roots, stems, old leaves (those present before the start of incubation) and new leaves (those that grew during the incubation) were separated from each other using a razor blade.

The separated tissues were stored at -80 °C until analysis.

Plant tissues were placed in a freeze-drier for 72 h, and a 0.2 g aliquot of the dried plant material was ground in liquid nitrogen using a mortar and pestle. The pulverized tissue samples were subsequently extracted and cleaned as described above for A. thaliana cells. The hydroponic media was collected, filtered with GF/F filters,

acidified to pH 3 with HCL, and extracted using an HLB cartridge as previously described.

2.2.4 Structural elucidation using mass spectrometry

Instrument analysis was first performed on a Waters ACQUITY ultra- performance liquid chromatography (UPLC) in tandem with a Waters Micromass triple quadrupole (TQD) mass spectrometer equipped with an electrospray ionization

(ESI) interface (Waters, Milford, MA) to determine the rate of uptake and the concentration of the parent compound. Separation was achieved on an ACQUITY

UPLC HSS T3 column (2.1 mm × 100 mm, 1.7 µm, Waters) at 40 °C. Mobile phase A consisted of water containing 5% methanol and acidified using 0.001% formic acid.

Mobile phase B was composed of pure methanol. The solvent gradient program, in reference of mobile phase A, was as follows, 0-2 min 95%; 2-3min 30%; 3-3.25 min

5%; 3.25-4.5 95%. The flow rate was 0.3 mL min-1 and the injection volume was 5 µL.

The mass data were acquired using Intellistart® (Waters) in multiple reactions monitoring (MRM) and in the positive ESI mode. Proposed metabolites were scanned in selective ion reaction (SIR). Mobile phase A and B consisted of acidified water and methanol (0.1% formic acid) respectively. The solvent gradient program, in reference of mobile phase A, was as follows, 0-5 min 95%, 5-9 min 50%, 9-10 min 0%, 10-12 min 95%. The flow rate was 0.4 mL and the injection volume was 5 µL. The specific instrument settings were: capillary voltage 1.7kV, collision gas (Aragon, 99.9%), dwell time 0.022 s, source temperature 150 °C, desolvation temperature 450 °C, desolvation gas 900 L h-1 and cone gas 50 L h-1. The cone voltage (V) and the collision energy (V)

for each standard can be found in SI table 1.

Accurate mass data were obtained using an Agilent 1200 series HPLC (with

UV detector) coupled to an Agilent 6210 time-of-flight high-resolution mass spectrometer (TOF-HRMS) with ESI/APCI mixed ion source. The separation was achieved on a Thermo Scientific Hypersil Gold C18 column (2.1mm x 100mm, 3 um).

Mobile phase A consisted of water containing 0.1% formic acid. Mobile phase B was composed of acetonitrile 0.1% formic acid. The solvent gradient ran from 5% to 100%

B in 18 minutes at 0.3mL/min flow rate. Samples were analyzed at the High

Resolution Mass Spectrometry Facility in the Chemistry Department at the University of California, Riverside. Raw data files were obtained and converted to mzXMLfiles using ProteoWizard MSConvert and analyzed using MZmine 2 open software (Pluskal,

2010). Candidate metabolites were proposed based on the presence of unique peaks in the treatment that were absent in the controls (Figure S1). Identification uncertainty was determined using the Warwick mass accuracy calculator by comparing theoretical m/z to the observed m/z.

To structurally identify sulfamethoxazole metabolites, the uncertainties in confirmation were evaluated against the metabolite identification criteria as outlined in

Schymanski et al. (2014). Using the criterion, Level 1 structures are those with direct confirmation against authentic standards. Level 2 structures are probable structures based on library spectrum data, literature data, and experimental information and Level

3 structures are tentative structures derived from strong MS/MS information for the proposed structures, but the position of substitutions could not be determined with

certainty (Table 1). Structural identification for sulfamethoxazole metabolites was determined from accurate mass information and fragmentation patterns received using the QTOF mass analyzer (Figure S2). The information was compared to mass spectra libraries and/or the literature on known human metabolites of sulfamethoxazole.

Further, identified metabolites were scanned in the selective ion reaction, multiple reaction monitoring and MS scan modes using Targetlynx™ software (Waters) with comparison against authentic standards when available (Figure S3). Results were further compared with literature reporting common enzymes in the metabolism of other xenobiotics in plants (Badenhorst et al., 2013; Vanderford and Snyder, 2006;

Zhou et al., 2015) to determine the most likely pathways and metabolite structures of sulfamethoxazole.

The relative fractions of individual metabolites were determined on the Waters

UPLC-TQD MS/MS in the selective ion reaction scan mode. Authentic standards of sulfamethoxazole and N4-acetylsulfamethoxazole (S296) were used as an example to verify the identity of proposed structures as well as to quantify these compounds.

Retention times, accurate mass, and fragmentation patterns were used for validation.

2.2.5 Data Analysis and Quality Control

Individual peaks were detected and integrated using TargetLynx XS software

(Waters). Data were analyzed and graphed using the Prism 6 GraphPad software (La

Jolla, CA). Results were calculated as the mean ± standard error (SE), and a Student's t-test was conducted to assess systematic differences between multiple groups and two groups (α=0.05).

All treatments in the A. thaliana cell incubation experiment were conducted in triplicate, and all treatments in the hydroponic cultivation experiment were conducted in quadruple (to account for potential loss of plants). Calibration curves were prepared with standards of sulfamethoxazole, sulfamethoxazole-d4 and N4- acetylsulfamethoxazole and the linear range with r2 ≥ 0.99 was used to ensure accuracy. A limit of detection of 3 ng mL-1 and a limit of quantification of 5 ng mL-1 for sulfamethoxazole were determined based on a signal to noise ratio of 3 and 10, respectively. Whenever possible, authentic standards, online mass spectra databases, and mz calculators were used to verify the identity of the proposed metabolic products.

2.3 Results and Discussion

2.3.1 Kinetics of parent, extractable and non-extractable residues

The metabolism of sulfamethoxazole in Arabidopsis thaliana cells was validated using a range of controls. No sulfamethoxazole was detected in the media or cell blanks, and there was no detectable disappearance of sulfamethoxazole in the cell- free media, suggesting the absence of contamination or abiotic transformation.

Moreover, no significant difference was seen in the cell mass between the chemical- free control and the treatments indicating that sulfamethoxazole did not affect the growth of A. thaliana under the experimental conditions. In the non-viable cell control, it was found that sulfamethoxazole was adsorbed to the cell matter, but the fraction did not contribute significantly to the dissipation of sulfamethoxazole from the media (P >

0.05). In contrast, in the live cell treatments, sulfamethoxazole dissipated appreciably

from the media, with the average concentration decreasing from 246 ± 1.74 ng mL-1 initially to 176 ± 5.23 ng mL-1 after 96 h of incubation (Figure 1). Concentrations were therefore not adjusted for recovery or loss to adsorption to cell matter or the surfaces of the flask. As there was no significant difference between measured initial concentrations of the cell-free flasks, and viable and non-viable flasks, we assumed that adsorption to the container was insignificant.

Concurrent to the dissipation in the medium, sulfamethoxazole was detected in the A. thaliana cells, and the level was the highest at 3 h sampling point decreasing thereafter. The presence of sulfamethoxazole in the live cells provided direct evidence of its uptake into A. thaliana cells. The level of sulfamethoxazole in the cell matter decreased after reaching the maxima at 3 h, Fitting a decrease in the sulfamethoxazole level in the cell to a first-order decay model yielded a half-life of 19.4 h (r2 = 0.94).

This was in comparison to a biological half-life of 10 h in humans for sulfamethoxazole (Kaplan et al., 1973). The decrease of sulfamethoxazole in the live

A. thaliana cells suggested active metabolism.

The use of 14C labeled sulfamethoxazole enabled determination of the fractions of sulfamethoxazole and its metabolites that were incorporated into the cell matter, which could not be characterized using traditional extraction and analytical methods. A rapid increase in the bound residue fraction was observed during the 96 h cell cultivation, while the increase in the extractable residue form in the cells was more gradual (Figure 2). During the incubation, the fraction in bound residues increased steadily to 53 ± 10% at 96 h, clearly suggesting that A. thaliana cells were capable of

effectively metabolizing and then sequestering sulfamethoxazole and its metabolites in the cell system. In contrast, the fraction of the extractable residues was relatively low, ranging from 0% to 22 ± 1.6%. Extractable residues in plant metabolism are thought to contain Phase I and Phase II metabolites, including conjugates, while Phase III metabolism results in the incorporation or sequestration of metabolites into the cell wall (Sandermann, 1994). Therefore, formation of bound residues may be regarded as detoxification of a xenobiotic in plants. Several previous studies also demonstrated that plant cells and whole plants were capable of metabolizing PPCPs, transforming them into more polar intermediates and sequestering them in their cell walls or vacuoles (Fu et al., 2017a; Huber et al., 2009, 2012).

2.3.2 Metabolism of Sulfamethoxazole in A. thaliana cells

In A. thaliana cells, tentative metabolism pathways of sulfamethoxazole were derived by combining spectra data, knowledge of human metabolism of sulfamethoxazole and its degradation in water systems (Figure 4). In the proposed metabolism pathways, sulfamethoxazole underwent Phase I metabolism including oxidation and hydroxylation reactions, which was followed by Phase II metabolism through acetylation and rapid conjugation with glucuronic acid, amino acids, and glutathione (Figure 3 and 4). The end products of Phase II metabolism were then further sequestered likely through incorporation into cell walls and other cell components, resulting in the formation of non-extractable bound residues.

4-Nitroso-sulfamethoxazole (designated as S268) is a known metabolite in human metabolism. The knowledge of its formation in mammalian livers, structure,

and properties (Cribb et al., 1993) was used to detect its presence in the HPLC-TOF

MS scans. Subsequent UPLC-MS/MS scans suggested the rapid formation of this intermediate. Individuals with low production of N-acetyltransferase were previously reported to exhibit a high production of hydroxylamine sulfamethoxazole and nitroso- sulfamethoxazole. In humans, these metabolites were found to be responsible for the adverse side effects associated with the consumption of sulfonamides, such as skin rashes or hives (Cribb et al., 1993). N4-Acetyl-5-OH-sulfamethoxazole (S313) was also previously shown to form by cytochrome-P450 oxidation in mammalian livers

(Zhou et al., 2009). In higher plants, it was likely formed through oxidation reactions mediated by cytochrome-P450 enzymes, a superfamily of enzymes in both plants and mammals (Gonzalez and Nebert, 1990).

N4-Acetylsulfamethoxazole (S296), sulfamethoxazole-glucuronide (S430) and

N4-sulfamethoxazole-glutathione conjugate (S606) metabolites were all previously found in human metabolism of sulfamethoxazole (Cribb et al., 1993). N4- acetylsulfamethoxazole was detected in A. thaliana cells and confirmed using its authentic standard. The glucose and glutathione conjugates were detected by comparing the exact mass and fragmentation patterns to proposed spectra libraries for each compound (TMIC, 2017). However, observed difference in the fragmentation patterns indicated that conjugation location differed from in those observed for human metabolism. A similar pattern was observed in previous studies concerning the plant metabolism of pharmaceuticals (Fu et al., 2017b).

The proposed amino acid conjugate in A. thaliana cells is, to the best of our knowledge, the first evidence for their occurrence in higher plants. Conjugation with amino acids has been considered a detoxification pathway for other pharmaceuticals

(Fu et al., 2017a; Marsik et al., 2017). The structure proposed here for leucylsulfamethoxazole (S385) was, in part, based on a (M+ H) m/z of 132.0765, 223.1135

and 255.1677 showing distinct fragments of C6H13NO2 C12H18N2 and C10H12N3O3S

(Table S2). The position of the amino acid on the benzene ring was selected based on

optimum stable formation (Carney and Giuliano, 2011).

In the tentative metabolism pathways in A. thaliana cells, sulfamethoxazole underwent Phase I oxidation, forming 4-nitroso-sulfamethoxazole (S268) followed by phase II conjugation with leucine or glutathione. Based on the signal strength, a relatively high level of the N5-leucyl-sulfamethoxazole conjugate (S385) was detected at the 3 h sampling point but remained at trace levels for most of the incubation duration, with the exception of the 48h sampling point. The glutathione conjugate

(S606) appeared quickly (3 h into incubation), spiked at 48h, and decreased to a non- detectable level by the end of the cultivation (Figure 3).

Conjugation with glucuronic acid (S430) was also observed to form quickly

(3h), and direct glycosylation of sulfamethoxazole has also been observed in mammals

(van der Ven et al., 1995). Another pathway appeared to be acetylation of the sulfamethoxazole amine followed by rapid oxidation to form the S296 and S313 metabolites (Figure 3), with N4-acetylsulfamethoxazole being the predominant metabolite at the end of incubation. It has been shown that this acetylation pathway

predominates in human metabolism for detoxification (Spielberg, 1996). Future research should be conducted to determine if enzymes similar to those seen in human metabolism of sulfamethoxazole actively participate in its metabolism in plants, such as CYP2C9 and N-acetyltransferases 1 and 2.

2.3.3 Uptake, Translocation and Transformation in Cucumber Seedlings

Similar metabolites were found in cucumber seedlings grown in the nutrient solution containing sulfamethoxazole. Metabolites from Phase I and Phase II metabolism were similarly detected and extensive Phase III sequestration was further observed (Table 2). When cucumber plants were exposed to 1.0 µg L-1 sulfamethoxazole in nutrient solution, sulfamethoxazole was taken up into the plant, with accumulation primarily in the roots, old leaves, and stems, and absent in the new leaves (Table 2). When 14C activity was used for calculation, a translocation factor

(TF), i.e., the ratio of the concentration in leaves/stems over that in the roots, was estimated to be 0.32. While this value was higher than that reported by Dodgen et al.

(2015) (based on the parent form) it suggested that sulfamethoxazole is not readily translocated in the plant after entering the root. This was consistent with models used to predict the behavior, e.g., diffusion through cell membranes, of similar pharmaceutical compounds (Trapp, 2009). These models suggested that polar compounds (log Kow 0 to 1), such as sulfamethoxazole, would have translocation factors from the solution to the xylem ranging from 0.25 to 0.5.

Bioconcentration factors were calculated for the roots (BCFR) and the shoots

(BCFS) as the ratio of specific radioactivity in the tissue over that in the growth media.

The mean BCFR and BCFS were determined to be 1.59 and 0.53, respectively. The calculated BCFR and BCFs were similar to those found for other vegetable species in

Dodgen et al. (2015). These studies together suggested that while plants are capable of taking up sulfamethoxazole, it mainly remains in the root with limited potential for translocation to the other organs of the plant (Miller et al., 2016; Wu et al., 2015,

2012).

After the 7 d cultivation, sulfamethoxazole parent and metabolites identified in the A. thaliana cell incubation were similarly scanned in cucumber samples from the hydroponic cultivation experiment, including plants tissues and blank control without plants. The proposed metabolites were not detected in the control treatment or the cucumber hydroponic solution, indicating that the transformation occurred within the cucumber seedlings following uptake. All of the metabolites proposed for A. thaliana cells were detected in the cucumber seedlings. However, due to the lack of authentic standards for most of the metabolites or low signal (S/N < 3), we did not attempt to quantify individual metabolites in the cucumber plants. In a previous study, Chen et al.

(2017) investigated the uptake, metabolism, and elimination of sulfamethoxazole in

Brassica rapa chinensis and Ipomoea aquatica. In that study, no metabolites of sulfamethoxazole were detected in plant tissue. This could be attributed to a number of factors, such as, differences in extraction protocols, instrument analysis or extensive phase III metabolism that decreased the level of metabolites below the limit of detection. Our findings were in line with previous research conducted with the related

sulfa-drug sulfamethazine, in which N-acetyl-sulfamethoxazole and hydroxy- sulfamethoxazole were detected in Zea mays L. plants (Michelini et al., 2012).

The total amount of extractable and non-extractable residues in the cucumber plants ranged from 94% to 80% indicating that some mineralization (conversion to

CO2) occurred. This rate of mineralization in the plant cultivation system was higher than that of sulfamethoxazole in soils(Höltge and Kreuzig, 2007a). Because plant

14 respiration (the release of CO2) may contribute to the loss of C after mineralization, this finding further highlights the detoxification prowess of higher plants (Dodgen et al., 2014).

2.4 Conclusion

Results from this study demonstrated that 14C tracing, high-resolution mass spectrometry, and cell and hydroponic cultures may be used in a complementary manner to obtain a complete depiction of plant metabolism of emerging contaminants.

The antibiotic sulfamethoxazole was taken up and metabolized extensively by A. thaliana cells and cucumber seedlings. The Phase I metabolism involved the oxidation of the amine group, which was followed by Phase II reactions including conjugation with glutathione, and direct conjugation of the parent compound with glucuronic acid and leucine. The finding that sulfamethoxazole may be directly conjugated with biomolecules in higher plants merits further investigation, as conjugates may become deconjugated upon ingestion. Therefore, consideration of such conjugates may offer a more accurate assessment of risk for such chemicals, for example, through the dietary

intake. The potential for deconjugation may be particularly significant for antibiotics, due to an increased prevalence of antibiotic-resistant bacteria that pose a severe threat to the effectiveness of treatment for infections in healthcare worldwide.

Tables

Table 1: Summary of mass spectra information and proposed structures of sulfamethoxazole and its metabolites

2 n)

Name

r (mir t ID m/z Obs. m/z Calc. Mass difference Predicted formula Fragment (m/z) Confidence level

3 S 4 NS 8 N NO H 10.57 5 11 6 H 6 SMX H C H ±0.002 6 Level 1 Level C 254.0552 254.0521 155.9934 127.0593 108.0362 AS, HR AS, 10 C MS, MRM MS, C Sulfamethoxazole

S S

4 3 S 2

O O OS

- 3 3 3 2 N N N NO N 6 9 9 2 4 2.06 MS, SIR MS, H S268 ±0.02 2013) H H H H - Nitroso 6 4 10 Level 2b Level 10 10 268.1158 268.1392 252.1170 216.2067 152.0517 132.1073 famethoxazole C C C C C HR (Bonvin, et.al, Sul

OS OS 2 2

3 NOS N N e acetyl S 8 13

8 4 OH - - H S296 H H H 8 ±0.004 8 4 Level 1 Level N4 12 AS, HR AS, 11.60 296.0751 296.0705 279.1728 180.1797 163.1293 129.0783 C C C MS, MRM MS, C sulfamethoxazol

S S

5 3 S

3 OH

8 -

O O

5 3 3 O 3 O - 2 OS 5 5 N N

N 14 11 4 H CH 4.76 MS, SIR MS, 6 - S313 CH 2009) - H H H - ±0.004 Acetyl C 7 - Level 2b Level 313.0781 313.0732 299.2023 279.1739 254.0648 194.117 124.0893 12 10 C N4 HR C C sulfamethoxazole (ZhouSF al., et

S S S 2

5 4 3

N O O O

2 O O 4 2 3 8 OS O 12 5 N N N

H 18 leucyl 15.49 3 H 24 15 13 - H 2 MS, SIR MS, 6 S385 H C - H H H ±0.031 C3 Level 3 Level C - C 12 - 385.1852 385.1546 325.2122 301.1429 283.1326 255.1677 223.1135 124.0897 16 12 10 C HR C C C sulfamethoxazole

S S S

S S

9 9 7 6 7 3

O O O

3 3 3 3 NO N N N N 14 9 19 20 13 6.76 MS, SIR MS, S430 H H - H H H ±0.006 Level 3 Level 12 10 glucuronide 430.0893 430.0836 419.3306 391.3031 316.2214 251.1917 16 15 16 C C HR C C C Sulfamethoxazole (Rieder,1973)

3 3 2

S S S S 3

7 5 4

S 2 O

O O O 3 11 onjugate 4 4 3 O OS 7 5 N O N N N 6 13 16.71 H 20 16 12 MS, SIR MS, 6 S606 N CH - H ±0.006 H H H - C 25 Level 2b Level 606.0894 606.0951 555.3787 476.3320 391.3031 302.1659 255.2408 124.0893 10 16 12 10 H HR C sulfsulfamethoxazole C C C - 20 glutathione c glutathione C (Cribb1991) al., et N4

AS: confirmed by authentic standard; MRM: Multiple Reaction Monitoring; SIR: Selective Ion Reaction MSL: Comparison with mass spectra library; HR-MS: High resolution mass spectrometry; tr retention time obtained from HPLC TOF-MS/MS analysis aSchymanski E. et al., 2014 14-Amino-N-(5-methyl-1,2-oxazole-3-yl)benzenesulfonamide), 2N-(5-Methyl-3-isoxazolyl)-4- nitrobenzenesulfonamide, 3N-{4-[5-Methyl-1,2-oxazol-3-yl)sulfamoyl]phenyl}ethanimidic acid, 4 N-(4- {[5-(hydroxymethyl)-1,2-oxazol-3-yl]sulfamoyl}phenyl)ethanimidic acid, 52-[2-Amino-5-(5-methyl-3- isoxazolylaminosulfonyl)phenylamino]-4-methylvaleric acid, 6N-(5-Methylisoxazole-3-yl)-N-(beta-D- glucopyranuronosyl)-4-aminobenzenesulfonamide, 72-amino-4-{[(1R)-1-[(carboxymethyl)carbamoyl]-2- {[({4-[(5-methyl-1,2-oxazol-3-yl)sulfamoyl]phenyl}amino)sulfinyl]sulfanyl}ethyl]carbamoyl}butanoic acid

Table 2: Detection of 14C-sulfamethoxazole in cucumber seedlings after 7 d growth in nutrient solution (nominal % of spiked 14C activity)

Extractable Bound Parent Parent Residues Residues Compound Compound (%) (%) (ng/g) (%) Roots 1.09 ± 0.38 20.93 ± 2.80 3155 ±1439 1.06 ± 0.40 Stems 0.91 ± 0.17 1.83 ± 0.51 2486 ± 202.2 0.52 ± 0.29 Old Leaves 1.56 ± 0.04 2.95 ± 0.85 2780 ±1341 0.82 ± 0.39 New Leaves 1.67 ± 0.11 1.48 ± 0.43 N.D. N.D Media 56.02 ± 3.40 4658 ± 1626 46.6 ± 16.3

Figures

Figure 1. Sulfamethoxazole in the medium and Arabidopsis thaliana cells. (A)

Kinetics of sulfamethoxazole in cell growth media. (B) Kinetics of sulfamethoxazole

in viable and non-viable cells. Error bars represent the standard error of triplicate.

Figure 2. Distribution of 14C activity in media and Arabidopsis thaliana cells after treatment with 14C-sulfamethoxazole (nominal % of initially spiked radioactivity)

Figure 3. Kinetics of proposed sulfamethoxazole metabolites in Arabidopsis thaliana cells (based on peak area)

Figure 4. Proposed metabolism pathways of sulfamethoxazole in Arabidopsis thaliana cells. Blue (Phase I) and Red (Phase II) circles represent the altered functional group. Dashed arrow: Proposed structures; solid arrow: Verified structured

Figure 5. Representative chromatogram of the metabolite N4- acetylsulfamethoxazole in cucumber shoots and Arabidopsis thaliana cells

Supplementary Information

Table 1. Optimized MRM conditions for the analysis of the chosen PPCPs by UPLC-TQD

MS/MS (CV: cone voltage in V; CE: collision energy in eV)

Compound MRM MRM1 CV/CE MRM2 CV/CE (Precursor (Quantification) (Qualification) mass) Sulfamethoxazole 253.9681 92.0480 34/28 155.9518 34/14 Sulfamethoxazole-d4 258.0128 96.0797 32/26 159.9900 32/16 N4- 296.0228 134.0380 32/24 65.0087 32/38 acetylsulfamethoxazole

Figure 1. HPLC-TOF HRMS TIC Scans

A +ESI TIC control Arabidopsis thaliana cell culture

B +ESI TIC treated Arabidopsis thaliana cell culture

C +ESI TIC control cucumber seedlings

D +ESI TIC treated cucumber seedlings

Figure 2: HPLC-QTOF HRMS Mass Spectra

A +ESI mass spectrum Sulfamethoxazole

B +ESI mass spectrum 4-Nitroso-sulfamethoxazole

C +ESI mass spectrum N4-Acetylsulfamethoxazole

D +ESI mass spectrum N4-Acetyl-5-OH-sulfamethoxazole

E +ESI mass spectrum leucylsulfamethoxazole

F +ESI mass spectrum sulfamethoxazole-glucuronide

G mass spectrum N4-sulfamethoxazole-glutathione conjugate

Figure 3. N4-Acetylsulfamethoxazole SIR and MRM scans; UPLC TQD-MS/MS

Chapter 3: Formation of biologically active benzodiazepine metabolites

in Arabidopsis thaliana cell cultures and vegetable plants under

hydroponic conditions

Abstract

The use of recycled water for agricultural irrigation comes with the concern of exposure to crops by contaminants of emerging concern (CECs). The concentration of

CECs in plant tissues will depend on uptake, translocation and metabolism in plants.

However, relatively little is known about plant metabolism of CECs, particularly under chronic exposure conditions. In this study, metabolism of the pharmaceutical diazepam was investigated in Arabidopsis thaliana cells and cucumber (Cucumis sativus) and radish (Raphanus sativus) seedlings grown in hydroponic solution following acute (7 d)/high concentration (1 mg L-1), and chronic (28 d)/low concentration (1 µg L-1) exposures. Liquid chromatography paired with mass spectrometry, 14C tracing, and enzyme extractions, were used to characterize the metabolic phases. The three major metabolites of diazepam - nordiazepam, temazepam and oxazepam - were detected as

Phase I metabolites, with the longevity corresponding to that of human metabolism.

Nordiazepam was the most prevalent metabolite at the end of the 5d incubation in A. thaliana cells and 7 d, 28 d seedling cultivations. At the end of 7d cultivation, non- extractable residues (Phase III) in radish and cucumber seedlings accounted for 14% and

33% of the added 14C-diazepam, respectively. By the end of 28 d incubation, the non-

extractable radioactivity fraction further increased to 47% and 61%, indicating Phase III metabolism as an important destination for diazepam. Significant changes to glycosyltranferase activity were detected in both cucumber and radish seedlings exposed to diazepam. Findings of this study highlight the need to consider the formation of bioactive transformation intermediates and different phases of metabolism to achieve a comprehensive understanding of risks of CECs in agroecosystems.

Key words: Contaminants of emerging concern; pharmaceuticals; benzodiazepines; diazepam; metabolism in plants

3.1 Introduction

Global climate change has resulted in shifts in precipitation patterns, causing stress on freshwater resources, especially in arid and semi-arid regions (Trenberth, 2011;

Walther et al., 2002). In many of these areas, demand for water has led to increasing use of municipally treated wastewater (TWW) (Lazarova et al., 2001; Tal, 2006). Agriculture has been one of the primary targets for TWW reuse with water districts and governments promoting the adoption of recycled water for irrigation (Chen et al., 2013; Elgallal et al.,

2016; Marcus et al., 2017). However, the use of TWW for irrigation may come with potential risks, as TWW is known to contain a wide variety of human pharmaceuticals

(Carballa et al., 2004; Kolpin et al., 2002; Xia et al., 2005).

The use of pharmaceutical compounds has increased with population growth and economic development, resulting in over 1500 compounds currently in circulation (Guo et al., 2016). Their widespread consumption has led to their occurrence in TWW as well as in TWW-impacted surface water (Cunha et al., 2017; Xiang et al., 2018). For many of these pharmaceuticals, there is limited knowledge about their potential chronic effects in the environment (Archer et al., 2017; Bourdat-Deschamps et al., 2017). Further, many of these compounds can transform in the environment, resulting in the formation of transient or recalcitrant transformation products, many with unknown fates and effects in environmental compartments (Fu et al., 2017b).

Diazepam belongs to the class of psychoactive compounds known as benzodiazepines, one of the most prescribed classes of pharmaceuticals (Bachhuber et al.,

2016). Diazepam is one of the most commonly detected pharmaceuticals in TWW, with

concentration ranging from ng L-1 to low µg L-1 (Kosjek et al., 2012). This is likely due to its extensive use and low removal efficiency (~ 8%) during secondary wastewater treatment (Verlicchi et al., 2012).

In humans, diazepam is primarily metabolized via phase I oxidative metabolism by demethylation to nordiazepam (major pathway), or hydroxylation to temazepam

(minor pathway), and then further oxidized to oxazepam (de Angelis et al., 1974).

Oxazepam undergoes phase II metabolism via rapid glucuronidation and then excretion via urine (Sonne, 1993). The three primary metabolites of diazepam are psychoactive compounds, and each is a prescribed pharmaceutical for treating psychological conditions and alcohol withdrawal symptoms (Clarke and Nicholson, 1978; Malcolm et al., 1989).

Both oxazepam and nordiazepam have been commonly detected in TWW, often at µg L-1 levels (Kosjek et al., 2012). However, there is little knowledge about the occurrence, formation, and fate of such metabolites outside the wastewater treatment systems (Boxall et al., 2012).

Several studies have focused on the uptake and accumulation of pharmaceuticals in agricultural plants as a result of TWW irrigation (e.g., Carter et al., 2014; Goldstein et al., 2014; Wu et al., 2014, 2013). These studies have demonstrated the capacity of higher plants to take up these compounds; however, until recently, relatively little consideration has been given to their metabolism in plants (Fu et al., 2017a; He et al., 2017; Huber et al., 2012, 2009). Recent studies have shown that higher plants can metabolize xenobiotics similarly to humans with phase I modification reactions followed by phase II conjugation reactions using detoxification enzymes that function as a ‘green liver’ (Sandermann,

1994). In higher plants, phase I and phase II reactions are followed by a phase III sequestration, resulting in the formation of bound residues (Sandermann, 1994). Many of these studies have also highlighted a chemical-specific and species-specific nature of plant metabolism of pharmaceuticals.

In this study, we examined the uptake and biotransformation of diazepam in higher plants. Arabidopsis thaliana cells were used for an initial kinetic evaluation and metabolic profiling (Huber et al., 2009; van Poecke and Dicke, 2004). Cucumber

(Cucumis sativus) and radish seedlings (Raphanus sativus) were then used under hydroponic conditions to understand metabolism of diazepam and its effect on selected metabolic enzymes in whole plants.

3.2 Materials and Methods

3.2.1 Chemicals and solvents

Analytical standards of non-labeled diazepam, nordiazepam, oxazepam, temazepam, and oxazepam-glucuronide were purchased from Sigma-Aldrich (St. Louis,

MO). Diazepam-d5 was purchased from Fisher Scientific (Fair Lawn, NJ) and 14C- labeled diazepam (50-60 mCi/mmol, N-methyl-14C) was purchased from American

Radiolabeled Chemicals (Saint Louis, MO). Standards were prepared in HPLC grade methanol and stored at -20 °C before use. HPLC grade acetonitrile and methanol were used for extraction along with ultrapure water. Mobile phases were prepared using

Optima™ grade methanol and deionized water (18 MΩ). All solvents were purchased from Fisher (Fair Lawn, NJ).

3.2.2 Incubation experiments using Arabidopsis thaliana cells

PSB-D A. thaliana cell line (CL84840) was purchased from the Arabidopsis

Biological Resource Center (ARBC) at Ohio State University (Columbus, OH) and cultured in a liquid culture suspension at 25 °C and 130 rpm in the dark. Cell cultures were maintained in accordance with the ARBC maintenance protocol (Gehring et al.,

2018). The A. thaliana seed culture was produced by inoculating 7 mL of cell culture into

43 mL fresh growth media, followed by 96 h cultivation at 25 °C on a rotary shaker (130 rpm) in the dark. After 96 h, 3 mL of the seed culture was inoculated into 27 mL fresh growth media to create an approximate initial cell density of 3.3 g (dry weight g L-1).

Flasks were spiked with 30 µL of a stock solution of diazepam (1.0 mg mL-1) and 10 µL of a 14C-diazepam stock solution (0.01 mCi mL-1) to yield an initial concentration of 1 µg mL-1 and a specific radioactivity of 7.4 × 103 dpm mL-1 with an initial methanol content of 0.13% (v/v). Simultaneously, autoclaving cell suspension flasks prepared control treatments before chemical spiking (non-viable cell control), flasks containing diazepam without cells (medium control), and flasks containing living cells without diazepam

(background control). Control treatments were used to determine adsorption, abiotic degradation, and potential toxicity to cells. Flasks were incubated for 120 h in triplicate and sacrificed at 0, 6, 12, 24, 48 and 96 h for sampling and analysis.

At each sampling time point, samples were collected and centrifuged at 13,000 g for 15 min in 50 mL polypropylene tubes. The supernatant was collected and stored at

-20 °C until further analysis. Cells were immediately stored at -80 °C and then freeze- dried for 72 h. After drying, each sample was spiked with 50 µL of 10 mg L-1 diazepam-

d5 as a surrogate for extraction-recovery calibration and extracted using a method from

Wu et al. (2012), with minor modifications. Briefly, cells were sonicated (50/60 Hz) for

20 min with 20 mL methyl tert-butyl ether and then 20 mL of acetonitrile and centrifuged at 13,000 g for 15 min. The supernatants were combined and concentrated to near dryness under nitrogen at 35 °C and then reconstituted in 1 mL of methanol. The cells were then extracted with 20 mL acidified deionized water (pH 3) and the supernatant was combined with the methanol extract for clean-up. Prior to clean-up, 100 µL of cell material extract and growth media were combined with 5 mL liquid scintillation cocktail I (R. J. Harvey

Instruments, Tappan, NY) to measure the radioactivity in the extractable form on a

Beckman LS500TD Liquid Scintillation Counter (LSC) (Fullerton, CA).

Clean-up was carried out using solid phase extraction (SPE) with 150 mg Waters

Oasis© HLB cartridges that were preconditioned with 7 mL methanol and 14 mL deionized water. Samples were loaded onto cartridges and then eluted with 20 mL methanol under gravity. The eluate was dried under nitrogen and further recovered in 1.5 mL methanol:water (1:1, v/v). After re-suspension extracts were transferred to microcentrifuge tubes and centrifuged at 12,000 g in a tabletop d2012 Micro-Centrifuge

(SciLogex, Rocky Hill, CT). Samples were further filtered through a 0.22-µm polytetrafluoroethylene membrane (Millipore, Carrigtwohill, Cork, Ireland) into 2 mL glass vials and stored at -20 °C before analysis. Extraction of growth media was done after adjusting the solution to pH 3 using HCl, and followed by SPE with Waters HLB cartridges, as described above. The extraction recoveries for the tissues and media were

88 ± 7% and 80 ± 14%, respectively.

After extraction, the cell matter was air dried, and a 10-mg subsample was removed and combusted on a Biological Oxidizer OX-500 (R. J. Harvey Instruments,

Tappan, NY) to determine the radioactivity in the non-extractable form. The evolved

14 CO2 was captured in 15 mL Harvey Carbon-14 Cocktail II (R.J. Harvey Instruments,

Tappan, NY) and analyzed on a LSC.

3.2.3 Whole plant hydroponic cultivation

Hydroponic cultivations were carried out using cucumber (Cucumis sativus cv. sumter) and radish (Raphanus sativus, cv. cherry belle) seedlings. Seeds were purchased from Lowes (Murrieta, CA) and germinated in a commercially labeled organic potting soil in a growth chamber (22°C 16 h day/20°C 8 h night cycle; a photosynthetic photon

2 1 flux density of 200 µmol/m s , relative humidity of 75-80%). After the appearance of the first true leaf, uniform seedlings were selected, rinsed with distilled water, and individually placed in amber jars containing 900 mL hydroponic solution (Oasis® 16-4-

17 hydroponic fertilizer 4.16 g L-1). After 3 d of adaption, plants were exposed to diazepam by spiking with 100 µL of the above stock solutions to reach a nominal concentration of 1 mg L-1 and an initial specific radioactivity of 2.5 × 103 dpm L-1. The cultivation lasted for 7 d.

A parallel treatment (low concentration/chronic contact) with an initial diazepam concentration at 1 µg L-1 and a specific radioactivity of 2.5×102 dpm was included to simulate more realistic exposure levels and to validate the high level treatments. The cultivation lasted for 28 d with the culture solution renewed every 3 d. Plant blanks

(hydroponic solution in jars with plants but without diazepam) and treatment blanks

(spiked culture solution in jars without plants) were placed alongside the treatment jars.

At the end of 7 d or 28 d cultivation, the seedlings were removed from the jars.

Before sample preparation, roots were rinsed thoroughly with distilled water. Harvested plants were separated into below ground biomass (roots) and above ground biomass

(shoots). Flowering buds from cucumbers were also separated to observe any potential for accumulation in fruits. Tissues were freeze-dried and then stored at -80 °C until analysis. A 0.2-g aliquot of the dried plant tissue was ground to a fine powder using a mortar and pestle. Samples were extracted and prepared as described above. Samples of hydroponic solution were filtered with 0.7 µM GF/F filter and acidified to pH 2 with 1 M

HCl, followed by SPE extraction using Oasis HLB cartridges.

3.2.4 Glycosyltransferase activity assay

To determine glycosyltransferase activity, fresh tissues of cucumber and radish plants were frozen in liquid nitrogen and homogenized with 2 mL 50 mM phosphate- buffered saline (pH 7.0) containing 1% soluble polyvinylpyrrolidone and 5 mM ethylenediaminetetraacetic acid (EDTA). The homogenate was centrifuged at 13,000 g and 4 °C for 20 min. The activity of glycosyltransferase was measured immediately by mixing 100 µL of supernatant with 0.95 mL of the reaction mixture containing 50 mM

PBS (pH 7.0), 2 mM MgCl2, 2 mM uridine 5'-diphosphoglucose, 3.125 mM 4- nitrophenyl β-D-glucuronide, 3.125 salicin and 0.95 mL of 1 mM 2,4,5-trichlorophenol

(TCP). The assay mixture was incubated at 30 °C for 30 min, and then stopped by adding

10 µL of phosphoric acid. After centrifugation at 13,000 g for 5 min, the supernatant was

collected and diluted (1:4; v/v) with HPLC grade acetonitrile and 0.1% trifluoroacetic acid (TFA).

The enzyme activity was determined using an Agilent 1200 series HPLC paired with UV detector and a Thermo Scientific Acclaim™ 120 C18 5-µm column (4.6 × 250 mm). An isocratic flow was set with 1 mL min-1 70:30 mobile phase A (acetonitrile and

0.1% TFA) and mobile phase B (water and 0.1% TFA) for 10 min. The TCP-glucoside was detected at 205 nm. A six-point (400 µM- 2000 µM) TCP standard calibration curve was used to determine activity.

3.2.5 Structural elucidation using mass spectrometry

Instrument analysis was performed on a Waters ACQUITY ultra-performance liquid chromatography (UPLC) combined with a Waters Micromass Triple Quadrupole

(TQD) mass spectrometer equipped with electrospray ionization interface [ESI (Waters,

Milford, MA)]. Separation was achieved using an ACQUITY UPLC C18 column (2.1 mm × 50 mm, 1.7 µm, Waters) at 40 °C. Mobile phases A and B consisted of deionized water (18 MΩ) and methanol, respectively, each acidified using formic acid (0.2%). The solvent gradient program, with respect to mobile phase A, was as follows: 0-0.5 min

80%; 0.5-2.5 min 45%; 2.50- 3.5 min 10%; 3.5-4.0 80%; followed by equilibration for

1.00 min. The flow rate was 0.4 mL min-1, and the injection volume was 5 µL. Mass data were acquired using MassLynx® (Waters) multiple reactions monitoring (MRM) in the

ESI positive mode. The specific instrument settings were: capillary voltage 0.69 kV, collision gas (Aragon, 99.9%), dwell time 0.022 s, source temperature 150 °C,

7 8

desolvation temperature 450 °C, desolvation gas 900 L h-1 and cone gas 50 L h-1 (Table

S1).

3.2.6 Data analysis and quality control

All treatments in the A. thaliana cell incubation experiment were conducted in triplicate, and all hydroponic cultivations were conducted using four replicate jars containing individual plants to account for potential loss of plants. Calibration curves

(ranging from 5 to 1000 ng mL-1) with standards of diazepam, diazepam-d5, nordiazepam, temazepam, oxazepam and oxazepam-glucuronide were used for quantification with the r2 values of at least 0.99 for all analytes. A limit of detection

(LOD) of 1 ng mL-1 and a limit of quantification (LOQ) of 3 ng mL-1 for diazepam and its metabolites (except oxazepam-glucuronide) were determined through preliminary experiments. For oxazepam-glucurnonide the LOD was 3 ng mL-1 and the LOQ was 5 ng mL-1. LODs and LOQs were calculated based on a signal to noise ratio of 3 and 10, respectively.

Individual peaks were detected and integrated using TargetLynx XS software from MassLynx platform (Waters). Data were analyzed with StatPlus (Walnut, CA) and graphed using Prism 6 GraphPad software (La Jolla, CA). Results were calculated as the mean ± standard deviation (SD) (n=3). The Student's t-test was used to test significant differences in the extractable and non-extractable radioactivity and glycosyltransferase activity at α = 0.05. Systematic differences in the concentration of diazepam in plant tissues were assessed using one-way ANOVA with Fisher’s Least Significant Difference post-hoc (α = 0.05).

3.3 Results and discussion

3.3.1 Kinetics of diazepam in A. thaliana cell cultures

Active plant metabolism of diazepam was validated using a range of controls. No diazepam was detected in the non-treated media or the cell blanks, and there was no significant degradation of diazepam in the cell-free media, suggesting no contamination or significant abiotic transformation. Moreover, no significant difference (p > 0.05) was seen in cell mass between the chemical-free control and the treatments, indicating that diazepam did not inhibit the growth of A. thaliana. Furthermore, no significant amount of diazepam was adsorbed to the cell matter in the non-viable cell control. In contrast, diazepam dissipated appreciably from the media containing viable cells, with the average concentration decreasing from 698 ± 41.5 to 563 ± 8.93 ng mL-1 after 120 h of incubation, a decrease of nearly 20% (Figure 1a).

Parallel with the dissipation in the medium, diazepam was detected in the A. thaliana cells, with the highest level appearing after 48 h and a substantial decrease thereafter (Figure 1b). The decrease in diazepam level in the cell fit a first-order decay model and yielded a half-life of about 68 h (r2 = 0.97). This half-life was in comparison to a biological half-life of 48 h in humans (Dhillion, 1982), indicating a moderate persistence in plant cells.

3.3.2 Metabolism of diazepam in A. thaliana cells

Out of the four known diazepam metabolites only nordiazepam and temazepam were detected in the A. thaliana cells over the 120 h incubation. Temazepam was

detected first, with the highest concentration (645 ± 50.4 ng g-1) being observed at 12 h, which was followed by a decrease to 58.6 ± 17.0 ng g-1 at the end of the 120 h cultivation. Nordiazepam gradually increased over the 120 h incubation time from 128 ±

61.0 ng g-1 at 6 h to 535 ± 92.0 ng g-1 at the end of incubation (Figure 1c). These results correlated with their behavior in the human body, as nordiazepam displayed one of the most prolonged biological half-lives of the benzodiazepine family (50 to 120 h), while temazepam had a significantly shorter half-life (8 to 20 h) (Yasumori et al., 1993). The parallels observed between human and plant metabolism in this study and others (e.g., Fu et al., 2017a; Huber et al., 2012, 2009; LeFevre et al., 2015; Marsik et al., 2017) is intriguing, as it indicates that we may be able to use the knowledge of biologically active metabolites formed during human metabolism as a guide to study their formation and longevity in environmental compartments such as higher plants.

The complementary use of 14C labeled diazepam facilitated the determination of the fraction of diazepam and its metabolites that were incorporated into the cell matter

(bound residue fraction), which could not be determined using traditional extraction and analytical methods. We observed that the radioactivity in the media decreased while the extractable and bound residue fractions increased over the 120 h incubation (Figure 2).

The extractable radioactivity in the viable cells increased to 113 ± 31 dpm g-1 at 120 h

(Figure 2b). The bound residues (non-extractable radioactivity) increased steadily to a final level of 1120 ± 224 dpm g-1 (Figure 2c), indicating that A. thaliana cells were capable of metabolizing and then sequestering diazepam and its metabolites, likely in

vacuoles and cell walls. The formation of these bound residues is commonly regarded as a detoxification pathway of xenobiotics in higher plants.

3.3.3 Uptake and translocation of diazepam in cucumber and radish seedlings

Diazepam was found in the cucumber and radish seedlings following a 7 d cultivation at the higher concentration and a 28 d of cultivation following treatment at the lower concentration (Figure 3). After treatment with 1 mg L-1 with diazepam for 7 d, a significantly higher concentration of diazepam was observed in the roots as compared to the shoots in radish seedlings (p < 0.05) whereas diazepam was more evenly distributed throughout the entire plant of cucumber seedlings (Figure 3a). However, after the 28 d cultivation following the lower concentration treatment, this pattern appeared to be different for both plant species. In the radish plants, diazepam was more evenly distributed in the roots, but was significantly lower in the shoots (p < 0.01). In the cucumber plants there was a significantly higher concentration in roots (p < 0.01) and a significantly lower concentration in the shoots (Figure 3b, p < 0.05). These differences may be due to variations in metabolism between the two species, as well as dynamic changes as a function of contact time in both plant growth and its ability to metabolize and translocate diazepam.

The bioconcentration factors for the roots (BCFR) and shoots (BCFS) were

14 calculated as the ratio of C activity in plant tissue (Ctissue) to that in the media (Cmedia):

(1)

For the radish plants cultivated for 7 d the average BCFR and BCFS were 0.49 and

1.13, respectively. For the 7 d cucumber plants, BCFR and BCFS were 0.78 and 3.70, respectively (Table S2). The higher BCF values found for the shoots were in disagreement with Wu et al., (2013). It is possible that the discrepancy between the studies was due to the use of 14C-labeling in the current study, as both extractable and non-extractable parent compound and its metabolites were included in the calculation of

BCFs in this study, whereas only the extractable form of the parent compound was used for calculation in the previous study. It may be argued that the use of 14C accounted for both parent and metabolites in extractable and non-extractable forms, and therefore reflected the overall accumulation of diazepam in plants (Dodgen et al., 2014; Fu et al.,

2017b; Trapp et al., 1990).

The translocation factor (TF) was similarly calculated using the ratio of 14C activity in the shorts (Cshoots) to that in the roots (Croots):

(2)

For the 7 d cultivation experiment, TF values for radish and cucumber plants were

3.5 and 4.7, respectively. For the 28 d cultivation experiment, the TFs increased for both plants to 5.9 and 5.0, respectively. These TF values were much greater than that reported in Wu et al., 2013. However, these results were similar to the predicted values for weakly basic compounds (Table S3) under weakly basic conditions (pH 7.81) (Trapp, 2009).

Again, the higher TFs observed in this study may be similarly attributed to the inclusion of the non-extractable residues and metabolites in the calculations.

3.3.4 Metabolism of diazepam in cucumber and radish seedlings

Similar metabolites to those in A. thaliana cells were found in seedlings grown in the nutrient solution spiked with diazepam, with nordiazepam being predominant (Figure

4). In the 7 d and 28 d cultivation experiments, temazepam was found to be the second major metabolite in the leaves of the cucumber seedlings, and the level was higher in the

7 d cucumber seedlings than the 28 d plants (Figure 4a,b). Oxazepam was detected in the leaves of both plant species after the 7 d cultivation (Figure 4a,c). The higher accumulation of diazepam and the biologically active metabolites in the leaves may have ecotoxicological ramifications; for example, many insects consume leaves, even if they are not edible tissues for humans (Pennington et al., 2017a).

Our results were in agreement with recent findings in Carter et al. (2018), in which they observed the formation of nordiazepam, temazepam and oxazepam in radish and silverbeet plants exposed to diazepam and chlordiazepoxide. They similarly showed nordiazepam to be the major metabolite with oxazepam and temazepam constituting a much smaller fraction at the end of 28 d cultivation in soil. However, in that study, the authors did not track the formation of these metabolites over time or influence of treatment concentrations.

Phase III metabolism appeared to increase from the 7 d to 28 d cultivation for both radish and cucumber seedlings (Table 1). Between the plant species, the cucumber seedlings had a greater fraction of non-extractable radioactivity in comparison to the

radish seedlings (Table 1). In the 7 d cultivation experiment, the mass balances came to

99.3% for the cucumber plants but only 58.1% for the radish seedlings (Table 1). Due to the multiple water changes (Figure S1), a complete mass balance was not attainable for the 28 d cultivation experiment. However, when a proxy mass balance (sum of roots and shoots) was calculated for both species, a similar pattern was observed. A total of 83.0% of the added 14C radioactivity was calculated for the cucumber treatments while the fraction was 61.3% for the radish plants. This could be due to increased mineralization in

14 the growth media and respiration of CO2 through plant in the radish cultures. As mineralization is viewed as the final stage of detoxification (Dodgen et al., 2014), it is likely that the radish plant was more efficient in their ability to detoxify diazepam than cucumber plants. The Brassicaceae family, which includes the common radish, has been shown to be effective for phytoremediation due to their possession of genes that increase tolerance to stressors (Peer et al., 2006) and activation of enzymes capable of extensive biotransformations (Piotrowska-Długosz, 2017).

3.3.5 Diazepam-induced changes to glycosyltransferase activity

The activity of glycosyltransferase was measured in the control seedlings as well as seedlings exposed to diazepam for the 7 d and 28 d cultivation experiments (Figure 5).

Glycosyltransferase catalyzes the transfer of sugars, such as glucuronic acid, to many types of acceptor molecules, including xenobiotics (Gachon et al., 2005). The conjugation of glucuronic acid with oxazepam is the major detoxification pathway of diazepam in humans (Sonne, 1993).

No detectable level of oxazepam-glucuronide was observed in radish or cucumber seedlings for either the 7 d or 28 d cultivation. However, there was a significant difference in the glycosyltransferase activity in radish seedlings treated with diazepam for

7 d and 28 d, although a distinct pattern in the changes of the enzyme activity was absent

(Figure 5). For the 7 d cultivation experiment, a significant decrease in glycosyltransferase activity was observed in the shoots of radish seedlings when compared to the control (Figure 5c, p < 0.01). In contrast, no significant change in glycosyltransferase activity was observed in the shoots of cucumber seedlings when exposed to diazepam (Figure 5a). In the 28 d cultivation experiment, only the cucumber seedlings exhibited significant differences in the enzyme activity, with an increase in activity detected in the shoots (p < 0.01) and a decrease in the cucumber buds (Figure 5b, p < 0.01).

Even though we did not detect oxazepam-glucuronide in the exposed plants, changes in the glycosyltransferase activity indicated that conjugation might have occurred with the parent and its metabolites, including those not examined in this study, or at levels below our detection capability. In addition, it may be postulated that rapid phase III metabolism may have limited the accumulation of such conjugates in the plant tissues, making the conjugates transient metabolites. In previous studies, glycosyltransferase was observed to catalyze the detoxification of ibuprofen in Phragmite australis during a 21 d exposure (He et al., 2017). Further, the formation of a glucose conjugate has been considered to be a major detoxification pathway for several environmental contaminants (e.g., Ando et al., 2015; Fu et al., 2017a, 2017b). These

studies together suggest the importance of phase II metabolism in the metabolic fate of pharmaceuticals in higher plants.

3.4 Conclusions

Agricultural irrigation using TWW may pose unintended risks, such as exposure to pharmaceuticals by humans through dietary intakes and unexpected ecological consequences following consumption of pharmaceuticals by terrestrial organisms such as insects. In this study, the pharmaceutical diazepam taken up and metabolized by

Arabidopsis thaliana cells, as well as cucumber and radish seedlings. The uptake resulted in extensive phase I metabolism, as demonstrated by the formation of the three major diazepam metabolites, nordiazepam, temazepam and oxazepam. However, the three plant species displayed notable metabolic differences. Further, exposure to diazepam resulted in alteration of glycosyltransferase that is involved in phase II metabolism. The observed interaction between a plant’s ability to accumulate and metabolize diazepam and diazepam’s apparent ability to affect a plant’s detoxification enzymes warrants further consideration, as it may be indicative of other previously unobserved metabolites. Results from this study a present new insight on plants’ ability to take up and metabolize pharmaceuticals, including their ability to create other biologically active compounds.

This and other studies highlight the need to consider plant metabolism and bioactivation in multiple crops and under realistic environmental conditions when assessing the safety of TWW reuse for agricultural irrigation.

Tables

Table 1. Extractable and non-extractable radioactivity in radish and cucumber plants

cultivated for 7d and 28d. The values are shown as mean ± SD (n=3). (a) Significant

differences (α = 0.05, student t-test) relative to the inter-plant tissues. (b) Significant

differences (α = 0.05, student t-test) relative to intra-plant tissues

Radioactivity Media Shoots Roots Buds 7 d Extractable 30.8% ± 3.8 12.9% ± 10a 9.35% ± 3.6 N.A. Radish 7 d Non- Extractable N.A. 2.0% ± 0.4a 2.95% ± 1.4 N.A. Radish 28 d Extractable N.A. 2.23% ± 1.0b 11.6% ± 1.0b N.A. Radish 28 d Non- Extractable N.A. 37.3% ± 6.4 10.4% ± 4.0 N.A. Radish 7 d Extractable 21.9% ± 12 34.0% ± 7.5ab 10.0% ± 5.1b N.A. Cucumber 7 d Non- Extractable N.A. 28.6% ± 1.09ab 4.65% ± 2.3b N.A Cucumber 28 d Extractable N.A. 13.3% ± 10b 10.3% ± 4.8b 0.96% ± 0.4b Cucumber 28 d Non- Extractable N.A. 30.3% ± 8.4b 13.5% ± 9.5b 14.5% ± 2.5b Cucumber

Figures

Figure 1. Diazepam in the medium and Arabidopsis thaliana cells. (A) Kinetics

of diazepam in cell growth media (B) Kinetics of diazepam in viable and non-

viable cells. C) Kinetics of nordiazepam and temazepam in viable cells. The values

are shown as mean ± SD (n=3)

Figure 2. Radioactivity in Arabidopsis thaliana cells. (A) Radioactivity in cell growth media (B) Extractable radioactivity in viable and non-viable cells. (C)

Non-extractable radioactivity in viable and non-viable cells. The values are shown as mean ± SD (n=3)

Figure 3. Diazepam in radish and cucumber plants. (A) Concentration of diazepam in radish and cucumber seedlings cultivated for 7 d. (B) Concentration of diazepam in radish and cucumber seedlings cultivated for 28 d. The values are shown as mean ± SD

(n=3). *p < 0.05 **p < 0.01 ANOVA (one-way).

Figure 4. Diazepam metabolites in radish and cucumber seedlings. (A) Concentration of diazepam metabolites in cucumber seedlings cultivated for 7 d. (B) Concentration of diazepam metabolites in cucumber seedlings cultivated for 28 d. (C) Concentration of diazepam metabolites in radish seedlings cultivated for 7 d. (D) Concentration of diazepam metabolites in radish seedlings cultivated for 28 d. The values are shown as mean ± SD (n=3).

Figure 5. Glycosyltransferase activity in radish and cucumber seedlings. (A) glycosyltransferase activity in cucumber seedlings cultivated for 7 d. (B) glycosyltransferase activity in cucumber seedlings cultivated for 28 d. (C) glycosyltransferase activity in radish seedlings cultivated for 7 d. (D) glycosyltransferase activity in radish seedlings cultivated for 28 d. The values are shown as mean ± SD

(n=3). (a) Significant differences (α = 0.05, student t-test) relative to the intra-plant tissues. (b) Significant differences (α = 0.05, student t-test) relative to inter-plant tissues

Supplementary Information

Table 1: Optimized MRM conditions for the analysis of the chosen analytes by UPLC

TQD MS/MS (CV: cone voltage in V; CE: collision energy in eV)

Table 2: Bioconcentration Factors and Translocation Factors for cucumber and radish seedlings during 7d and 28d cultivation

BCFR BCFS TF

Cucumber 0.78 3.70 3.5 (7d) Cucumber N/A N/A 5.9 (28d) Radish (7d) 0.49 1.13 4.7

Radish (28d) N/A N/A 5.0

Table 3: Properties of diazepam and it major metabolites

1,2,3 1 Name CAS No Chemical Pka Log Kow Structure Formula

Diazepam 439-14-5 C16H13ClN2O 3.4 2.82

Nordiazepam 1088-11-5 C15H11ClN2O 2.85 2.79

Temazepam 860-50-4 C16H13ClN2O2 1.6 2.16

Oxazepam 604-75-1 C15H11ClN2O2 1.7 2.01

Oxazepam- 6801-81-6 C21H19ClN2O8 -0.87 1.37 Gluconoride

1O’Neil, M.J., 2006; 2Wu et al., 2015; Wishart DS, et al., 20183

Figure 1: Percent nominal radioactivity in the hydroponic solutions during 28 d cultivation and plant free controls

Chapter 4: Bioaccumulation, metabolism and biochemical effects of

contaminants of emerging concern in the earthworm species Eisenia

fetida

Abstract

Contaminants of emerging concern (CECs) present in recycled water and biosolids pose a potential threat to soil dwelling organisms when land applied. However, the fate of CECs in these organisms is relatively poorly understood. In this study we assessed the uptake, metabolism, and impacts of three pharmaceuticals, i.e., sulfamethxazole, diazepam, and naproxen, and one cosmetic preservative, i.e., methyl paraben in earthworms (E. fetida) exposed in an artificial soil. Liquid chromatography paired with mass spectrometry and enzyme assays were used to characterize sorption, uptake, transformation, and biochemical effects of the CEC exposure in the artificial soil matrix and earthworms, including alterations to antioxidant enzymes associated with oxidative stress (i.e., super oxide dismutase, catalase, and glutathione-S-transferase). The presence of earthworms did not significantly affect sorption and Kd values ranged from

(0.71 ± 0.2 to 11.0 ± 4.7) for all four CECs. Sulfamethoxazole, diazepam and naproxen were accumulated in earthworm tissues up to 14 d, followed by a decrease in their levels to 21 d. The primary metabolite of sulfamethoxazole, N4-acetylsulfamethoxazole, was detected in earthworm tissues after 1 d of exposure. The N4-acetylsulfamethoxazole concentration peaked at 3 d and decreased thereafter. N4-acetylsulfamethoxazole was also found in the soil where the concentration peaked at 7 d. The major metabolites of

naproxen and methyl paraben, i.e., o-desmethylnaproxen, and p-hydroxybenzoic acid, were detected in the soils, but not in the earthworm tissues, suggesting active metabolism and excretion. Exposure to the CECs resulted in up-regulation of antioxidant enzymes.

Activity of glutathione-S-transferase increased continuously from 3 d of exposure, superoxide dismutase increased within 1- 3 d and catalase activity was enhanced by 21 d.

Collectively, the enzyme assays indicated that exposure to CECs caused oxidative stress in E. fetida. This study highlights the need to consider the role of, and effects on terrestrial invertebrates when understanding risks from CECs in agroecosystems.

Key words: Contaminants of emerging concern, Ecotoxicity, Eisenia fetida, Metabolism

4.1 Introduction

Water scarcity has led to continuously increasing use of municipally treated wastewater (TWW) in agro-environments, especially in arid and semi-arid regions

(Lazarova et al., 2001; Tal, 2006). Similarly, the use of municipal biosolids to improve soil health is increasing (Lu et al., 2012). The land application of TWW and biosolids can introduce contaminants of emerging concern (CECs) into terrestrial environments (Petrie et al., 2015). Consequently, a range of CECs have been detected in agricultural soils

(Walters et al., 2010). Literature pertaining to the effects of CECs in terrestrial ecosystems is, however, limited (Boxall et al., 2012; Carter et al., 2014; Petrie et al.,

2015; Wu et al., 2015). The majority of previous research has been concerned with the fate and effects of CECs in plants (Christou et al., 2017; C. Sun et al., 2018; Wu et al.,

2014), and only a few studies have considered CECs in terrestrial invertebrates (Carter et al., 2014; Liu et al., 2010; Pennington et al., 2018; 2017).

One of the most important invertebrates in agricultural fields is earthworm.

Earthworms ameliorate agricultural soil structure through the formation of new aggregates and macropores; improving soil tilth, aeration, infiltration, and drainage

(Lavelle, 1988). Furthermore, earthworms consume plant litter, recycle organic matter and aid in nutrient cycling (Curry and Schmidt, 2007). Earthworms dominate soil fauna with an average biomass of 10 – 200 g m-2 (Laskowski et al., 1988). Due to their ecological importance and abundance, earthworms are a good candidate for ecotoxicity testing (Davies et al., 2003). Herein we sought to understand some of the potential consequences of CECs exposure in earthworms. For this study, we selected the

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earthworm species Eisenia fetida as the test organism due to their widespread use in the scientific literature and extensive habitat range.

For CECs, we considered three pharmaceuticals, i.e., naproxen, diazepam, and sulfamethoxazole, and one cosmetic preservative, i.e., methyl paraben. These four compounds were selected due to their range of physicochemical properties and frequent detections in the environment (Focazio et al., 2008; Lu et al., 2018; Silva et al., 2011).

Naproxen is a commonly consumed nonsteroidal anti-inflammatory drug that has been often found in TWW and biosolids (Straub and Stewart, 2009). Diazepam is a psychoactive compound from one of the most commonly prescribed classes of pharmaceuticals of wastewater treatment plants. Diazepam has also been frequently detected in TWW due to poor removal efficiency (Bachhuber et al., 2016; Verlicchi et al.,

2012). Sulfamethoxazole is an antibiotic and has garnered significant scientific interest due to the growing concern over antibiotic resistance (Białk-Bielińska et al., 2014).

Methyl paraben is a preservative that is widely used in various cosmetic products, is amongst the most frequently detected parabens in TWW and biosolids, and is a known endocrine disruptor (Błędzka et al., 2014).

Several studies have examined the fate and toxicity of these compounds in aquatic organisms (e.g., Jones et al., 2002; Quinn et al., 2008; Yamamoto et al., 2011). The observed adverse effects on aquatic organisms have raised concerns about the unintended consequences from widespread consumption and, ultimate release of the CECs into the aquatic environment (Daughton and Ternes, 1999). With the increasing use of TWW and biosolids for agriculture, it is crucial to also understand the effects of these compounds on

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terrestrial organisms.

In this study, we carried out laboratory experiments to assess the potential uptake, biotransformation, and biochemical effects of CECs in earthworms. Eisenia fetida was exposed to the four CECs in an artificial soil, and kinetics of the parent compound, uptake and metabolite formation were evaluated. Changes in enzymes associated with oxidative stress (i.e., superoxide dismutase, glutathione-S-tranferase, and catalase) and lipid peroxidation (determined by the formation of malondialdehyde) were assessed as biochemical markers of potential toxicity.

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4.2 Materials and Methods

4.2.1 Chemicals and solvents

Diazepam, naproxen, methyl paraben, nordiazepam, o-desmethylnaproxen, p- hydroxybenzoic acid, and naproxen-d3 were purchased from Sigma-Aldrich (St. Lewis,

MO). Sulfamethoxazole was purchased from MP Biomedicals (Santa Ana, CA).

Diazepam-d5 was purchased from Fisher Scientific (Fair Lawn, NJ), sulfamethoxazole- d4 and N4-acetylsulfamethoxazole were purchased from Santa Cruz Biotechnology

(Santa Cruz, CA), and methyl paraben-d4 was purchased from Toronto Research

Chemicals (Toronto, Canada). Standards were prepared in HPLC grade methanol and stored at -20 °C before use. HPLC grade acetonitrile and methanol were used for extraction along with ultrapure water. Mobile phases were prepared using Optima™

LC/MS grade methanol and deionized water (18 MΩ). All solvents were purchased from

Fisher (Fair Lawn, NJ).

4.2.2 Test Soil

An artificial soil was used in this study to limit factors, which was in accordance with OECD guideline 317 (OECD 317, 2010). Fine grain quartz sand was purchased from Carolina Biological (Burlington, NC). Peat moss was purchased from Four Winds

Trading (Wheat Ridge, CO). Kaolinite clay was purchased from Pantai Chemical USA

(Alpharetta, GA). The different components were homogenized by mixing for 30 min, moistened to 50% with DI water, and adjusted to pH 6.0 with powdered calcium carbonate (Fisher, NJ).

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4.2.3 Test Organism

The earthworms Eisenia fetida was purchased from Uncle Jim’s Worm Farm

(Spring Grove, PA). Prior to the experiment, the worms were cultured in a medium of cow manure and peat (1:1), moistened with DI water and maintained at the room temperature (21 ± 1 °C). The worms were fed weekly with dehydrated potato flakes

(Laura J. Carter et al., 2014). Adult earthworms (E. fetida) with fully developed clitellum and weighing 200 - 500 mg were selected for the following experiments.

4.2.4 Bioaccumulation Experiment

Preliminary experiments were carried out to assess any potential mortality from the test compounds on E. fetida (Table S1). The uptake and bioaccumulation of the test compounds followed OECD guideline on “Bioaccumulation in Terrestrial Oligochaetes.”

Tests were performed in glass jars painted black and then white to reduce light and heat absorption. Jars contained 150 g ± 0.5 (d.w.) of artificial soil with 3 worms in each container. The worms were allowed to acclimate to the test conditions for 24 h before exposure to the test compounds. The incubation was carried out at room temperature.

Prior to spiking, earthworms were removed from jars and soils were spiked with different volumes of the standard stock solutions (1 mg mL-1) to arrive at initial concentrations of 70, 50, 275, and 200 ng g-1 for sulfamethoxazole, diazepam, naproxen and methyl paraben, respectively. Soils were moistened with deionized water, homogenized by stirring, and the earthworms were then reintroduced. Concentrations were selected based on environmental concentrations previously reported in the literature

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(McClellan and Halden, 2010; Sabourin et al., 2012; Walters et al., 2010). Moisture content of 50% (w/w) was selected based on preliminary experiments and maintained during the incubation. Jars containing the spiked soil without E. fetida (non-earthworm control) and non-spiked soil with E. fetida (background control) were prepared and maintained simultaneously. At the start of the incubation, individual mature worms were maintained in separate jars to assess treatment-induced weight changes, if any.

Samples were taken at 0 h, 1 d, 3 d, 7 d, 14 d, and 21 d of incubation. At each sampling time point, four treatment and four control jars were harvested for a total of twelve worms per time point per treatment. Worms were collected, rinsed with deionized water and placed in Petri dishes with a moistened paper towels for 24 h to purge their gut content. They were then weighed, frozen in liquid nitrogen and stored at -80 °C until extraction.

The earthworms were homogenized with 8 mL acetonitrile:H2O (70:30; acidified to pH 3 using acetic acid) for 5 min using a Kinematica™ Polytron PT 10/35 GT

Benchtop Homogenizer (Bohemia, NY). The CECs were extracted from the homogenized using 10 min of sonication, followed by 15 min of centrifugation at 15000 g. The supernatants were collected, dried under nitrogen, and reconstituted using 1.5 mL methanol:H2O (80:20, v/v). The reconstituted extracts were placed in LC-vials for analysis.

Porewater was collected using 20 g (wet weight) of soil by centrifugation at

15000 g for 20 min, after which 2 mL of water was withdrawn and further centrifuged at

12000 g for 15 min. The resulting supernatant was used for instrument analysis. The soil

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was extracted by vortexing 10 g (w.w.) of soil with 10 mL acetonitrile:H2O (70:30, v/v, pH 3) for 5 min, followed by sonication for 20 min. Samples were centrifuged at 15000 g for 20 min, and the supernatant was collected, dried under nitrogen and reconstituted in

1.5 mL methanol:H2O (80:20, v/v). All extracts were filtered using 0.2 µm PTFE syringe filters before instrument analysis. Extraction efficiencies were assessed using deuterated standards and are given in the Supplementary Information (Table S2).

4.2.5 Calculation of Adsorption Coefficient, Bioconcentration Factor, and

Bioaccumulation Factor

The distribution coefficient (Kd) was calculated for each time point has the ratio of the individual CEC in the soil (CSorbed) to the concentration in the soil porewater

(CDissolved).

!" ! ( ) !"#$%& ! �! = !" (1) ! !"##$%&'! !"

The bioconcentration factor was calculated has the ratio of the concentration of the individual CEC in the earthworm tissue (Ctissue) to that of the concentration in the soil porewater (Cporewater).

!" ! !"##$% ! �� = !" (2) ! !"#$%&'$# !"

The bioaccumulation factor was calculated has the ratio of the concentration of the individual CEC in the earthworm tissue (Ctissue) to that the concentration in the soil porewater (Cporewater).

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!" ! ( ) !"##$% ! �� = !" (3) ! ( ) !"#$ !

4.2.5 Assay of Antioxidant Enzyme Activities in E. fetida

Earthworm tissues (0.20 g ± 0.05) were frozen in liquid nitrogen and then homogenized with 2 mL of 50 mM potassium phosphate buffer (PBS) (pH 7.4) with 1% polyvinylpyrrolidone (PVP) and 1 mM ethylenediaminetetraacetic acid (EDTA) using a

Kinematica™ Polytron PT 10/35 GT Benchtop Homogenizer (Bohemia, NY). The homogenate was then centrifuged at 12 000 g for 20 min at 4 °C (Song et al., 2009). The resulting supernatant was used for enzyme activity assays as described below.

The activities of glutathione-S-transferase (GST), catalase (CAT) and superoxide dismutase (SOD) were determined as in Sun et al. (2014). To determine GST (EC

2.5.1.18) activity, 100 µL of supernatant was combined with 2 mL of a reaction mixture containing 50 mM PBS (pH 7.4), 5 mM glutathione (reduced), and 1 mM 1-Chloro-2,4,- dinitrobenzene (CDNB) dissolved in 96% ethanol. The GST activity was measured at

340 nm for 3 min and the concentration was calculated using the GSH-CDNB adduct synthesis (ε=9.6 mM-1 cm-1). The CAT activity was determined by combining 200 µL of supernatant with 3 mL reaction mixture containing 10 mM H2O2 in 50 mM PBS buffer

(pH 7.4). The concentration was calculated by following the consumption of H2O2 at 240 nm for 3 min (ε=39.4 mM-1 cm-1). The activity of SOD was determined by combining

100 µL supernatant with 3 mL reaction mixture containing 50 mM PBS buffer (pH 7.4),

13 mM methionine, 75 µM nitro blue tetrazolium (NBT), 2 µM riboflavin, and 0.1 mM

EDTA. The mixture was illuminated for 15 min at a light intensity of 5,000 lux for 15

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min. One unit of SOD activity was defined as the concentration of enzyme required to cause 50% inhibition of NBT when monitored at 560 nm.

The protein content was used to standardize enzyme activity and determined by combining 5 mL of Coomassie Brilliant Blue G-250 reaction mixture and 100 µL of supernatant and use to standardize enzyme activity. Concentration was calculated from a six-point standard curve (0.01mg L-1 – 1.0 mg L-1) using bovine serum albumin monitored at 595 nm (Wang et al., 2015).

4.2.6 Assay of Lipid Peroxidation

The detection of malondialdehyde (MDA) was used to determine lipid peroxidation, as MDA results from lipid peroxidation of polyunsaturated fatty acids

(Davey et al., 2005). Earthworm tissues (0.2 ± 0.05 g) were frozen using liquid nitrogen and homogenized with 5 mL 0.1% trichloroacetic acid using a benchtop homogenizer, followed by centrifugation at 12000 g for 15 min. A 1.0 mL aliquot of the supernatant was combined with 4 mL of 20% trichloroacetic acid containing 0.5% thiobarbituric acid, incubated at 95 °C for 30 min and then cooled in an ice bath. The mixture was centrifuged at 12 000 g for 10 min, and monitored at an absorbance of 532 nm and a non- specific absorbance at 600 nm. The MDA concentration was calculated using its extinction coefficient ε=155 mM-1 cm-1 (Wang et al., 2015).

4.2.7 CEC Quantification using Mass Spectrometry

Instrument analysis was performed on a Waters ACQUITY ultra-performance liquid chromatography (UPLC) combined with a Waters Micromass triple quadrupole

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(TQD) mass spectrometer equipped with an electrospray ionization interface (ESI)

(Waters, Milford, MA). Separation was achieved using an ACQUITY UPLC C18 column

(2.1 mm × 100 mm, 1.7 µm, Waters) at 40 °C. Mobile phases A and B consisted of deionized water (18 MΩ) and methanol, respectively, each acidified using 0.1% formic acid. The solvent gradient program, with respect to mobile phase A, was as follows: 0-0.5 min, 80%; 0.5-9.0 min, 50%; 9.0-10.0 min, 0%; 10.0-12.0 min, 80%; followed by equilibration for 1.00 min. The flow rate was 0.4 mL min-1, and the injection volume was

5 µL. Mass data were acquired using Masslynx® (Waters) multiple reactions monitoring

(MRM) in the ESI positive mode. The specific instrument settings were: capillary voltage 1.72 kV, collision gas (Aragon, 99.9%), dwell time 0.016 s, source temperature

150 °C, desolvation temperature 500 °C, desolvation gas 1000 L h-1 and cone gas 50 L h-

1. Cone voltage and collision energy are listed in the SI (Table S3).

4.2.8 Data analysis and quality control

All treatments in the E. fetida incubations experiments contained four replicates and mortality, if any, was assessed immediately upon stoppage. Standard calibration curves (ranging from 5 to 1000 ng mL-1) with r2 values of at least 0.98, were made from standards of diazepam, naproxen, methyl paraben, sulfamethoxazole, nordiazepam, o- desmethylnaproxen, p-hydroxybenzoic acid, N4-acetylsulfamethoxazole, diazepam-d5, naproxen-d3, sulfamethoxazole-d4, methyl paraben-d4 and used for quantification for all analytes. A limit of detection (LOD) of 1 ng mL-1 and a limit of quantification (LOQ) of

5 ng mL-1 were determined for all analytes, except for p-hydroxybenzoic acid that had a

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LOD of 3 ng mL-1 and an LOQ of 5 ng mL-1. LOD and LOQs were calculated from a signal to noise ratio of 3 and 10 respectively.

Compound peaks were detected and integrated using TargetLynx XS software

(Waters). Data were analyzed and graphed with StatPlus (Walnut, CA) and Prism 8

GraphPad software (La Jolla, CA). Results were calculated as the mean ± standard error

(SE), and a Student’s t-test or ANOVA (one-way) with a Tukey-Kramer post-hoc was used to assess the systematic difference between groups (α=0.05).

4.3 Results and Discussion

4.3.1 Distributions of CECs among soil, porewater and earthworm

The concentrations of CECs were monitored in three phases, soil, soil porewater, and earthworm tissue, in both the presence and absence (non-earthworm control) of earthworms throughout the 21 d incubation. To determine the potential effect of earthworms’ presence on the partitioning of the four CECs amongst the soil and soil porewater the distribution coefficient (Kd) was calculated at each time point and the differences between earthworm treatment and the non-earthworm controls were compared (Table 1). No significant differences in the Kd values (P > 0.05) were observed between the earthworm treatment and non-earthworm controls for any of the CECs, indicating that earthworms did not significantly affect the association of these CECs to the solid phase of the artificial soil.

-1 For diazepam, the Kd values were calculated to range between 0.84 to 6.56 mL g throughout the incubation. These Kd values were lower than those previously reported for

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diazepam in batch and field sorption measurements using agriculture soils (8.3 to 59.8

-1 mL g ) (Carter et al., 2018) but were in agreement with the low Kd values observed for pharmaceuticals in sandy soils (Chefetz et al., 2008). The low Kd values indicated that diazepam was not strongly adsorbed to the solid matrix of the artificial soil. The Kd for

-1 naproxen ranged between 0.87 to 11.0 mL g throughout the incubation. The low Kd values were consistent with those previously reported for naproxen in sandy soils [0.49

-1 mL g (Teijón et al., 2013)] The Kd values for sulfamethoxazole were similarly very low throughout the incubation, ranging from 0.71 to 1.75 mL g.-1 These values were consistent with those previously reported in the literature for grassland soils (1.3 mL g -1) and arable land soils (2.9 mL g -1), indicative of its high mobility in the soil environment

(Höltge and Kreuzig, 2007b). The derived Kd values, for methyl paraben in the earthworm treatment (2.99 mL g -1) and non-earthworm controls (4.00 mL g -1) could be calculated only for the initial sampling point as it rapidly disappeared from the soil and soil porewater. This may be due to rapid biodegradation in the soil and/or rapid metabolism in E. fetida (Lu et al., 2018; Mincea et al., 2010)(Lu et al., 2018; Mincea et al., 2010).

To verify active uptake of CECs by earthworms, a range of controls were used, including soil blanks and non-earthworm controls. None of the parent CECs were detected in the earthworm or soil blanks. However, degradation of both methyl paraben and sulfamethoxazole was observed in the non-earthworm soil, indicating that microbial and/or abiotic degradation of these compounds occurred in the media (Figure S1). The parent compounds of diazepam, sulfamethoxazole, and naproxen were detected in

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earthworms throughout the 21 d incubation, with the concentration of both diazepam and naproxen increasing to 53.8 ± 24 ng g-1 and 110 ± 25 ng g-1, peaking at 14 d. These results suggested that both diazepam and naproxen were being taken up and accumulated in E. fetida (Figure 2a,b). Sulfamethoxazole, on the other hand, appeared to have a relatively stable concentration in the earthworm tissues throughout the incubation (Figure

2b). However, this could be due to active metabolism of sulfamethoxazole in earthworm instead of limited uptake or accumulation. Methyl paraben was not quantifiable in earthworm tissues and was rapidly lost in the artificial soil.

For each of the three quantifiable CECs in the earthworm tissues (i.e., sulfamethoxazole, naproxen and diazepam), the bioconcentration factor (BCF) and bioaccumulation factor (BAF) were calculated and compared for each time point. No significant differences in the BAF were observed for sulfamethoxazole or naproxen over the course of the incubation (P > 0.05). However, the BAF for diazepam did significantly increase over time (P < 0.05), indicating that the increased exposure time to soil porewater resulted an increased concentration of diazepam in the earthworm tissues, likely due to slower metabolism or time needed for reach equilibrium. The BCF for diazepam and naproxen did not significantly change throughout the incubation period (P

> 0.05). For sulfamethoxazole the a significant difference in BCF was observed between the 3 d and the 14 d sampling points, but no clear pattern in BCF values over time was discernable.

A significant difference between BAF the BCF values were observed for sulfamethoxazole at 7 d (P < 0.05). There was a trend towards a significantly higher BCF

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than BAF for diazepam throughout the incubation. The trend may be due to increased uptake of the compounds from the soil porewater by E. fetida, which was consistent with several previous studies that showed dermal absorption via water to be the primary route for uptake of contaminants by worms ( Carter et al., 2014; Jager et al., 2003; Kure et al.,

2009). However, due to a lack of quantifiable replicates (s/n < 10) in soil or soil porewater statistical significance could not be assessed. Further research is necessary to understand the exposure pathways for polar CECs for invertebrates such as in earthworms in soils.

Intriguingly, it was also observed that all three quantifiable CECs displayed an similar pattern where there was an initial increase in BAF or BCF up to 3 or 7 d, followed by a decrease at 7 or 14 d, and followed by increases again till the end incubation (Figure

S1). This pattern may be indicative of early uptake and metabolism, followed by an insufficient response from detoxification enzymes, resulting in storage and accumulation of the compounds in the earthworm tissues, as was previously observed in aquatic organisms (Burkina et al., 2015). To the best of knowledge, this was the first time the

BCF and BAF have been calculated in earthworms for naproxen, diazepam and sulfamethoxazole.

While many studies have considered the bioconcentration/accumulation of different CECs in plants and earthworms (e.g.,Carter et al., 2014, 2016; Dodgen et al.,

2015; Macherius et al., 2014; Wu et al., 2013), very few studies have considered the changes to these values over time. The results from this study showed that it may be valuable to evaluate the BCF or BAF of CECs in earthworms over time, as there may be

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an increased potential for biomagnification at a given time point. However, more research is needed to determine if these differences persist for a longer time scale and if they occur in different soils or for other CECs.

4.3.2 Metabolism of CECs in E. fetida

Out of four major metabolites examined in this study (i.e., nordiazepam, N4- acetylsulfamethoxazole, o-desmethylnaproxen, and p-hydroxybenzoic acid) only N4- acetylsulfamethoxazole was quantifiable in the earthworm tissues. N4- acetylsulfamethoxazole was seen to increase to a peak concentration of 4.39 ± 0.4 ng g-1 at 3 d and then decreased to 2.62 ± 0.01 ng g-1 by the end of the 21 d incubation (Figure

3). Further, N4-acetylsulfamethoxazole was also detected in the earthworm soil with the highest concentration observed at 7 d (Figure 3c), indicating that earthworms was capable of metabolizing these CECs and excreting the transformation product into its surrounding environment. N4-acetylsulfamethoxazole is the primary metabolite of sulfamethoxazole in humans and has been previously detected in wastewater effluent, environmental samples and plant tissue (Dudley et al., 2018; Radke et al., 2009). However this was, to the best of our knowledge, the first time that N4-acetylsulfamethoxazole has been observed to form in earthworms.

The continued observation of the formation of N4-acetylsulfamethoxazole, an acetyl conjugate, in the environment is of considerable interest because conjugates have the potential to maintain the biological activity of the parent compound (Fu et al., 2017b).

Further, because researchers traditionally only quantify parent compounds during environmental assessments, the formation and accumulation of conjugates implies that

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there may be an underestimation of environmental exposure to CECs such as pharmaceuticals and further incomplete environmental risk assessment of CECs (Chen et al., 2013). This is of particular concern for antibiotics, because of the rise of antimicrobial resistance (Ventola, 2015).

The major metabolite of methyl paraben, p-hydroxybenzoic acid, was observed in all soil samples, including the non-treated controls (Figure 3c, Figure S3). This was likely due to the endogenous p-hydroxybenzoic acid in sphagnum peat (Hardy et al., 2009).

However, the concentration of p-hydroxybenzoic acid was higher in the spiked earthworm treatments than in the blank controls or non-earthworms chemical controls indicating that E. fetida was also capable of taking up and metabolizing methyl paraben and excreting of p-hydroxybenzoic acid into the soil. This was consistent with previous contact tests in which 70% of the initial methyl paraben was found to be metabolized to p-hydroxybenzoic acid and phenol within 48 h in E. fetida (Mincea et al., 2010).

The transformation products o-desmethylnaproxen and nordiazepam were not detected (S/N < 10) in earthworm tissues, but o-desmethylnaproxen was quantifiable in earthworm-CEC treated soils during the 21 d incubation (Figure 3b), indicating active uptake, metabolism, and excretion. The quantification of the major metabolites for naproxen, sulfamethoxazole and methyl paraben, o-desmethylnaproxen, N4- acetylsulfamethoxazole, and p-hydroxybenzoic acid suggested a trend in the capabilities of E. fetida to take up, metabolize and excrete then transformation products of some

CECs in the soil environment.

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Previous studies used radiolabeling and LC-FTMS analysis to assess potential metabolism of carbamazepine, diclofenac and fluoxetine in E. fetida but were unable to verify the presence of metabolites in earthworm tissues (Carter et al., 2014). Results from this and other studies indicated that metabolism may be chemical-specific. To the best of knowledge, this was the first study to identify and quantify metabolites of these CECs in

E. fetida dwelling in a soil.

4.3.3 Effects of CECs on earthworm antioxidant enzymes

Activities of several vital antioxidant enzymes were determined after exposure to

CECs. A significant increase in the activity of glutathione-S-transferase (GST) in the treatment samples over the controls was observed starting after 3 d into the incubation (P

< 0.05), and the GST activity continued to increase until the end of the 21 d incubation

(Figure 4a). This observation suggested that increased exposure time resulted in increased oxidative stress because glutathione is considered a critical antioxidant that acts to maintain redox homeostasis and signaling in cells (Saint-Denis et al., 1998). Further,

GST is a crucial enzyme family for the detoxification of xenobiotics during Phase II metabolism (Stenersen et al., 1979). Thus, the observed increase in GST activity may indicate that there was a formation of additional Phase II metabolites. However, the detection of these potential metabolites was not attempted due to a lack of authentic standards. High GST activity was also observed at 0 h for both the controls and treated samples. However, this increase in activity is likely due to the initial stress of the earthworm being transferred into the test media, and the effect dissipated within the first day of exposure.

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No significant difference in catalase (CAT) activity was observed between the treatment and controls until the end of the exposure period (Figure 4b). At the 21 d time point a significant increase was seen in the treatment samples (P < 0.05), indicating that extended exposure to CECs likely resulted in increased production of hydrogen peroxide in earthworm tissue (Saint-Denis et al., 1998). However, an increase in the CAT activity was also found in the control at 0 h. The increase in CAT activity was, again, likely due to the initial stress of the earthworms being transferred to different environmental conditions and the difference dissipated within 24 h.

A significant increase in superoxide dismutase (SOD) was observed at 1 d and 3 d

(P < 0.01). However, no significant differences were observed between the treatment and controls after 3 d (Figure 4c). This trend was in accordance with SOD has the first line of defense against reactive oxygen species (Mittler, 2002). SOD acts to catalyze the superoxide radical into oxygen molecules or hydrogen peroxide (Davies, 2008). As an increase in CAT was observed at the later time point it was likely that SOD activity increased initially, resulting in an increased production of hydrogen peroxide, which was then detoxified by CAT.

Previous studies examining the biochemical effects of CEC exposure in earthworms showed somewhat similar trends. For example, a study exploring the biochemical and genetic toxicity of triclosan in E. fetida showed a dose-dependent hormesis effect over time for both CAT and GST activity, with increasing activity being observed after a 2 d exposure at low doses and an inhibition of enzyme activity being observed after 14 d at high doses. Further, similar studies considering oxidative stress in

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E. fetida exposed to herbicides showed an increase in enzyme activities at lower concentrations and a suppression of enzyme activities at high concentrations (Song et al.,

2009; Zhang et al., 2013). Thus, it is likely that the lower, environmentally relevant, concentrations of CECs used in this study resulted in the observed increase in enzyme activities while these concentrations were not high enough to cause an inhibition in enzyme activity.

4.3.4 CEC-induced lipid peroxidation

Malondialdehyde (MDA) content was used has a proxy for oxidative damage as it is produced from the lipid peroxidation of polyunsaturated fatty acids (Davey et al.,

2005). The only significant increase in MDA content was observed at 7 d (P < 0.001), after which there was no discernible trend in lipid peroxidation production. Previous studies exploring MDA production in earthworms exposed to the antibacterial triclosan displayed increasing production of MDA with increasing concentrations of triclosan (Liu et al., 2010). Thus, it was possible that the reduction in MDA content at the later time points was due to the low initial concentrations used in this study, and the observed increases in metabolism and detoxification reactions may have also served to alleviate the oxidative damage.

4.4 Conclusions

Many regions in the world are seeing increased use of TWW and biosolids.

However, agricultural irrigation using TWW and the land application of biosolids may pose unintended ecological consequences, including exposure to contaminants of

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emerging concern by terrestrial organisms. In this study, three pharmaceuticals, sulfamethoxazole, naproxen and diazepam, were found to be taken up by the earthworm species Eisenia fetida. Upon uptake, earthworms appeared to be capable of metabolizing these pharmaceuticals; however, the extent of these transformations appeared to be highly chemical-specific. Exposure to CECs also resulted in the up-regulation of antioxidant enzymes GST, CAT, and SOD, indicating oxidative stress. Further, the increase in GST suggested the potential for the formation of phase II metabolites that warrant further investigation in future research. Results from this study provided new insights into the adverse effects of CECs exposure in terrestrial organisms, highlighting the need to consider these trace contaminants when assessing the safety of irrigation with TWW and biosolid application in the agroecosystems.

Acknowledgements:

The authors would like to thank the U.S. Environmental Protection Agency STAR program (Award No: R835829) and the National Science Foundation- the Water SENSE

Integrative Graduate Education Research Training program (No. DGE-1144635) for funding support

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Tables

Table 1: Dissociation constants (Kd) for selected compounds in artificial soil containing earthworms and non-earthworm controls. The values are shown as mean ± SE (n=3).

N.D. = Non-determinable

0 d 1 d 3 d 7 d 14 d 21 d Diazepam Earthworm 4.0 ± 0.3 2.98 ± 1.9 1.34 2.60 ± 1.1 0.84 ± 0.1 N.D. Control 5.43 ± 0.9 2.41 6.18 ± 4.2 4.8 ± 2.2 N.D. 6.56 ± 1.9 Sulfamethoxazole Earthworm 1.60 ± 0.2 0.90 ± 0.3 0.49 ± 0.14 1.71 ± 0.06 1.02 ± 0.05 0.71 ± 0.2 Control 1.47 ± 0.2 1.75 ± 0.5 0.75 ± 0.21 1.08 ± 0.06 1.02 ± 0.05 0.71 ± 0.2 Naproxen Earthworm 1.60 ± 0.2 7.69 ± 3.3 8.8 ± 5.2 0.87 ± 0.5 1.77 ± 1.0 3.96 ± 1.2 Control 1.97 ± 0.4 6.49 ± 2.5 11.0 ± 4.7 2.0 ± 0.6 1.84 ± 0.4 5.86 ± 1.6 Methyl Paraben Earthworm 2.99 ± 0.2 N.D. N.D. N.D. N.D. N.D. Control 4.0 ± 1.2 N.D. N.D. N.D. N.D. N.D.

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Table 2: Bioconcentration factors (BCF) and Bioaccumulation factors (BAF) of selected compounds in E. fetida. The values are shown as mean ± SE (n=3). (a) Significant differences relative to time point (α = 0.05, ANOVA one-way, post hoc Tukey-Kramer).

(b) Significant differences of BAF to BCF (α = 0.05, student t-test). N.D.= non- determinable.* standard error could not be calculated due to lack of quantifiable compound concentration in replicates.

1 d 3 d 7 d 14 d 21 d Diazepam BAF 2.21 ± 1.0a 3.04 ± 1.9a 0.56* 3.95 ± 1.3a 1.32 ± 0.3a BCF 5.15 ± 2.0 10.1* 6.19* 27.7 ± 14.2 N.D. Sulfamethoxazole BAF 2.54 ± 0.2 3.1 ± 0.4 2.94 ± 0.39b 5.44 ± 1.8 3.7 ± 0.5 BCF 2.10 ± 0.5 1.57 ± 0.6a 3.38 ± 0.73b 4.33 ± 1.6a 3.23 ± 0.6

Figures

Naproxen BAF 0.40 ± 0.1 0.28 ± 0.05 0.82 ± 0.3 1.55 ± 0.4 0.73 ± 0.02 BCF 3.32 ± 2.8 1.9 ± 1.0 1.1 ± 0.6 2.6 ± 0.4 4.42 ± 1.9

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Figure 1:

Kinetics of CECs in three matrices: soil, porewater and earthworms. (A) Diazepam. (B)

Sulfamethoxazole (C) Naproxen (D) Methyl paraben. The values are shown as mean ±

SE (n=3).

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Figure 2: Kinetics of N4-acetylsulfamethoxazole in earthworm tissue. The values are shown as mean ± SE (n=3).

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Figure 3: Kinetics of metabolites in earthworm and non-earthworm soils. (A) N4- acetylsulfamethoxazole. (B) o-desmethylnaproxen. (C) p-hydroxybenzoic acid. The values are shown as mean ± SE (n=3).

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Figure 4: Activity of antioxidant enzyme in earthworms exposed to CECs over time (A) glutathione-S-transferase (GST); catalase (CAT); superoxide dismutase (SOD). The values are shown as mean ± SE (n=3). * p-value < 0.05 ** p-value < 0.01 (student t-test).

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Figure 5: Malondialdehyde (MDA) content in E. fetida. The values are shown as mean ±

SE (n=3). *** p-value < 0.001 (student t-test).

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Supplemental Information

Table 1: Mortality assessment of earthworms exposed to selected contaminants of emerging concern for 21 d

Sample 7d 14d 21d Naproxen 5 ng g-1 9/10 10/10 10/10 50 ng g-1 11/10 10/10 11/10 500 ng g-1 10/10 11/10 10/10 5000 ng g-1 10/10 10/10 10/10 Diazepam 5 ng g-1 10/10 9/10 9/10 50 ng g-1 10/10 10/10 10/10 500 ng g-1 10/10 9/10 9/10 5000 ng g-1 10/10 9/10 9/10 Sulfamethoxazole 5 ng g-1 10/10 11/10 10/10 50 ng g-1 10/10 12/10 12/10 500 ng g-1 10/10 10/10 10/10 5000 ng g-1 9/10 10/10 10/10 Methylparaben 5 ng g-1 10/10 10/10 10/10 50 ng g-1 10/10 8/10 8/10 500 ng g-1 10/10 10/10 10/10 5000 ng g-1 10/10 12/10 11/10 Mixture 5ng g-1 10/10 10/10 10/10 50ng g-1 11/10 10/10 10/10 500 ng g-1 10/10 11/10 11/10 5000 ng g-1 10/10 10/10 11/10 Control 8/9 10/10 10/10 10/10 10/10 10/10 10/10 10/10 10/10 9/10 9/10 10/10

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Table 2: Extraction recovery rate for parent compounds; mean ± standard deviation (n=3)

Parent Compound Soil Earthworm Diazepam 87.6 ± 29.4 43.3 ± 12.9 Sulfamethoxazle 40.8 ± 7.0 52.4 ± 8.3 Naproxen 11.1 ± 2.4 20.3 ± 4.6 Methylparaben 70.9 ± 10.9 84.8 ± 18.6

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Table 3: Optimized MRM conditions for the analysis of the chosen analytes by UPLC

TQD MS/MS (CV: cone voltage in V; CE: collision energy in eV)

Compound MRM MRM1 CV/CE MRM2 CV/CE (Precursor (Quantification) (Qualification) mass) Diazepam 285.0268 192.9736 54/44 153.8944 54/46 Diazepam-d5 290.0957 198.0478 48/30 154.0864 48/28 Nordiazepam 271.0398 140.0378 52/28 165.0721 52/28 Methylparaben 153.0154 120.9931 26/10 58.9521 26/16 Methylparaben-d4 157.0104 124.9919 30/12 58.9836 30/14 4-Hydroxybenzoic 138.9998 106.9807 26/8 77.0002 26/16 acid Naproxen 231.0581 170.0437 24/24 115.0309 24/52 Naproxen-d3 233.9928 173.0388 22/24 144.1676 22/40 o-desmethylnaproxen 217.0467 171.0642 26/14 184.9744 26/8 Sulfamethoxazole 253.9681 92.0480 34/28 155.9518 34/14 Sulfamethoxazole-d4 258.0128 96.0797 32/26 159.9900 32/16 N4- 296.0228 134.0380 32/24 65.0087 32/38 acetylsulfamethoxazole

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Table 4: Properties of contaminants of emerging concern and their major metabolites

Name Activity Chemical CAS Molecula Pka1,2 Log

2 Formula ID r Weight K ow

Diazepam Benzodiazepine C16H13ClN2O 439-14- 284.7 3.4 2.82 5

Diazepam-d5 Benzodiazepine C16H8D5ClN2O 65854- 289.8 3.4 2.82 76-4

Nordiazepam Benzodiazepine C15H11ClN2O 1088- 270.7 2.85 2.79 and Metabolite 11-5

Methylparaben Preservative C8H8O3 99-76-3 152.2 8.5 1.96

Methylparaben-d4 Preservative C8H4D4O3 362049- 156.2 8.5 1.96 51-2

4-Hydroxybenzoic Metabolite C7H6O3 99-96-7 138.1 4.54 1.58 acid

Naproxen Anti- C14H14O3 52079- 230.3 4.15 3.18 inflammatory 10-4

Naproxen-d3 Anti- C14H13D3O3 958293- 233.3 4.15 3.18 inflammatory 77-1 o-desmethylnaproxen Metabolite C13H12O3 52079- 216.2 4.34 2.71 10-4

Sulfamethoxazole Antibiotic C10H11N3O3S 723-46- 253.3 6.1 0.89 6

Sulfamethoxazole-d4 Antibiotic C10H7D4N3O3S 1020719 257.3 6.1 0.89 -86-1

N4- Antibiotic C12H13N3O4S 21312- 295.3 5.68 0.86 acetylsulfamethoxazo 10-7 le 1Wu et al 2014 2HSDB 2018

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Figure 1: Bioaccumulation factors (ng g-1) and bioconcentration factors (g mL-1) in E.

Fetida cultivated in artificial soil. Error bars represent standard error of the mean (n=3).

(A) Diazepam (B) Naproxen (C) Sulfamethoxazole.

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Figure 2: Kinetics of CECs in porewater. (A) Diazepam in earthworm and non- earthworm porewater. (B) Sulfamethoxazole in control and earthworm treated porewater. (C) Naproxen in control and earthworm treated porewater. (D) Methyl paraben in control and earthworm treated porewater. The values are shown as mean ± SE (n=3).

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Figure 3: Concentration of hydroxybenzoic acid in media control soil. Error bars represent standard error of the mean (n=3).

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Chapter 5: Summary of Findings and Future Work

5.1 Summary of Findings

Currently we are experiencing a series of global trends that are creating unique challenges for the future of sustainable development. These trends include shifting precipitations patterns, rising temperatures, growing human populations, and rapid urbanization. In order to meet these challenges, traditionally under-utilized resources, such as treated wastewater (TWW) and biosolids, will have to be harnessed. These resources are derived from wastewater treatment plants and contain biologically active, pseudo-persistent, trace chemicals referred to as contaminants of emerging concern

(CECs). Land application of TWW and biosolids for agriculture and landscaping has the potential to introduce CECs into terrestrial ecosystems, from where they could accumulate, be metabolized and/or cause adverse effects in terrestrial organisms. This dissertation has described the ability of terrestrial plants and invertebrates to take up and metabolize CECs and highlighted the potential for these trace contaminants to induce biochemical changes in non-target terrestrial organisms. The findings of this project, overall conclusions, and recommendations for future work are summarized below.

5.1.1 Metabolism of sulfamethoxazole in Arabidopsis thaliana cells and cucumber seedlings

In arid and semi-arid areas, TWW reuse is becoming increasingly prevalent for agricultural irrigation. However, irrigation with TWW has the potential to introduce

CECs including antibiotics into agroecosystems. One of the most commonly prescribed

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and environmentally relevant antibiotics is sulfamethoxazole. However, little is known about the fate of sulfamethoxazole in terrestrial plants. In this study, sulfamethoxazole was observed to be taken up and actively metabolized by Arabidopsis thaliana cells into six transformation products. The transformation products included oxidation of the amine group, producing Phase I metabolites, which was followed by conjugations with glutathione, glucuronic acid and leucine, producing Phase II metabolites. Phase III metabolism (formation of bound-residues) was assessed by determining the mass balance of 14C-sulfamethoxazole in A. thaliana cells and cucumber seedlings. Non-extractable

14C-sulfamethoxazole increased over time in both A. thaliana cells and cucumber

(Cucumis sativus) seedlings, indicating that Phase III metabolism significantly contributed to the fate of sulfamethoxazole in A. thaliana cells and cucumbers. Further, in

A. thaliana cells and cucumber seedlings, the mass balances were calculated to range from 80-120% and 80-94%, suggesting a minor role of mineralization. The results from this study highlighted the potential of terrestrial plants to transform pharmaceuticals, forming both bioactive Phase I metabolites and Phase II conjugates, and store them as in the form of bound residues as Phase III metabolism.

5.1.2 Formation of biologically active benzodiazepine metabolites in Arabidopsis thaliana cell cultures and vegetable plants under hydroponic conditions

Plant uptake of CECs from TWW reuse and biosolid application has been documented in agroecosystems. Previous studies suggested that plants were capable of metabolizing CECs after uptake. However, these studies often reported different results even with the same CECs, likely due to the use of different plant species and/or different

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laboratory conditions. In this study, the metabolism of the benzodiazepine diazepam was explored in three different plant species, A. thaliana, cucumber (Cucumis sativus), and radish (Raphanus sativus). The plants were exposed to diazepam in laboratory under three different laboratory exposure conditions that included a 6 d cell culture, an acute (7 d)/high concentration (1 mg L-1) hydroponic cultivation, and a chronic (28 d)/low concentration (1 µg L-1) hydroponic cultivation. 14C-Diazepam was incubated concurrently with non-labeled diazepam to determine the fractions of extractable and non-extractable radioactivity to quantify Phase III metabolism. Diazepam was taken up and metabolized in all plant species under the different exposure conditions. A. thaliana cells actively transformed diazepam into temazepam and nordiazepam via Phase I metabolism. This metabolism mimicked human metabolism, as temazepam and nordiazepam are the minor and major metabolites, respectively, formed during human metabolism of diazepam. Intriguingly, both of these metabolites are bioactive and prescribed pharmaceuticals in their own right, alluding to a potential for increased risk from consumption not considered in previous studies. The fraction of non-extractable residue increased over the 6 d incubation, indicating extensive Phase III metabolism over time in A. thaliana cells. In cucumber and radish seedlings, a similar Phase I metabolism pattern was observed, with nordiazepam being the most prevalent metabolite at the end of the 7 d and 28 d cultivations. However, significant differences in phase III metabolism were observed between the radish and cucumber plants. For example, after the acute exposure, diazepam mass balance was 99.3% for cucumber seedlings but only 58.1% for radish seedlings, indicating increased mineralization in the radish system. Diazepam

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induced changes in the regulation of glycosyltransferase activity in both cucumber and radish seedlings, indicating the formation of Phase II metabolites. The results from this study showed that exposure conditions and plant species can influence the metabolism of diazepam, and formation of bioactive transformation intermediates and different phases of metabolism should be considered in order to achieve a comprehensive understanding of risks of CECs in agroecosystems.

5.1.3 Bioaccumulation, metabolism and biochemical effects of contaminants of emerging concern in earthworm Eisenia fetida

Exposure of terrestrial invertebrates to CECs will likely increase with increasing

TWW reuse and biosolid application. However, currently there is limited information on the fate and effects of CECs in terrestrial organisms. In this study, the earthworm E. fetida was exposed to three pharmaceuticals, i.e., sulfamethoxazole, diazepam, and naproxen, and one preservative, i.e., methyl paraben, for 21 d in an artificial soil. Methyl paraben did not accumulate in the earthworm tissue, likely due to its rapid degradation in the soil. The other CECs showed an accumulation in earthworm tissues from the soil/soil porewater. The presence of E. fetida did not significantly affect the adsorption of these

CECs to the soil. The presence of primary metabolites (i.e., N4-acetylsulfamethoxazole, o-desmethylnaproxen, p-hydroxybenzoic acid) in the treated soil suggested that E. fetida were capable of actively metabolizing the three pharmaceuticals and excreting the metabolites. However, the metabolism was chemical-specific, and only N4- acetylsulfamethoxazole was detected in earthworm tissues. Exposure to the four CECs also resulted in the up-regulation of several antioxidant enzymes, including glutathione-

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S-transferase, superoxide dismutase, and catalase, and an increase in malondialdehyde, indicating oxidative stress in the exposed E. fetida. Results from this study highlighted the need to consider the role of, and effects on terrestrial invertebrates when understanding risks of CECs in agroecosystems.

5.5.4 Overall Conclusions

Our findings illuminate the complexity of the interactions between contaminants of emerging concern and terrestrial organisms. The dissertation highlights the ability of terrestrial organisms to take up and transform CECs through metabolism, which results in both bioactivation and detoxification of the target contaminants. This project also demonstrates the ability of CECs to alter the biochemistry of the studied terrestrial organisms by changing the regulation of enzymes associated with oxidative stress and metabolism. The use of cell cultivations, hydroponic studies, and artificial soil allowed us to examine the metabolism and effects of CECs in terrestrial organisms with limited confounding factors. However, it is highly likely that similar studies conducted in soils may show low rates of uptake and different patterns in metabolism. Our research suggests that scientifically sound understanding of fate of, and risks from, CECs in the environment cannot solely rely on the assessment of the parent compound and/or only consider the potential for human exposure to CECs. One must also consider the potential for the formation of metabolites and the consequences of exposure to all non-target organisms in order to better understand the fate and risks of CECs in terrestrial environments. The results have potential implications for policy makers and other

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stakeholders attempting to assess the risks for the land application of treated wastewater and biosolids.

5.2 Future Research

The results presented in this dissertation suggest the highly chemical-, species-, and research technique- specific nature of the environmental fate of CECs. For example, cell cultures often form amino acid conjugates while whole plants form sugar conjugates during xenobiotic metabolism. The differences in CEC metabolism imposed by treatments or species warrant further investigation. Additionally, more toxicological data are needed on the effects of these and other compounds in terrestrial invertebrates, especially for those of agricultural importance. From the research conducted in this dissertation, future research should focus on the impacts of exposure and the potential for transformation of CECs under different conditions and in multiple species. Future studies should place emphasis on experimentation using biosolids and TWW with inherent compounds and field conditions to improve environmental relevance. Future risk assessments should be conducted by taking into account the formation of biologically active and conjugated metabolites, and with regard to the potential toxicity of CECs in non-target terrestrial organisms.

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