ENVIRONMENTAL APPLICATION AND IMPLICATION OF ENGINEERED NANOMATERIALS IN SOIL, WATER , AND PLANT SYSTEMS By Jianzhou He A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Crop and Soil Sciences Environmental Toxicology Doctor of Philosophy 2020 ABSTRACT ENVIRONMENTAL APPLICATION AND IMPLICATION OF ENGINEERED NANOMATERIALS IN SOIL, WATER , AND PLANT SYSTEMS By Jianzhou He Nanotechnology is promising to develop novel engineered nanomaterials (ENMs) for industrial and consumer products, environmental engineering, and agricultural production. Specifically, ENMs can be used to remove contaminants from wastewater and to produce plant care agrochemicals (e.g., nanopesticides). The rapid ly growing use of nano - enabled products is leading to increased release of ENMs into agroecosystems and their uptake by organisms and crop plants, thus posing potential threats to environmental and human health. To maximize environmental benefits and minimize potential risks of ENMs, this dissertation research focused on both environmental application and implication of ENMs in soil, water , and plant systems. The first study systematically investigat ed the photodegradation of antibiotic cephalexin (as an emerging contaminant in water) by novel ZnO nanowires under simulated sunlight through a combination of experimental and theoretical techniques including kinetic batch degradation, free radical trappi ng experiments, degradation product identification by mass spectrometry, and molecular computation . The ZnO nanowires substantially increased the degradation efficiency of cephalexin , especially under circumneutral and alkaline condition s Anions ( HCO 3 ) and natural organic matter inhibited the p hotodegradation of cephalexin by scavenging 2 ) . Finally, c ephalexin photodegradation pathways included hydroxylation, demethylation, decarboxylation, and dealkylation. Overall, the ZnO nanowires can be potentially used for removing antibiotics from contaminated water. The second study examined the effect of biochar amendment s on soil sorption of Ag NPs and Ag + , their soil - bound fractions, and their uptake and translocation by radish es grown in a loamy sand soil spiked with 1 mg kg 1 Ag NPs or Ag + . Biochar amendment had no significant effect on Ag uptake by radish es , which also did not differ with and without Ag addition . Results of Ag sorption, soil - bound fractions, and soil pore water concentrations showed that the bioavailable Ag at the environmentally - relevant concentration (1 mg/kg) was low, which may partially explain the non - s ignificant effect of biocha r amendment on the Ag uptake. This study demonstrated that the accumulation risk of Ag NPs and Ag + ions in vegetable crops at environmentally - relevant concentrations is very low. The third study addressed the role of stomata in the internalization of Ag NPs using abscisic acid (ABA) - responsive ecotypes (Ler and Col - 7) and ABA - insensitive mutants ( ost1 - 2 and scord7 ) of Arabidopsis thaliana in batch sorption experiments. The sorbed amount of Ag NPs was much lower in the leaves of the L er and Col - 7 ecotypes treated with 10 µM ABA than that in the ABA - free control, mainly due to ABA - induced stomatal closure. However, sorption of Ag NPs by the leaves of the ost1 - 2 and scord7 mutants did not change with and without ABA. Microscopic images s howed that internalized Ag NPs were located in the cell membrane, cytoplasm and plasmodesmata. Clearly stomata play an important role in the internalization of ENMs in plants, which has broader implications in foliar application of nanopesticides and minim izing ENMs contamination of food crops. Overall, the sustainable use of nanotechnology will depend on improved knowledge on and proper management of its benefits and potential risks. Copyright b y JIANZHOU HE 2020 v T his dissertation is dedicated to my parents, Taiming He and Qiongyao Zhang, my love, Dandan Peng, and my beloved daughter, Michelle Peng He. I love you all ! vi ACKNOWLEDGMENT S First and foremost, I would like to express my sincere gratitude to my advisor Dr. Wei Zhang for his continuous guidance and support of my Ph.D. study, and for his enthusiasm, knowledge, commitment, patience, and encouragement . In the past four years, Dr. Zhang has been acting not only as a major advisor but also as a mentor for my life. This dissertation would not have been completed without his support and help , which is especially invaluable to me during this COVID - 19 pandemic . I am also very grateful to the other members of my dissertation committee, Dr. Hui Li, Dr. Elliot Ryser, and Dr. Sheng Yang He, for their insightful comments and guidance for my doctoral study . I would also like to thank my group members, colleagues, and friends at Michigan State U niversity , Drs. Yuanbo Li, Yingjie Zhang, Cheng - Hua Liu, Sangho Jeon, Zeyou Chen, Feng Gao, Xun Qian, Shuai Zhang, Yike Shen, Qi Yuan, Ya - Hui Chuang, Anping Peng, Bingyu Wang, Xiaojie Hu, Fengling Liu, Shuang Liu, as well as Geoff Rhodes, Gayathri Gunathil aka, Yu Zhang, Hongyu Dang, Jinfeng Li, and many others, for their help, support, and friendship. I also want to acknowledge the help specifically from Drs. Dengjun Wang, Yaozhong Zhang, and Li Zhang . Finally, I would like to thank my parents, Taiming He a nd Qiongyao Zhang, my parents - in - law, Shuyun Peng and Qizhen Ran, my brothers, Xin He and Jianghu Peng , for their unconditional support and unreserved love . My deepest gratitude is owed to my wife, Dandan Peng, for her love, support, and companionship for the past eight years. Special thank s to my beloved daughter, Michelle Peng He. vii TABLE OF CONTENTS LIST OF TABLES ................................ ................................ ................................ ......................... ix LIST OF FIGURES ................................ ................................ ................................ ........................ x CTION AND OBJECTIVES ................................ ................................ ... 1 Nanomaterials and Nanotechnology ................................ ................................ ............... 2 Environmental Application of Nanotechnology ................................ ............................. 5 Environmental Implication of Nanotechnology ................................ .............................. 6 Research Objectives ................................ ................................ ................................ ........ 8 NANOWIRES UNDER SIMULATED SUNLIGHT ................................ ................................ ... 10 Abstract ................................ ................................ ................................ ......................... 11 Introduction ................................ ................................ ................................ ................... 12 Materials and Methods ................................ ................................ ................................ .. 15 Chemicals ................................ ................................ ................................ .................. 15 Synthesis and Characterization of ZnO Nanowires ................................ .................. 16 Photodegradation Experiments ................................ ................................ ................. 17 Trapping Experiments ................................ ................................ ............................... 19 Analytical M ethods ................................ ................................ ................................ .... 19 Theoretical Calculation ................................ ................................ ............................. 20 Results and Discussion ................................ ................................ ................................ .. 20 Characterization of ZnO Nanowires ................................ ................................ ......... 20 Photodegradation Efficiency and Reusability ................................ ........................... 24 Reactive Species in Photodegradation ................................ ................................ ...... 28 Effect of Solution pH ................................ ................................ ................................ .. 30 Effect of Bicarbonate ................................ ................................ ................................ . 32 Effect of SRNOM ................................ ................................ ................................ ....... 33 Identification of Intermediates and Transformation Pathways ................................ . 34 Conclusion ................................ ................................ ................................ ..................... 37 - BOUND FRACTIONS OF SILVER NANOPARTICLES AND IONS BUT NOT THEIR UPTAKE BY RADISH AT AN EN VIRONMENTALY RELEVANT CONCENTRTAION ................................ ........................ 38 Abstract ................................ ................................ ................................ ......................... 39 Introduction ................................ ................................ ................................ ................... 40 Materials and Methods ................................ ................................ ................................ .. 43 Chemicals, Biochar and Soil ................................ ................................ ..................... 43 Radish Growth Experiment ................................ ................................ ....................... 44 Ag Sorption Experiment ................................ ................................ ............................. 47 BCR Sequential Extraction ................................ ................................ ........................ 48 Analytical Methods ................................ ................................ ................................ .... 49 viii Data Analysis ................................ ................................ ................................ ............. 49 Results and Discussi on ................................ ................................ ................................ .. 50 Radish Biomass ................................ ................................ ................................ .......... 50 Uptake of Ag by Radish ................................ ................................ ............................. 53 Ag Sorptio n to Soils ................................ ................................ ................................ ... 57 Soil - bound Ag Fractions ................................ ................................ ............................ 60 Conclusion ................................ ................................ ................................ ..................... 63 NANOPARTICLES BY ARABIDOPSIS T HALIANA ................................ ................................ .. 65 Abstract ................................ ................................ ................................ ......................... 66 Introduction ................................ ................................ ................................ ................... 67 Materials and Method s ................................ ................................ ................................ .. 69 Plant Materials ................................ ................................ ................................ .......... 69 Ag NPs Suspension ................................ ................................ ................................ .... 70 Stomata Assays ................................ ................................ ................................ .......... 70 Sorption Experimen ts ................................ ................................ ................................ 71 Data Analysis ................................ ................................ ................................ ............. 71 Visualization of Sorbed Ag NPs ................................ ................................ ................. 72 Results and Discussion ................................ ................................ ................................ .. 73 Ag NPs Characterization ................................ ................................ ........................... 73 Stomatal A perture Assay ................................ ................................ ........................... 75 Kinetics and Equilibrium Sorption of Ag NPs ................................ ........................... 77 Visualization of Sorbed Ag NPs ................................ ................................ ................. 81 Conclusion ................................ ................................ ................................ ..................... 84 CHAPTER V CONCLUSIONS AND FUTURE WORK ................................ ............................ 85 Conclusions ................................ ................................ ................................ ................... 86 Future Work ................................ ................................ ................................ .................. 87 APPENDICES ................................ ................................ ................................ .............................. 88 A PPENDIX A: Supplementary Materials to CHAPTER II ................................ .......... 89 A PPENDIX ................................ ....... 103 A PPENDIX ................................ ........ 112 REFERENCES ................................ ................................ ................................ ........................... 127 ix LIST OF TABLES Table 2.1. Zeta potentials and hydrodynamic diameters of ZnO nanowires under various experimental conditions. ................................ ................................ ................................ ............... 23 Table 2.2. Fitted parameters for degradation kinetics of cephalexin in all experiments 27 Table 2.3. Dissolution of ZnO nanowires (20 mg L 1 ) after 3 - hrs experiment under different solution pH 28 Table 3.1. Radish growth in soils in the absence (blank) and presence of spiked AgNO 3 and Ag NPs without (0%) or with (0.1% and 1%) biochar amendment* 46 Table A1. Experimental conditions in selected photocatalytic degradation studies using ZnO photocatalysts in literature . 91 Table A2. Frontier electron densities on atoms in cephalexin calculated by Gaussian 09 program at B3LYP/6 - 311 + G* level with a minimum energy. 92 Table A3. Photodegradation Kinetics Data . 100 Table B1. Fitted parameters of the sorption isotherms using the Langmuir model*. 103 Table B2. Fractions of Ag + and Ag NPs in the soil samples collected from the radish - growing pots on Day 35*. 104 Table B3. Fractions of Ag + and Ag NPs in the soil samples collected from the radish - free pots on Day 35*. 105 Table B4. AgNO 3 Sorption Data . 110 Table B5. Ag NPs Sorption Data . 111 Table C1. Measured stomatal aperture width and width - to - length ratio of Arabidopsis thaliana treated with opening buffer in the absence (control) or presence of 10 µM abscisic acid (ABA ). . 113 Table C2. Ag NPs Sorption Kinetics Data . 122 Table C3. Ag NPs Sorption Equilibrium Data . x LIST OF FIGURES Figure 1.1. Publications trend in engineered nanomaterials research. Web of Science resu lts for ................................ ................................ ................................ ............................... 4 Figure 2.1. Representative (a) scanning electron microscopy, (b) transmission electron microscopy images, and (c) X - ray diffraction pattern of ZnO nanowires. ................................ ... 21 Figure 2.2. Observed (a) and fitted (b) photodegradation kinetics of cephalexin by ZnO nanowires (20, 40, and 80 mg L 1 ) at pH 7. 2 under simulated sunlight. Error bars represent the standard deviations ( n = 3). Solid lines were fitted to initial parts of kinetics that followed the first - order degradation. ................................ ................................ ................................ ................................ ... 25 Figure 2.3. Effect of various free radical scavengers on the photodegradation of cephalexin by ZnO nanowires (20 mg L 1 ) at solution pH of 7.2 under simulated sunlight. Error bars represent the standard deviations ( n = 3). Solid lines were fitted to the first - order kinetic equation. .......... 29 Figure 2.4. Effects of initial solution pH (a), bicarbonate (b) and Suwannee River natural organic matter (SRNOM) (c) on the photodegradation of cephalexin by ZnO nanowires (20 mg L 1 ) under simulated sunlight. Error bars represent the standard deviations ( n = 3). So lid lines were fitted to the first - order kinetic equation. ................................ ................................ ................................ ..... 31 Figure 2.5. Proposed photocatalytic degradation pathways of cephalexi n by ZnO nanowires under simulated sunlight irradiation. ................................ ................................ ................................ ...... 36 Figure 3.1. Dry biomass of radish roots (A) and shoots (B) harvested on Day 35 in each pot (two plants) in the absence (blank) and presence of spiked AgNO 3 and Ag NPs without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations ( n = 3) . Different letters indicate the significant difference in means at p < 0.05. Brackets indicate comparisons between the AgNO 3 and Ag NPs treatments as well as between the treatment groups with or without Ag addition, respectively. Figure 3.2. Total Ag concentrations in the roots (A) and shoots (B) of radish plants grown in soil pots in the absence (blank) and presence of spiked AgNO 3 and Ag NPs without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations ( n = 3). Different letters indicate the significant difference in means at p < 0.05. Brackets indicate the comparisons between the AgNO 3 and Ag NPs treatments as well as between the treatment groups with or without silver addition, respectively. Figure 3.3. Root concentration factor (A) and translocation factor (B) of Ag in either AgNO 3 or Ag NPs in radish plants grown in soil pots without (0%) or with (0.1% and 1%) biochar amendment. xi Error bars are the standard deviations ( n = 3). Different letters indicate the significant difference in means at p < 0.05. Brackets indicate comparisons b etween the AgNO 3 and Ag NPs treatments. 56 Figure 3.4. Measured (symbols) and fitted (lines) sorption isotherms of Ag in either AgNO 3 or Ag NPs by the soil samples without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations from two replicates. Figure 3.5. Soil - bound fractions of Ag in either AgNO 3 or Ag NPs in the soil samples without (0%) or with (0.1% and 1%) biochar amendment measured by the BCR extraction method. Figure 4.1. Representative TEM image of Ag NPs (A), particle size distribution of Ag NPs measured from the TEM images (B), aggregation kinetics of 10 mg L 1 Ag NPs suspension at pH 6.5 in DI water (C), and summarized TEM primary particle size, hydrodynamic diameter ( D h ) and - potential of Ag NPs (D). Figure 4.2. Stomatal aperture width of Arabidopsis thaliana treated with openin g buffer in the absence (control) or presence of 10 µM abscisic acid (ABA). Ler and Col - 7 are ecotypes, and ost1 - 2 and scord7 are corresponding ABA - insensitive mutants. Results are shown as mean ± standard deviation ( n = 36 63 stomata). Symbol (*) indicate s a statistically significant difference in means between the control and 10 µM ABA treatments ( p Figure 4. 3 . Sorbed amount of Ag NPs at various concentrations of 1, 5, and 10 mg L 1 by the leaves of Arabidopsis thaliana Ler (A) , ost1 - 2 (B) , Col - 7 (C) , and scord7 (D) treated in opening buffer without (control) or with abscisic acid (10 µM ABA) for 1 h. Error bars represent the standard deviations ( n = 3). Symbol (*) indicates statistically significant difference in means betw een the control and 10 µM ABA treatment ( p Figure 4.4 . Relation between the reduction percentages of the sorbed Ag NPs and the change of stomatal aperture induced by the presence of 10 µM ABA . For each Arabidopsis genotype, the sorbed amount reduction (%) was calculated for 1, 5, and 10 mg L 1 of the Ag NPs exposure . Figure 4.5. Representative TEM images of Ag NPs in the leaves of Arabidopsis scord7 mutant after the sorption experiment (10 mg L 1 Ag NPs). B lank: no Ag NPs exp osure; C ontrol: opening buffer without abscisic acid (ABA) treatment for 1 h before 10 mg L 1 Ag NPs exposure; 10 µM ABA: opening buffer with 10 µM ABA treatment for 1 h before 10 mg L 1 Ag NPs exposure. Red a rrows indicate the spots of Ag NPs at the subce llular locations. Figure A1. Chemical structure (a) of cephalexin and species distribution (b) as a function of solution pH (p K a 1 = 2.5 and p K a 2 = 7.2). Figure A2. Structural formula of cephalexin optimized by Gaussian 09 program. Figure A3. Energy dispersive X - ray spectroscopy for elemental analysis. Elements C and Ir are background, confirming that the powders are primarily ZnO. xii Figure A4 . Zeta potential of ZnO nanowires (20 mg L 1 ) suspended in background solution (5 mM NaCl) as a function of solution pH. Figure A5. Removal efficiency of cephalexin by ZnO nanowires (20 mg L 1 ) under simulated sunlight at pH 7.2 in 5 cycles (90 min reaction time each cycle). Figure A6. 1 , [cephalexin] = 100 µg L 1 , [NaCl] = 5 mM, pH = 7.2, and irradiation time = 180 min). Figure A7. LC chromatograms and m/z of degradation products of cefalexin by ZnO nanowires under simulated sunlight Figure B1. Representative transmission elec tron microscopy image (a) and aggregation kinetics (b) of Ag NPs (10 mg L 1 ) dispersed in deionized water. The Ag NPs suspension was stable at a time frame of 4 h. Figure B2. Representative scanning electron microscopy images of the biochar used in this study. 107 Figure B3. Measured pH values of soil samples collected from the radish - growing and radish - free pots on Day 35. Figure B4. The MINTEQ calculation showed that no precipitation occurs (blue text indicates undersaturation) during hydroxylammonium chloride extraction. Simulation condition: [Ag + ] = 0.05 mg L 1 (assuming maximum dissolution, 1 mg silver/k g soil × 1 g soil ÷ 20 ml = 0.05 mg L 1 ); [NH 2 OH·HCl] = 0.5 M; pH = 2; T = 25 °C. Figure C1. Representative confocal laser scanning microscopy images of Arabidopsis thaliana leaves. Images were obtained by scanning leaf discs (3 mm × 3 mm) of Ler, ost1 - 2 , Col - 7, and scord7 treated with opening buffer without (control) or with 10 µM abscisic acid (10 µM ABA) solution for 1 h. Scale bar is 20 µm. As observed, the ost1 - 2 and scord7 mutants of Arabidopsis did not respond to ABA - induced stomatal closure and exhibited constitutively larger apertures, while the stomata of Ler and Col - Figure C 2. Stomatal width - to - length ratio of Arabidopsis thaliana treated with opening buffer in the absence (control) or presence of 10 µM abscisic acid (ABA). Results are shown as mean ± standard deviation ( n = 36 84 stomata). Symbol (*) indicates a statistically significant difference in means between the control and 10 µM AB A treatments ( p Figure C 3. Sorption kinetics of 2 mg L 1 Ag NPs on the scord7 leaves. Error bars represent the standard deviations ( n = 2). . 117 Figure C 4 . Surface distribution analysis of Ag NPs in the leaves of Arabidopsis Col - 7 ecotype after the sorption experiment (10 mg L 1 Ag NPs exposure) characterized by SEM - xiii Ag NPs exposure, denoted as blank; (B) opening buffer treatment for 1 h before 10 mg L 1 Ag NPs exposure, denoted as control; (C) opening buffer with 10 µM ABA treatment for 1 h before 10 mg L 1 Ag NPs exposure, denoted as 10 µM ABA. Note that the stomata aperture in the SEM images could not represent the opening or closure status due to artifacts during the sample preparation (e.g., freeze - dr ying). More bright dots were found surrounding the stomatal opening of C than B plausibly due to the ABA induced sto matal closure minimizing the internalization of sorbed Ag NPs. . 118 Figure C 5 . Distribution of Ag NPs in selected micro - area on Col - 7 leave after the sorption experiment (10 mg L 1 Ag NPs). Upper: SEM images at different magnifications; Bottom: elemental mapping of carbon, oxygen and silver. Mapping images indicate the Ag is widely distributed in the selected micro - area. .. 119 Figure C6 . Subcellular TEM images of the leaves of Arabidopsis Col - 7 ecotype (A C) and scord7 mutant (D F) after the sorption experiment (10 mg L 1 Ag NPs). B lank: no Ag NPs exposure; C ontrol: opening buffer without abscisic acid (ABA) treatment for 1 h before 10 mg L 1 Ag NPs exposure; 10 µM ABA: opening buffer with 10 µM ABA treatment for 1 h before 10 mg L 1 Ag NPs exposure. The scale bar is 500 nm. It is difficult to identify any possible Ag NPs in the chloroplasts because of noisy background. .. 12 0 Figure C7 . Subcellular TEM images of the leaves of Arabidopsis Col - 7 after the sorption experiment (10 mg L 1 Ag NPs) in the c ontrol treatment (i.e., opening buffer without ABA for 1 h ) . The scale bars are 500 nm in A, C, E, F, and 200 nm for B and D . 1 INTRODUCTION AND OBJECTIVES 2 Nanomaterials and Nanotechnology Nanomaterials are materials with at least one dimension sized between approximately 1 to 100 nanometers (nm, = 10 9 m) 1 , originating from either natural, incidental, or engineered sources . Naturally occurring nanomaterials may come from volcanic ash, ocean spray, forest fires, rock weathering , etc . Nanomaterials can also be generated i ncidentally as byproducts of mechanical or industrial processes (e.g., combustion, vehicle engine exhausts, and welding fumes). In contrast, e ngineered nanomaterials (ENMs) are deliberately designed and produced for their unusual, tunable properties or functions 2 . D espite the ubiquity of natural , incidental, and engineered nanomaterials, only in the past decades have their impacts on the environment been extensively studied 2 . This progress mainly resulted from multiple advances in analytic tools and techniques , for example, the development of scanning tunneling microscop y which has provided us with much better understanding of the distinct behaviors of ENMs. Nanotechnology manipulates matter at the nanoscale to explore distinct properties and phenomena. Since the launch of the National Nanotechnology Initiative (NNI) in 2000, three main development al stages of nanotechnology have been envisioned, and we are now at the beginning stage translating nanoscale science to marketable technology and products ( Nano 3 ) 3 . Wo rldwide t he market value of nanotechnology - incorporated products was $1 trillion in 2013, and was projected to be over $3 trillion by 2020 4 and about one order of magnitude more by 2030 3 . I n 2005 the Woodrow Wilson International Center for Scholars and the Project on Emerging Nanotechnologies initiated the Nanotechnology Consumer Products Inventory, which was revised in 2013, documenting 1814 nano - enabled consumer products from 622 companies in 32 countries 5 . A very recent search of the scientific literature shows that since 2000 over 8 000 articles on the topic of ENMs were published with an exponential increase (Figure 1.1) , and is expected to grow 3 substantially in next ten years and beyond . Nanotechnology will likely continue to grow exponential ly by vertical science - to - technology transition and horizontal expansion to other fields (e.g., environmental field). 4 Figure 1 .1. Publications trend in engineered nanomaterials research. Web of Science results for 5 Environmental Application of Nanote chnology N anotechnology for e nvironmental application is one of the most promising approaches to address environmental problems in air, water, and soil. Among them, water treatment using ENMs is a main focal point. The world is facing water scarcity globally, due to shortages and pollution of fresh water resources 6, 7 . More alarmingly, a n increasing number of emerging contaminant s (e.g., pharmaceuticals) are found in rivers, lakes, groundwater aquifers, and oceans 8 - 16 , which raises substantial concerns on their impact to ecosystem and human health . P harmaceuticals are not readily removed by conventional wastewater treatment plant s (WWTPs) 16 - 19 . As a result , pharmaceuticals are increasingly being detected at ng L 1 µg L 1 levels in aquatic environments 15, 20, 21 , which may pose acute and chronic toxicities to aquatic organisms 22 - 24 . In addition, p harmaceuticals ubiquitous ly foun d in reclaimed wastewater and sewage sludge could be released into agricultural soils via crop irrigation and land application of sewage sludge, followed by assimilat ion and transform ation in food crops 25 - 28 . Thus, the h uman population may be potentially exposed to pharmaceutical residues or their metabolites via consumption of contaminated food crops. Paltiel et al 29 showed that volunteers consuming reclaimed wastewater - irrigated produce had higher urinary concentration s of carbamazepine and its metabolites compared to those consuming fresh water - irrigated produce. T herefore , rising public health and environmental concerns necessitate the development of effective and robust methods to remove pharmaceuticals from water . R e cent advances in nanotechnology can be harnessed to develop next - generation water treatment techniques 22 . Several ENMs (e.g., TiO 2 , ZnO, Bi x O y , g - C 3 N 4 , GO) may be used to remove pharmaceuticals from wastewater through adsorption, photocatalysis, or membra ne filtration. In particular, photocatalysis by ENMs has been intensively studied for treating water 6 and wastewater due to its low cost and high efficiency 30 - 36 . In fact there have been o ver 8000 publications on photocatalytic water treatment since 2000 3 5 . I t has shown that photocatalysis assisted with ENMs exhibits remarkable performance for advanced, robust, and multifunctional water treatment. However, many obstacles still need to be addressed in order to optimize photocatalysis, including effective se paration of electron - hole pairs (e /h + ) and subsequent generation of reactive radicals, and recovery and separation of ENMs catalysts. Environmental Implication of Nanotechnology Like many other emerging technologies, environmental impacts of nanotechnology must be considered . For example, nanotechnology m ay pose risks to environment and human health 37 - 39 , especially with the rapid production and widespread application of ENMs . Th erefore, it is crucial to assess the downstream impact of ENMs release on ecosystems and human health, thereby developing mitigation strategies. ENMs are increasingly incorporated into many industrial and consumer products, such as textile, paint, clothing , sunscreen, cosmetics, antimicrobial agents, medicine, food additives, and pesticides 5, 40, 41 . The production, use , and disposal of ENMs will lead to their inevitable dissemination into the environment. E nvironmental discharge of ENMs can occur through multiple pathways, including direct use of ENMs or ENM - containing products, sewage treatment, and landfills. Gottschalk and Nowack 42 concluded that sewage sludge, wastewater, and waste incineration of ENMs - containing products are three major flows of EMMs from a life - cycle perspective. Detailed evidence showed that both colloidal and ionic silver were steadily leached from certain socks during wa ter washing 43 . U nder natural weather ing conditions, researchers revealed that TiO 2 nanoparticles (NPs) were detached from facade paints and then carried by runoff into natural receiving water 44 . In fact , Ag NPs was detected at 100 ng L 1 in the effluent of a 7 wastewater treatment plant in the US 45 . The predicted environmental concentrations of Ag NPs in surface water, wastewater effluents, and sewage sludge were 0.09 80 ng L 1 , 0.016 1 , and 1.29 6.24 mg kg 1 , respectively 46, 47 . The annual inc rease of ENMs ranged from 1 ng kg 1 for 1 for TiO 2 NPs for sludge - treated soils in Europe and the US 46 . Crop plants are prone to ENM s exposure as a result of intentional application of nanopesticides and nanofertilizers, irrigation with reclaimed water, sewage sludge application, and atmospheric deposition 48, 49 . Extensive studies proved that ENMs in soil and water could be potentially taken up by crop plants and subsequently enter into the human food cha in 50 - 57 For instance, agglomeration of Cu NPs was observed in cells of mung bean ( Phaseolus radiatus ) and wheat ( Triticum aestivum ) 58 . Aggregates of CeO 2 NPs can be taken up by corn plants via the apoplastic pathway and translocated further to vascular t issues 59 . G eneral ly , ENMs can enter plant roots via the apoplastic or symplastic pathways 60 - 66 . It was thought that ENMs could be restricted stopped by the Casparian strip barrier following the apoplastic pathway 67 . However , the former restriction may be overcome by endocytosis 62, 68, 69 , physical wound and disease damage to roots , and the emergence of new secondary roots 64, 66 . It is believed that r educing the bioavailability of ENMs in soil is critical to minimizing their migration from soil to plant. S oil components (e.g., humic acid) can affect ENMs properties and subsequently their assimilation by plants 70 . Recently, soil amend ed with biochar was shown to effectively limit plant uptake of ENMs 71 - 74 , due to strong sorption capacity of biochar to many organic and inorganic cont aminants (including ENMs) . However, the applicability of biochar amendment to realistic exposure level of ENMs in soil has not been tested. This knowledge gap 8 limit s our ability to assess the risks of ENMs via human dietary exposure and the feasibility of biochar amendment as a mitigation strategy. Additionally, it should be noted that the development of plant protection products (e.g., nanopesticides) ha s recently drawn immense attention. For example, over one hundred pesticides containing Ag NPs have been registered in the last decade 75 - 77 . Ag and TiO 2 NPs are being used in fungicides as active ingredients 78, 79 , and silica NPs as carriers 80 . Application of nano - enabled agrochemicals can lead to direct conta ct between ENMs and the plant surface. E arlier studies have shown that ENMs likely enter plants by diffusion through the cuticle wax barrier and/or through openings in cuticle and stomata 68, 81, 82 . For example, Larue et al. 49 provided the first evidence that Ag NPs were entrapped by the cuticle and penetrated into the leaves through stomata. Although several studies addressed the potential of EN M s entry through stomatal openings, no studies have specifically investigated the role and contribution of stom ata in the sorption and internalization of EN M s by plant leaves. Research Objectives Objective 1: Evaluate photocatalytic degradation of cephalexin by ZnO nanowires. Photocatalysis assisted with novel ENMs (e.g., ZnO nanowires) may be promising for removing pharmaceuticals from water. Here we hypothesize that ZnO nanowires can effectively degrade antibiotics in water under sunlight with the degradation efficiency affec ted by water chemistries, such as solution pH, inorganic an ions, and natural organic matter. To test this hypothesis, we systematically investigate d the photocatalytic degradation of the antibiotic cephalexin using ZnO nanowires under simulated sunlight an d aimed to learn more about the efficiency, influencing factors, and mechanism of photocatalytic degradation of cephalexin . 9 Objective 2: Investigate the uptake of silver nanoparticles and ions by radish in soil with biochar amendment. Biochars often have s trong sorption capacity to many organic and inorganic contaminants (including ENMs), and thus could potentially decrease crop uptake of ENMs from soil. Here w e hypothesize that biochar can reduce the uptake of silver nanoparticles (Ag NPs) and ions by a ve getable crop ( radish ). However, biochar effectiveness may be different when plants are exposed to Ag NPs at environmentally relevant levels, compared to the previous literature using unrealistically high concentrations . Hence, the objective of this study was to evaluate the feasibility of soil biochar amendment s to decreas e radish uptake of Ag NPs and ions at environmentally relevant concentrations . Objective 3 : Explore the role of stomata in foliar sorption of silver nanoparticles by Arabidopsis thaliana . Foliar uptake is regarded as a significant pathway for the internalization of ENMs in to plant s, in addition to root uptake, because of the possible role of stomata openings . Here w e hypothesize that stomata openings contribute substantially to the interna lization of ENMs into leaves. T he objective of this study was to assess the sorption of Ag NPs by leaves using abscisic acid (ABA) - responsive ecotypes and ABA - insensitive mutants of Arabidopsis thaliana . The three objectives are addressed in the following three chapters in this dissertation. Objective 1 is addressed in Chapter II, Objective 2 in Chapter III, and Objective 3 in Chapter IV. The dissertation concludes with Chapter V that summarizes the findin gs and suggests future research directions. 10 PHOTOCATALYTIC DEGRADATION OF CEPHALEXIN BY ZnO NANOWIRES UNDER SIMULATED SUNLIGHT 11 Abstract Increasing concentrations of anthropogenic antibiotics and their metabolites in aqueous environments ha ve caused growing concerns over the proliferation of antibiotic resistance and potential adverse impacts to agro - environmental quality and human health. Photocatalysis using novel en gineered nanomaterials such as ZnO nanowires may be promising for removing antibiotics from water. However, much remains to be learned about the efficiency and mechanism for photocatalytic degradation of antibiotics by ZnO nanowires. This study systematica lly investigated photodegradation of cephalexin using ZnO nanowires under simulated sunlight. D egradation efficiency of cephalexin was substantially increased in the presence of ZnO nanowires especially at P hotodegradation followed the first - order kinetics with degradation rate constants ( k ) ranging between 1.19 × 10 1 2.52 × 10 1 min 1 at 20 80 mg L 1 ZnO nanowires. Radical 2 ) predominantly contributed to the photodegradation of cephalexin. With the addition of HCO 3 (1 5 mM) or Suwannee River natural organic matter (SRNOM, 2 10 mg L 1 ), the k values were substantially decreased by a factor of 1.8 70 to 1.69 × 10 3 6.67 × 10 2 min 1 , probably due to the screening effect of HCO 3 or SRNOM sorbed on ZnO nanowires and scavenging of free radicals by free HCO 3 or SRNOM in solution. Based on mass spectrometry and molecular computation, the pathways for photodegradation of cephalexin included hydroxylation, de methylation, decarboxylation, and dealkylation. Overall, the se novel ZnO nanowires have the potential to be used for removing antibiotics from contaminated water. 12 Introduction Antibiotics are widely used to treat or prevent bacterial infections. Cephalexin, in particular, is a semisynthetic cephalosporin drug that has been extensively produced and used in human s administered to animals and humans are excreted as either parent compounds or biotransformed metabolites 8, 83 , and are subsequently discharged to the environment. As a result, a plethora of antibiotics have been detected in wastewater, surface water, an d seawater at ng L 1 to µg L 1 level s 15, 21, 23 . The elevated environmental concentrations of antibiotics have raised public concern over the ir potential adverse impact to agro - environmental quality , human health , livestock, and ecosystems 24, 84, 85 . For example, when assessing risk quotients (RQ) of antibiotics at environmental concentrations, invertebrate s 08) 23 . There is also a general consensus that antibiotics in the environment facilitate the development and proliferation of antibiotic resistant bacteria and antibiotic resistance genes 20, 86 , even at concentrations much lower than minimum inhibitory co ncentrations 87 . In addition, treated wastewaters and biosolids containing antibiotics are widely used in agricultural production, resulting in accumulation and transformation of antibiotics in crop plants 25 - 27, 88 . Thus, sensitive and susceptible human p opulations may be disproportionally impacted by exposure to antibiotics and their metabolites through dietary intake of contaminated food crops 29, 88 . Therefore, it is becoming increasingly important to develop effective methods for removing antibiotics f rom contaminated waters. R emoval of antibiotics by wastewater treatment plants (WWTPs) is highly variable , depending on wastewater characteristics, treatment processes, operational conditions of WWTPs, and properties of the individual antibiotics 17, 18 . F or example, cephalexin concentration s can be up 13 to 64 µg L 1 in WWTP influents 16 and 5 µg L 1 in WWTP effluents 89 . The removal efficiency of cephalexin in conventional WWTPs varies between 9% 96% 16, 19 . Advanced tertiary treatment methods may be used for removing antibiotics at WWTPs, such as activated carbon adsorption, ozonation, nanofiltration, and reverse osmosis 17 . Enhanced removal, albeit not for all compounds nor for all WWTPs, has been observed with the advanced treatment methods 17 . In comparison, photocatalysis has recently received tremendous research attention for removing antibiotics from water due to its low cost and high efficiency 30 - 33 . One of the most critical aspects in photocatalysis (e /h + ) when catalysts are irradiated with photons at an energy level greater or at least equal to their band gap energy. Notably, zinc oxide (ZnO) nanomaterials can be excited by a broad solar spectrum 90 . They als o have excellent quantum efficiency 91 and environmental friendliness 92 . Thus, ZnO nanomaterials have been studied for photocatalytic degradation of organic dyes (e.g., methylene blue and rhodamine 6G), phenolic compounds (e.g., chlorophenol and fluorophe nol), and pharmaceuticals (e.g., ampicillin, caffeine, cloxacillin, resazurin, and tetracycline) 33, 91, 93 - 98 . P hotodegradation efficiency of organic contaminants by ZnO nanomaterials depends on a number of factors including the dosage, structure, and mor phology of ZnO nanomaterials, initial contaminant concentration, light wavelength, irradiation intensity, solution pH, temperature, inorganic species, and natural organic matter (NOM) 91, 98 . Specifically, ZnO nanomaterials have various structures and morp hologies such as nanospheres, nanobelts, nanosheets, nanorods, nanowires, nanodumbbells, nanoflowers, and nanotetrapods 91, 97 , resulting in various photocatalytic efficiencies due to the difference in specific surface area, surface - to - volume ratio, and ex ternal mass transfer rate 91, 93, 95, 99 . In particular, ZnO nanowires may have enhanced catalytic properties due to their high surface - to - volume ratio, and strong defect emission 95, 97, 99 . 14 ZnO nanowires may be promising as large - scale, cost - effective, a nd highly efficient photocatalysts and can be more easily separated and recovered especially when embedded on substrates 95, 100 - 102 . Nonetheless, only a limited number of studies have investigated the photodegradation of organic contaminants by ZnO nanowi res 95, 102 a nd to our best knowledge , no study has examined the photodegradation of cephalexin by ZnO nanowires. Therefore, more work is needed to develop better ZnO nanowire - based treatment systems for degradation of cephalexin and other antibiotics from contaminated water. As mentioned above water chemistry such as solution pH, anions (e.g., HCO 3 , Cl , and NO 3 ), and NOM may substantially influence the photodegradation of cephalexin. Solution pH may inhibit or enhance photodegradation by influencing dissociation and charge speciation of ionizable contaminants and contaminant sorption onto catalyst surfaces 91, 9 8, 103 . Similarly, NOM and anions (e.g., HCO 3 ) could compete for sorption sites on photocatalyst surfaces and/or scavenge reactive species, thus decreasing photodegradation efficiency 104 - 108 . However, it was also reported that the presence of HCO 3 actua lly enhanced the photodegradation of sulfamethoxazole 32, 36 . Hence, the effect of water chemistry on the photodegradation of cephalexin warrants further investigation. Therefore, this study aimed to investigate the photodegradation of cephalexin by ZnO na nowires under simulated sunlight at a range of water chemistries. The tested ZnO nanowires were synthesized via a hydrothermal method and characterized using transmission electron microscopy (TEM), scanning electron microscopy (SEM) with energy - dispersive X - ray spectroscopy (EDS), X - ray diffraction (XRD), and a Zetasizer. Photodegradation efficiency and kinetics, and the effects of water chemistries (i.e., solution pH, HCO 3 , and NOM) were systematically examined, and underlying photodegradation mechanisms (including pathways) 15 were identified using both molecular computation and mass spectrometry. This study could provide useful information for improved understanding on the photodegradation of cephalexin by ZnO nanowires and for future development of novel t reatment systems that incorporate ZnO nanowires. Materials and Methods Chemicals Cephalexin (99.7%, analytical grade), potassium dichromate (99.5%), zinc acetate dihydrate (99.9%), tert - butanol (99.5%), p - benzoquinone (98%), and methanol (HPLC grade) were purchased from Sigma - Aldrich (St. Louis, MO). Sodium bicarbonate, sodium chloride, sodium hydroxide and ethylenediaminetetraacetic acid disodium salt (EDTA - 2Na) were of analytical grade and obtained from J.T. Baker (Philipsburg, NJ). Acetonitrile (HPLC gra de) and hydrochloric acid (37%) were purchased from EMD Chemicals (Gibbstown, NJ). Ethanol (100%) was purchased from Fisher Scientific Inc. (Hampton, NH). Chemical structure and species distribution of cephalexin are shown in Supplementary Figure A1 of App endix A. Suwannee River natural organic matter (SRNOM, 2R101N) was acquired from the International Humic Substances Society and contains 50.70% C, 3.97% H, 41.41% O, 1.27% N and 1.78% P. Reagents were used without further purification. All solutions were p repared in Milli - Millipore, United States Filter Corporation, CA) and then adjusted to the desired pH using 0.1 M HCl or 0.1 M NaOH. To prepare the stock solution of SRNOM, 0.2 g of SRNOM was first dissolved in 100 mL of Milli - Q water and stirred overnight. The SRNOM solution was adjusted to pH 7.2 before filtering through a 0.45 - µm cellulose acetate membrane to remove any insoluble materials. Total organic carbon (mg C L 1 ) of the SRNOM stock solution was quantified using an Aurora 1030 TOC analyzer (OI Analytical, Xylem, UK). 16 Synthesis and Characterization of ZnO Nanowires ZnO nanowires were synthesized using a modified hydrothermal method as previously described 109, 110 . Briefly, a ZnO seed solution was prepared by addi ng 0.1 M Zn(CH 3 COO) 2 2 O in ethanol and stirring at 58 °C for 2 h. The ZnO seed solution was then mixed with 0.5 M NaOH solution at a 1:2 volume ratio and transferred to a Teflon - lined autoclave, followed by heating at 150 °C for 24 h. Afterwards, the mix ture was centrifuged at 10,000 rpm for 10 min at room temperature. The pellets were thoroughly rinsed with deionized (DI) water and dried in an oven at 60 °C for 12 h. The prepared ZnO nanowires were imaged by TEM (JEOL 2200FS, Tokyo, Japan) and SEM (JEOL 7500F, Tokyo, Japan) equipped with an Oxford EDS at an accelerating voltage of 5.0 kV. The XRD pattern of the ZnO nanowires was obtained on a Bruker D8 DaVinci diffractometer ( Bruker Corporation , Billerica, MA ) equipped with Cu X - ray radiation operating at 40 kV and 40 mA. The ZnO nanowire s were placed in a PVMA sample holder and rotated at 5º/min. Peak intensities were obtained by counting with the Lynxeye detector every 0.02º/0.5 s. A Zetasizer (Nano ZS, Malvern, UK) was used to measure zeta potential and hydrodynamic diameter of the 3 1 ). Measurements were repeated 3 times for at least two separate samples at 25 °C. To measure ZnO nanowire d issolution, aliquots of ZnO nanowire suspensions were filtered using Amicon Ultra 3k ultrafiltration unit s (Millipore, Burlington, MA). The filtrates were measured for Zn 2+ ion concentration using atomic absorption spectroscopy (AAnalyst 400, PerkinElmer, Waltham, MA). 17 Photodegradation Experiments A series of photodegradation experiments were carried out in a SUNTEST XLS+ Solar simulator (Atlas Material Testing Technology LLC, Mount Prospect, IL) equipped with an air - cooled 1700 W xenon lamp. A daylight fil ter with a cutoff wavelength < 290 nm was used to simulate sunlight, and the temperature was maintained at 25 ºC using a SunCool unit. The irradiation intensity was set to 700 ± 5 W m 2 , similar to that used in other studies 111, 112 . All photodegradation experiments were conducted in triplicate. To assess the effect of catalyst dos e on photodegradation efficiency, ZnO nanowires were added to 50 mL of a 100 µg L 1 cephalexin solution at 20, 40, or 80 mg L 1 . Background solution ionic strength was 5 mM NaCl and the initial solution pH was 7.2. The initial cephalexin concentration of 100 µg L 1 was slightly higher than average environmental cephalexin concentrations (ng L 1 to µg L 1 ), and somewhat above the reported maximum concentration of 64 µg L 1 in influent at a WWTP 16, 19, 89 . However, the level was far lower than the cephalexin 1 ) typically used in previous studies 30, 113 - 115 . With increasing contaminant concentrations, the photodegradation efficiency would be expected to increase until reaching the optimal contaminant concentration, and then decrease at higher contaminant concentrations 91, 98 . Therefore, we selected the low initial concentration to better assess the photodegradation efficiency at environmentally - releva nt concentrations. Prior to exposure to simulated sunlight, the suspension was placed in the dark for 30 min to reach the adsorption - desorption equilibrium. After initiating the photodegradation experiments, aliquots (0.5 mL) of the suspensions were withdr awn at various time intervals ( 5, 10, 25, or 30 min) and then immediately quenched with methanol. The samples were centrifuged at 13,000 rpm for 15 min, and the supernatant was collected and stored at 4 °C in amber glass vials before analysis. At each samp ling 18 point the suspension was shaken vigorously by hand to minimize sedimentation during the photodegradation experiments. No visible sedimentation was observed, likely due to the stabilization of ZnO nanowire aggregates after initial aggregation and the s hort sampling intervals (5 30 min). To evaluate the performance of ZnO nanowires for cephalexin degradation in environmentally - relevant conditions, a wide range of water chemistries was examined. Various 3 1 ) concentrations were adjusted in the suspensions containing 20 mg L 1 ZnO nanowires and 100 µg L 1 cephalexin. Parallel control experiments were also conducted under identical conditions without ZnO nanowires or with Zn 2+ ions (~10 mg L 1 , as suming the maximum dissolution) in place of ZnO nanowires. Additional experiments using recycled ZnO nanowires were performed to evaluate the reusability and stability of ZnO nanowires using experimental condition s and procedures identical to those of the kinetic experiments. At the end of each cycle (90 min irradiation time per cycle), the cephalexin concentration in the solution was measured and then replenished to 100 µg L 1 , followed by the next light irradiation. Degradation efficiency was calculated b y dividing the difference between the initial and final concentration with the initial concentration in each cycle. The photodegradation kinetics of cephalexin were fitted using a first - order kinetic model: where C 0 and C t represent the cephalexin concentrations at time 0 (i.e., after reaching the adsorption - desorption equilibrium in dark) and the sampling time t , respectively, and k is the first - order degradation rate constant (min 1 ). One - way ANOVA analysis was performed to test the statistical significance in mean values of k using the least significant difference (LSD) test with . 19 Trapping Experiments To identify reactive radicals responsible for cephalexi n degradation, trapping experiments were performed under experimental conditions identical to those in the photodegradation experiments, except that specific scavengers were added to the suspensions before exposure to simulated sunlight. Ethylenediaminetet raacetic acid disodium (EDTA - 2Na, 0.1 mM) was used as a scavenger for the photo - induced holes (h + ) 32 , tert - 116 , potassium dichromate (K 2 Cr 2 O 7 , 50 µM) for the photo - induced electrons (e ) 117 , and p - 2 ) 118 . Analytical Methods Cephalexin concentrations in the samples were determined using a Shimadzu prominence high - performance liquid chromatography system (Shimadzu, Columbia, MD) coupled with an AB Scie x 4500 QTrap triple quadrupole mass spectrometer (SCIEX, Foster City, CA) (LC - QTrap - MS/MS). An Agilent Eclipse Plus C18 column (50 mm × 2.1 mm, 5 µm, Agilent, Santa Clara, CA) was used for chromatographic separation prior to mass spectrometry analysis. The binary mobile phase consisted of water (phase A) and acetonitrile (phase B) with 0.1% (v/v) formic acid. The degradation products were identified using LC - QTrap - MS/MS under positive ionization mode, in combination with an enhanced mass scan as a survey sc an, information dependent acquisition criteria, and an enhanced product ion scan to obtain the MS/MS fragment patterns. The full approach for identifying cephalexin oxidation products can be found in Chuang et al. 119 . Details on analytical method and prod uct identification procedures are given in Supplementary Text A1 of Appendix A. 20 Theoretical Calculation To help identify the reactive sites for free radical attack and possible photodegradation pathways, molecular orbital calculation of cephalexin with the optimal conformation 34 . The frontier electron densities (FED) of the highest occupied molecular orbital (HOMO) and the lowest unoccupied molecular orbital (LUMO) of e ach atom in cephalexin were calculated to theoretically predict locations of reactive sites for free radical attack. The positions with higher values of 2FED HOMO 2 are more susceptible to electron extraction, whereas those with higher values of FED HOMO 2 + F ED LUMO 2 are more likely to be attacked by free radicals 34 . Results and Discussion Characterization of ZnO Nanowires Representative SEM, TEM, and XRD pattern of ZnO nanowires are shown in Figure 2.1. The average diameter of ZnO nanowires was 39 ± 10 nm and their length varied between 56 1,454 nm with a mean value of 328 nm measured from randomly selected ~100 ZnO nanowires using ImageJ 1.52a software (Figure 2.1a and b). The sharp diffraction peaks in Figure 2.1c confirmed good crystallinity of the ZnO nanowires, and the intense peaks that originated from (100), (002), (101), (102), (110), (103), (200), (112), and (201) can be well indexed to the hexagonal wurtzite Zn O structure in accordance with the standard JCPDS card (no. 36 - 145). The strong intensity of (002) peak indicated the preferable growth of ZnO nanowires along the c - axis orientation. Additionally, no diffraction patterns of any other impurities were detect ed, and no other elements except Zn and O were found on the EDS spectra (Supplemental Figure A3), confirming that the synthesized ZnO nanowires were primarily pure ZnO hexagon that is favored as a photocatalyst 120 . 21 Figure 2 . 1 . Representative (a) scanning electron microscopy, (b) transmission electron microscopy images, and (c) X - ray diffraction pattern of ZnO nanowires. 22 Table 2.1 shows the zeta potentials and hydrodynamic diameters of ZnO nanowires in suspensions of varying water chemistries. As solution pH increased, the zeta potentials became more negative. The surface charge of ZnO nanowires reversed from positive to n egative at pH 6.8 which is defined as the point of zero charge (pH pzc ) for ZnO nanowires (Supplemental Figure A4). Accordingly, prior to charge reversal, the hydrodynamic diameters of ZnO nanowires increased with increasing pH, due to decreasing electrosta tic repulsion between positively charged ZnO nanowires. After the solution pH increased beyond pH pzc of 6.8, the hydrodynamic diameters of ZnO nanowires decreased, resulting from increasing electrostatic repulsion between negatively charged ZnO nanowires ( Table 2.1). The largest hydrodynamic diameter (5192 ± 481 nm) was observed near pH pzc , indicating the absence of any repulsive electrostatic interaction 121 . The addition of HCO 3 a minor influence on the zeta potentials and hydrodynamic diame ters of ZnO nanowires. However, 1 ) decreased the surface charge of ZnO nanowires sharply (Table 2.1), leading to a decrease in hydrodynamic diameter compared to the SRNOM - free suspensions of ZnO nanowires. This can be attri buted to both increased electrostatic and steric repulsion by the adsorbed SRNOM on the surface of ZnO 1 did not produce size measurements met the analysis quality criteria of the Zetasizer. Therefore, the measured size should have been primarily contributed by ZnO nanowires, and not by SRNOM. We did not attempt to distinguish the contribution of the adsorbed SRNOM layer to the aggregate size of ZnO nanowires, which was beyond the scope of thi s study. 23 Table 2. 1 . Zeta potentials and hydrodynamic diameters of ZnO nanowires under various experimental conditions. a This measurement did not meet the quality criteria using dynamic light scattering. 24 Photodegradation Efficiency and Reusability Figure 2.2 shows the photocatalytic degradation kinetics of cephalexin in the absence and presence of ZnO nanowires under simulated sunlight at pH 7.2. A negligible amount of cephalexin was degraded by hydrolysis and photolysis, as well as by the presence of Zn 2+ ions (10 mg L 1 ) over 180 min under simulated sunlight. However, the addition of ZnO nanowires (i.e., 20, 40, or 80 mg L 1 ) significantly enhanced the photodegradation of cephalexin ( p < 0.0 5, Table 2.2) , with over 96% of the cephalexin degrad ed within 25 min and complete decay within 45 min in the presence of 20 mg L 1 ZnO nanowires. Liu et al. 30 investigated the photocatalytic activity of CdSe quantum dots (500 mg L 1 ) on cephalexin (15 mg L 1 ) under UV irradiation, and only observed a degradation efficiency of 70.34% after 60 min. However, comparing the photodegradation efficiency of different catalysts is often very challenging due to differences in experimental conditions such as catalys t - to - contaminant ratios ranging between 30 500 in previous studies (Supplemental Table A1) 114, 122 - 126 . Additionally, turnover number and turnover frequency are difficult to estimate because determining the number of surface - active sites is a challenge 12 7 . Nonetheless, this study revealed that ZnO nanowires could effectively degrade cephalexin at trace - level concentrations (< 100 µg L 1 ) that are about two orders of magnitude lower than commonly 1 ) in previous studies 30, 113 - 115 . Thus, ZnO nanowires could be well suited to degrade cephalexin from environmental water. 25 Figure 2 .2. Observed (a) and fitted (b) photodegradation kinetics of cephalexin by ZnO nanowires (20, 40, and 80 mg L 1 ) at pH 7.2 under simulated sunlight. Error bars represent the standard deviations ( n = 3). Solid lines were fitted to initial parts of kinetics tha t followed the first - order degradation. 26 Cephalexin d egradation followed the first - order kinetic model (Figure 2.2b) and the fitted k values are summarized in Table 2.2. The k values for cephalexin degradation remarkably increased from 6.00 × 10 4 min 1 for hydrolysis and 4.00 × 10 4 min 1 for photolysis to 1.19 × 10 1 2.52 × 10 1 min 1 in the presence of ZnO nanowire (20, 40, or 80 mg L 1 ). This observation is congruent with other studie s 32, 33 . For example, the photodegradation rate of sulfamethoxazole was 2.04 × 10 4 min 1 by photolysis and increased to 0.023 min 1 by photocatalysis using g - C 3 N 4 32 . Additionally, in comparison the k values for cloxacillin (0.029 min 1 ), amoxicillin (0. 018 min 1 ), and ampicillin (0.015 min 1 ) using a UV/ZnO system 33 were 10 times lower than the observed k values in this study. Thus, our ZnO nanowires had a high photodegradation efficiency. The reusability test results are shown in Supplemental Figure A5 . The ZnO nanowires were capable of completely degrad ing cephalexin (100 µg L 1 ) within 90 min in the first two use cycles, with > 94% degradation efficiency achieved in the fifth cycle. The slight decrease in efficiency was likely due to the dissolution of ZnO nanowires, as evidenced by the detected Zn 2+ concentration of 2.79 mg L 1 after 5 cycles (Table 2.3). Interestingly, there were no visible morpholog ical changes in the ZnO nanowires after 180 min irradiation as shown by the TEM images (Supplementary Figure A6). Therefore, these results suggest that the ZnO nanowires possess satisfactory stability for cephalexin degradation over at least 5 cycles under simulated sunl ight. 27 Table 2. 2 . Fitted parameters for degradation kinetics of cephalexin in all experiments. Experimental Sets Degradation rate constant ( k ) (min 1 ) R 2 Hydrolysis * 6.00 (0.00) × 10 4 d 0.83 Photolysis ** 4.00 (1.73) × 10 4 d 0.83 10 mg L 1 Zn 2+ 1.07 (0.05) × 10 3 d 0.98 20 mg L 1 ZnO nanowires 1.19 (0.02) × 10 1 a 0.99 40 mg L 1 ZnO nanowires 2.33 (0.12) × 10 1 b 0.99 80 mg L 1 ZnO nanowires 2.52 (0.18) × 10 1 c 0.99 No scavenger 1.19 (0.02) × 10 1 a 0.99 0.1 mM EDTA - 2Na 3.13 (1.00) × 10 4 c 0.65 10 mM tert - butanol 3.59 (0.42) × 10 3 d 0.98 50 µM K 2 Cr 2 O 7 5.55 (0.20) × 10 2 b 0.98 1 mM p - benzoquinone 6.56 (1.00) × 10 3 d 0.70 pH = 5.0 8.33 (2.08) × 10 4 b 0.95 pH = 6.2 4.13 (0.59) × 10 3 b 0.96 pH = 7.2 1.19 (0.02) × 10 1 a 0.99 pH = 8.3 2.15 (0.11) × 10 1 c 0.97 pH = 9.2 3.03 (0.08) × 10 1 d 0.99 0 mM HCO 3 1.19 (0.02) × 10 1 a 0.9 9 1 mM HCO 3 6.67 (0.35) × 10 2 b 0.98 2 mM HCO 3 5.45 (0.18) × 10 2 c 0.99 5 mM HCO 3 1.69 (0.26) × 10 3 d 0.82 0 mg L 1 SRNOM 1.19 (0.02) × 10 1 a 0.99 2 mg L 1 SRNOM 4.64 (0.39) × 10 2 b 0.99 5 mg L 1 SRNOM 3.07 (0.40) × 10 3 c 0.99 10 mg L 1 SRNOM 2.60 (0.35) × 10 3 c 0.96 * The degradation experiments were performed in the dark without ZnO nanowires. ** The degradation experiments were performed under simulated sunlight without ZnO nanowires. Numbers in parentheses are standard deviations from three replicates. In e ach treatme nt group (e.g., ZnO nanowire dosage, different scavengers , varying solution pH, bicarbonate concentration, and NOM concentration) different letters indicate the significant difference in means of degradation rate constants by the LSD post - hoc comparison te st at p < 0.05 . 28 Tabl e 2. 3 . Dissolution of ZnO nanowires (20 mg L 1 ) after 3 - hrs experiment under different solution pH. Reactive Species in Photodegradation In photocatalysis several reactive species (e.g., h + , and O 2 ) may be produced and react with cephalexin 32, 116, 117 . The photodegradation rates of cephalexin were significantly reduced by adding a h + scavenger (0.1 mM EDTA - tert - butanol), and O 2 scavenger (1 mM p - benzoquinone), respectively ( p < 0.05, Table 2.2 and Figure 2.3). Further compa rison showed that the inhibitory effect of tert - butanol was lower than that of EDTA - 2Na. Knowing that the combination of h + with OH 2 are the main reactive species contributing to the photocatalytic degradation of cephalexin. A ddition of the e scavenger (50 µM K 2 Cr 2 O 7 ) remarkably suppressed the photodegradation of cephalexin but with a lower inhibition effect than other three scavengers ( p < 0.05, Table 2.2 and Figure 2.3). Thus, photo - induced electrons (e ) had a smaller contribution to the photodegradation of cephalexin. 29 Figure 2. 3 . Effect of various free radical scavengers on the photodegradation of cephalexin by ZnO nanowires (20 mg L 1 ) at solution pH of 7.2 under simulated sunlight. Error bars represent the standard deviations ( n = 3). Solid lines were fitted to the first - order k inetic equation. 30 Effect of Solution pH As shown in Figure 2.4a and Table 2.2, the photodegradation rate of cephalexin was significantly accelerated and the k value increased from 8.33 × 10 4 min 1 to 3.03 × 10 1 min 1 as the pH increased from 5.0 to 9.2 ( p < 0.05) . A similar trend was observed for the degradation of amoxicillin, ampicillin, and cloxacillin by ZnO under UV irradiation 33 , which was thought to be collectively controlled by the attractive and repulsive forces between catalyst surfaces and antibiotics. Along this line of thought, at a solution pH less than the pH pzc of 6.8, the attraction between positively charged ZnO nanowires (Table 2.1 and Supplementary Figure A4) and the zwitterionic and/or anionic species of cephalexin (Supplemental Figure A1b) would have enhanced the photodegr adation of cephalexin. At a solution pH greater than pH pzc , the repulsion between anionic cephalexin and negatively charged ZnO nanowires would have limited photocatalytic degradation. Surprisingly, the opposite trends were observed in this study, suggesti ng that electrostatic interaction had little effect on the photodegradation of cephalexin by ZnO nanowires. A previous study showed that solution pH can affect the generation of radical species 103 . At a lower pH the h + holes were the major oxidation speci es, whereas at a neutral or conditions, the increased OH reacting with h + 32, 33, 103 . As hydroxy production at higher pH contributed to greater cephalexin degradation. Additionally, the dissolution of Zn 121 , which could partly explain the negligible cephalexin degradation at pH 5.0 and 6.2 in addition to 31 Figure 2. 4 . Effects of initial solution pH (a), bicarbonate (b) and Suwannee River natural organic matter (SRNOM) (c) on the photodegradation of cephalexin by ZnO nanowires ( 20 mg L 1 ) under simulated sunlight. Error bars represent the standard deviations ( n = 3). Solid lines were fitted to the first - order kinetic equation. 32 Effect of Bicarbonate Cephalexin degradation decreased with increasing bicarbonate concentration (1, 2, and 5 mM) at pH 7.2 as shown in Figure 2.4b. The k value decreased from 1.19 × 10 1 min 1 at 0 mM bicarbonate to 6.67 × 10 2 1.69 × 10 3 min 1 at 1 5 mM bicarbonate ( p < 0.05 , Table 2.2 ). Based on speciation calculations using the p K a values (i.e., p K a 1 = 6.3 and p K a 2 = 10.3) for a H 2 CO 3 /HCO 3 /CO 3 system 128 , at pH 7.2 the predominant species are HCO 3 (88.8%) and H 2 CO 3 (11.2%). HCO 3 3 ) 107 , which would considerably decrease the photocatalytic efficiency of ZnO 3 106 . Additionally, inorganic ions including HCO 3 can be adsorbed on the surface of TiO 2 in the order of HPO 4 > HCO 3 > SO 4 > NO 3 > Cl , even when TiO 2 is negatively charged 129 . Therefore, the inhibition effect of co - existing HCO 3 on cephalexin degradation might be partly explained by the formation of a HCO 3 layer on the surface of the ZnO nanowires which further impairs the approach of cephalexin to the ZnO nanowires 130 . Similarly, Torki and Faghihian 31 revealed that HCO 3 inhibited the photodegradation of cephalexin when using synthesized Fe 3 O 4 @PPY NiO, Fe 3 O 4 @PPY NiS, and Fe 3 O 4 @PPY NiO NiS as catalysts under visible light. In contrast, several researchers found that HCO 3 had a negligible or even enhanced effect on photocatalytic degradation of antibiot ics. Hu et al. 36 and Song et al. 32 both showed that the addition of bicarbonate (HCO 3 ) enhanced the photocatalysis of sulfamethoxazole by TiO 2 and g - C 3 N 4 , respectively. These observations may be due to the 3 with the ani line group of sulfamethoxazole ( k = 6×10 8 M 1 s 1 ). Likewise, a very recent paper by Qian et al. 115 3 radicals can selectively react with the electron - rich groups of cephalexin, such as the primary amine (Supplemental Fig ure A1a), which would significantly increase the oxidation of cephalexin 33 by thermally - activated persulfate. Conversely, a negligible effect of HCO 3 on the photodegradation of sulfadiazine and sulfamerazine by g - C 3 N 4 w as also observed by Song et al. 32 . Clearly, whether bicarbonate plays an inhibit ory or enhanc ing role in the degradation of antibiotics is dependent on the type of dominant radical species and the chemical properties of the antibiotic. In addition to bicarbonate (HCO 3 ), other inorganic ani ons such as Cl and NO 3 are also ubiquitous in water. Similarly, it is hypothesized that Cl and NO 3 may inhibit the photodegradation of cephalexin by preferential adsorption on the photocatalyst surface, competitive reaction with holes (h + ), and consumi 98, 131 . Nonetheless, this hypothesis needs to be confirmed in future studies. Effect of SRNOM Figure 2.4c presents the effect of SRNOM on the degradation efficiency of cephalexin by ZnO nanowires under simula ted sunlight. As expected, the k value decreased from 1.19 × 10 1 min 1 in the absence of SRNOM to 4.64 × 10 2 2.60 × 10 3 min 1 with the presence of 2 10 mg L 1 SRNOM ( p < 0.05, Table 2.2), which is consistent with previous studies 32, 104, 105 . As SRNOM concentration increased from 0 to 10 mg L 1 , the degradation efficienc y dramatically decreased from near 100% to 40.9%. This could be attributed to the quenching effect of SRNOM on the major reactive 2 ) 106 . NOM is a well - known scavenge r of reactive radicals especially at high concentrations 104 . Additionally, NOM is capable of attenuating light and consuming most of photons 104, 108 . SRNOM sorbed on the ZnO nanowire surface occupied the reactive sites as evidenced by the more negative zeta potentials for the ZnO nanowires (Table 2.1), which further inhibited the approach of cephalexin molecules onto the surface of the ZnO nanowires 105 . It should be noted that NOM can also ac t as a photosensitizer to enhance photolysis. NOM was observed to promote indirect photolysis of caffeine, sulfamethoxazole, and diuron 132 . 34 Dissolved organic matter (DOM) was excited to triplet states ( 3 DOM*) that are highly reactive with organic pollutan ts 133 . For example, 3 DOM* effectively facilitated the photo - transformation of 17 - estradiol 134 and 2,4 - dihydroxybenzophenone 135 . Identification of Intermediates and Transformation Pathways FED calculation s w ere used to predict the positions of free radical attac k , which w ere s selectively attack positions within a molecule (i.e., intramolecular selectivity) 136 . As given in the Supplemental Table A2, C23 ha d the highest value of 2FED HOMO 2 , followed by C18 and C17, and then C19, N24, and C20. Thus, these positions would be prone to electron extraction. The FED HOMO 2 + FED LUMO 2 values decreased in the order 34 . Thus, these result s could provide a theoretical basis for predicting the chemical structures of intermediates and photocatalytic degradation pathways in combination with LC - QTrap - MS/MS Photodegradation products of cephalexin were ana lyzed by LC - MS full scan and enhanced product ion scan. Their structures were obtained in the ChemSpider database or constructed by ChemDraw software (ChemOffice programs, PerkinElmer, Waltham, MA) following molecular formula calculation based on the MS sp ectra and m/z values. The proposed structures of products and degradation pathways are depicted in Figure 2.5. According to the FED theory and identified products, the photocatalytic pathway of cephalexin by ZnO nanowires mainly included hydroxylation, dem ethylation, decarboxylation, and dealkylation. Photocatalytic product 1 (P1, 2 - lactam ring opening through hydroxylation and demethylation via substitution of methyl with hydroxyl addition. Previous studies have reported similar degradation pathways for amoxicillin and cephalosporins 137, 138 . 35 Despite the highest values of 2FED HOMO 2 and/or FED HOMO 2 + FED LUMO 2 at C23 and C18 of the phenyl ring, an earlier study by Hsu et al. 138 cephalexin is unlikely to be oxidized before the core structure (Supplemental Figure A1a). ly hinder and block the reactive sites in the core structure. Thus, demethylation at C3 with hydroxyl addition was more likely, similar to the degradation of sulfamethoxazole 139 . Intermediate P2 (m/z = 370.72) was generated by losing one O atom at the C9 position. Decarboxylation and dealkylation led to the formation of P3 (m/z = 327.88), which has been documented in degradation of cephalexin and other pharmaceuticals 30, 137, 140 . Subsequently, the oxidation product P4 (m/z = 192.72) and small fragment P7 (m/z = in P3. Hsu et al. 138 phenylglycine, the precursor of P7, was released as a byproduct after 3 h of cephalexin oxidation by MnO 2 . Furthermore, dehydration with the loss of H 2 O an d oxygen can form the intermediates P5 (m/z = 174.74) and P6 (m/z = 158.72), respectively. In this study seven products were tentatively proposed based on the MS spectra and m/z values in the photocatalytic degradation pathways, which were justified by cal culat ed theoretical FED and reported pathways in the literature. Due to the lack of authentic standards, these intermediates were not confirmed and the transformation pathway was thus speculative. Nonetheless, this approach has been widely used in other st udies 30, 115, 138 . Furthermore, cephalexin and its transformation products could be oxidized into smaller molecules by attack from free radicals, which should be explored in future studies. As the transformation products may be also bioactive against micr oorganisms, future work should examine the bioactivity of photodegradation intermediates . 36 Figure 2. 5 . Proposed photocatalytic degradation pathways of cephalexin by ZnO nanowires under simulated sunlight irradiation. 37 Conclusion In this study, laboratory - scale photocatalytic degradation of cephalexin under simulated sunlight using ZnO nanowires was invest igated for efficiency, water chemistry effect, and degradation pathways. D egradation efficiency of cephalexin substantially increased with increasing solution pH (particularly at alkaline condition) in the presence of ZnO nanowires, indicating ZnO nanowire s have excellent photocatalytic activity for cephalexin. Photodegradation of cephalexin decreased with increasing HCO 3 and NOM concentrations, due to scavenging of 2 ) that are largely responsible for the degradation of cephalexin. Therefore, effective strategies will need to be designed when using ZnO nanowires in water and wastewater treatment at elevated levels of HCO 3 and/or NOM. For example, ZnO nanowires may be embedded into membrane reactors that can be used for advanced water and wastewater treatments. Future research should focus on developing feasible engineering designs that incorporate ZnO nanowires for the removal of contaminant s from bot h water and wastewater as well as assessing the bioactivity of transformation products during photodegradation . Author Contribution: Jianzhou He and Wei Zhang conceived and designed the research experiments; Jianzhou He conducted the experiments, analyzed the data and wrote the manuscript; Yaozhong Zhang and Junghoon Yeom provided the ZnO nanowires and the SEM characterization; Yang Guo performed the theoretical calculation; Geoff Rhodes provided assistance on the LC - MS/MS method; Junghoon Yeom, Hui Li, an d Wei Zhang reviewed and edited the manuscript. 38 BIOCHAR AMENDMENT CHANGES SOIL - BOUND FRACTIONS OF SILVER NANOPARTICLES AND IONS BUT NOT THEIR UPTAKE BY RADISH AT AN ENVIRONMENTALY RELEVANT CONCENTRTAION 39 Abstract Plant uptake of silver nanoparticles (Ag NPs) and ions (Ag + ) largely depend s on their exchangeable and soil - bound fractions, which may be influenced by biochar amendment. This study investigated the effects of biochar amendment (0.1% and 1.0%) on soil sorption o f Ag NPs and Ag + , their soil - bound fractions, and their uptake and translocation by radish grown in a loamy sand soil spiked with 1 mg kg 1 Ag NPs or Ag + . Sorption of Ag + to the soils was much greater than that of Ag NPs, mainly because negatively charged soil particles attract Ag + ions but repel negatively charged Ag NPs. Biochar amendment at 1.0% (by weight) significantly decreased the reducible fraction of Ag + ions in soils with and without radish plants and increased the oxidisable fraction of Ag NPs in soils with radish plants. Biochar amendment had no significant effect on Ag uptake by radish plants ( p > 0.05), probably due to low exchangeable Ag fractions in all experimental treatments. In this short - term experiment (35 days), the addition of 1 mg kg 1 Ag NPs or Ag + did not substantially elevate the level of Ag in radish roots (0.05 ± 0.02 1.06 ± 0.98 mg kg 1 ) and shoots (0.01 ± 0.00 0.03 ± 0.01 mg kg 1 ), compared to the blank control ( p > 0.05). Radish uptake of Ag NPs and Ag + at the environmentally - r elevant concentration was low with root concentration factors between 0.03 ± 0.03 to 0.29 ± 0.21 and root - to - shoot translocation factors between 0.08 ± 0.10 to 0.89 ± 1.21 , which may partially explain the non - significant effect of biochar amendment on Ag u ptake. 40 Introduction Silver (Ag) nanoparticles (NPs) are one type of the most widely used engineered nanomaterials and are increasingly incorporated into industrial and commercial products 5, 141 . It is generally believed that the production, use , and disposal of Ag NPs will lead to their dissemination into agricultural soils 42, 43, 47 . Soils are thus considered a major sink or reservoir for Ag NPs 46 , and the predicted concentrations of Ag NPs in soils were 0.02 µg kg 1 and 0.1 µg kg 1 in realist ic and high emission scenarios, respectively 47 . Also the annual increase of Ag NPs in soils was estimated at 22.7 ng/kg in Europe and 8.3 ng kg 1 in the US, with a more rapid increase in sludge - treated soils (1.58 µg kg 1 in Europe and 0.66 µg kg 1 in the US). This is of particular concern because the concentrations of Ag NPs in sewage sludge can be as high as 1.68 mg/kg in Europe and 1.55 mg kg 1 in the US 46 . More importantly, if the released Ag NPs in soils accumulate in food crops and enter the human f ood chain, dietary exposure to Ag may threaten human health. A number of studies have found that Ag in either particulate or ionic form can be taken up by plants 50 - 57 . For example, when duckweed ( Landoltia punctata ) was exposed to 10 mg L 1 of total Ag co ncentration in the form of either Ag NPs, Ag 2 S NPs, or AgNO 3 for 24 h in a hydroponic system, Ag was detected on and in the roots of duckweed 142 . Using single - particle inductively coupled plasma mass spectrometry and transmission electron microscopy energy - dispersive spectroscopy, Li et al. 143 detected Ag - containing NPs in the leaves of soybean and rice after exposure to Ag NPs (30 mg L 1 for soy bean and 1 mg L 1 for rice) and Ag + ions (0.5 mg L 1 ) in hydroponic systems. Similarly, Ag NPs were taken up by the roots and subsequently translocated in the shoots of Arabidopsis thaliana cultivated in ¼ Hoagland's solution containing 2 mg L 1 Ag NPs 54 . When exposed to 70 mg kg 1 of Ag NPs or Ag + ions spiked into a Templeton silt loam, silverbeet ( Beta vulgaris L. ssp. maritima (L.) Arcang. ) and spinach ( Spinacia oleracea L. ) could 41 1 in the edible parts, which may pose human healt h risks 144 . Dimkpa, et al. 145 also found Ag NPs in the shoots of wheat ( Triticum aestivum L.) after root exposure to 2.5 mg kg 1 Ag NPs or Ag + ions in a sand growth matrix, suggesting root uptake and translocation of Ag in wheat. Nonetheless, the relativ e contribution of Ag NPs and Ag + ions to Ag plant uptake has not yet been determined. Cvjetko et al. 51 found that equal amounts of Ag NPs or Ag + accumulated in the roots and leaves of tobacco ( Nicotiana tabacum ), consistent with another study on nine vegetables 144 . In contrast, several studies have documented higher Ag contents in the roots after exposure to Ag NPs than to Ag + ions in hydroponics at total Ag concentrations of 50 1 10 mg L 1 53, 146 - 148 . Fu rthermore, the ionic Ag form tends to have higher translocation efficiency from roots to shoots than the nanoparticle form 149 . However, Yang et al. 146 reported that the translocation factor for 109 Ag in 109 Ag NPs was remarkably higher than that for 107 Ag in 107 AgNO 3 in hydroponic Hewitt solutions. Given these aforementioned conflicting results, more research is needed on the relative significance of Ag NPs and Ag + ions uptake by plants. In general, t he negatively charged plant cell membran e attract s Ag + ions, thus facilitating their entry into plant cells through ion channels or by passive diffusion. Ag NPs could be potentially taken up by plant roots via the apoplastic or symplastic pathways and translocated to shoots through the vascular s and plasmodesmata 62 - 66 . Nonetheless, the size exclusion of plant roots may be overcome by endocytosis 62, 68, 69 , and additional entry points due to physical and disease damage to r oots and emergence of new secondary roots 64, 66 . Overall, plant uptake of Ag NPs largely depends on the bioavailab ility of Ag in soils 144, 150, 151 . The enhanced aggregation and/or interaction with soil moieties, e.g., iron oxides or organic matters (OM), could decrease the bioavailability of Ag NPs in soils. For example, the presence of soil OM increased the amount of Ag retained in the solid 42 phase an d decreased the dissolved Ag levels 152 . Natural OM decreased the accumulation of Ag in wheat by strongly binding to dissolved Ag + 150 . Of particular importance is, therefore, to determine the fractions and bioavailability of Ag NPs associated with soils. Indeed, a well - established three - step sequential extraction method (known as the BCR method) has been widely used to determine the fractions and bioavailability of heavy metals in soils and sediments 153, 154 . It was suggested that the exchangeable fractio n, the Fe/Mn oxides - bound fraction, and the OM - bound fraction are potentially bioavailable to plants at varying degrees 151, 155, 156 . Recently, Zhang, Musante, White, Schwab, Wang, Ebbs and Ma 151 successfully applied the BCR method to estimate the soil f ractions of cerium oxide nanoparticles in two soil types to assess their bioavailability to radish ( Raphanus sativus L). However, to our best knowledge little has been done to examine the soil - bound fractions of Ag NPs so as to inform their bioavailability to plants 151, 155, 157 . Reducing the mobility and bioavailability of Ag NPs in soils is critical to minimizing their migration from soils to plants. Soil amendment s with biochar may be an effective strategy to minimize the entry of nanoparticles into the human food chain 71 - 74 , and are being promoted as a sustainable agricultural practice . L 1 ) in rice ( Or yza sativa L.) grown hydroponically were significantly suppressed by 2% (w/v) biochar, due to decreased bioavailability of Ag NPs 71 . Similarly, the addition of rice husk - derived biochar to a hydroponic system (3% wt) lowered the Cu content in the roots an d shoots of wheat ( Triticum aestivum Cu 2+ to biochar and blockage of Cu access to plant roots 74 . Nonetheless, most previous studies exposed plants to unrealistically high concen trations of Ag NPs in hydroponic (up to 1,000 mg L 1 ) or soil systems (up to 10,000 mg kg 1 in soils) 51, 71, 144, 158 , and their results may not be applicable in realistic scenarios. This knowledge gap may limit our ability to assess the risks of 43 Ag NPs v ia human dietary exposure and the feasibility of biochar amendment as a mitigation strategy. Therefore, this study aimed to examine the effect of biochar amendment on the sorption of Ag NPs and Ag + ions to soils and their soil - bound fractions, and further investigate the uptake and accumulation of Ag NPs and Ag + ions at an environmentally - relevant Ag level in soils, as characterized by total Ag concentrations in plant roots and shoots, as well as the root concentration factor (RCF) and root - to - shoot translo cation factor (TF) . The spiked concentration of Ag NPs or Ag + ions (1 mg kg 1 ) was close to environmentally - relevant levels 159, 160 . Although much greater than the predicted concentrations of Ag NPs in soils, it was comparable to that in sewage sludge 46 . This Ag level was also needed for accurate measurement in the soil, root and shoot samples. Additionally, the BCR sequential extract ion method and soil pore water extraction were used to gain more insight in to the bioavailability of spiked Ag NPs and Ag + in radish - growing and radish - free soils. Materials and Methods Chemicals, Biochar and Soil Polyvinylpyrrolidone (0.6%) stabilized Ag NPs (1000 mg L 1 ) were obtained from the US Research Nanomaterials, Inc. (Texas, US). Silver nitrate (AgNO 3 ) was purchased from Fisher Scientific Inc. (Hampton, US). The morphology of Ag NPs was characterized by transmission electron microscopy (TEM, JEOL 2200FS, Japan). The size and surface charge were measured by dynamic light scattering using a Zetasizer (Nano - ZS, Malvern Instruments, UK ). TEM imaging confirmed that the Ag NPs were spherical (Supplementary Figure B1a in Appendix B) with a mean diameter average hydrodynamic diameter of 63.7 nm with a polydispersity index of 0.15. Aggregation 44 kinetics indicated that the Ag NPs were stable (Figure B1b) mainly due to steric repulsion induced by polyvinylpyrrolidone coated on the surface of Ag NPs 161 . Acetic acid, hydroxylammonium chloride, hydrogen peroxide, ammonium acetate, and nitric acid of analytical reagent grade were used. The biochar was produce d by Biogenic Reagents LLC. (Michigan, US) from pinewood via pyrolysis at 650 °C for about 30 min in a rotary reactor. As per the proximate analysis , the fixed carbon, ash , and volatile matter contents (by weight) were 90%, 5% and 4%, respectively. The ele mental content of biochar was 76.9% C, 0.2% N, and 1.2% H by weight. The s pecific s urface area and p ore volume were determined to be 543.6 m 2 g 1 and 0.3 cm 3 g 1 , respectively 162 . Its morphology was examined using scanning electron microscopy (SEM, JEOL 7500F, Japan) and exhibited a highly porous structure and rough surface (Supplementary Figure B2). The tested soil was collected at a - dried , and passed through a 2 - mm sieve before use. The soil was classified as a loamy sand cont aining 74.6% sand, 24.6% silt, and 0.78% clay as determined by a laser particle size analyzer (Malvern Mastersizer 2000E, UK). It had a pH of 7.8, OM of 2.3%, and cation exchange capacity (CEC) of 7.9 meq/100g . Biochar amendment (0.1% and 1.0%) had an insi gnificant influence on the soil pH (Supplementary Figure B3), OM, and CEC 162 . The background Ag concentration in soil was measured at 0.26 mg kg 1 . Radish Growth Experiment Radish plants were grown in soil pots spiked with Ag (Ag NPs or AgNO 3 ) and fifteen selected pharmaceuticals (acetaminophen, caffeine, carbadox, carbamazepine, estrone, lamotrigine, lincomycin, monensin, oxytetracycline, sulfadiazine, sulfamethoxazole, triclosan, - estradiol) to mimic the co - con tamination with pharmaceuticals, engineered nanoparticles, and/or heavy metals 163 . This paper only report s the study on Ag, with 45 the work involving pharmaceuticals reported in a companion paper 162 . The addition of pharmaceuticals did not change the bioma ss or Ag uptake of radish plants. The experimental design is summarized in Table 3.1 . To evenly distribute Ag in the soil, about 1 kg of soil was first spiked with Ag NPs or Ag + along with the fifteen pharmaceuticals in acetone and then thoroughly mixed in a fume hood 162, 164 . After solvent evaporat ing , the spiked soil portion and a predetermined amount of biochar were mixed with a predetermined amount of clean soil in a clean m otorized concrete mixer to homogenize the spiked Ag and pharmaceuticals into the soil. The resultant soil samples had 1 mg kg 1 of added Ag (Ag NPs or Ag + ) with 0.1 or 1 mg kg 1 of each pharmaceutical and contained 0%, 0.1% or 1.0% of the biochar by weight . The biochar - free and biochar - amended soils in the absence of Ag and pharmaceuticals were used as the blank c ontrol. 46 Table 3 .1 . Radish growth in soils in the absence (blank) and presence of spiked AgNO 3 and Ag NPs without (0%) or with (0.1% and 1%) biochar amendment*. Treatment Biochar Ag NPs Ag + 15 Pharmaceuticals Radish growth pots Radish - free pots blank 0.0% / / / 1 0 0.1% / / / 1 0 1.0% / / / 1 0 AgNO 3 0.0% 1 mg kg 0.1 mg kg 3 2 0.1% 1 mg kg 0.1 mg kg 3 2 1.0% 1 mg kg 0.1 mg kg 3 2 Ag NPs 0.0% 1 mg kg / 1 mg kg 3 2 0.1% 1 mg kg / 1 mg kg 3 2 1.0% 1 mg kg / 1 mg kg 3 2 *Note: In total 33 pots included 21 pots with the radish plants and 12 pots without the radish plants. The radish - free pots were designed to sample soils for pore water extraction. The radish pots had 2 radish seedlings. High concentration (1 mg kg ) of e ach pharmaceutical was co - added with Ag NPs, whereas low (0.1 mg kg ) pharmaceutical concentration was added with AgNO 3 . 47 The prepared soil samples were transferred to plastic pots (14.6 cm diameter × 10.8 cm height) at ~ 1 kg per pot. Five radish seeds (Burpee & Co., Warminster, PA) were directly sowed in each pot. After germination, plants were thinned to two seedlings per pot. Radish plants were grown in a climate - controlled greenhouse with natural sunlight at 25/21 °C day/night. Soil moisture level wa s maintained at 60% of the predetermined water - holding capacity by adding deionized (DI) water daily to compensate for evapotranspiration water loss. After 35 days, radish plants were sampled and washed with DI water to remove any attached soil particles. Two plants in each pot were divided into root and shoot samples, weighed, freeze - dried, and ground in to fine powders. All plant samples were kept in a desiccator until further use. Meanwhile, the soil samples in the radish - growing and radish - free pots were collected, and soil pore water was extracted immediately using a centrifugation - filtration method described previously 162, 164 . Briefly, aliquots (25 g) of the soil sample w ere placed on a layer of glass wool in a 20 - mL disposable plastic syringe. The syringe was inserted in to a 50 - mL polypropylene Corning tube, which was centrifuged at 4,000 × g for 40 min. The resulting pore water was stored at 4°C after passing through a 0.45 - µm membrane filte r, and the soil samples were freeze - dried for further Ag analysis. Ag Sorption Experiment Batch sorption experiments were conducted in duplicate to examine the sorption of Ag NPs and Ag + to the soils with different biochar amendment (0%, 0.1%, and 1.0%). S oil samples of 0.5 g were placed in 50 - mL polypropylene Corning tubes, followed by adding 25 mL of 5 mM KNO 3 containing a series of Ag NPs concentrations (0, 1.1, 2.2., 5.6, 11.1, 22.2, 55.6, 111, and 222 mg L 1 ) or AgNO 3 concentrations ( 0, 1, 2, 5, 10, 20, 50, 100, and 200 mg L 1 ), respectively . The resultant mixtures were sha ken for 24 h at room temperature in the dark, centrifug ed at 2000 × g 48 for 5 min and filt ered through 0.45 - µm membrane filters (Millipore, US). All filtrates were kept at 4 °C prior to analysis. BCR Sequential Extraction To examine the effect of biochar amendment on soil - bound Ag fractions in the soil samples and to further link the extractable fractions with the uptake of Ag, the well - established BCR method was used to extract Ag fro m the soil samples collected from the radish - growing and radish - free pots on Day 35 according to Wang et al. 165 . First, to measure the exchangeable fraction (f1) 1 g of soil was mixed with 20 mL of 0.1 M acetic acid in a shaker at 30 ± 5 rpm and 25 °C for 16 h. The extract was centrifuged at 3000 × g for 20 min, and the supernatant was collected. The residue solid was washed by shaking for 15 min with 20 mL of DI water, followed by centrifugation. Afterward, the Fe/Mn oxide - bound reducible fraction (f2) was extracted by adding 20 mL of 0.5 M hydroxyla mmonium chloride to the residue solids from the first step, and then the mixture was shaken at 30 ± 5 rpm and 25 °C for 16 h. The extract was again separated and the residue was washed as in the previous step. It is noted that Ag (assuming 1 mg kg 1 in the soil completely dissolved into the aqueous phase) would not precipitate with 0.5 M hydroxylammonium chloride as confirmed by the MINTQ calculation (Supplementary Figure B4). Third, in order to measure the OM - bound oxidisable fraction (f3) , 5 mL of 30% hyd rogen peroxide was added to the residue from the second step to oxidize OM. The mixture was digested for 1 h at 25 °C and for another hour at 85 °C, with occasional shaking. Then a second 5 mL aliquot of H 2 O 2 was added, followed by digestion for 1 h at 85 °C. Afterwards the residue was extracted with 25 mL of 1 M ammonium acetate solution (pH of 2.0 adjusted by HNO 3 ) by shaking at 30 ± 5 rpm and 25 °C for 16 h. The extract was collected and the residue was washed again. Finally, to measure the residual frac tion (f4) the soil residue from the third step was digested with 10 mL of HNO 3 49 (70%). All extraction experiments were conducted in triplicate for each sample. Because Ag NPs are not stable in the BCR extraction reagents and background Ag in soils would be also extracted, this study could not differentiate the Ag sources and species in the extracts. All Ag forms were measured as total Ag concentration using an atomic absorption spectrometer (AAS) after acid digestion. Analytical Methods Before analys i s, the filtrates from the sorption experiments, BCR extracts, soil pore water samples, and soil samples were digested with concentrated HNO 3 at 115 ºC 166 . The radish shoot and root samples were digested with HNO 3 and H 2 O 2 ( v:v = 4:1) at 115 ºC 143 . Afterwards, total Ag concentration in the digested samples w as determined using a flame AAS (AAnalyst 400, PerkinElmer, Waltham, MA). Due to the low Ag levels in the soil pore water samples and the radish root and shoot samples, the total Ag concentration in these samples w as measured by a graphite furnace AAS (PinAAcle 900T, PerkinElmer, Waltham, MA) after acid digestion. Data Analysis The sorbed Ag NPs or Ag + by the soil in the batch sorption experiments was calculated from the difference between the initial and final Ag concentrations using Eq. (3.1). (3.1) where is the amount of Ag sorbed to the soils (mg g 1 ), and are the i nitial and final equilibrium Ag concentrations in solution (mg L 1 ), respectively, is the solution volume (L), and is the mass of the soils (g). The Ag sorption isotherms were fitted with the Langmuir model in Eq. (3.2), using the nonlinear fitting f unction in MATLAB R2017b software (The MathWorks Inc., Natick, MA). (3.2) 50 where is the maximum Ag sorption capacity (mg g 1 ), and is the Langmuir constant related to the affinity of binding sites (L mg 1 ). Th e RCF and TF values were calculated using Eqs. (3.3) and (3.4), respectively. (3.3) (3.4) where , , and are the Ag concentrations in the radish root, shoot, and soil samples ( mg kg 1 , dry weight), respectively. One - way ANOVA analysis was performed to test the statistical significance in means among experimental treatments using the least significant differe nce (LSD) test with SPSS 22.0 software for Windows (IBM Corp., Armonk, NY) at p 0.05 level. Results and Discussion Radish Biomass In all three treatments (i.e., blank, Ag NPs, and AgNO 3 ) the absence or presence of biochar (i.e., 0%, 0.1%, and 1.0%) had no significant effect on the root and shoot biomass of the radish plants in each pot (two plants) ( p > 0.05, Figure 3.1). Intriguingly, the presence of 1 mg kg 1 Ag NPs and Ag + p 5), but had no significant effect on the radish shoots ( p > 0.05). It is likely that the trace Ag levels in soil pore water (below the detection limit of the graphite furnace AAS, 0.5 µg L 1 , as discussed more in detail in Sections below) enhance d root gro wth, but were not high enough to translocate upward to influence shoot growth. Indeed, a very recent study also observed stimulation of the root biomass of rice ( Oryza sativa L.) but no significant effect on shoot biomass when grow n in a hydroponic system 1 Ag NPs and Ag + for 14 days 146 . However , at higher exposure levels dose - dependent phytotoxicity of Ag NPs and Ag + has been observed to negatively influence 51 seed germination, root elongation, and/or biomass growth 71, 145, 147, 148, 158, 167 . To date no consensus has been reached regarding the phytotoxicity of Ag NPs and/or Ag + , which is mainly attributed to the differences in exposure scenarios, physicochemical properties of nanoparticles, and plant species in various experimental settings. T he increased root biomass was unlikely due to the presence of pharmaceuticals, as the level of pharmaceuticals differed by 10 fold between the Ag NPs (co - added with 1 mg kg 1 of each pharmaceutical) and AgNO 3 (co - added with 0.1 mg kg 1 of each pharmaceutic al) treatments (Table 3.1), but there was no significant difference between the Ag NPs and AgNO 3 treatments for either the radish root or shoot biomass. 52 Figure 3.1. Dry biomass of radish roots (A) and shoots (B) harvested on Day 35 in each pot (two plants) in the absence (blank) and presence of spiked AgNO 3 and Ag NPs without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations ( n = 3) . Different letters indicate the significant difference in means at p < 0.05. Brackets indicate comparisons between the AgNO 3 and Ag NPs treatments as well as between the treatment groups with or without Ag addition, respectively. 53 Uptake of Ag by Radish There were no significant differences in total Ag concentration in the radish roots and shoots for all treatments as a result of add ing biochar, Ag NPs or AgNO 3 (Figure 3.2). Specifically, t he biochar amendment (0.1% and 1.0%) had no significant effect on Ag uptake by the radish plants ( p > 0.05) . This result contradict s the study of Abbas et al. 71 who report ed that biochar addition (2% w/v) decreased the uptake of Ag in rice ( Oryza sativa L.) in a hydroponic system containing a 0 mg L 1 , which was at least 5 order s of magnitude greater than the Ag concentrations in the soil pore water (< 0.5 µg L 1 for the total Ag concentration of 1 mg kg 1 in the soil) in this study. Furthermore, the addition of 1 mg kg 1 Ag ( Ag NPs or AgNO 3 ) did not substantially increase the uptake or accumulation of Ag in radish roots (0.05 ± 0.02 1.06 ± 0.98 mg kg 1 ) and shoots (0.01 ± 0.00 0.03 ± 0.01 mg kg 1 ) , compared to the blank ( 0.03 ± 0.00 0.49 ± 0.05 mg kg 1 for the roots, and 0.01 ± 0.00 0.02 ± 0.01 mg kg 1 for the shoots at a background Ag of 0.26 mg kg 1 ) ( p > 0.05). As such, the insignificant effect of the various biochar amendment rates on Ag uptake likely resulted from the low level of Ag (1 mg kg 1 ) in the soil, most of which was not readily bioavailable to the radish plants (discussed in more detail in Sections below). Since the Ag concentration of 1 mg kg 1 used in this study was similar to the measured background Ag concentration (0.26 mg kg 1 ), but about 4 order s of magnitude grea ter than the predicted Ag concentration in soils due to the release of engineered Ag NPs 47 , the Ag NPs released into soils are unlikely to result in a realistic increase of Ag in food crops, indicating a low food safety risk from the released Ag NPs. 54 Figure 3.2. Total Ag concentrations in the roots (A) and shoots (B) of radish plants grown in soil pots in the absence (blank) and presence of spiked AgNO 3 and Ag NPs without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations ( n = 3) . Different letters indicate the significant difference in means at p < 0.05. Brackets indicate the comparisons between the AgNO 3 and Ag NPs treatments as well as between the treatment groups with or without silver addition, respectively. 55 Compared to the shoots, the radish roots had notably greater concentrations of Ag in all treatments (Figure 3.2). The calculated RCF values varied from 0.03 ± 0.03 to 0.29 ± 0.21, suggesting low root uptake of Ag NPs and Ag + ions. The calculated TF values ranged from 0.08 ± 0.10 to 0.89 ± 1.21 (Figure 3.3), suggesting low translocation of Ag NPs and Ag + ions from the radish roots to shoots. Similarly, Ag was predominantly retained in roots with minor or even no translocation to shoots in previous studies 51, 55 . No significant difference was observed in the RCF and TF values between the AgNO 3 and Ag NPs treatments at all biochar amendment rates (Figure 3.3). This is in agreement with previous studies on the uptake of Ag NPs an d Ag + by tobacco ( Nicotiana tabacum ) 51 and by nine edible plants 144 , i.e., the uptake and accumulation of Ag NPs and Ag + were equally efficient. However, Rawat et al. 168 reported that the TF value of Cu 2+ was greater than that of CuO NPs in bell pepper plants, and Yang et al. 146 found that the TF value of Ag + was significantly lower than that of Ag NPs in rice plants. In this study , the similar RCF and TF values of Ag NPs and Ag + in the radish plants were likely due to their low and comparable concentra tions in the soil pore water, limit ing bioavailability to the radish plants. 56 Figure 3.3. Root concentration factor (A) and translocation factor (B) of Ag in either AgNO 3 or Ag NPs in radish plants grown in soil pots without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations ( n = 3). Different letters indicate the significant difference in means at p < 0.05. Brackets indicate comparisons between the AgNO 3 and Ag NPs treatments. 57 Ag Sorption to Soils Figure 3.4 presents the sorption isotherms of Ag NPs and Ag + ions by the soil samples without (0%) and with (0.1% and 1.0%) biochar amendment. As summarized in Supplementary Table B1, the Langmuir model fitted the experimental data well with R 2 > 0.95. Biochar amendment (0.1% and 1.0%) had an insignificant influenc e on Ag sorption by the soils (Fig ure 3. 4). The soil samples had notabl y higher sorption capacities for Ag + than for Ag NPs (Figure 3.4). Likewise, Wang et al. 169 found that the sorption of Ag + to ten soils was greater than that for Ag NPs. This is becaus e negatively charged soil particles attract Ag + ions but repel negatively charged Ag NPs. Ag NPs in soils (pH of 7.8) would be more negatively charged than the measured zeta .0 49 and its surface charge become s more negative with increasing pH. In fact, s orption of Ag + ions correlated with the content of negatively charged soil OM and clay, and sorption of Ag NPs with the level of positively charged iron/aluminum oxide at circ umneutral pH 161, 169 - 172 . Interestingly, for soil with 1.0% biochar the values were similar for Ag NPs and Ag + , but the value for Ag NPs was much lower than for Ag + . Thus, in that soil sample Ag NPs were more likely to attach to the biochar surface via low - affinity interactions (such as weak attachment due to the second energy minima) 173 . Furthermore, it was reported that metal ions may be reduced after sorption to biochar 72, 174 - 177 . However, this study was unable to explore the transformation between Ag + and Ag 0 , i.e., dissolution of Ag NPs to Ag + ions and reduction of Ag + to Ag 0 by electron - donating moieties (e.g., phenolic moieties) in biochar 175 - 177 . The Ag tr ansformation should be explored in future studies by employing Fourier transformed infrared spectroscopy (FTIR), X - ray photoelectron spectroscopy (XPS), or X - ray absorption fine structure spectroscopy (XANES). Finally, despite the greater sorption capacity of Ag + than Ag NPs to soils with and without biochar, Ag + and Ag 58 NPs uptake was not significantly different (Figure 3.2 and Figure 3.3), which may be explained as follows. Ag sorption capacity is primarily dictated at the high end of Ag concentrations , wh ereas no substantial difference was observed at low Ag concentration range (Figure 3.4), which is more relevant to the spiked Ag concentration (1 mg kg 1 ) in soils. In combination with the low Ag concentrations in soil pore water and the low exchangeable A g fraction, no difference in the uptake of Ag + and Ag NPs is therefore expected. 59 Figure 3.4. Measured (symbols) and fitted (lines) sorption isotherms of Ag in either AgNO 3 or Ag NPs by the soil samples without (0%) or with (0.1% and 1%) biochar amendment. Error bars are the standard deviations from two replicates. 60 Soil - bound Ag Fractions As sho wn in Figure 3.5, Supplementary Table B2 , and Table B3, the highest proportion of followed by the OM - - bound reduci ble the large st proportion of spiked Ag (> 61.9% for Ag NPs and > 58.3% for Ag + ions) was in the reducible fraction, followed by the residual fraction (13.2% e fraction (5.1% 23.8%) > exchangeable fraction (0 0.09%), except for radish - growing soil with 1.0% biochar spiked with AgNO 3 and the radish - free soil without biochar spiked with Ag NPs. In those two soil samples the residual fraction was the greatest for Ag + (45.6%) in the former, and similar to the residual fraction for Ag NPs (41.8%) in the latter. It is plausible that the sp iked Ag + ions and Ag NPs were associated with the surface of soil Fe/Al oxides and clays 169, 170 , which could be readily extracted by hydroxylammonium chloride at pH 2 as the reducible fraction. Our results agree with several earlier studies on other meta llic ions and nanoparticles. For example, the majority of Pb was associated with Fe/Mn oxides in soils collected from Guizhou, China 178, 179 . Zhang et al. 151 found that the exchangeable fraction of cerium oxide nanoparticles was the lowest and the residu al fraction was dominant in both loamy sand and silty loam soils. Nonetheless, Choleva et al. 157 showed that both Au NPs and ions were predominantly associated with soil OM (the oxidisable fraction) and the reducible fraction bound to metal oxides was min or (<10%). Compared to the biochar - free treatment , biochar amendment (1.0%) significantly decreased the reducible fraction and increased the residual fraction of Ag + ions, but only increased the oxidisable fraction of Ag NPs ( p < 0.05, Figure 3.5, and Supp lementary Table B2) in soils with radish plants . Conversely, without radish plants biochar amendment (1.0%) decreased the 61 reducible fraction of Ag + and the residual fraction of Ag NPs (Figure 3.5 and Supplementary Table B3). It appears that radish plants m odulated the changes in the soil - bound fractions of Ag + and Ag NPs induced by biochar amendment (1.0%). Biochar addition may increase negatively charged surface sites in soils and release dissolved OM into soil water 180 - 182 , which may scavenge Ag + from Fe/Mn oxides and increase repulsion between Ag NPs and silicate minerals, leading to decreased reducible and residual fractions of Ag + and Ag NPs, respectively. Furthermore, radish root exudates (including organic acids, carbohydrates, and root mucil age) may also facilitate the release of Ag + from Fe/Mn oxides via complexation with Ag + , or the release of Ag NPs from the residual fraction in the rhizosphere soil 145, 147 . 62 Figure 3.5. Soil - bound fractions of Ag in either AgNO 3 or Ag NPs in the soil samples without (0%) or with (0.1% and 1%) biochar amendment measured by the BCR extraction method. 63 Finally, the exchangeable fraction of Ag was very low for all treatments (Figure 3.5, Supplementary Table B2, and Table B3). The Ag concentration for all extracted pore water 1 ). This low Ag concentration in soil pore water was expected, as Ag NPs and Ag + ions can be transformed to Ag 2 S 152 , which has very lower solubility ( K sp = 5.9 × 10 51 ) in water. Ag was still detected in the radish plants , especially in the roots (Figure 3.2A), suggesting that the radish plants could take up the trace level of Ag in soil pore water. It is thought that f ast exchange reactions may occur betwe en the reactive (e.g., reducible and oxidisable) and available (i.e., exchangeable) pools 183 , thus both the reactive and available fractions are potentially bioavailable to plants 151, 155, 156 . T he combined reducible and oxidisable as potentially bioavailable to the radish plants over time. T he release of plant root exudates might also slowly release Ag from the reducible and oxidisable fractions into the ex changeable fraction, and thus continuously supply a trace level of Ag in soil pore water for root uptake. Since the soil - bound fractions of Ag measured by the BCR method could only represent the maximum potential bioavailability, correlation of Ag uptake a nd accumulation with specific fractions may be difficult to achieve, especially at environmentally - relevant concentrations 184, 185 . Conclusion This study has several important findings that are relevant to assessing the risks of Ag NPs to food safety, and the effect of biochar amendment on Ag fractions and plant uptake of Ag NPs and Ag + ions in soils. Biochar amendment significantly changed the soil - bound fractions of Ag NPs and Ag + ions, as modulated by the radish plants. In any case, the readi ly bioavailable fraction (i.e., the exchangeable fraction) was very low. Thus, plant uptake of Ag at environmentally relevant concentrations is predominantly controlled by low uptake from soil pore water and a 64 continuous supply of Ag from the reactive frac tions. Additionally, at the low Ag level the biochar amendment primarily changed the reducible, oxidisable and residual fractions, suggesting a long - term effect on mass transfer between various soil - bound Ag fractions and subsequent uptake of Ag by plants, which may be different from results at high Ag levels. In contrast to previous studies using unrealistic high concentrations of Ag NPs 52, 54 - 57 , this short - term study (35 days) showed that Ag uptake and accumulation in food crops will be low at environme ntally - relevant concentrations. This study was limited in that only one Ag concentration (1 mg kg 1 ), one crop (radish) and one soil type were used, and molecular Ag species were not differentiated. Future work should use other advanced analytical techniqu es (e.g., FTIR, XPS, XANES, and single particle inductively coupled plasma mass spectrometry) to spe ciate the bioavailable fractions of Ag NPs and Ag + in diverse soil, water , and plant systems. Author Contribution: Jianzhou He, Yuanbo Li, and Wei Zhang conceived and designed the research experiments; Jianzhou He, Yuanbo Li, and Haonan Qi conducted the experiments; Jianzhou He and Haonan Qi analyzed the samples for Ag; Jianzhou He analyzed the data and wrote the manuscript; Hui L and We i Zhang reviewed and edited the manuscript. 65 STOMATA FACILITATE FOLIAR SORPTION OF SILVER NANOPARTICLES BY ARABIDOPSIS THALIANA 66 Abstract Application of nanopesticides may substantially increase the uptake and accumulation of engineered nanomaterials (ENMs) in food crops. However, mechanis ms for internalization of ENMs into plant leaves are unclear . This study investigated the role of stomata in the internalization of s ilver nanoparticles (Ag NPs) using abscisic acid (ABA) - responsive ecotypes (Landsberg erecta [Ler] and Col - 7) and ABA - insensitive mutants ( ost1 - 2 and scord7 ) of Arabidopsis thaliana in batch sorption experiments, in combination with microscopic visualizati on . Compar ed with those of the ABA - free control, stomatal apertures were significantly lower for the Ler and Col - 7 ecotypes ( p the ost1 - 2 and scord7 mutants, after exposure to 10 - µM ABA for 1 h. S orption kinetics showed t hat the m aximum sorption of Ag NPs was achieved after 60 min of equilibration. The amount of Ag NPs sorbed by the leaves of Ler and Col - 7 ecotypes treated with 10 µM ABA w as much lower than that in the ABA - free control, mainly due to ABA - induced stomatal c losure. In contrast there was no significant difference in foliar sorption of Ag NPs by the ost1 - 2 and scord7 mutants with and without ABA. Ag NPs were widely attached to the Arabidopsis leaf surface, and localized at the cell membrane, cytoplasm and plasm odesmata, as revealed by scanning electron microscopy and transmission electron microscopy, respectively. These results highlight the important role of stomata in the internationalization of ENMs in plants and may have broader implications in foliar applic ation of nanopesticides and minimizing contamination of food crops by ENMs. 67 Introduction Nanotechnology has been strongly advocated for improving agricultural productivity and sustainability through optimization of agrochemical applications (e.g., nanofe rtilizers , nanopesticides , and controlled release) , delivery of genetic materials, and intelligent surveillance of plant disease 186 - 188 . Specifically, the development of nanopesticides ha s drawn immense attention from scientific communities and agr o chemical companies. For example, due to its antimicrobial and insecticidal properties 189 , silver nanoparticles (Ag NPs) have been used in > 100 pesticides registered in the last decade 75 - 77 . The rapidly growing use of nano - enabled products in agriculture will exacerbate the release of engineered nano material s ( ENMs ) into soil and water systems 42, 47, 190 , which is estimated to be much higher than the combined release from all other sources 76 . Th us , there is an increasing concern over the risks of nanope sticides (such as Ag NPs) used on food crops. It is important to study the transfer and internalization of ENMs to food crops due to potential human exposure and food safety risks . Crop plants are prone to elevated exposure of ENMs from plant protection an d fertili zer products, crop irrigation with reclaimed water, biosolid application, and atmospheric deposition 48, 76 , leading to the uptake and accumulation of ENMs in plants 48, 49, 65, 191, 192 . As a result, the human population may be exposed to ENMs vi a consumption of food crops 65, 192 - 194 . Root uptake of ENMs has been well studied 50, 52, 57, 64, 66, 67, 146, 148, 195, 196 , with ENMs taken up via the apoplastic and symplastic pathways and transported to aerial portion s through the vascular system 65 . Moreover, the size exclusion limit of the root membrane 62 - 66 may be overcome by endocytosis 62, 68, 69 and additional entry points coming from physical and disease damage to roots as well as emergence of new secondary roots 64, 66 . 68 In contrast , studies on foliar uptake of ENMs have been limited 49, 82, 197, 198 . It is thought that ENMs may first attach to the leaf surface and then penetrate the waxy cuticle and/or stomata on the epidermis 66, 199 . However, most ENMs are not small enough to pass through the small pores (< 5 nm) in the cuticle 200 . Rather they may enter through stomata where the opening size is in micrometers 49, 199, 201 - 203 . Previous studies have speculated that plant stomata could serve as points of entry for ENMs 82, 1 97 . For example, Ce was detected in the leaf, flower, stem, and root of cucumber ( Cucumis sativus ) a fter foliar exposure to aerosol and aquasol of cerium dioxide nanoparticles ( CeO 2 NPs ) , indicating uptake through the leaves and further translocation to other plant parts 204 . Larue et al. 49, 198 conjectured that Ag NPs and TiO 2 nanoparticles may be internalized into lettuce leaves via stomata after foliar exposure. However, Birbaum et al . 205 exposed maize leaves to CeO 2 NP s aerosol or aquasol , and found that the sorbed amount ( Ce g 1 leave ) was independent of stomata l opening or closure . Although previous researchers have hypothesized the potential uptake of ENMs through stomatal openings, no study has directly shown the differential internalization of ENMs by plant leaves when stomata are manipulated under otherwise similar experimental conditions. For this purpose, regulation of stomatal closure by the plant hor mone abscisic acid (ABA) and related signal transductions 206 may be harnessed to elucidate the role of stomata in the internalization of ENMs using ABA - responsive plant ecotypes and ABA - insensitive mutants . The ABA - insensitive plant mutants are best chara cterized in Arabidopsis thaliana 207, 208 . It was shown that the ost1 - 2 and scord7 mutants of Arabidopsis do not respond to ABA - induced stomatal closure and exhibit constitutively larger apertures 206, 209 . Also Arabidopsis thaliana was frequently chosen a s the model plant in plant biology and nanotoxicity studies 65, 148, 206, 209 - 211 because findings from Arabidopsis are generally applicable to other plant species 68 . 69 Therefore, this study aimed to investigate the role of stomata in the sorption (includin g surface attachment and internalization) of Ag NPs to plant leaves using the model plant Arabidopsis thaliana . The ABA - responsive ecotypes of Arabidopsis thaliana (i.e., Landsberg erecta [Ler ] and Col - 7) and their corresponding ABA - insensitive mutants (i.e., ost1 - 2 and scord7 for Ler and Col - 7, respectively) were used to assess the role of stomata in the internalization of Ag NPs through batch sorption experiments combined with confocal laser scanning microscopy (CLSM), scanning electron microscopy (SEM) with energy dispersive spectrometer (EDS), and transmission electron microscopy (TEM). We hypothesized that the sor ption of Ag NPs with ABA treatment would be lower than that without ABA for th e Ler and Col - 7 ecotypes, but similar for the ost1 - 2 and scord7 mutants. This study provided direct evidence on the internalization of Ag NPs through stomatal entry. Materials and Methods Plant Materials Seeds of the Ler, Col - 7, ost1 - 2 , and scord7 were sterilized in 70% ethanol for 10 min and in 50% bleach for 5 min, respectively, washed three times with autoclaved deionized (DI) water, and then dispersed into 0.2% agar before sowing. The seeds were directly sown in sterilized Arabidopsis - growth mix packed in nursery pots (equal parts of Suremix, medium vermiculate and perlite) 212 , placed in a tray, and grown in a growth chamber that was well - maintained at 22 ºC with a 12/12 h photoperiod under a light intensity of 100 µE/m 2 /s . The plants were watered three times a week with one - half strength Hoagland solution to maintain a water height of 1 cm above the bottom of the tray. F ully expanded leaves from 5 - week - old plants were used f or all experiments . 70 Ag NPs Suspension A stock sus pension of Ag NPs (1000 mg/L) stabilized with polyvinylpyrrolidone (PVP , 0.6% in solution ) was purchased from the US Research Nanomaterials, Inc. (Texas, US). The stock solution was filtered with Amicon Ultra - 10kDa filters (Millipore, US) to remove free PV P from the solution. The retained Ag NPs were then washed repeatedly and dispersed in DI water to obtain the working suspension (Supplementary Text C1 in Appendix C). The working suspension of Ag NPs was ultrasonicated (Fisher Scientific, FS20) for 5 min b efore use. The morphology and size of Ag NPs was characterized by transmission electron microscopy (TEM, JEOL 2200FS, Japan), and its size and zeta ( ) potential by a Zetasizer Nano - ZS ( Malvern Instruments, UK ). Detailed information on sample preparation and characterization can be found in Supplementary Text C2. Stomata Assays Leaf discs (~ 3 mm × 3 mm) were cut from the leaves of the 5 - week - old mature plants with a scalpel and immersed in lidded 6 - well plates containing opening buffer (25 mM MES and 10 mM KCl, pH 6.15) without (the control treatment) or with 10 µM ABA (the 10 µM ABA treatment). To preserve the stomatal aperture status in Arabidopsis plants, the leaves were not further treat ed with any other st omatal opening buffer. The plates were cultured in the growth chamber for 1 h before observation with CLSM . CLSM images were taken at an excitation wavelength of 488 nm and emission wavelengths of 505 - 525 nm in a field of view of 318.2 µm. Briefly, the dis cs were placed on glass microscope slides, and then scanned to a depth of ~10 µm using an Olympus FluoView 1000 CLSM at three random locations to generate Z - scans. The Z - scans were stacked to produce images of the leaf surface for stomata analysis. Three d iscs from each treatment were used and three locations per disc were imaged by CLSM. The width of the stomatal aperture was measured using software ImageJ from 36 63 stomata in each treatment , and 71 the width - to - length ratio of the stomatal aperture was simi larly measured on 36 84 stomata per treatment . Sorption Experiments Batch experiments were conducted to investigate the sorption of Ag NPs by whole Arabidopsis thaliana leaves treated with the opening buffer without (the control treatment) or with 10 µM AB A (the 10 µM ABA treatment). First, the kinetic sorption experiment was performed to estimate the time to reach sorption equilibration. Briefly, about 0.4 5 g of whole scord7 leaves from the control treatment were immersed in an Ag NP suspension of 2 mg L 1 (pH 6.5) up to 12 0 min. For the equilibrium sorption experiments, about 0.45 g of whole leaves (i.e., Ler, Col - 7, ost1 - 2 and scord7 ) were placed in 60 - mL glass jars each containing 25 mL of Ag NPs solution at three concentrations of 1, 5, and 10 mg L 1 . T he blank experiment was performed using DI water without Ag NPs. The mixture of leaves and Ag NPs w as equilibrated at room temperature in a horizontal shaker (160 rpm) for predetermined 60 min. Afterwards, the leaves were separated from the solution using a tweezer. The final concentration of Ag NPs in the remaining suspension was determined using an atomic absorption spectroscope (AAS, AAnalyst 400, PerkinElmer) after nitric acid digestion. Data Analysis The sorbed concentration of Ag NPs to the leaves was calculated by the difference between the initial and final concentrations of Ag NPs using Eq. (4.1) : (4.1) where is the sorbed concentration of Ag NPs to the leaves (mg kg 1 ), and are the initial and equilibrium concentrations of Ag NPs in the suspension (mg L 1 ), respectively, is the 72 volume of the suspension (mL), and is the fresh weight of leaves (g). All sorption experiments were conducted in at least du plicate. One - way AN OVA analysis was performed to identify statistically significant differences in measured parameters using the least significant difference (LSD) test. All statistical analyses were conducted using SPSS 22.0 software for Windows (IBM Corp., Armonk, NY), wit h the differences of means considered significant at p . Visualization of Sorbed Ag NPs To determine the localization of Ag NPs in Arabidopsis thaliana leaves, Col - 7 and scord7 leaves were prepared for microscopic observation after exposure to 10 mg L 1 of Ag NPs. For SEM the leaves were freeze - dried (Electron Microscopy Sciences, Model EMS750X, Hatfield, PA) and mounted on aluminum stubs using High vacuum carbon tabs (SPI Supplies, West Chester, PA). The samples were coated with ~5 nm - thick osmium using an NEOC - AT osmium chemical vapor deposition coater (Meiwafosis Co., Ltd., Osaka, Japan) before examination with a JEOL 7500F SEM (JEOL Ltd., Tokyo, Japan) equipped with energy dispersive X - ray spectroscopy (Oxford Instruments, High Wycomb, Bucks, Eng land). TEM s ample preparation followed a standard procedure as previously described 143, 146 . Briefly , the leaves were cut into small pieces , which were then prefixed in 2.5% glutaraldehyde and 2% paraformaldehyde for 24 h, washed three times (20 min/each) with 0.1 M cacodylate buffer (pH 7.4), and post - fixed in 1% osmium tetroxide for 2 h. After again rinsing three times (20 min/each) in 0.1 M cacodylate buffer (pH 7.4), the specimens were dehydrated in an acetone series (30%, 50%, 70%,80%, 90%, 95%, and 1 00%), with each step maintained for 30 min. Afterwards, specimens were sequentially infiltrated in different mixtures of acetone and Spurr resin (ratio = 3:1, 2:2, 1:3, and 0:1) and finally embedded in silicone molds. The polymerization was performed 73 at 60 °C for 48 h. The prepared specimens in Spurr resin were sectioned (70 nm thick ness ) using a PowerTome XL ultramicrotome (RMC Boeckeler, US) equipped with a diamond knife. The ultra - thin sections were placed on 200 - mesh copper grids and then imaged with a JEOL 1400 TEM at an accelerating voltage of 100 kV. Our preliminary tests showed that sample staining with uranyl acetate and lead citrate introduced numerous black spots that interfered with the identification of Ag NPs. Therefore, the specimen was not st ained. Results and Discussion Ag NPs Characterization As shown in Figure 4.1, the primary particles of Ag NPs were near - spherical in shape with an average diameter hydrodynamic diameter ( D h ) of 36.1 nm for the Ag NP aggregates, and a polydispersity index of 0.24. The D h value of the Ag NP aggregates was about twice that of the primary particl e size measured by the TEM, probably due to slight aggregation of Ag NPs in suspension. The time - resolved D h measurements indicated that Ag NPs were stable at pH 6.5 in DI water over 3 h. 74 Figure 4.1. Representative TEM image of Ag NPs (A), particle siz e distribution of Ag NPs measured from the TEM images (B), aggregation kinetics of 10 mg L 1 Ag NPs suspension at pH 6.5 in DI water (C), and summarized TEM primary particle size, hydrodynamic diameter ( D h ) and - potential of Ag NPs (D). 75 Stomatal Apertur e Assay The s tomatal aperture s for the different genotypes of Arabidopsis thaliana are presented in Figure 4.2 , Supplementary Table C1, Figure C1 , and Figure C2 . As shown in the CLSM images (Figure C1), ABA (10 µM) induced normal stomata closure for the Ler and Col - 7 ecotypes , but the stomata for the ABA - insensitive mutants ( ost1 - 2 and scord7 ) failed to close . This observation was expected as the stomata of the ecotypes would respond to the regulation of ABA, but those of the mutants would be insensitive to ABA 209 . T he width of the stomatal aperture measured using ImageJ decreased from 4.03 ± 0.72 µm and 4.45 ± 0.66 µm in the control treatment to 2.62 ± 0.89 µm and 2.58 ± 0.85 µm in the presence of 10 µM ABA for the Ler and Col - 7 ecotypes, respectively ( p , Figure 4.2 and Table C1 ) . However, the stomatal width of the ost1 - 2 and scord7 mutants remained statistically unchanged after ABA exposure . It is worth noting that the exposure to 10 µM ABA did no t fully close the stomata of the Ler and Col - 7 ecotypes (Figure 4.2, Supplementary Table C1, Figure C1, and Figure C2), which is consistent with previous observations on partial stomatal closure of Col - 0 and Col - 7 in response to ABA 206, 209 . The measured stomatal width - to - length ratios similarly demonstrated the lower stomatal aperture in the presence of 10 µM ABA than that without ABA for the Ler and Col - 7 ecotypes, but the non - response of the ost1 - 2 and scord7 mutants to the ABA treatment (Supplementary Table C1 and Figure C2). 76 Figure 4.2. Stomatal aperture width of Arabidopsis thaliana treated with opening buffer in the absence (control) or presence of 10 µM abscisic acid (ABA). Ler and Col - 7 are ecotypes, and ost 1 - 2 and scord7 are corresponding ABA - insensitive mutants. Results are shown as mean ± standard deviation ( n = 36 63 stomata). Symbol (*) indicates a statistically significant difference in means between the control and 10 µM ABA treatments ( p 77 Ki netics and Equilibrium Sorption of Ag NPs As shown in Supplementary Figure C3, sorption of Ag NPs to the scord7 leaves increased rapidly at the beginning and then reached the maximum by about 60 min. This was much shorter than the equilibrium time (14 h) used in the previous sorption experiment of Ag NPs by the leaf discs (diameter of 4 mm) of ficus tree ( Ficus ben jamina ) 213 . The equilibrium sorption amounts of Ag NPs to Arabidopsis leaves were shown in Figure 4. 3 . As expected, the treatment of 10 µM ABA for 1 h significantly decreased the sorbed amount of Ag NPs by the Ler and Col - 7 ecotypes at all exposure concen trations ( p < 0.05, Figure 4. 3 a and c) except for Col - 7 at 10 mg L 1 of Ag NPs (Figure 4. 3 c) , but did not change the sorption of Ag NPs by the ost1 - 2 and scord7 mutants (Figure 4 .3 b and d). Compared to the control treatments, the mean sorbed Ag NPs decreas ed by 85.8%, 23.0%, and 62.8% for the Ler and 14.2%, 21.3% and 16.9% (statistically non - significant) for the Col - 7 at 1, 5, and 10 mg L 1 Ag NP in the presence of 10 µM ABA, respectively. This is mainly due to the different stomatal responses to ABA between the Arabidopsis ecotypes and mutants 207, 208 . T he presence of ABA (10 µM) triggered stomatal closure in the Ler and Col - 7 ecotypes and led to smaller stomatal apertures than those of th e control treatments (Figure 4.2), resulting in decreased sorption of Ag NPs (Figure 4. 3 ). In contrast, the stomata of ost1 - 2 and scord7 remained open in the presence of ABA (Figure 4.2) 206, 209 , result ing in no difference in the amount of sorbed Ag NPs b etween treatments with and without ABA . It is unclear why the extent of decrease in the sorption of Ag NPs upon exposure to ABA was less for the Col - 7 than for the Ler. Nonetheless, there was no statistical difference between the control and ABA treatments for the Col - 7 at 10 mg L 1 Ag NPs (Figure 4.3c), which may need more replications to determine any statistical difference. 78 Figure 4. 3 . Sorbed amount of Ag NPs at various concentrations of 1, 5, and 10 mg L 1 by the leaves of Arabidopsis thaliana Ler (A) , ost1 - 2 (B) , Col - 7 (C) , and scord7 (D) treated in opening buffer without (control) or with abscisic acid (10 µM ABA) for 1 h. Error bars repres ent the standard deviations ( n = 3). Symbol (*) indicates statistically significant difference in means between the control and 10 µM ABA treatment ( p 79 The reduction of sorbed Ag NPs correlated with the change in stomatal aperture induced by the presence of 10 µM ABA (Figure 4.4). The data points were distributed along the 1:1 line, indicating that the decreased amount of Ag NPs sorbed by Arabidopsis leaves was directly due to the decreased stomatal aperture (Figure 4.2). Again, decreased Ag NPs s orption from stomata closure was more pronounced for the Ler than the Col - 7 (Figure 4.4). By using the ABA - responsible Arabidopsis ecotypes and ABA - insensitive mutants this study provides compelling evidence that stomata facilitate the internalization of A g NPs into plant leaves, which supports previous findings on Ag NPs and other ENMs (e.g., CeO 2 , TiO 2 , and Au) 49, 68, 143, 205, 214 . For example, a recent study examining the translocation of CeO 2 NPs applied to cucumber leaves suggested that small CeO 2 NPs can penetrate the leaf through hydathodes and stomata 204 . It is worth mentioning that after penetrating the wax cuticle and leaf epidermis , ENMs can eas ily access the downward phloem vascular system and be transported to other parts of the plant 66, 2 03, 215, 216 . Thus, stomata - facilitated internalization of ENMs could be an important route for plant uptake of ENMs. 80 Figure 4.4 . Relation between the reduction percentages of the sorbed Ag NPs and the change of stomatal aperture induced by the presenc e of 10 µM ABA . For each Arabidopsis genotype, the sorbed amount reduction (%) was calculated for 1, 5, and 10 mg L 1 of the Ag NPs exposure . 81 Visualization of Sorbed Ag NPs S urface - attached and internalized Ag NPs were characterized for the leaves of Arabidopsis Col - 7 ecotype and the scord7 mutant by SEM - EDS and TEM , as shown in Figure 4.5, and Supplementary Figure s C 4, C5, and C6 in Appendix C. No silver was detected in the SEM (Figure C 4 a and d) for the blank (i.e., no Ag NPs exposure). However, bright dots were clearly observed on the surface of Col - 7 leaves in the control and 10 µM ABA treatments (Supplementary Figure C 4 b and c), which was verified as elemental Ag by the EDS spectra (Supplementary Figure C 4 e and f). Ag was di stributed randomly and widely on the Col - 7 leaf surface (Supplementary Figure C 5 ). It is interesting to note that more Ag NPs were found on the leaf surface in the presence of 10 µM ABA (Supplementary Figure C4c), likely because the closure of stomata decr eased the internalization of Ag NPs and consequently increased the amount of Ag NPs available for surface attachment. It is reported that surface roughness facilitates the deposition of ENMs 217, 218 . However, since the SEM - imaged areas were near stomata a nd other surface structures were not well resolved, the distribution pattern of attached Ag NPs could not be discerned on certain structural features of the Arabidopsis leaves, such as grooves and veins with high roughness 219 . Ag NPs were localized subcel lularly in the Col - 7 and scord7 leaves for the control and 10 µM ABA treatments (Figure 4.5, Supplementary Figure C6 and Figure C7) . The TEM images of chloroplast appeared to have black spots (such as plastoglobuli ) found in all treatments (Figure C6), whi ch obscured any possible identification of Ag NPs. Thus, we focused on other subcellular locations. As shown in Figure 4.5, for the scord7 leaves, Ag NPs internalized through the stomatal openings were found in the cytoplasm ( Figure 4.5 c, d and e), attached to the cell membrane (Figure 4.5f ) , and near or entrapped within the plasmodesma (Figure 4.5c) . Similarly, Ag NPs were also found at the cell membrane, and near or within the plasmodesma in the Col - 7 leaves in the c ontrol 82 treatment (Figure C7) . Our observation is consistent with earlier studies documenting that Ag NPs were detected in the parenchyma cells of lettuce leaves 49 and associated with the cell walls in soybean and rice leaves 143 after foliar exposure. The role of p lasmodesma as a co nduit for inter - cellular transfer (Figure 4.5 c and Figure C ) has previously been reported. Indeed, gold nanoparticles (15 nm) and Ag NPs (20 nm) were accumulated in the plasmodesmata of the roots of woody poplar 220 and Arabidopsis thaliana 148 , respec tively. Clearly, plasmodesma is sufficiently large (20 50 nm) 221 to allow passage of Ag NPs measuring 20.9 nm in diameter from one cell to another , leading to entry into the symplastic pathway. In addition to these subcellular locations discussed above, p olystyrene particles were found to accumulate in the guard cells of Vicia faba leaves 82 . Kurepa et al. 68 further revealed that TiO 2 Alizarin red S nanoconjugates were enclosed in the vacuoles of stomatal guard cells in Arabidopsis thaliana . Finally , the internalized Ag NPs existed as single particles or aggregate clusters in the Arabidopsis leaves (Figure 4.5 and Figure C 7 ). Nonetheless, our work was conducted in vitro in a relatively short time frame (1 h sorption experiment ) and could not reveal in - situ transformation of Ag NPs. In vivo studies over longer time showed that Ag NPs could be oxidized or dissolved, and Ag + ions could complex with glutathione 49 , or be reduced to 0 Ag 143, 222 . 83 Figure 4.5. Representative TEM images of Ag NPs in the leaves of Arabidopsis scord7 mutant after the sorption experiment (10 mg L 1 Ag NPs). B lank: no Ag NPs exposure; C ontrol: opening buffer without abscisic acid (ABA) treatment for 1 h before 10 mg L 1 Ag NPs exposure; 10 µM ABA: opening buffer with 10 µM ABA treatment for 1 h before 10 mg L 1 Ag NPs exposure. Red a rrows indicate the spots of Ag NPs at the subcellular locations. 84 Conclusion The highlight of this study was the use of the ABA - responsive Arabidopsis ecotypes (i.e., Ler and Col - 7) and the ABA - insensitive mutants (i.e., ost1 - 2 and scord7 ) to investigate the role of stomata in the internalization of Ag NPs . The sorbed amount of Ag NPs by the leaves positively correlated with the stomatal aperture, that is, when ABA triggered stomatal closure in the Ler and Col - 7 ecotypes , the amount of Ag NPs sorbed to the Arabidopsis leaves decreased. In contrast, no difference in Ag NPs sorption was observed for the ost1 - 2 and scord7 leaves with or without ABA due to similar stomata apertures. The internalized Ag NPs were associated with the cell membrane, cytoplasm and plasmodesmata of the leaves. Since internalized ENMs canno t be removed by washing, f uture studies are needed to investigate the risks associated with human consumption of ENMs - contaminated crop plants and fresh produce through foliar inputs. Furthermore, the important role of stomata in internalization of Ag NPs also has implications for the delivery of pesticides, nutrients, and other active ingredients by ENMs via the stomata pathway. Author Contribution: Jianzhou He and Wei Zhang conceived and designed the research experiments; Jianzhou He conducted the experi ments and performed sample analysis, analyzed the data, and wrote the manuscript; Li Zhang provided assistance on the Arabidopsis growth and the CLSM measurements; Sheng Yang He, Elliot Ryser, Hui Li , and Wei Zhang reviewed and edited the manuscript. 85 CHAPTER V CONCLUSIONS AND FUTURE WORK 86 Conclusions The promise of nanotechnology to enhance ecosystem and agricultural sustainability critically depends on science - based management of its benefits and risks. This dissertation research demonstrated the potential application of ENMs to treat contaminated water, as well as the uptake of ENMs by plants and underlying mechanisms. Its main findings are as follows: 1. The novel ZnO nanowires exhibited high photodegradation efficiency for cephalex in at circum neutral and alkaline pH with satisfactory reusability over at least five cycles. Cephalexin was degraded through hydroxylation, demethylation, decarboxylation, and dealkylation. I norganic anions (HCO 3 ) and natural organic matter decreased the 2 ). 2. B iochar amendment had no significant effect on the uptake of Ag by radish plants , which did not differ with or without Ag addition (1 mg/kg) . The Ag concentration in soil exchangeable fractions and soil pore water exhibited low bioavailability of Ag at environmentally - relevant concentrations (e.g., 1 mg/kg), which may partially explain the non - s ignificant effect of biochar amendment on Ag uptake. Therefore, the risk of Ag N Ps and Ag + ions accumulat ing in vegetable crops at environmentally - relevant concentrations appear to be very low . 3. Stomata facilitated the internalization of Ag NPs, as revealed by the greater amount of Ag NPs sorbed to ABA - respons ive Ler and Col - 7 ecotype leaves in the ABA - free control than those treated with 10 µM ABA. However, Ag NPs sorption by the ost1 - 2 and scord7 mutant leaves was similar with and without ABA. The sorbed Ag NPs were widely attached to the leaf surface, localized near the cell membrane , and in the cytoplasm and plasmodesmata. 87 Future Work This dissertation research points to several new research directions. Considering the high efficiency of ZnO nanowires and the inhibitory effect of HCO 3 and NOM on the photodegradation of cephalexin, future work should be directed to develop effective eng ineering solution s for water enriched with HCO 3 and NOM . To avoid the need of separating ZnO nanowires from treated water, ZnO nanowires may be embedded into membrane reactors for advanced water and wastewater treatments. The identified photodegradation p athways suggest that parent compound cephalexin is not completely removed but transformed into other product compounds. As these transformed products may be also bioactive, future work should test the bioactivity of transformed intermediates produced durin g photodegradation. It is also plausible that ZnO nanowires could be used as catalysts to photodegrade other organic contaminants (including antibiotics), which should be further investigated. In soil - water - plant systems, uptake of Ag NPs by radish plants at environmentally - relevant concentrations appeared to be low, mainly due to low bioavailability of Ag in soil pore water. Future work should extend this type of study to more vegetable and soil types, assisted with advanced analytical techniques (e.g., FT IR, XPS, XANES, and single particle inductively coupled plasma mass spectrometry) to molecularly speciat e Ag in diverse soil, water , and plant systems. Finally, recognizing that stomata facilitate the internalization of ENMs, future research could develop stomata manipulation strategies to enhance the internalization of beneficial ENMs and minimize the accumulation of harmful ENMs in plants so as to impr ove agricultural productivity, food safety, and the human health. 88 APPENDICES 89 A PPENDIX A: Supplementary Material s to CHAPTER II Text A1. LC - QTrap - MS/MS Method Cephalexin was analyzed using a LC - QTrap - MS/MS system. The sample injection volume 1 . After a 2 min pre - equilibration with 100% phase A, the gradient program for phase B was as follows: the phase B increased to 40% from 0.01 to 1.0 min, then to 70% fr om 1.0 to 2.0 min, maintained at 70% from 2.0 to 3.2 min, then increased to 100% from 3.2 to 3.5 min, and finally maintained at 100% until 4.5 min. The mass spectrometer was set to positive polarity. The curtain gas pressure, ion spray voltage, temperature , and entrance potential were 20 psi, 5000 V, 700 °C, and 10 V, respectively. The multiple reaction monitoring (MRM) mode was used with the transitions of one precursor ion m/z 348.0 to three product ions of m/z 190.5, m/z 174.0, and m/z 158.0 for qualific ation and quantification. The retention time for cephalexin was 2.56 min. All the samples were quantified using matrix - matched standard curves. The LC - QTrap - MS/MS was used to identify cephalexin degradation products under positive ionization mode, using th e combination of an enhanced mass scan (EMS) as survey scan, information dependent acquisition (IDA) criteria, and an enhanced product ion (EPI) scan. The EMS scan for MS spectrum was performed at a scan rate of 10,000 Da/s from m/z 50 to 600. The decluste ring potential (DP), entrance potential (EP), and collision energy (CE) were respective 60 V, 10 V, and 10 V. The IDA threshold was selected at 1000 cps to trigger the EPI scan, following the same scanning rate, DP voltage, and EP voltage with those in the EMS scan, while the CE was set up at 35 V. The collision energy spread (CES) was selected at 15 V. Thus, the averaged MS/MS spectrum was acquired with CE at 20, 35 and 50 V. The MasterView software in PeakView 2.2 package was utilized to process the obtai ned LC - QTrap - MS/MS data. The intensity of precursor ion > 3 (default threshold of the ratio of 90 precursor ion in cephalexin degradation sample to that in cephalexin - free control) was regarded as positive result. The MS spectra of precursors and their corres ponding EPI - triggered MS/MS spectra were then linked to Formula Finder in MasterView to obtain related molecular formulas. The calculated formula was used to search candidate chemicals and their structures in ChemSpider database (http://www.chemspider.com/ ). Then possible structure was compared to parent cephalexin for similar moieties. If candidate structure could be assigned to the fragments of mass spectra, degradation product was considered to be tentatively identified. If no appropriate chemical struct ure was obtained from the ChemSpider database, ChemDraw software (ChemOffice programs, PerkinElmer, Waltham, MA) was used to construct the possible structures based on MS spectra, m/z value, theoretical frontier electron densities calculation, and literatu re report. It should be noted that the tentatively identified intermediates in our study were not further confirmed due to the lack of authentic standards. 91 Table A1. Experimental conditions in selected photocatalytic degradation studies using ZnO photoca talysts in literature. 92 Table A2. Frontier electron densities on atoms in cephalexin calculated by Gaussian 09 program at B3LYP/6 - 311 + G* level with a minimum energy. 93 Figure A1. Chemical structure (a) of cephalexin and species distribution (b) as a function of solution pH (p K a 1 = 2.5 and p K a 2 = 7.2). 94 Figure A2. Structural formula of cephalexin optimized by Gaussian 09 program. 95 Figure A3. Energy dispersive X - ray spectrosco py for elemental analysis. Elements C and Ir are background, confirming that the powders are primarily ZnO. 96 Figure A4. Zeta potential of ZnO nanowires (20 mg L 1 ) suspended in background solution (5 mM NaCl) as a function of so lution pH. 97 Figure A5. Removal efficiency of cephalexin by ZnO nanowires (20 mg L 1 ) under simulated sunlight at pH 7.2 in 5 cycles (90 min reaction time each cycle). 98 Figure A6. 1 , [cephalexin] = 100 µg L 1 , [NaCl] = 5 mM, pH = 7.2, and irradiation time = 180 min). 99 Figure A7. LC chromatograms and m/z of degradation products of cefalexin by ZnO nanowires under simulated sunlight. Cephalexin 100 Table A 3 . Photodegradation Kinetics Data Experiments Replicates Cephalexin concentration (µg L ) over time Initial 5 min 15 min 25 min 45 min 65 min 95 min 125 min 155 min 180 min Hydrolysis 1 138.60 130.72 138.88 137.08 133.48 128.76 127.49 134.92 125.56 125.97 2 134.16 132.68 126.08 127.64 132.00 124.15 123.60 129.55 125.09 118.98 Photolysis 1 129.65 125.16 128.06 129.08 126.81 126.16 122.09 126.16 118.93 128.27 2 133.41 125.56 122.09 131.27 123.96 126.60 122.48 122.17 124.12 125.24 3 130.04 125.95 124.41 125.87 126.65 126.44 119.01 122.58 121.85 130.04 10 mg L Zn 2+ 1 104.27 106.78 105.07 104.32 104.68 96.70 93.61 91.42 89.01 85.56 2 106.39 107.77 105.82 104.63 104.80 97.16 94.96 89.71 89.92 89.95 3 105.07 104.90 104.22 104.83 102.37 99.59 90.00 90.19 89.44 87.10 20 mg L ZnO nanowires 1 108.40 51.33 2.69 4.41 0.52 0.14 0.32 0.05 0.02 0.02 2 112.13 54.66 16.01 4.53 0.73 0.17 0.04 0.04 0.04 0.03 3 115.28 49.15 16.68 4.42 0.70 0.17 0.03 0.05 0.02 0.03 40 mg L ZnO nanowires 1 95.92 29.19 4.57 0.26 0.04 0.05 0.02 0.03 0.03 0.02 2 97.44 31.83 3.24 0.17 0.05 0.02 0.03 0.03 0.03 0.03 3 99.77 32.20 4.20 0.32 0.02 0.02 0.03 0.03 0.05 0.03 80 mg L ZnO nanowires 1 42.36 7.23 0.69 0.12 0.05 0.03 0.02 0.02 0.02 0.03 2 35.93 8.05 0.93 0.11 0.05 0.03 0.03 0.04 0.01 0.03 3 38.11 7.38 0.69 0.04 0.03 0.02 0.37 0.03 0.03 0.02 EDTA - 2Na (h + ) 1 137.94 135.62 133.30 131.01 136.68 136.63 137.94 137.15 135.34 131.51 2 143.73 132.23 139.81 137.63 143.73 141.41 141.96 134.76 135.55 134.02 3 141.63 139.45 139.24 139.98 141.63 141.44 140.10 136.29 135.62 130.38 tert - butanol 1 119.59 114.57 109.64 97.94 95.89 86.56 78.23 72.46 64.09 63.47 2 113.09 117.18 106.17 99.91 93.18 86.77 79.43 74.88 67.46 68.09 3 120.12 110.50 106.84 98.42 98.76 84.93 80.77 75.36 67.08 65.14 101 Table A3 K 2 Cr 2 O 7 (e ) 1 137.54 63.21 24.81 12.60 4.61 1.88 0.78 0.33 0.27 0.20 2 129.81 60.60 23.45 11.56 4.26 1.61 0.62 0.24 0.17 0.10 3 133.47 59.02 23.45 12.48 3.58 1.30 0.49 0.22 0.17 0.12 p - benzoquinone 2 ) 1 216.83 152.61 125.58 105.27 103.86 98.61 98.92 103.08 101.94 106.86 2 206.73 152.72 121.67 107.84 103.21 99.55 103.41 106.00 100.05 105.98 3 196.29 150.99 122.68 115.37 103.25 94.60 97.71 104.84 99.35 106.54 pH = 5.0 1 95.18 88.38 89.14 90.11 92.90 88.45 87.12 84.46 82.35 81.82 2 99.06 93.62 92.58 93.02 91.98 90.60 88.80 88.47 85.07 85.80 3 96.40 91.08 92.19 96.98 94.79 92.28 88.50 88.02 89.68 88.08 pH = 6.2 1 75.58 77.17 82.40 73.18 62.25 60.10 40.53 45.30 44.22 39.22 2 67.97 75.36 81.92 74.53 61.90 55.47 44.08 40.32 40.25 33.78 3 76.68 80.11 81.75 70.91 61.56 56.60 44.86 40.81 37.85 32.30 pH = 8.3 1 122.12 42.14 2.41 0.02 0.02 0.01 0.01 0.01 0.01 0.01 2 130.43 38.66 2.58 0.08 0.02 0.02 0.01 0.02 0.01 0.01 3 122.74 38.45 3.63 0.14 0.02 0.01 0.01 0.01 0.01 0.01 pH = 9.2 1 112.82 25.62 1.03 0.05 0.07 0.04 0.02 0.04 0.09 0.05 2 103.01 23.84 0.65 0.10 0.06 0.06 0.03 0.04 0.04 0.05 3 103.42 24.55 0.97 0.05 0.01 0.02 0.02 0.02 0.03 0.12 1 mM HCO 3 1 127.27 64.54 29.58 15.86 5.85 2.47 0.43 0.08 0.03 0.01 2 117.36 60.13 24.45 10.16 1.86 0.50 0.07 0.02 0.01 0.02 3 119.81 61.97 27.03 15.39 4.47 1.58 0.35 0.07 0.03 0.02 2 mM HCO 3 1 100.29 68.98 39.96 25.00 11.90 5.79 2.04 0.70 0.30 0.19 2 123.38 64.57 30.24 18.05 6.78 2.77 0.74 0.19 0.07 0.04 3 109.46 61.39 24.85 13.64 4.59 1.82 0.40 0.12 0.05 0.04 5 mM HCO 3 1 65.37 83.10 76.31 72.46 73.85 65.59 64.35 58.84 53.57 50.13 2 71.96 78.93 76.22 75.04 72.37 63.69 59.08 55.21 51.16 48.52 3 75.85 79.66 79.25 69.61 69.59 62.71 60.04 57.92 53.29 53.31 102 2 mg L SRNOM 1 122.87 100.97 68.59 51.00 19.37 5.06 1.09 0.20 0.14 0.09 2 117.24 100.66 75.37 51.42 20.74 5.74 1.00 0.28 0.20 0.01 3 116.21 99.37 72.80 56.15 25.26 7.39 1.55 0.63 0.51 0.12 5 mg L SRNOM 1 92.25 98.43 89.23 87.18 79.84 50.69 67.27 58.27 52.07 50.53 2 90.79 94.10 87.69 88.47 83.50 76.13 68.88 64.32 61.15 54.70 3 92.65 97.29 91.13 87.02 82.00 73.07 66.35 61.74 62.11 53.12 10 mg L SRNOM 1 94.94 94.88 88.99 85.06 83.65 71.71 70.48 52.60 72.14 61.22 2 93.50 96.75 90.13 83.52 55.77 70.10 75.37 70.79 68.74 58.56 3 87.89 92.58 95.88 82.85 65.19 71.29 67.34 64.55 63.47 44.03 103 A PPENDIX B: Supplementary Material s Table B1. Fitted parameters of the sorption isotherms using the Langmuir model*. Sorbate Sorbent q max (mg g ) k L (L mg ) R 2 soil + 0% biochar 5.1 (0.2) a 2.4 (0.2) × 10 2 a 0.97 AgNO 3 soil + 0.1% biochar 4.7 (0.0) a 3.1 (0.5) × 10 2 a 0.98 soil + 1.0% biochar 4.7 (0.6) a 3.5 (0.8) × 10 2 a 0.98 soil + 0% biochar 2.1 (0.1) A 2.1 (0.3) × 10 2 A 0.99 Ag NPs soil + 0.1% biochar 2.1 (0.1) A 2.3 (0.2) × 10 2 A 0.95 soil + 1.0% biochar 4.2 (2.3) A 9.2 (0.9) × 10 3 A 0.97 *Results with the same letter in a column are not significantly different among treatments by the LSD post - hoc comparison test at the 5% level. Numbers in parentheses are standard deviations from two replicates. 104 Table B2. Fractions of Ag + and Ag NPs in the soil samples collect ed from the radish - growing pots on Day 35*. Treatment Biochar f1: exchangeable (%) f2: reducible (%) f3: oxidisable (%) f4: residual (%) 0% 0.10 4.72 25.1 70.1 blank 0.1% 0.08 4.67 18.8 76.5 1.0% 0.22 6.45 29.5 63.9 0% 0.01 ± 0.00 a 73.4 ± 14.9 a 8.9 ± 3.9 a 18.1 ± 18.8 a AgNO 3 0.1% 0.03 ± 0.03 a 77.6 ± 11.2 a 7.6 ± 2.4 a 14.7 ± 8.8 a 1.0% 0.00 ± 0.00 a 39.2 ± 6.4 b 15.1 ± 8.9 a 45.6 ± 3.7 b 0% 0.02 ± 0.01 A 76.6 ± 4.6 A 9.2 ± 1.8 A 14.2 ± 2.8 A Ag NPs 0.1% 0.05 ± 0.04 A 66.5 ± 6.3 A 16.2 ± 4.1 AB 17.3 ± 2.7 A 1.0% 0.02 ± 0.01 A 65.4 ± 6.2 A 18.9 ± 5.8 BC 15.7 ± 3.6 A *Results are expressed as means ± standard deviations ( n = 3). The same letter in a column represents that means are not significantly different among different biochar amendments (0%, 0.1%, and 1.0%) by the LSD post - hoc comparison test at the 5% level. 105 Table B3. Fractions of Ag + and Ag NPs in the soil sample s collected from the radish - free pots on Day 35*. Treatment Biochar f1: exchangeable (%) f2: reducible (%) f3: oxidisable (%) f4: residual (%) 0% 0.03 ± 0.00 a 73.4 ± 1.8 a 5.1 ± 17.0 a 21.5 ± 0.0 a AgNO 3 0.1% 0.07 ± 0.00 a 64.8 ± 4.5 ab 16.7 ± 1.7 a 18.4 ± 2.9 a 1.0% 0.04 ± 0.01 a 58.3 ± 4.1 bc 23.8 ± 9.5 a 17.9 ± 13.7 a 0% 0.01 ± 0.00 A 49.3 ± 8.8 A 8.8 ± 6.7 A 41.8 ± 2.3 A Ag NPs 0.1% 0.09 ± 0.08 A 69.9 ± 17.5 A 16.9 ± 11.9 A 13.2 ± 5.6 B 1.0% 0.03 ± 0.00 A 61.9 ± 16.9 A 20.7 ± 8.3 A 17.4 ± 8.6 B * Results are expressed as means ± standard deviations ( n = 2). The same letter in a column represents that means are not significantly different among different biochar amendments (0%, 0.1%, and 1.0%) by the LSD post - hoc comparison test at the 5% level. 106 Figure B1. Representative transmission electron microscopy image (a) and aggregation kinetics (b) of Ag NPs (10 mg L 1 ) dispersed in deionized water. The Ag NPs suspension was stable at a time frame of 4 h. 107 Figure B2. Representative scanning electron microscopy images of the biochar used in this study. 108 Figure B3. Measured pH values of soil samples collected from the radish - growing and radish - free pots on Day 35. 109 Figure B4. The MINTEQ calculation showed that no precipitation occurs (blue text indicates undersaturation) during hydroxylammonium chloride extraction. Simulation condition: [Ag + ] = 0.05 mg L 1 (assuming maximum dissolution, 1 mg silver/kg soil × 1 g soil ÷ 20 ml = 0.05 mg L 1 ); [NH 2 OH·HCl] = 0.5 M; pH = 2; T = 25 °C. 110 Table B4. AgNO 3 Sorption Data AgNO 3 Initial concentration (mg L ) 0% biochar - Rep. 1 0% biochar - Rep. 2 0.1% biochar - Rep. 1 Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) 1 0.51 0.07 0.05 0.51 0.06 0.05 0.50 0.12 0.04 2 0.50 0.09 0.09 0.50 0.09 0.10 0.51 0.15 0.09 5 0.50 0.22 0.24 0.51 0.23 0.24 0.50 0.29 0.23 10 0.51 1.02 0.44 0.50 1.14 0.44 0.51 1.03 0.44 20 0.50 4.27 0.78 0.51 2.88 0.84 0.50 4.51 0.77 50 0.50 18.63 1.56 0.51 17.90 1.58 0.50 18.82 1.56 100 0.50 48.59 2.56 0.50 49.63 2.50 0.51 49.14 2.52 200 0.51 119.89 3.96 0.50 121.19 3.92 0.50 124.79 3.74 Initial concentration (mg L ) 0.1% biochar - Rep. 2 1.0% biochar - Rep. 1 1.0% biochar - Rep. 2 Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) 1 0.51 0.12 0.04 0.50 - 0.02 0.05 0.50 - 0.02 0.05 2 0.50 0.14 0.09 0.50 0.02 0.10 0.50 0.01 0.10 5 0.50 0.32 0.23 0.51 0.12 0.24 0.50 0.17 0.24 10 0.51 0.87 0.45 0.50 0.89 0.46 0.50 0.83 0.45 20 0.50 3.83 0.80 0.51 3.70 0.81 0.50 3.11 0.84 50 0.51 17.20 1.61 0.50 16.84 1.65 0.50 17.02 1.64 100 0.51 44.45 2.74 0.51 45.28 2.69 0.50 45.28 2.73 200 0.51 122.49 3.83 0.51 125.37 3.67 0.51 117.77 4.06 111 Table B5. Ag N Ps Sorption Data Ag NPs initial concentration (mg L ) 0% biochar - Rep. 1 0% biochar - Rep. 2 0.1% biochar - Rep. 1 Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) 1.11 0.51 0.06 0.05 0.51 0.06 0.05 0.51 0.02 0.05 2.22 0.50 0.44 0.09 0.50 0.44 0.09 0.50 0.47 0.09 5.56 0.50 3.10 0.12 0.51 2.83 0.14 0.51 2.11 0.17 11.12 0.51 6.99 0.20 0.50 6.89 0.21 0.50 5.51 0.28 22.25 0.51 12.45 0.48 0.50 15.25 0.35 0.51 14.02 0.41 55.62 0.50 33.96 1.07 0.51 37.51 0.89 0.50 33.98 1.08 111.24 0.51 83.87 1.35 0.51 86.53 1.22 0.50 87.13 1.20 222.49 0.51 186.54 1.78 0.51 191.55 1.53 0.50 187.29 1.75 initial concentration (mg L ) 0.1% biochar - Rep. 2 1.0% biochar - Rep. 1 1.0% biochar - Rep. 2 Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) Soil (g) C e (mg L ) Q s (mg g ) 1.11 0.51 0.06 0.05 0.50 0.02 0.06 0.51 0.02 0.06 2.22 0.50 0.65 0.08 0.51 0.75 0.07 0.51 0.41 0.09 5.56 0.50 2.87 0.13 0.51 2.46 0.15 0.50 2.23 0.17 11.12 0.50 5.02 0.30 0.51 5.75 0.27 0.51 5.27 0.29 22.25 0.51 13.41 0.44 0.50 14.00 0.41 0.50 13.80 0.42 55.62 0.51 32.53 1.14 0.50 36.00 0.98 0.50 35.06 1.02 111.24 0.50 90.06 1.06 0.51 88.16 1.14 0.50 84.89 1.31 222.49 0.50 183.10 1.97 0.51 173.24 2.44 0.50 182.78 1.99 112 A PPENDIX Text C1. PVP Removal from Ag NPs Stock Silver nanoparticles (Ag NPs) with PVP coating were purchased from the US Research Nanomaterials, Inc. (Texas, US). The measured PVP concentration in solution was 0.6%. Ultrafiltration (Amicon Ultra - 10kD membrane filter, Millipore, US) was used to remove free PVP in the aqueous suspension. The filter device was centrifuged at 5,000 × g for 25 min to separate the Ag NPs from solution . The retained Ag NPs were rinsed with DI water, followed by centrifugation. The washing was repeated for 3 times, and the Ag NPs were recovered and re - suspended in DI water. The resultant concentration of Ag NP suspension was determined using atomic absorption spectroscopy (AAnalyst 400, PerkinElmer). Text C2. Ag NPs Characterization The size and morphology of Ag NPs (US Research Nanomaterials, Inc., Texas, US) were determined using transmission electron microscopy (TEM, JEOL 2200FS, Japan) at an accelerating voltage of 10 kV. One drop of Ag NPs sample was spotted onto a 100 - mesh carbon film coated copper grid and dried in the air. The TEM size distribution was c alculated using Image J by randomly selecting ~150 particles from 15 TEM micrographs. Time - resolved dynamic light scattering (DLS) was used to characterize the aggregation kinetics of Ag NPs suspension at pH 6.5 in DI water by measuring hydrodynamic diamet er of a 3 - mL Ag NP suspension ( 10 mg L 1 ) over 3 h using a Zetasizer (Nano - ZS, Malvern Instruments, of Ag NPs suspension was measured with the Zetasizer . All measurements were conducted at least in triplicate s using freshly prepared suspensions at room temperature. 113 Table C1. Measured stomatal aperture width and width - to - length ratio of Arabidopsis thaliana treated with opening buffer in the absence (control) or presence of 10 µM abscisic acid (ABA ). Plants Stomatal aperture width (µm) control 10 µM ABA Ler 4.03 ± 0.72 ( n = 42) 2.62 ± 0.89 ( n = 45) ost1 - 2 5.69 ± 0.92 ( n = 54) 5.68 ± 0.78 ( n = 54) Col - 7 4.45 ± 0.66 ( n = 58) 2.58 ± 0.85 ( n = 63) scord7 4.56 ± 0.83 ( n = 36) 4.29 ± 0.84 ( n = 37) Plants Stomatal width - to - length ratio control 10 µM ABA Ler 0.40 ± 0.09 ( n = 51) 0.29 ± 0.08 ( n = 43) ost1 - 2 0.45 ± 0.09 ( n = 57) 0.48 ± 0.11 ( n = 58) Col - 7 0.41 ± 0.09 ( n = 84) 0.30 ± 0.08 ( n = 84) scord7 0.46 ± 0.05 ( n = 36) 0.44 ± 0.06 ( n = 36) 114 Figure C1. Representative confocal laser scanning microscopy images of Arabidopsis thaliana leaves. Images were obtained by scanning leaf discs (3 mm × 3 mm) of Ler, ost1 - 2 , Col - 7, and scord7 treated with opening buffer without (control) or with 10 µM abscisic acid (10 µM ABA) solution for 1 h. Scale bar is 20 µm. As observed, the ost1 - 2 and sco rd7 mutants of Arabidopsis did not respond to ABA - induced stomatal closure and exhibited constitutively larger apertures, while the stomata of Ler and Col - 115 116 Figure C 2. Stomatal width - to - length ratio of Arabidop sis thaliana treated with opening buffer in the absence (control) or presence of 10 µM abscisic acid (ABA). Results are shown as mean ± standard deviation ( n = 36 84 stomata). Symbol (*) indicates a statistically significant difference in means between the control and 10 µM ABA treatments ( p 117 Figure C 3. Sorption kinetics of 2 mg L 1 Ag NPs on the scord7 leaves. Error bars represent the standard deviations ( n = 2). 118 Figure C 4 . Surface distribution analysis of Ag NPs in the leaves of Arabidopsis Col - 7 ecotype after the sorption experiment (10 mg L 1 Ag NPs exposure) characterized by SEM - exposure, denoted as blank; (B) opening buffer treatment for 1 h before 10 mg L 1 Ag NPs exposure, denoted as control ; (C) opening buffer with 10 µM ABA treatment for 1 h before 10 mg L 1 Ag NPs exposure, denoted as 10 µM ABA. Note that the stomata aperture in the SEM images could not represent the opening or closure status due to artifacts during the sample preparation (e.g., freeze - dr ying). More bright dots were found surrounding the stomatal opening of C than B plausibly due to the ABA induced stomatal closure minimizing the internalization of sorbed Ag NPs. 119 Figure C 5 . Distribution of Ag NPs in selected micro - area o n Col - 7 leave after the sorption experiment (10 mg L 1 Ag NPs). Upper: SEM images at different magnifications; Bottom: elemental mapping of carbon, oxygen and silver. Mapping images indicate the Ag is widely distributed in the selected micro - area. 120 Figu re C6 . Subcellular TEM images of the leaves of Arabidopsis Col - 7 ecotype (A C) and scord7 mutant (D F) after the sorption experiment (10 mg L 1 Ag NPs). B lank: no Ag NPs exposure; C ontrol: opening buffer without abscisic acid (ABA) treatment for 1 h before 10 mg L 1 Ag NPs exposure; 10 µM ABA: opening buffer with 10 µM ABA treatment for 1 h before 10 mg L 1 Ag NPs exposure. The scale bar is 500 nm. It is difficult to identify any possible Ag NPs in the chl oroplasts because of noisy background. 121 Figure C7 . Subcellular TEM images of the leaves of Arabidopsis Col - 7 after the sorption experiment (10 mg L 1 Ag NPs) in the c ontrol treatment (i.e., opening buffer without ABA for 1 h ) . The scale bars are 500 nm in A, C, E, F, and 200 nm for B and D. 122 Table C2. Ag NPs Sorption Kinetics Data Time (min) Fresh leaves (g) C oncentration ( C e , mg L ) Sorbed amount ( Q e , mg kg ) Rep. 1 Rep. 2 Rep. 1 Rep. 2 Rep. 1 Rep. 2 initial concentration = 2.098 mg L , volume = 25 mL 5 0.41 0.40 1.95 1.91 9.12 11.50 10 0.42 0.43 1.77 1.78 19.26 18.45 20 0.43 0.44 1.69 1.67 23.73 24.65 40 0.41 0.41 1.59 1.57 30.78 32.22 60 0.40 0.41 1.56 1.58 33.27 31.93 90 0.42 0.42 1.57 1.59 31.64 30.64 120 0.41 0.41 1.63 1.64 28.70 27.97 123 Table C 3 . Ag NPs Sorption Equilibrium Data Plants Ag NPs exposure Treatments Replicates Fresh leaves (g) Concentration ( C 0 , mg L ) Concentration ( C e , mg L ) Sorbed amount ( Q e , mg kg ) Ler 1 mg L control 1 0.4526 1.25 0.41 46.25 control 2 0.4642 1.25 0.50 40.25 control 3 0.4560 1.25 0.73 28.36 10 µM ABA 1 0.4679 1.25 1.07 9.69 10 µM ABA 2 0.4580 1.25 1.15 5.31 10 µM ABA 3 0.4539 1.25 1.23 1.29 5 mg L control 1 0.4555 4.55 2.38 118.90 control 2 0.4677 4.55 2.28 120.93 control 3 0.4653 4.55 2.05 133.91 10 µM ABA 1 0.4578 4.55 2.72 99.52 10 µM ABA 2 0.4475 4.55 3.01 85.94 10 µM ABA 3 0.4624 4.55 2.66 102.20 10 mg L control 1 0.4690 9.38 6.07 176.24 control 2 0.4637 9.38 5.80 193.14 control 3 0.4627 9.38 6.56 152.49 10 µM ABA 1 0.4778 9.38 8.42 50.14 10 µM ABA 2 0.4599 9.38 8.06 71.56 10 µM ABA 3 0.4631 9.38 8.04 72.36 124 ost1 - 2 1 mg L control 1 0.4940 1.25 0.67 29.22 control 2 0.4890 1.25 0.85 20.62 control 3 0.4540 1.25 0.75 27.61 10 µM ABA 1 0.4530 1.25 0.87 21.27 10 µM ABA 2 0.4630 1.25 0.85 21.89 10 µM ABA 3 0.4870 1.25 0.87 19.78 5 mg L control 1 0.4800 4.55 2.79 91.58 control 2 0.4660 4.55 2.07 132.74 control 3 0.4590 4.55 2.07 134.66 10 µM ABA 1 0.4540 4.55 2.51 112.35 10 µM ABA 2 0.4520 4.55 2.26 126.24 10 µM ABA 3 0.4810 4.55 2.30 116.96 10 mg L control 1 0.5747 14.6 10.22 194.70 control 2 0.6063 14.6 10.20 185.40 control 3 0.6053 14.6 9.83 200.96 10 µM ABA 1 0.5499 14.6 10.21 203.76 10 µM ABA 2 0.5662 14.6 9.83 214.95 10 µM ABA 3 0.6015 14.6 10.47 175.73 125 Col - 7 1 mg L control 1 0.4581 1.81 1.16 35.65 control 2 0.4606 1.81 1.25 30.25 control 3 0.4527 1.81 1.23 32.32 10 µM ABA 1 0.4692 1.81 1.26 29.16 10 µM ABA 2 0.4701 1.81 1.29 27.62 10 µM ABA 3 0.4615 1.81 1.30 27.48 5 mg L control 1 0.4581 6.86 5.41 78.73 control 2 0.4723 6.86 5.15 90.23 control 3 0.4768 6.86 5.27 83.30 10 µM ABA 1 0.4697 6.86 5.57 68.70 10 µM ABA 2 0.4642 6.86 5.59 68.22 10 µM ABA 3 0.4766 6.86 5.68 61.62 10 mg L control 1 0.4292 12.0 8.45 207.52 control 2 0.4272 12.0 8.97 177.71 control 3 0.4295 12.0 9.37 153.71 10 µM ABA 1 0.4462 12.0 9.27 153.56 10 µM ABA 2 0.4539 12.0 9.72 126.28 10 µM ABA 3 0.4618 12.0 8.90 168.18 126 scord7 1 mg L control 1 0.4292 1.44 0.53 52.89 control 2 0.4315 1.44 0.50 54.34 control 3 0.4164 1.44 0.56 52.96 10 µM ABA 1 0.4378 1.44 0.56 50.03 10 µM ABA 2 0.4449 1.44 0.60 47.43 10 µM ABA 3 0.4399 1.44 0.62 46.72 5 mg L control 1 0.4623 4.86 3.85 54.30 control 2 0.4447 4.86 3.46 78.71 control 3 0.4566 4.86 3.52 73.37 10 µM ABA 1 0.4369 4.86 3.32 87.67 10 µM ABA 2 0.4334 4.86 3.41 83.52 10 µM ABA 3 0.4368 4.86 3.51 76.81 10 mg L control 1 0.5637 14.6 9.67 222.65 control 2 0.5626 14.6 9.75 219.65 control 3 0.5615 14.6 9.54 229.50 10 µM ABA 1 0.5373 14.6 9.85 225.24 10 µM ABA 2 0.5304 14.6 10.05 218.91 10 µM ABA 3 0.5442 14.6 9.93 218.99 127 REFERENCES 128 REFERENCES 1. 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