The Timescale of Mutarotation of Monosaccharides and Equilibrium

Home , Mutarotation

Part I: The Timescale of Mutarotation of Monosaccharides and Equilibrium of p- nitrophenol and p-nitrophenoxide ion in Sodium bis(2-ethylhexyl) sulfosuccinate/ Isooctane Microcemulsions Part II: The Toxicity of Triclocarban in Ceriodaphnia dubia ; Accumulation of Triclosan and Triclocarban in Anacharis, Corbicula, Pimephales, and Sediment in a Microcosms

By Cecelia Rose Stuckert

April, 2011

I Certificate of Acceptance

This thesis presented in partial fulfillment of the requirements for the degree of Master of Science in Biochemistry By Cecelia Rose Stuckert Has been accepted by the graduate faculty of The College of Science and Mathematics Colorado State University – Pueblo

Graduate Advisor (Dr. Chad Kinney) Date

Committee Member (Dr. Scott Herrmann) Date

Committee Member (Dr. Del Wayne Nimmo) Date

Committee Member (Dr. Sandra Bonetti) Date

Graduate Director (Dr. Melvin Druelinger ) Date II Part I: The Timescale of Mutarotation of Monosaccharides and Equilibrium of p- nitrophenol and p-nitrophenoxide ion in Sodium bis(2-ethylhexyl) sulfosuccinate/ Isooctane Microcemulsions Part II: The Toxicity of Triclocarban in Ceriodaphnia dubia ; Accumulation of Triclosan and Triclocarban in Anacharis, Corbicula, Pimephales, and Sediment in a Microcosms

An abstract of a thesis presented to the graduate faculty of The College of Science and Mathematics Colorado State University – Pueblo

In partial fulfillment of the requires for the degree of Biochemistry

By Cecelia Rose Stuckert

III Dedication

This thesis is dedicated to my wonderful husband, Stephen W. Stuckert aka ‘Pete’. The most magnificent, supportive, and benevolent man I have ever had the privilege to know and love. Without his overwhelming support I would have never made it through the tough times of the education process. He continues to keep me smiling and moving forward and for that I am forever grateful.

IV Acknowledgements

Thank you, God for all the blessings that have been bestowed in my life.

I offer my sincere gratitude to Dr. Annette Gabaldon, Dr. Melvin Druelinger, and Dr. Don Mykles for giving me the opportunity to continue my education in the NIH Bridges program.

Thank you to all my family and friends for their support and encouragement.

I offer my sincere gratitude to Dr. Sandra Bonetti for her assistance on Part I of this thesis.

I offer my sincere gratitude to Dr. Chad Kinney, Dr. Scott Herrmann, and Dr. Del Wayne Nimmo for all the financial support, expertise, and encouragement for the completion of this thesis. Without their help Part II would have never been possible.

Thank you, USGS of Denver, Dr. Stephen Werner and Dr. Edward Furlong for allowing me to use their facility to run my samples when the instrumentation at CSU-Pueblo was out of commission.

I offer my sincere gratitude to all the Biology and Chemistry staff and faculty of CSU- Pueblo and CSU-Fort Collins that I had the privilege to work with the last 3 years.

I offer my sincere gratitude to Kent Bowman and JBI Construction for allowing me to print my thesis on their copy machines.

V ABSTRACT

Part I: The Timescale of Mutarotation of Monosaccharides and Equilibrium of p- nitrophenol and p-nitrophenoxide ion in Sodium bis(2-ethylhexyl) sulfosuccinate/

Isooctane Microcemulsions

Reverse micellular dispersions were used for the groundwork for the reaction between the enzyme glucosidase and p-nitrophenyl-β-D-glucopyranoside the theoretical end products, glucose and p-nitrophenol, were monitored using 1H NMR and UV-Visible spectroscopy. The reverse micelles were created with sodium bis(2-ethylhexyl)sulfosuccinate (AOT) and isooctane.

Hypothesis I: The timescale of the mutarotation of glucose and other sugar molecules can be monitored with 1H NMR. Hypothesis II: When p-nitrophenol is entrapped in reverse micelles there can be an effect on the extinction coefficient. Specific Aim I: Monitor the mutarotation of glucose and other sugar molecule using 1H NMR. Determine if the Michaelis-Menten kinetic reaction rate be solved based on the mutarotation of the anomeric carbon. Specific Aim II:

Determine if UV-Vis spectroscopy can be used to solve for the extinction coefficient of p- nitrophenol in reverse micelles of multiple sizes.

During an 8 hour assay using 1H NMR analysis of glucose and mannose the reaction rate of alpha verse beta mutarotation was determined. The data for glucose did not reflect the accepted final concentration of the beta glucose anomer (65%) in an aqueous environment. The final concentration of the beta glucose anomer when entrapped in reverse micelles was 48% . The forward reaction rate of glucose was 0.0015 mM / min. The reverse reaction rate of glucose and reverse and forward reaction rate for mannose were equivalent at 0.0008 mM / min. The data collected during an 8 hour assay for other sugars, D-galactosamine Hydrochloride and N-acetyl-

D-glucosamine were inconclusive.

UV-Visible Spectroscopy analysis was used to determine the extinction coefficient of 1 mM p-

nitrophenol and p-nitrophenoxides in multiple AOT concentrations (0.1, 0.2, 0.3, 0.4, and 0.5M) and for various w0 values (6, 12, and 30). The extinction coefficient was determined for the

wavelength of maximum absorbance, 410 nm. This project laid the groundwork for another student to continue and it is suggested that in the future the UV-Vis assays be conducted at a fixed concentration of p-nitrophenol for each w0 value in order to generate data that can easily compared.

Part II: The Toxicity of Triclocarban in Ceriodaphnia dubia ; Bioaccumulation of

Triclosan and Triclocarban in Anacharis, Corbicula, Pimephales, and Sediment in a

Microcosm

Many anthropogenic organic contaminants (AOCs) have become more scrutinized in the last 10 years for their presence in many natural environments. The presence of many such contaminants in surface waters has been tracked back to the effluent of Waste Water Treatment

Plants (WWTPs).Two chemicals, triclosan (TCS) and triclocarban (TCC), have received more attention do to their pervasiveness in many aquatic and terrestrial environments. Hypothesis I :

The LC50 of TCC can be determined for Ceriodaphnia dubia in an environment similar to that found in WWTP effluent receiving streams. Hypothesis II: The bioaccumulation of TCC and

TCS can be determined when freshwater organisms found in North American waterways are placed together in microcosms with water fortified with TCC and TCS. This research had two specific aims regarding the paucity of information in regards to the aquatic toxicology of TCC and bioaccumulation of TCC and TCS in species that can be found in receiving waters connected to WWTPs. Specific Aim I: Determine the toxicity and effects of TCC on the Ceriodaphnia dubia , water flea, using a 7 day static renewal toxicology assay. Specific Aim II: Using

VII microcosms and fresh water species representative of those found in streams receiving WWTP effluent to determine bioaccumulation of TCC and TCS during a 10 day study. Ceriodaphnia dubia were used to determine the median lethal concentration, LC50, and the median effect concentration, EC50, of TCC during a 7 day static renewal test. The LC50 of the C.dubia was determined to be 0.03ng/L TCC. The EC50 is 0.01ng/L. These values, particularly the EC50 determined in this study, are surprisingly low considering the EPA recently published a no observable effect concentration, NOEC, for C.dubia of 1.46 µg/L and EC50 of 3.2 µg/L TCC in a

48 hour static test. The data from this study indicate a much lower NOEC and EC50 values for

Ceriodaphnia dubia.

Microcosms were used to determine the partitioning and bioaccumulation of TCS and

TCC in a simulated freshwater environment that contained Corbicula (freshwater bivalve),

Pimephales (fish), the plant Anacharis, and sediment over a 10 day period. The concentrations of TCC and TCS used in two microcosms, 100 ng/L and 250 ng/L, were based on literature values of the compounds found in WWTP effluent and assuming the effluent makes up 10% and

25% of the receiving streams, respectively. Two other microcosms were water controls (RECON and TEG). RECON water was used to create an EPA moderately-hard water environment and

TEG was the carrier used to place TCC and TCS into solution due to their low water solubility.

On a daily basis one liter of water was removed from each microcosm and renewed with control or fortified water to simulate water renewal in a receiving stream. Using liquid chromatography – mass spectrometry (LC/MS) the concentrations of TCC and TCS were measured in the organisms and sediment. TCC and TCS were observed in all microcosms and baseline specimen revealing the pervasiveness of both chemicals. However, the quantity of TCC and TCS were reduced in the

Corbicula and Pimephales in the control microcosms and was found to accumulate within the sediment and Anacharis of the microcosm. In the fortified treated microcosms, TCC

VIII bioaccumulated in the Corbicula and Anacharis. In the Pimephales the amount of TCC in the fortified microcosms was either close to the baseline amount or lower. The highest amount of accumulation in either of the fortified microcosms was in the sediment, approximately 35-45% of the TCC that was added. TCS in the fortified microcosms accumulated in the Anacharis,

approximately 6-7 % compared to baseline Anacharis. The concentration of TCS in the

Corbicula was reduced in the fortified specimen. TCS in the Pimephales accumulated. Both

TCC and TCS were detected in the sediment in all the microcosms.

IX TABLE OF CONTENTS Dedication IV Acknowledgements V Abstract VI List of Figures XII List of Tables XIII Part I Introduction 1 Review of Literature 1 Microemulsions: What are microemulsions? 1 Reverse Micelles: How they were discovered 3 Sugar Molecules in RMs 7 P-Nitrophenol 11 Objective 13 Materials and Methods 14 Materials Used in Investigations of Sugar Mutarotation and p-Nitrophenol in Microemulsions 14 AOT, surfactant purification 15 Preparation of Reverse Micelles or Microemulsions 17 Monosaccharide mutarotation using Nuclear Magnetic Nuclear Magnetic Resonance Spectroscopy, NMR Spectroscopy for mutarotation studies in RMs 20 P-nitrophenol / p-nitrophenoxides using Behavior of p-nitrophenol in Reverse Micelles: UV-Vis Spectroscopy 23 Results and Discussion 24 Sugar Molecules in Reverse Micelles 24 UV-Visible Spectroscopy 31

X Part II Introduction 37 Triclocarban (TCC) 41 Triclosan (TCS) 43 Environmental Exposure to TCS and TCC 44 Ceriodaphnia dubia 48 Fathead Minnow (Pimephales promelas) 51 Asian Clams (Corbicula fluminea) 52 Anacharis / Elodea 53 Materials and Methods Chemicals and Materials used in Research 54 Toxicology – Effects of triclocarban on C. dubia 55 Aquatic Microcosm: Bioaccumulation and Partitioning of Triclosan (TCS) and Triclocarban (TCC) 62 Solid and Semisolid Sample Preparation 66 Microcosm Water Sample Preparation 68 Liquid Chromatography / Mass Spectroscopy (LC/MS) 68 Results and Discussion 70 Toxicology- Triclocarban 70 Aquatic Microcosm: Bioaccumulation and Partitioning of Triclosan and Triclocarban 76 References 86 Appendices 104 Mannose NMR Spectra A-i N-acetyl-D-glucosamine NMR Spectra A-ix D-galactosamine Hydrochloride NMR Spectra A-xxii C.dubia data collection sheets Test 1 A-xxxiv Test 2 A-xlii C.dubia data combined A-xlix XI LIST OF FIGURES Part I Figure 1. Normal Micelle, Reverse Micelle and Lipid bilayer formations 5 Figure 2. Surfactant, AOT 7 Figure 3. Solvent, isooctane 7 Figure 4. Chemical structure of D-mannose 10 Figure 5. Chemical structure of α-N-acetyl-D-glucosamine 11 Figure 6. Chemical structure of α-D-galactoseamine Hydrochloride 11 Figure 7. Expansion of NMR spectrum in region where H-1 Signals for the glucose monomers in AOT/isooctane RMs α-D-glucose 5.3 – 5.2 ppm and β-D-glucose 4.7 – 4.6 ppm 23 Figure 8. Mutarotation of glucose 23 Figure 9. Mutarotation studies of D-glucose, [AOT] = 0.5 M,

[glucose]D2O = 0.043 mM 26 Figure 10. Mutarotation studies of mannose, [AOT] = 0.5 M,

[mannose]D2O=0.043 mM 27 Figure 11. Mutarotation studies of N-acetyl-D-glucosamine, [AOT] = 0.5 M,

[NADG]D2O = 0.043 mM 28 Figure 12. Mutarotation of D-galactosamine hydrochloride, [AOT] = 0.5 M,

[DGHCl]D2O = 0.043 mM 29 Figure 13. Absorbance vs Wavelength for p-nitrophenol in [AOT] = 0.5 M 33 Figure 14. PNP’s extinction coefficient at wavelength = 410 nm in multiple [AOT] 36 Figure 15. PNP’s extinction coefficient verses concentration of PNP

Variable w0 values 36 Part II Figure 16A. Chemical Stucture of Triclocarban (TCC) 39 Figure 16B. Chemical Structure of Triclosan (TCS) 39 Figure 17. Grid with Color Coded Solutions for static renewal toxicology study 59 XII Figure 18. Sample Data Sheet and Data used for Toxicology Analysis of n = 10 C.dubia in TCC 61 Figure 19. Microcosm Jar with Sand on the Bottom 63 Figure 20. Microcosm Set up in Incubator Upon Initial Onset of Experiment 65 Figure 21. Anacharis with Debris on its Surface being Removed from one of the Microcosms 65 Figure 22. Dionex ASE-100 Instrument 67 Figure 23. C.dubia Daily Total Average of Young per Adult per Treatment 72 Figure 24. C.dubia from 0.03 µg/L giving birth 2-3 days late and dying in process, no live young to report 75 Figure 25 A. C.dubia in RECON and TEG controls that were healthy 75 Figure 25 B. C.dubia in 1.0 µg/L TCC concentration after 24 hours 75 Figure 26. TCC and TCS in the Water Samples for TEG Control and 100 ng/L microcosms 81 Figure 27. Amount of TCC and TCS in Sediment 82 Figure 28. Amount of TCC and TCS in Anacharis 83 Figure 29. Amounts of TCC and TCS in Pimephales 84 Figure 30. Amounts of TCC and TCS in Corbicula 85

LIST OF TABLES Part I Table 1- Materials used in Investigations using AOT Microemulsions 15

Table 2- w0 Values, Concentrations and Volumes Calculations used in 1H NMR Monosaccharide Mutarotation studies 19 Table 3-Controls used in mutarotation studies 21

Table 4-Mutarotation of Glucose in RMs, w0 = 12, [AOT] = 0.5 M 26

Table 5-Mutarotation of Mannose in RMs, w0 = 12, [AOT] = 0.5 M 27

Table 6-Mutarotation of N-acetyl-D-glucosamine in RMs, w0 = 12, [AOT] = 0.5 M 28 XIII Table 7-Mutarotation of D-galactoseamine Hydrochloride in RMs,

w0 = 12, [AOT] = 0.5 M 29 Table 8- Reaction Rates for Glucose and Mannose 31

Table 9-Extinction Coefficient of p-nitrophenol per w0 Value 34

Part II Table 10 – Scientific literature TCS and TCC concentrations found in WWTP influent/ effluent, or freshwater ways (such as rivers or streams connected to WWTP effluent) 50 Table 11- Chemicals, Specimen and Materials used in Toxicology and Bioaccumulation investigations 56 Table 12- Reagents added to MilliQ Water to create 1L EPA Moderately Hard Water- RECON water 57 Table 13- LC/MS Pump Flow for Data Generation 69 Table 14- Quantifying and Characteristic Ions Monitored for Each Analyte and Internal Standard 69 Table 15-Daily average of young per adult of C.dubia for RECON and TEG controls 73 Table 16- TCS in microcosm samples in ng/kg dry weight (mean±s); n=3 77 Table 17- TCC in microcosm samples in ng/kg dry weight (mean±s); n=3 77 Table 18- Mass Balance Accounting of TCC within the Microcosm 78 Table 19-Mass Balance Accounting of TCS within the Microcosm 78

XIV Part I: Introduction

Microemulsions: What are microemulsions?

Microemulsions are stable isotropic, colorless dispersions of three (3) to five (5) components. The components consist of polar and non-polar liquid phases and a wide range of surfactants (Fendler et al., 1973; Shome et al., 2007). The hydrophilic and hydrophobic areas create a spherical droplet structure that may be used as chemical reactor due to posessing interfacial properties at a nanoscale level (Lopez-Quintela, 2003;

Lopez-Quintela, et al., 2004). In 1981, microemulsions were defined by Danielson and

Lindman as a system of single isotropic and thermodynamically stable liquid solutions

(oil and non-ionic surfactants) that consist of water with or without electrolyte

(Constantinides and Scalart, 1997). More recently, microemulsions have been described as water-in-oil (w/o) or oil-in-water (o/w) droplets that are disordered and within these systems there is an important entropy contribution to the free energy of the system

(Wennerstrom, et al., 2006; Constantinides and Scalart, 1997). A valuable property of microemulsions is the typically low interfacial tension that is found at the liquid-liquid interfaces of the spherical droplet that is formed (Lopez-Quintela, 2003; Wennerstrom, et al., 2006).

Multiple or single surfactants can be used to create microemulsions. Some surfactants that have been used to create microemulsions are ionic and some are neutral amphiphilic substances. For example, sodium bis(2-ethylhexyl) sulfosuccinate (AOT or

Aerosol-OT) and cetyltrimethylammonium bromide (CTAB) are both ionic surfactants with the former being anionic and the latter is cationic. Nonionic surfactants can also be used to create microemulsions. Some examples of these are Triton X-100,

poly(oxyethylene)5 nonylphenol ether (NP-5), poly(oxyethylene)9 (NP-9), poly(oxyethylene)12 nonylphenol ether (NP-12), and pentaoxyethylene-glycol-nonyl- phenyl ether (Igepal-C0520) (Lopez-Quintela et al., 2004). The microemulsions that are created using either ionic or non-ionic surfactants can make a micelle or reverse micelle depending on the type and concentration of surfactant relative to water (Fendler et al.,

1972; Prince 1975). Reverse micelles and micelles have been shown to be beneficial in evaluating reactions that occur with charged ions. Ions can be concentrated in a minute area and the reactions with other molecules, such as enzymes, can be monitored (Shome et al., 2007). Furthermore, when microemulsions have been used in analysis of organic reactions, microemulsions overcome solubility problems or heterogeneity issues because they can solubilize both polar and non-polar substances. The ability to compartmentalize and concentrate reactants suggests that microemulsions may provide a better method of monitoring reactions at a nanoscale level (Lopez-Quintela et al., 2004).

The core size of a microemulsion or reverse micelle system (RM) is related to the molar ratio of water to surfactant molecules in the solution, or w0 = [water]/ [surfactant].

Based on the relative amounts of surfactant and water used, the size of the microemulsion and its water pool can vary and may alter the thermodynamics of the system. For example, in studies using CdS, AgCl, and ZnS in AOT/isooctane RMs the particle sizes of the compounds rarely exceeded the size of the reverse micelle aqueous inner core. The study suggests a surfactant-stabilization of nanoparticle droplets (Lopez-Quintela et al.,

2004). Studies using controlled, model systems such as microemulsions, have grown in popularity since their discovery in the late 1970s. In the last 10 years there have been

over 5000 publications using microemulsions indicating a limitless number of opportunities for use in investigations of constrained environments.

Reverse Micelles: How and when they were discovered

During the latter part of the 1970s, the research conducted by E. J. Fendler with micelles led to the search for new model systems to study phenomena such as enzyme activity at lipid-water interfaces. Micelles are microemulsions that form when lipids with a hydrophilic head and a hydrophobic tail assemble in highly aqueous systems. They congregate with the polar heads on the outside and tails on the inside, imitating the outer membrane of biological cells. Micelles were being used in conjunction with 13C NMR and 1H NMR to determine the kinetics of enzymes entrapped in the hydrophobic oil pool of the microemulsions. Unfortunately, the enzymatic activity was found to decrease when enzymes were entrapped in the synthetic micelles. The disappointment associated with this finding led to the use of a new membrane model system the reverse micelle microemulsions.

In Fendler‟s reverse micelles, the detergents mainly used to create the microemulsions were anionic surfactants. The surfactants used were alkylammonium carboxylate surfactants in a non-polar solvent such as cyclohexane, carbon tetrachloride, benzene or dichloroethylene. Formation of RMs was monitored with proton magnetic resonance (Fendler, 1976). When a limited amount of water was added to the surfactants the formation of a macro-homogeneous environment or reverse micelle, was created in accordance with the induced interactions of charges on the molecules (Chang et al., 2000;

Desfosses et al., 1991). The percentage of the non-polar solvent, surfactant and water determine the size of a reverse micelle. The reverse micelle size can be monitored using

carbon tagged and / or proton Nuclear Magnetic Resonance (NMR) spectroscopy, 13C

NMR and/or 1H NMR, ultraviolet –visible spectroscopy (UV-Vis), or fluorescence spectroscopy (Fendler, 1976; Chen et al., 2002; Dorovska-Taran et al.,1993; Walde et al.,1988).

In reverse micelles, the polar head group of the surfactant congregates towards the center which is also referred to as the water pool (Chen et al., 2006). It is within this area that enzymes or molecules can be located and interact with a multitude of molecules within the water pool or with the head group or side chains of the detergent or surfactant

(Boicelli et al., 1985; Fendler, 1976; Eggers et al., 1988; Stahla et al., 2008; Hebrant et al.,1993). As noted before, the detergents can be anionic, cationic, or nonionic (Chen et al., 2006; Shome et al., 2007). Nonetheless the molecular assemblies that are formed are very similar to those found at the outer or inner membrane (normal micelles and reverse micelles, respectively) of cells found in biological systems (Figure 1). These closed assemblies give researchers a model control system by which to investigate and model the activity of molecules and enzymes in the cellular environment (Desfosses et al., 1991;

Eggers et al., 1988; Singh and Aruna., 1995).

Using reverse micelles as models for biological systems has grown in popularity in the medical research field. The ability to study enzymes and enzymatic behavior in biomimetic systems is an aid to researchers in determining inhibitors, active sites, molecular specificity, kinetic activity, or cellular interactions that may occur in vivo

(Bansal-Mutalik et al., 2004; Chen et al., 2006; Eggers et al., 1988; Fendler, 1976;

Boicelli et al., 1985; Luchter-Wasylewska,and Iciek, 2004; Stahla et al., 2008; Desfosses et al., 1991; Hebrant et al., 1993; Singh and Aruna.,1995).

Lipid Bilayer Figure 1. Normal Micelle (NM) and Reverse Micelle (RM) formation exhibits the similarity to formations of lipid bilayers found in biological systems.

(Bansal-Mutalik et al., 2004; Chen et al., 2006; Eggers et al., 1988; Fendler, 1976;

Boicelli et al., 1985; Luchter-Wasylewska, 2004; Stahla et al., 2008; Desfosses et al.,

1991; Hebrant et al., 1993; Singh and Aruna.,1995). In reverse micelles (RMs), it has been shown that counter ion exchange can be monitored or RMs can function as vessels for transportation of amino acids through membranes (Hebrant et al., 1993; Bansal-

Mutalik et al., 2004; Stahla et al., 2008). There is a wide range of possibilities for the use of RMs in the study of biomedical phenomena. Principally, RMs can be used to model the cellular functions of enzymes that function as catalysts in biological reactions.

These enzymatic reactions can occur in different environments within the cell to create amino acid linkages, sugar linkages, phosphate transfers and other molecules. Most importantly, the enzymatic reactions may be affected by the cellular location of the reactions and these environmental effects may be studied and more thoroughly described in microemulsions.

Our studies sought to lay the ground work for the study of β-glucosidase or exocellulase reactions in RMs. The predicted products of this enzymatic activity on the artificial substrate p-nitrophenyl-β-D-glucopyranoside are glucose and p-nitrophenol. To lay the groundwork for future cellulase studies, investigations were undertaken to determine the behavior of these products in reverse micelles under different conditions.

In Part I of this thesis, the kinetics of mutarotation for glucose and other monosaccharides in reverse micelles were investigated. The mutarotation of sugars was followed in reverse micelles using proton NMR spectroscopy or 1H NMR. Next the behavior of p-nitrophenol in reverse micelles under different conditions was studied. In these investigations, the

UV-Vis spectrum of p-nitrophenol in different size reverse micelles and using different amounts of AOT surfactant (Figure 2) in isooctane (Figure 3) was recorded. Each end product was contained in multiple surfactant concentrations of AOT / isooctane RMs. By establishing the behavior of glucose and p-nitrophenol in different types of reverse micelles without enzyme present, it would then be easier to follow the generation of these

products in reverse micelles by cellulolytic hydrolysis of p-nitrophenyl-β-D- glucopyranoside.

Figure 2. Surfactant, AOT.

Figure 3. Solvent, isooctane

Sugar Molecules in RMs

Glucose is an energy source that is extremely abundant in all living forms and is found in abundance in polymers such as glycogen, starch and cellulose. Glucose is obtained mainly by the hydrolysis of glucose-rich polysaccharides, namely starch and glycogen molecules. In these polysaccharides, glucose monomers link to one another creating long straight, branched, and elongated carbohydrate chains (Skelley and

Mathies, 2006; Macleod et al., 2006). Glycogen contains branched and elongated 7

glucose chains. The branches are alpha-1,6 linkages and the elongated chains are alpha-1,

4 linkages of glucose molecules. Starch molecules are the energy source found in plants have similarly linked glucose residue as glycogen, however with less branching.

Cellulose molecules contain 2000-26000 glucose molecules connected together with beta-1,4 linkages. Glycogen is a stored energy source in humans that can be found in the liver and the muscles. When energy is needed, glycogenlytic enzymes aid in the phosphorolysis of single glucose residues that will be used to drive adenosine triphosphate (ATP) production (Macleod et al., 2006; Lewis et al., 2006). Cellulose is not a usual energy source for animals; however fungi and bacteria have the ability to degrade cellulose into individual glucose molecules with the assistance of cellulases.

The orientation of glucose molecules in the cellulose main structural polysaccharide in plants is quite different from the glucose found in animal glycogen

(Lewis et al., 2006; Carta et al., 1992; Simons et al., 2005). This plant structural polymer has glucose molecules that have beta 1,4 linkages that allow the glucose to stack upon itself and hydrogen bond between the different glycan chains. Some examples where cellulose is found are tree trunks and stems of a leaf. The beta conformation at the anomeric carbon creates a molecule that cannot be used by animals for energy because animals do not synthesize cellulases that are capable of degrading this polymer (Carta et al., 1992; Simons et al., 2005). Cellulases are categorized as exo-β-glucosidases, cellobiosidases, and endo-gluconases.

When glucose enters an aqueous environment, it exists primarily in two conformations, that is as β-D-glucopyranose and as α-D-glucopyranose with the anomeric furanoses as minor components. The distribution of the pyranose isomers in the normal

solid state will be approximately 70% alpha and 30% beta. Investigators have shown that when glucose has reached thermal equilibrium in water under normal conditions, the percentages change dramatically and the beta isomer will have an approximate total concentration of 66% and the alpha will have 33% (Nelson and Cox, 2008) . This process of interconversion between the anomeric forms requires ring opening and is called mutarotation.

Mutarotation is a common phenomenon for other pentoses and hexoses and has been studied for L-arabinose, D-xylose, altrose, gulose, mannose, fructose, glucose, and galactose. The 66:33 ratio of glucose anomeric forms is related to the stability of the molecule in an environment with hydrogen bonding between molecules (Nelson and Cox,

2008; Silva et al., 2006). Mutarotation of glucose will continue however, the molar ratio between the anomeric conformations will remain in the ratio of 66:33 in a water environment. The mutarotation is also directly linked to the solvent that the glucose is placed in and how it interacts with the solvent, the pH of solution, and temperature of all solutions when they are mixed (Silva et al., 2006).

We undertook the investigation of D-glucose, mannose (Figure 4), N-acetyl-D- glucosamine (Figure 5), and D-galactosamine hydrochloride (Figure 6) in water and in

AOT reverse micelles. The RMs were created using isooctane and AOT. The monosaccharide was dissolved in deuterated water. This type of research is important because one can compare the rates of mutarotation in the reverse micelle to that in an aqueous system. The RM water pool may mimic the cytosolic side of the lipid bilayer of biological membranes. The mutarotation process may be affected by the size of the water

pool in accordance to the w0 value or molar ratio of the water to surfactant (Lopez-

Quintela, 2003; Luchter-Wasulewska et al., 2004).

When the glucose molecule is within the reverse micelle it should have free movement in the water pool area because glucose is so small. Others have used NMR spectroscopy and integration of the anomeric carbon proton signals to determine the ratio of glucose anomers in different environments and that were generated by biocatalysis

(Fendler, 1976; Carta et al., 1992; Lewis et al., 2006; Silva et al., 2006). From the NMR data they were able to determine the rate of change for enzyme catalysis and processes like mutarotation based on the chemical shifts and products that were noted in the spectra between aqueous and micellular systems using the pseudo first order Michaelis-Menten kinetic equations (Fendler, 1976; Silva et al., 2006). Therefore, it is concluded that in

AOT microemulsions the kinetic rate of glucose mutarotation can be determined when the glucose is in a water pool with or without any enzyme(s) present.

Figure 4. Chemical structure of D-mannose

O HO OH

H HO H3C NH

Figure 5. Chemical structure of α-N-acetyl-D-glucosamine.

Figure 6. Chemical structure of α-D-galactosamine hydrochloride.

P-nitrophenol

P-nitrophenol is an acid and an intermediate for some fungicides (Ellis et al.,

2006). The conjugate base of p-nitrophenol is the nitrophenoxide anion; these two species have different absorption maxima in the UV-Visible spectral range. The equilibrium between the neutral p-nitrophenol (PNP) and its anion, p-nitrophenoxide (PNP-), is dependent on the pH of the solution that contains the chemical. Environmental factors such as presence of other ionic species and ionic strength can also affect this equilibrium. 11

The aim of this research is to document the behavior of PNP in aqueous and RM systems so that this information can be used to analyze enzymatic mixtures (i.e. cellulase) where

PNP and its conjugate base are generated from enzymatic hydrolysis of the exocellulase substrate p-nitrophenyl-β-D-glucopyranoside.

It is likely that the behavior of PNP will change when placed in AOT RMs. The behavior of PNP in aqueous solutions and in AOT RM will be tracked by UV-Vis spectroscopy. The extinction coefficient will be determined in the PNP in aqueous solutions and in AOT RMs. Changes in PNP extinction coefficients due to environmental effects under the various conditions will be compared. The values will be compared to one another to determine if using AOT RMs has any effect on the extinction coefficient, or molar absorptivity values. This research will determine the equilibrium between the p- nitrophenol and the conjugate base, p-nitrophenoxide, in an aqueous environment versus within AOT RMs. The extinction coefficient will be determined for each application and compared to values obtained in RMs containing different AOT concentrations. The information garnered from the glucose and hexose mutarotation studies and from PNP studies in RMs will provide baseline information that may be used in following enzymatic hydrolyses of carbohydrate and associated polymers in RMs.

Objective

The purpose of this research was to provide (1) UV-Vis absorptivity data regarding the behavior of p-nitrophenol and its conjugate base and (2) anomer composition data for monosaccharides in AOT/isooctane RMs or microemulsions. The absorptivity data is needed as the groundwork for tracking the chomophoric phenol products that may form as a result of cellulase reactions performed in RMs using p-nitrophenyl-β-D- glucopyranoside as an artificial exo-cellulase substrate. The anticipated products of cellulase digestion of the PNP-glucoside are p-nitrophenol, p-nitrophenoxide ions and anomers of glucose. The monosaccharide (glucose) products are not chromophores and will need to be tracked by NMR methods. These products could be difficult to track in a

RM so this preliminary study was performed to determine the following:

1. The presence and amount of sugar anomers and rates of mutarotation of different

monosaccharides, including glucose, in RMs using 1H NMR Spectroscopy.

2. The extinction coefficient for the PNP- anion in RMs by means of UV-

Spectroscopy in RMs containing different concentrations of AOT. The effects of

the different surfactant concentrations and RM sizes on the extinction coefficient

for the PNP base will be determined.

Part I: Materials and Methods

Materials Used in Investigations of Sugar Mutarotation and p-Nitrophenol in

Microemulsions

P-nitrophenol (Sigma Aldrich, St Louis MI ) was used without any further purification and dissolved in de-ionized water. The monosaccharides, dextrose (Fisher

Scientific, Miami OK), mannose (Difco Laboratories, Lawrence KS), N-acetyl-D- glucosamine (Sigma Aldrich, St Louis MO), and D-galactosamine hydrochloride (Sigma

Aldrich, St Louis MO) were used without further purification and dissolved in deuterium oxide (Acros Organics, Fair Lawn, NJ) prior to NMR analyses. Methanol (Fisher

Scientific, Miami OK) was used to dissolve the sodium bis(2-ethylhexyl) sulfosuccinate

(Sigma Aldrich, St Louis MO) for purification. Activated charcoal (Fisher Scientific,

Miami OK) was used to extract impurities from the NaAOT or sodium bis(2-ethylhexyl) sulfosuccinate (Sigma Aldrich, St Louis MO) during purification. After the sodium bis(2- ethylhexyl) sulfosuccinate (Sigma Aldrich, St Louis MO) was purified it was dissolved in

2,2,4-trimethylpentane or isooctane (99.8% Sigma Aldrich, St Louis MO) in concentrations ranging from 0.1-0.5 M for microemulsion preparation prior to analysis by

1H NMR or UV-Vis spectroscopy. (Table 1)

Table 1 – Materials for Investigations in AOT Microemulsions. Compound, common name CAS or item number, Supplier, and all pertinent additional information or abbreviation

Sodium bis(2-ethylhexyl) CAS 577-11-7, Sigma Aldrich, Common name AOT, trade name docusate sulfosuccinate (Na AOT) sodium salt

Activated charcoal A-8164, Fisher Scientific , Norit Activated Charcoal

Methanol CAS 67-56-1, Fisher Scientific, Highgrade Methanol UN1230

2,2,4-Trimethylpentane CAS 540-84-1, Sigma Aldrich, Anhydrous 99.8% (isooctane)

D(+)galactosamine No. G-0500, Sigma Aldrich, 2-amino-2-deopxy-D-galctose; chondrosamine Hydrochloride (DGHCl) HCl

N-Acetyl-D-glucosamine No. A-8625, Sigma Aldrich, 2-acetamido-2-deoxy-D-glucose

D-mannose No. B171, Difco Laboratories, Inc.

Dextrose (D-glucose) CAS 50-99-7, Fisher Scientific p-nitrophenol (4-nitrophenol) CAS 100-02-7, Product number 48549, Supelco, Sigma Aldrich

Deuterium oxide (D2O) CAS 7789-20-0, Heavy water, Acros Organics

Methods: AOT, surfactant purification

Purification of AOT was required for this work. Throughout the literature there were various different procedures cited for the purification of the surfactant, AOT (Wei, et al.,

2002; Stahla et al., 2008; Menger and Yamada, 1979; Luchter-Wasylewska and Iciek,

2004). For the purification, a high grade methanol (Fisher Scientific, UN1230) was used.

The purification protocol from grad student Michelle Romanishan CSU-Fort Collins

(Debbie C Crans‟ lab) was as follows (Romanishan, 2009):

1. Na AOT was dissolved in a flask with a minimal amount of methanol. For

example, to make a 0.5 M purified AOT product, 20 g of AOT was placed in an

Erlenmeyer flask using a minimal amount of methanol or ethanol. Approximately

100 mL was just enough solvent so that all of the AOT dissolved. 15

2. Activated charcoal was added immediately after AOT dissolved in solvent.

Activated charcoal removed most of the impurities from the manufactured AOT.

3. The top of the flask was covered with Parafilm and placed on a stir plate for 3

hours.

4. The solution was gravity-filtered until it was clear and colorless (5-6 times).

5. The solution was placed in a round bottom flask, 250 mL, and rotovapped until

the solution cleared for approximately 30 minutes.

6. The clear, colorless product, purified Na AOT, was placed in a desicator for 7

days before using.

The actual purification process used followed the above protocol closely. The charcoal was added to the AOT solution containing 27.87 g of crude AOT in 25 mL of methanol and Parafilm placed on the top of the flask and the solution was stirred for the 3 hours. The AOT was purified when the activated carbon captured the impurities from the manufactured AOT. The last part of the purification process used a rotary evaporator to deplete as much residual methanol as possible from the AOT product, or to achieve product dryness. The equipment used was a BUCHI Rotovapor R-200, connected to a

BUCHI Heating Bath B-490, BUCHI Vac V-500, and a BUCHI Vacuum Controller V-

800. The drying process took approximately 30 to 45 minutes until the purified AOT was again a white, fluffy crystal. The purified AOT was then placed in a dessicator with dessicant to help keep the product dry. An AOT stock solution for use in experiments was made by placing purified AOT into 125 mL of a 0.5 M stock solution, using isooctane as

the solvent. This stock solution was then used to make multiple dilutions of the AOT detergent from 0.1 M to 0.5 M for use as surfactant in the RM experiments.

Preparation of Reverse Micelles or Microemulsions

Different monomeric sugars were entrapped in AOT/ isooctane microemulsions by mixing an aqueous monosaccharide solution with the surfactant solution. The vial that contained the RM components was vortexed for 20 seconds until the solution was homogenous. The aqueous (D2O) sugar solutions were initially 0.400 M and the initial

AOT/isooctane solutions were 0.1-0.5 M in AOT . Once microemulsions were prepared,

1H NMR data was collected and the mutarotation of the sugars was studied by observing and integrating the anomeric carbon 1H signals. Another substance analyzed in AOT/ isooctane microemulsions was p-nitrophenol or PNP, using a stock solution of phenol in distilled, deionized H2O and [AOT] = 0.1 M - 0.5 M. The extinction coefficient of p- nitrophenol in various RMs was observed by UV-Vis spectroscopy. Calculations of how much AOT and aqueous solution with appropriate solute were used to prepare the different sized microemulsions are found in Table 2. The desired value for the w0 or the molar ratio between the AOT and water was used to calculate the amount of

AOT/isooctane solution and water to be used in preparation of RMs.The equation for w0 is as follows:

[H O] w  2 (1) 0 [AOT]

The w0 values that were used in the micoemulsions were w0 = 6, 10, 12, 16, 20, and 30.

The next factor, the concentration of AOT used in experiments was factored in the above

equation. Several serial dilutions were made with the 0.5 M stock solution of AOT; the solutions were made to obtain concentrations of 0.1 M, 0.2 M, 0.3 M, and 0.4 M. All w0 values including each individual [AOT] were calculated and the [H2O] required for each was determined. A sample calculation using equation (1) and inserting w0 = 6 and [AOT]

= 0.5 is shown below:

[H O] [H O] w  2  6  2 [H O]  3 M (1A) 0 [AOT] 0.5 M 2

Using the initial molarity of water, 55.5 M, the amount of water needed to create a 3 M solution was calculated using the following equation:

M1V1  M 2V2 (2)

M1 equals the initial molarity of water, 55.5 M. M2 equal the ending molarity, in this sample case the ending molarity is 3 M. For the final volume or V2 in all equations was

1 mL. (This was the total volume that was used for both NMR spectroscopy and UV-Vis spectroscopy.) Therefore, the value for V1 or volume of aqueous solute solution was determined after rearrangement of equation 2 and solving for V1.

M 2V2 3 M (1 mL) V1  V1  V1  0.0541 mL (2A) M 1 55.5 M

In order to have a solution with w0 = 6 and [AOT] = 0.5 M, the solute (monosaccharide for NMR experiments or p-nitrophenol for UV-Vis experiments) had to be dissolved in

0.0541 mL of water. Stock solutions of glucose and other monosaccharides, 1 mL,

400 mM prepared. From this 1 mL, sugar stock solution only 0.0541 mL of the aqueous solution was used to prepare the RM, (w0=6, [AOT]= 0.5 M) thus altering the initial molarity of the glucose and AOT solutions. The final molarity of sugar or PNP solute or

AOT in the RM could be calculated from 2B equation. An example of a calculation for final glucose molarity in a RM is as follows:

M1V1  M 2V2  400 mM glucose (0.0541 mL)  M 2 (1 mL)  M 2  22 mM glucose (2B)

All calculations were made using a MS Excel spreadsheet. In Table 2 are presented values that were used in the experiments involving 1H NMR spectroscopy of multiple monosaccharides. A similar Table for the UV-Vis experiments involving PNP is found on page 33.

1 Table 2. w0 Values, Concentrations and Volumes Calculations used in H NMR Monosaccharide Mutarotation studies Amount of Amount of [AOT], [D2O], Monosaccharide AOT used Ending molarity w0 M M used, V1(mL) (mL) of glucose (mM) 12.000 0.500 6.000 0.108 0.892 0.043

20.000 0.500 10.000 0.180 0.820 0.072

30.000 0.500 15.000 0.270 0.730 0.108

Nuclear Magnetic Resonance Spectroscopy, NMR Spectroscopy for mutarotation studies in RMs

The rate of mutarotation of sugrs in RMs would be examined by collecting NMR spectra at different time points. NMR spectra were obtained using a INOVA NMR 300

(Varian) (CSU-Pueblo) and INOVA NMR 400 (Varian) (CSU-Fort Collins) both equipped with 6.1C VNMR software. The instrument was set up for 1H NMR spectroscopy. Prior to acquiring spectra, the NMR instrument used was locked on deuterated water. The number of transients or scans acquired for each individual experiment was 256 and it took the instrument approximately 15-20 minutes to complete this number of acquisitions. Once spectra acquisition was complete, the file was saved and the data acquisition was begun again, to collect data for the next time period. This process continued for eight hours or until 24 data files were collected.

After all acquisitions were completed the spectra were integrated in the area where the anomeric carbons‟ proton signals were located for each of the monosaccharides investigated. For the α-D-glucose molecule, the 1H doublet was located at 5.4 - 5.6 ppm, with a 1H doublet at 4.4 - 4.6 ppm for the β-D-glucose molecule. These peaks were integrated and the concentrations of the alpha versus the beta anomers of glucose were determined. In order to observe the mutarotation of the sugars in microemulsions, the

RMs were prepared immediately before the NMR analyses. The glucose molecule in an aqueous solution, specifically water, will continually mutarotate and will maintain a concentration of 2/3 β-glucose and 1/3 α-glucose under normal conditions (25º C).

Control samples were run and used to optimize NMR spectral parameters. The controls

were monosaccharide in D2O and AOT and isooctane with TSP, were analyzed and then the spectra were completed as listed in Table 3.

Table 3. Controls used in mutarotation studies Control Description # of scans

Surfactant TSP/ AOT / isooctane to verify spectra for AOT/ isooctane 256

Monosaccharide D2O with monosaccharide to determine area of spectra for 256

monosaccharide. Same sample was run 24 hours after the

initial scan to verify monosaccharide‟s location on spectra.

RM TSP/ AOT/ isooctane/monosaccharide. Verified the peaks 256

were not interfering with one another after they were in RM.

For each spectrum the H-1 signals for each glucose pyranose anomer were enlarged to magnify the specific location and to determine the amount of each anomer at each time point by integrating the specific spectral peaks. The sum of the anomeric carbon protons 100% and this was value was used to determine the percent of each anomer of glucose present at the specific time point. An example of the anomeric proton region in a 300 MHz 1H NMR spectrum is shown in Figure 7 where the [AOT] = 0.5 M and w0=12 (Figure 7). Unfortunately, when a larger w0 (>12)was used, there was interruption in the area of 4.7 - 4.6 ppm (β-anomer H-1) due to the HOD peak that concurrently increases in size with the deuterated water pool. After much trial and error, it was confirmed that using water suppression for these peaks did not give accurate results for determinating the percent of the beta isomer. Therefore, these w0 >12 samples were not used for data analysis. The NMR data from the w0 = 12 RMs with glucose were

used to determine the rate of mutarotation based on Michaelis-Menten kinetics rate calculations. An expression for the equilibrium constant for mutarotation of monosaccharides is as follows:

  monomer K  (3) s   monomer

The net reaction rate is the rate for the conversion of the alpha monomer to the beta monomer minus the rate for the reverse reaction and is expressed in equation 4 below.

rate  k1[  monomer] k1Ks[  monomer] (4)

In the equation k1 is the time it takes for the forward reaction and k-1 is the time for the reverse reaction. Both are expressed in units per minute. Figure 7 shows a FT-NMR spectrum (256 scans, accumulated over 20 minutes) of D-glucose and shows the anomeric carbon proton signals centered at 5.25 ppm for α-D-glucose and 4.66 ppm for the β-D-glucose. The integrals of these signals at different time points will be used to determine the kinetics of mutarotation. Figure 8 shows how the glucose molecule, as well as the sugar molecules used in this research, continually mutarotate while in an aqueous environment. Appendix I contains the NMR spectra acquired for the monosaccharides mannose, N-acetyl-D-glucosamine, and D-galactosamine hydrochloride.

Figure 7. Expansion of NMR spectrum in the region where H-1 in the sample containing for the glucose monomers in AOT/ isooctane RMs, α-D-glucose 5.3 - 5.2 ppm and β-D- glucose 4.7 - 4.6 ppm. (w0 = 12)

OH OH H k CH2OH H 1 OH H O H O HO H H HO H k HO H OH H -1 HO OH H OH H OH OH H OH OH H H H OH

Figure 8. Mutarotation of glucose

Behavior of p-Nitrophenol in Reverse Micelles:

UV-Vis spectroscopy

The purified AOT that was used in NMR experiments was also used in this section of the research. For UV-Vis analysis, a 1 mM solution of p-nitrophenol (pH 7) was used and the stock solution was stored at 34ºC. This solution had a shelf life of approximately 3 months. As a control, aqueous solutions were mixed and the UV-Vis spectra were recorded for the concentrations of AOT/isooctane with p-nitrophenol ranging from 0.1 - 0.5 M. Other controls were p-nitrophenol in DiH2O for the various w0 23

values that were used in the study. To create a microemulsion using the AOT detergent, a

0.5 M AOT stock solution was used and serial dilutions used to obtain each initial concentration of AOT, from 0.1 M to 0.5 M. The data for absorbance spectra at λ = 350 -

500 nm were used to determine the effect of [AOT] on PNP absorbance. The data were then used to determine the extinction coefficient for the p-nitrophenoxide anion at its maximum absorbance wavelength or λmax, 410 nm, for each AOT concentration (0.1 M -

0.5 M).

Part I: Results and Discussion

Sugar Molecules In Reverse Micelles

The alpha and beta anomeric H-1 signals of glucose molecules in RMs were found using 1H NMR spectroscopy and then integrated. These integrations were used to confirm the point at which the glucose molecule reached equilibrium with 50% of each anomer. The glucose molecule reached near equivalency (48:52) at 148 minutes (Table 4,

Figure 9). The NMR spectral experiments continued for 280 minutes and the concentration of the isomers remained the same for the duration of the experiment. The same sample was run the next day to determine if there was a change after 24 hours.

Glucose never reached the equilibrium of 2/3 β- glucose monomer, which is the amount documented in literature with glucose in water (Nelson and Cox, 2008; Silva et al.,

2006). When microemulsions were being used there appears to be a restriction of the molecule to reach the 65% concentration of the β- glucose monomer. This may demonstrate the effects that microemulsions have on the molecule and possible

interactions with the sulfoxide groups on the polar head group of AOT in the reverse micelle. It may also reflect the difference between water and D2O (Carta et al., 1992).

The other sugar molecules that were used in the mutarotation experiments were mannose (Table 5, Figure 10), N-acetyl-D-glucosamine (NADG, Table 6, Figure 11), and

D-galactosamine hydrochloride (DGHCl, Table 7, Figure 12). The spectra suggest that

1H NMR spectroscopy may be a suitable method for determining the mutarotation of sugars anomeric carbon. The signals from the anomeric carbon can be found in the spectra and therefore can be monitored with a timed assay. The signals that were found remain in approximately a 50:50 ratio through the entire assay for glucose. Mannose also had approximately a 50:50 ratio for the entire assay. However, NADG and DGHCl were changing at the same rate for the entire assay.

Regardless of the quality of the results, all monosaccharide data collected underwent a reaction rate calculation. The rates for NADG and DGHCl are the same for the alpha versus beta, meaning the molecules are changing at the same rate. Literature suggests that the mutarotation from these molecules can normally be monitored at the number two carbon in the ring and not the anomeric carbon (Chen et al., 2005; Hall, et al., 2007; Prag, et al., 2000). From the charts that were created there was no consistent pattern in these molecules at the anomeric carbon that could accurately determine the reaction rate. The figures created from the data generated from the 1H NMR spectra lead to the assumption that the molecules, NADG and DGHCl, will mutarotate in a pattern that is constant and equivilent with one another in RMs. even if they are sequestered in a small water pool.

Table 4. Mutarotation of Glucose in RMs, w0 = 12, [AOT]=0.5 M Time (minutes) alpha-glucose beta-glucose 115 79.46 20.54 153 60.04 39.96 258 49.81 50.19 408 48.07 51.93 464 50.84 49.16 518 52.05 47.95

Mutarotation of D-glucose w0 = 12 90 80

70 60 50 40 α-glucose 30

% % Monomer β-glucose 20 10 0 0 100 200 300 400 500 600 Time (minutes)

Figure 9. Mutarotation studies of D-glucose, [AOT] =0.5 M, [glucose] D2O= 0.043 mM

Table 5 . Mutarotation of Mannose in RMs, w0=12, [AOT] =0.5 M Total Minutes alpha-mannose beta-mannose 13 47.70 52.30 24 48.04 51.90 33 48.60 51.40 42 49.04 50.96 51 49.38 50.62 60 49.96 50.04 69 49.80 50.02 78 49.91 50.09 95 50.01 49.93 106 50.11 49.89 115 50.15 48.85 124 50.53 49.47 133 50.19 49.81 143 50.19 49.81 152 50.21 49.79 160 50.21 49.79 169 50.22 49.78 178 50.22 49.78 188 50.23 49.77 196 50.21 49.79 208 50.22 49.78 217 50.22 49.78 225 50.57 49.43

Mutarotation α-mannose and β-mannose, w0=12 53

50 α-mannose 49 % % Monomer β-mannose

47 0 50 100 150 200 250 Time (minutes)

Figure 10. Mutarotation studies of mannose, [AOT] =0.5 M, [mannose]D2O=0.043 mM

Table 6. Mutarotation of N-acetyl-D-glucosamine in RMs, w0= 12, [AOT] = 0.5 M

Total time alpha-NADG beta-NADG 56 50.65 49.35 72 49.62 49.97 81 49.97 50.03 92 48.87 50.13 102 45.91 54.09 111 47.05 52.95 121 49.95 52.05 130 47.24 52.76 141 48.87 51.13 151 51.52 48.48 160 45.11 54.89 165 46.98 53.02 169 47.68 52.32 180 48.76 51.24 189 47.71 52.29 198 46.90 53.10 209 45.57 54.43 219 48.86 51.14 227 44.37 55.63 236 51.13 48.87 245 45.58 54.42 254 46.26 53.74 262 47.12 52.88 272 44.40 55.60 280 45.38 54.62 289 47.20 52.80 297 52.96 52.96

Mutarotation of N-acetyl-D-glucosamine, w0=12 60 58 56 54 52 50 48 α-NADG 46

% % Monomer β-NADG 44 42 40 0 100 200 300 400 Time (minutes)

Figure 11. Mutarotation studies of N-acetyl-D-glucosamine, [AOT] = 0.5 M, [NADG]D2O = 0.043 mM

Table 7. Mutarotation of D-galactosamine Hydrochloride in RMs, w0=12, [AOT] = 0.5 M Total Minutes alpha-DGHCl beta-DGHCl 13 30.62 69.38 23 31.11 68.89 32 31.55 68.45 42 31.75 68.25 52 31.92 68.08 60 32.08 67.92 69 32.24 67.76 78 32.37 67.63 87 32.59 67.41 103 32.69 67.31 113 32.81 67.19 122 32.89 67.11 131 33.00 67.00 140 33.09 66.91 150 33.16 66.84 161 33.25 66.75 172 33.31 66.69 183 33.36 66.64 190 33.42 66.58 200 33.46 66.54 210 33.51 66.49 219 33.58 66.42 228 33.59 66.41 235 33.58 66.45

Mutarotation of D-galactosamine Hydrochloride, w0=12 80 70 60 50 40 α-DGHCl 30

% % Monomer 20 β-DGHCl 10 0 0 50 100 150 200 250 Time (minutes)

Figure 12. Mutarotation of D-galactosamine hydrochloride, [AOT] = 0.5 M, [DGHCl]D2O = 0.043 mM

The calculation for the forward reaction rate was solved for using the first part of equation 4.

rate  k1[  monomer] k1Ks[  monomer] (4)

The percentage of the monomer was determined using the integral data and the concentration of the monomer was solved using the concentration of the monomer in the

RMs. The final concentration was then divided by the time each data point was collected. After all data points were solved using the same method the average was determined for the entire experiment. The forward reaction rate (alpha glucose) was

0.0015 mM / min and the reverse reaction rate (beta glucose) was 0.0008 mM / min

(Table 8) in w0=12, [AOT] = 0.5 M RMs. When solving equation 4 using the forward and reverse reaction rate the overall reaction rate for glucose was 0.0007 mM / min.

These values differed from literature values of glucose mutarotation that have been suggested to be respectively, 0.17 M / min and 1.101 mM/ sec (Carta et al., 1992; Lewis et al., 2006). Mannose did not show a significant change from alpha to beta isomers and both forward and reverse reaction rates were identical (0.0002 mM/min). This differed from literature values of mannose using 1H NMR which were reported to be 0.00102 mM/ min in an aqueous environment instead of a RM (Anderson et al., 2008). The forward and reverse reaction rates for glucose and mannose are shown in Table 8. For this part of the project it would be suggested that if there is future work done with this assay, the researcher should focus on glucose and possibly use higher concentrations of glucose so the solvent peak from HOD does not interfere with the data analysis. Running assays with N-acetyl-D-glucosamine and D-galactosamine hydrochloride sugar molecules did not yield interpretable results for the mutarotation experiment. Therefore it 30

would be advisable to re-analyze these sugars using a more sensitive NMR instrument probe or a higher field instrument. Some other methods can also be used, for example,

13C NMR, fluorescence spectroscopy, or near-IR spectroscopy to determine the reaction rates for mannose, NADG, and DGHCl (Macleod et al., 2006; Skelley and Mathies,

2006; Lewis et al., 2006).

Table 8. Reaction Rates for Glucose and Mannose Sample Molar Ratio, w0 Forward Reaction, Reverse Reaction, α → β (moles per minute, β → α (moles per mM/min) minute, mM/min)

Glucose 12 0.0015 0.0008

Mannose 12 0.0002 0.0002

UV-Visible Spectroscopy

The second part of this research was designated to determine the extinction coefficient of p-nitrophenol in AOT/isooctane RMs. The extinction coefficient is calculated using the absorbance at 410 nm for the PNP anion. The wavelength, 410 nm, differed from documented wavelengths of 402 nm 405 nm, 407 nm, and 420 nm used in previous studies (Bessey and Love, 1952; Muresanu et al., 2005; Brouns et al., 2006).

When the p-nitrophenol is in the microemulsions the extinction coefficient was suppressed at low w0‟s and there is quite a difference between the aqueous solution versus those in the reverse micelles. The aqueous standards were mixed using H2O, instead of AOT/isooctane, in solutions equivalent to those of the highest concentrated

RMs (w0 = 6, 12, 30, [AOT] = 0.5 M). The maximum absorbance for the aqueous samples was 410 nm. The aqueous standards did relate to the documented absorbance at

407 nm. Using the reverse micelles the extinction coefficient is dramatically reduced. An example of this is shown in the UV-Vis spectroscopy data for [AOT] = 0.5 M in w0 = 6,

12, and 30 (Figure 13, Table 9).

This suggests that the p-nitrophenol in a reverse micelle has a lower molar absorbtivity and hence a lower extinction coefficient versus when the p-nitrophenol is in an aqueous solution. From the data that was acquired through the UV-Visible spectra, there did not seem to be as much of a change in spectra due to the different [AOT] in the

AOT RMs except at [AOT] = 0.5 M. The range of values was 1.037 – 1.910 for w0 = 6, for w0 = 12 the range was 0.784 - 2.519, and for w0 = 30 the range was 2.676 – 5.593 for

[AOT] 0.1, 0.2, 0.3, 0.4 M . The extinction coefficient was calculated at the wavelength of maximum absorbance, 410 nm, for the PNP anion. Table 9 shows the extinction coefficient of PNP-chromophore at each w0 and concentration of AOT.

Overall for p- nitrophenol there was a difference between aqueous solutions and microemulsions. The microemulsions decrease the extinction coefficient approximately ten times what can be found in the aqueous versus those in the microemulsions. The change from aqueous versus the microemulsions suggests that the molecules within the

RM water pool may be partially interacting with the surfactant interface. In the RM the p-nitrophenol entrapped within absorbs less light from the UV-Vis spectrometer. The RM was absorbing less light therefore reducing the absortivity measurement and extinction coefficient. Therefore, the PNP solution in RMs absorbs less light, which reduces both the absorbtivity value and the values of the extinction coefficient. In addition,

p-nitrophenol ↔ p-nitrophenoxide equilibrium may be perturbed in the RM. There was also a difference between literature values of the p-nitrophenoxide extinction coefficient in aqueous environments and the extinction coefficent that was determined through this research project (Brouns et al., 2006; Muresanu et al., 2005). The differences can be from the different concentrations of p-nitrophenol that were used in this research.

Unfortunately, there were no literature values to compare p-nitrophenol in microemulsions using the same environment that was used. The only literature data found used AOT/isooctane microemulsions with p-nitrophenol in a pH = 10 (Anderson et al.,

2008). The pH value in the literature was greater than what was used to complete this experiment, the pH did not exceed 7.6.

Wavelength vs Absorbance for [AOT] = 0.5M 1.80E+00

1.60E+00

1.40E+00

1.20E+00

1.00E+00 w0=6 AOT=0.5 8.00E-01 w0=12 AOT=0.5

Absorbance 6.00E-01 w0=30 AOT=0.5 4.00E-01

2.00E-01

0.00E+00 345 395 445 495

Wavelength, nm Figure 13. Absorbance vs Wavelength for p-nitrophenol in [AOT] = 0.5 M

Table 9. Extinction Coefficient of p-nitrophenol per w0 Value Extinction Coefficients for p-nitrophenol Ending Extinction Concentration of Concentration of p- Absorbance at coefficient -1 -1 w0 AOT nitrophenol (mM) λ = 410 nm (mMol cm ) 6 0.1 0.011 0.011 1.037

12 0.1 0.022 0.049 2.264

30 0.1 0.054 0.162 2.994

6 0.2 0.022 0.054 1.910

12 0.2 0.043 0.083 2.519

30 0.2 0.108 0.289 2.676

6 0.3 0.032 0.053 1.623

12 0.3 0.065 0.051 0.784

30 0.3 0.162 0.859 5.302

6 0.4 0.043 0.055 1.282

12 0.4 0.087 0.093 1.074

30 0.4 0.216 0.963 4.458

0.038

6 0.5 0.054 0.002 703.6*

1.028

12 0.5 0.108 0.111 1003*

5.593

30 0.5 0.270 1.510 2392*

*Aqueous standards

Unfortunately, it was discovered that there was an experimental flaw and error when this work was being completed. The data for the UV-Visible Spectroscopy should have all been completed using the same concentration for the p-nitrophenol. Although the

PNP stock solutions used were the same initial molarity for the entire study, the same amount of stock solution was not used for each w0 that was analyzed. However, this flaw was somewhat corrected by using calculated extinction coefficients based on final PNP concentrations. The AOT concentration effects on extinction coefficient were hard to tease out because w0‟s and PNP concentrations were both changing (Figure 14). The solved extinction coefficients at the wavelength of 410 nm (Figure 15) were compared to one another in regards to their w0 value. The data in Figure 14 suggests that high [AOT] there is a greater suppression of PNP absorbance. The data in Figure 15 suggest that the extinction coefficients are larger for the w0 =30 where the PNP is present in a large H2O pool and is less likely to encounter another PNP molecule or to bump into the surfactant interface.

[PNP] vs ԑ in Variable [AOT] 6.000

5.000

4.000 AOT = 0.1 M 3.000 AOT = 0.2 M AOT = 0.3 M 2.000

AOT = 0.4 M ԑ, ԑ, extinctioncoefficient 1.000 AOT = 0.5 M

0.000 0.000 0.050 0.100 0.150 0.200 0.250 0.300 [PNP], mM

Figure 14. PNP‟s extinction coefficient at wavelength = 410 nm in multiple [AOT].

[PNP] vs ԑ in different w0 values 6.000

5.000

4.000

3.000 w0 = 6 w0 = 12 2.000

w0 = 30 ԑ, ԑ, extinctioncoefficient 1.000

0.000 0.000 0.050 0.100 0.150 0.200 0.250 0.300 [PNP], mM

Figure 15. PNP‟s extinction coefficient versus concentration of PNP variable w0 values.

Part II: Introduction

Release of xenobiotic chemicals into streams and rivers has been under particular scrutiny since the New York Times Best seller, “Silent Spring” by Rachel Carson was released in the 1960‟s (Carson, 1962). This book greatly influenced the environmental movement which in part is credited with the formation of the U.S. Environmental

Protection Agency, USEPA, born on December 2, 1970. The USEPA relies on its own research as well as the research of scientists at universities and other government organizations to monitor land, water, and air. Recently, the use and disposal of pharmaceuticals and pesticides, aka anthropogenic organic contaminants or AOCs, used by consumers in the general public has received greater scrutiny because of their presence in the environment and possible harmful impacts.

Many AOCs are released into aquatic environments, either by their producer or via wastewater treatment plants (WWTPs) (Burks, 1982; Ahn et al., 2008; Arnot et al.,

2006; Lacey et al., 2008), AOCs have been measured in the influent as well as effluent of

WWTPs (Klein et al., 2010; Coogan et al., 2007; Chalew and Halden, 2009; Blanchard et al., 2001). Furthermore, many of these chemicals are detected in surface waters that receive treated wastewater effluent (Kolpin et al., 2002; TenEyck et al., 2007). However, the environmental effects of AOCs have not been widely investigated. This creates a concern regarding the many AOCs that are detected in surface waters that receive wastewater effluent and the aquatic life that is within these same waters (Naqvi et al.,

1993; US ASTM, 1993).

The lists of known toxins in wastewater that are regulated include pesticides,

PCBs, and asbestos, but the release of most pharmaceuticals, antimicrobial agents, and

many other AOCs are currently unregulated (US ASTM, 1993). Unfortunately, trace amounts (parts per billion or parts per trillon) of the AOC residuals released into surface waters can be circulated back into the drinking water, thus contaminating households indirectly by simply turning on the tap water supply (Benotti, et al., 2009; Blanset et al.,

2007).

Although researchers have begun to understand the environmental impacts of the release of such AOCs, little is known about long term environmental effects. The AOCs that are not currently regulated but detected in water samples from streams, rivers and tap water are being further studied for their toxicity (Blanset et al., 2007; Ciccotelli et al.,

1998; Wade et al., 1994). Experimentally AOC‟s can be determined a threat to the environment. If experimental results indicate a compound is lethal or being toxic to the aquatic organisms found within the affected area, there is a possibility the compound can become regulated (Arnot et al., 2006; Heidler et al., 2006; Johnson et al., 2007; Reg.

Tox. and Pharm., 1994). Some toxic effects accepted by the USEPA as negotiable AOCs that can be regulated are those that have been reported using reproduction and mortality rate of an organism (Reg.Tox. and Pharm, 1994; Arnot et al., 2006).

Recently the concern over the overuse of the antibacterial reagents triclocarban

(TCC) and triclosan (TCS) has increased (Coogan et al., 2007; Chalew and Halden, 2009;

Ishibashi et al., 2004; Miller et al., 2008; Morrall et al., 2004, Figure 16). The overuse of

TCS and TCC may interfere with the ability of naturally occurring organisms to fight infection and disease (Miller et al, 2008; McClellan and Halden, 2010; Coogan et al.,

2007). The concerns regarding the possible overuse of both chemicals starts with the

amount used on a yearly basis, which has been reported to be a staggering 4.5 x 105 kg per year each (Hodges et al., 2007).

A. B.

Figure 16.A. Triclocarban (TCC) B. Triclosan (TCS)

The concentrations of TCS used in products are normally within the range of 0.1 -

0.3 % by mass of the product (Chalew and Halden, 2009). The amount of TCC used in products, such as bar soap is approximate 2 % by mass (Heidler and Halden, 2008). A majority of TCC and TCS is released into the environment through land application of wastewater biosolids, but some of is also released in WWTP effluent into the connecting waterways (USEPA, 1989; Coogan et al., 2007, Heidler and Halden, 2007). Biosolids are WWTP sludge that meets regulatory requirements for terrestrial application to aid in the growth of plants in agriculture and restoration projects because it is a rich source of C and N (Kumar et al., 2010a; Reiss et al., 2009).

The biocides, TCC and TCS, were the subject of a literature review article by

Chalew and Halden (2009) that documents the toxicity threshold for both compounds in multiple organisms and compare this to available environmental occurrence data. The toxicity thresholds are the amount of TCC or TCS that create systemic effects including developmental toxicity of organisms in regards to their presence in the environment. The

organisms that were included in the review, include fish, algae, and crustaceans, which are expected to be in the freshwater environments where TCS and TCC have been detected. Interestingly, the amounts of TCS and TCC detected in WWTP effluent are similar. Chalew and Halden (2009) documented that the limit of toxicity threshold of

TCS in algae was similar to the quantity of TCS found in WWTP effluent (2-4 ng/L). It was also determined that TCC‟s toxicity threshold was higher for algae (4-5 ng/L). In fish the toxicity thresholds of TCS and TCC were found to be 4.5-6 µg/L and 3.5-5.5 µg/L respectively. Crustacea showed a higher toxicity threshold for TCS than for TCC, 3.5-6

µg/L and 1.5-4.5 µg/L, respectively (Chalew and Halden, 2009). However, in some studies reviewed by Chalew and Halden (2009) there is evidence that the toxicity thresholds have exceeded the published concentrations of WWTP effluent and the effects on the organisms are still being reviewed.

To date (2011) there are more research publications available in the scientific literature on TCS perhaps due to a wider range of products with lower concentrations than TCC. However, over the last 5 years there were an increasing number of papers on

TCC and its occurrence in the environment (Klein et al., 2010). An example of a research project that has raised concern over the environmental impact and some of the long term effects of TCS on zebrafish was documented by Orvos et al. (2002). Using zebrafish, Danio rerio, during a 48 hour static test, the median effective concentration,

EC50 , was found to be 280 µg/L and 360 µg/L using different embryo stages of the fish, stage ELE (eleutheroembryo) and E (embryo) respectively (Lammer et al., 2009; Orvos et al., 2002). The end point measured by Orvos et al. (2002) was abnormal fin lengths.

This type of effect is indication of TCS being an endocrine disruptor that interferes with

the natural growth process, in this case fin length for zebrafish. Also, TCC has indicated an endocrine disruption in female mudsnails. In mudsnails , Potamopyrgus antipodarum, the TCC specifically interrupted the ability for females to produce healthy embryos at exposure concentrations of 0.2, 1.6, and 10.6 µg/L over a 4 week period (Giudice and

Young, 2010). Nonetheless, there are limited available data on the environmental effects of TCS and TCC, but given the large quantities of TCC and TCS used on an annual basis, further research is warranted (Capdevielle et al., 2008; Ireland, 2010).

Triclocarban (TCC)

Since the late 1950s and early 1960s some antibacterial and pharmaceutical products have included the antimicrobial agent TCC (Heidler et al., 2006; Xia et al.,

2010). Initially, TCC was added to cosmetics and personal care products. In pharmaceuticals the chemical is added to a maximum of 1% of germicidal creams and ointments (Ash and Ash, 2004). Today, TCC is added to deodorant soaps, detergents, cleansing lotions, wipes and deodorants. It is added as a preservative in bar and liquid soap, including body washes (Ash and Ash, 2004). It is active against gram positive bacteria in amounts from 1-2% by mass of the product (McDonnell et al., 1998, USEPA,

2002; Giudice and Young, 2010). The TCC used in household products is generally washed down the drain and is not completely removed during wastewater treatment

(Higgins et al., 2009; Giudice and Young, 2010).

In 2002 it was alleged that there is minimal exposure of this chemical to aquatic life because the majority of this chemical, 94%, is removed from the WWTP‟s via sludge

treatment systems (USEPA, 2002). In surface water from WWTPs effluent the concentration of TCC has been predicted to be between 1.30 and 17.0 ng/L (USEPA,

2002). The concentrations reported by the TCC Consortium (2002) were on the low end compared to other studies that have measured the concentration of TCC to range from 48 to 540 ng/L in WWTP effluent (Hodges et al., 2007; Klein et al., 2010; Kumar et al.,

2009). Nonetheless, the agreement with all studies has shown there are trace amounts of

TCC in WWTP effluent. One concern about TCC is that it can undergo aerobic biodegradation to the carcinogenic and cytotoxic compounds, known as chloroanilines

(Giudice and Young, 2010). There are limited data on the bioaccumulation and effects of

TCC and chloroanilines in aquatic organisms and environments (Miller et al., 2008;

Giudice and Young, 2010; Higgins et al., 2009). This has caused much concern over the discharge concentrations of TCC and any potential hazards to aquatic organisms (Giudice and Young, 2010; Higgins et al., 2009). With the overwhelming use of this chemical, and detection of it in biosolids and WWTP‟s sludge, there remains little published regarding toxicology and bioaccumulation of TCC in aquatic environments connected to WWTPs where residual of the chemical can be found (Coogan et al., 2007; Higgins et al., 2009;

USEPA, 2002). A generalization that has been made in literature is that TCC and TCS are related with indications of both having endocrine disruption effects that can jeopardize aquatic environments when either chemical is released into the environment

(Coogan et al., 2007; USEPA, 2002). However, there is still more known regarding the presence, fate, and effects of TCS. The lack of scientific literature regarding the toxicology and environmental effects of TCC warrants concern for aquatic life in waters impacted by WWTP effluent.

Triclosan (TCS)

Starting in the late 1970s (Binelli, 2009) another antimicrobial agent TCS, was introduced for use in personal care and consumer products. Much like triclocarban, triclosan was assumed to be nontoxic to the environment and assumed to be a beneficial antibacterial agent; it has been widely used in many personal hygiene products (Glaser,

2004; Hodges et al., 2007). Triclosan is commonly used as a broad spectrum germicide.

Some products that contain TCC are soaps, toothpaste, cosmetics, pharmaceutical products, fabrics, plastics, textiles, dermatological creams, and many other products including clothing (Singer et al., 2002; Coogan et al., 2007; Binelli et al., 2009). In pharmaceuticals triclosan is used in dermatology products and for antibacterial applications in acne treatments. In toothpaste TCS is added at 1% weight. Research has shown TCS in concentrations ranging from 5 to 50 mg/L can cause an interruption in the ability of dental plaque species Streptococcus sanguis and Capnocytophaga gingivalis to grow and develop to maturity (Greenman et al., 1997). Triclosan is an ionizable chlorinated biphenyl ether (Orvos et al., 2002; Coogan et al., 2007) that inhibits activity of an enzyme enoyl-acyl carrier protein reductase in fungi and bacteria, stopping an essential step for membrane development (Ahn, 2008). Chronic exposure of natural bacteria to TCC in receiving streams has led to strains of TCS resistant bacteria (Coogan,

2007; Seaman et al., 2007).

Generally the TCS metabolite, methyl-triclosan (m-TCS) is detected in higher quantities in WWTP sludge compared to influent suggesting that TCS degrades to m-

TCS during wastewater treatment (Coogan et al., 2008). About 96% of the TCS used is washed down the drain and a majority of it is suspected to partition to biosolids during

wastewater treatment where it is detected from 0.4-30 µg/g (Lozano et al., 2010; Reiss et al., 2009). For example Kumar et al. (2009) reported 5370 ng/L of TCS in WWTP influent with only 180 ng/L in the effluent. The WWTP biosolids are normally applied to agricultural lands and the TCS that it contains has been shown to affect plants, earthworms, soil microorganisms, birds, and any other living organism that come into direct contact with the chemical (Reiss et al., 2009; Kinney et al., 2010). The study concluded that plants, specifically cucumber plants, in a biosolid amended environment have the highest sensitivity to TCS at concentrations as low as 0.280 µg/g (Reiss et al.,

2009).

Environmental Exposure to TCS and TCC

Triclosan and triclocarban primarily find their way into the environment through discharge of treated wastewater effluent or land application of biosolids, which are the end products of wastewater treatment (Orvos et al., 2002; Coogan et al., 2007; Kantiani et al., 2008). Throughout the US, there is generally a reduction in the aqueous concentration of TCS and TCC from WWTP influent to effluent (Kantiani et al., 2008;

Binelli et al., 2009; Klein et al., 2010). Generally 3-6% of the TCS in the influent leaves the WWTP in the effluent (Kantiani et al., 2008; Kumar et al., 2009). About 9% of the

TCC entering WWTPs is released in the effluent (Halden and Paull, 2004 and 2005;

Kumar et al., 2009). Much of the observed reduction in aqueous concentration during treatment can largely be attributed to partitioning into the organic carbon rich solids

(sludge or biosolid) during treatment (Klien et al., 2010; Coogan et al., 2007; Miller et al., 2008). 44

About 89% of the TCS and 78% of the TCC that are released into the environment is through land application of biosolids, primarily as an organic carbon- and nutrient- rich soil amendment on agricultural soil (Halden and Paull, 2004; Reiss et al.,

2009; Stasinakis et al., 2008). The remaining of TCC and TCS is discharged from

WWTPs‟ effluent into in the receiving streams and may eventually go through degradation into chloroanilines and m-TCS respectfully (Halden and Paull, 2005; Miller et al., 2008; Reiss et al., 2009; Son et al., 2009; Stasinakis et al., 2008). After many years of use, TCC and TCS persistence in the environment and the effects of either have been questioned. Research in the last 10 years has begun to evaluate the presence of the two bacteria fighting compounds in soils and sediment.

In sediment TCC is more persistent than TCS (Miller et al., 2008, 2010). Miller et al. (2008) measured the concentrations of TCC and TCS in sediment to be 1.6 ± 0.7 µg/g and 0.07 ± 0.01 µg/g, respectively (Miller et al., 2008). Another study evaluated WWTP sludge and pond sediment. The quantity of TCS found in the WWTP sludge was 0.515-

1.61 µg/g. The amount of TCS found in the pond, that was not affiliated with any WWTP effluent, was 0.581 µg/g (Kumar et al., 2010b). The high concentration found can be a direct result of the high use of TCS since the 1970s and the chemical seeping through the soil either through rainfall or wind erosion. Aquatic ecosystems sediment have also been evaluated for TCC (0.024-1.425 µg/g) and TCS (0.011 µg/g ) (Kumar et al., 2010b). The quantities of TCC and TCS measured in multiple environments illustrate how ubiquitous both compounds are in the total environment i.e., soil, sediment, and sediment associated with aquatic ecosystems.

The United States is not the only country raising concerns over the extensive use of TCS. More studies monitoring the concentrations of TCS and m-TCS found in waterways have been conducted abroad. For example in northeast Spain, Kantiani et al.

(2008) collected thirty-one water influent and effluent samples from eight different

WWTPs, sixteen river samples (that are WWTP effluent impacted), and twenty- two drinking water samples within the greater Barcelona area. An additional 26 samples of influent and effluent were collected from seven wastewater treatment plants along the

Ebro River. Kantiani et al., (2008) found that the amount of TCS was significantly reduced during wastewater treatment. The influent concentrations of TCS ranged from

200-12,500 ng/L, but traces were still present in the effluent, from 20-712 ng/L. Although generally observed to be produced during wastewater treatment (Kantiani et al., 2008;

Kumar et al., 2009 ). Kantiani et al., (2008) observed similar behavior for m-TCS during wastewater treatment with the influent ranging from 40-354 ng/L and in effluent m-TCS was below the detection limit up to 50 ng/L.

The TCS and m-TCS in wastewater effluent is much higher than the median effect concentration, EC50, values for the green alga, Selenastrum capricornutum, 70 ng/L and freshwater alga 3.40 ng/L as reported by Orvos, et al. (2002) and Tatarazako et al. (2004). The waterways that Kantiani et al (2008) tested were connected to WWTP effluent and the amount of TCS and m-TCS measured ranged from below the detection limit up to 285 ng/L. In the drinking water samples only two tested positive for TCS, however the quantity found was not reported.

Binelli et al., (2009), studied the in vitro exposure to TCS in zebra mussels,

Dreissena polymorpha, finding TCS at very low doses, 0.1-10 µM, caused severe DNA

injuries and hemocytes were affected . The amount of TCS that is observed in wastewater effluent impacted environments is similar to that quantity found to cause these DNA injuries in zebra mussels (Kantiani et al., 2008; Binelli et al., 2009). Triclosan creates an inhibition of the committed step in fatty acid synthesis in humans and wildlife and it has been found to have deleterious effects on aquatic organisms, raising the concern for its presence in WWTP effluent and in receiving streams (Heath et al., 1999; Liu et al.,

2002). To date TCS and TCC continue to be widely used and released into the environment (USEPA, 2003, 2009; Kantiani et al., 2008; Binelli et al., 2009).

The environmental exposure to TCC and TCS has led to bioaccumulation in animal and plant tissue (Orvos et al., 2002; Ahn et al., 2008). The organisms that have shown to accumulate TCC and TCS, such as algae, fish and earthworms create a potential pathway for biomagnification through the food chain (Reiss, et al., 2009; Coogan et al.,

2007). However, the potential biomagnification is not fully understood. The purpose of this research was to determine toxic effects and bioaccumulation of TCC and TCS on representative aquatic species. The research project tested two hypotheses: Hypothesis I :

The LC50 of TCC for Ceriodaphnia dubia is below the concentration of TCC observed in aquatic environments. Hypothesis II: TCC and TCS at environmentally relevant concentrations will accumulate in aquatic organisms. This research had two specific aims regarding the paucity of information in regards to the aquatic toxicology of TCC and bioaccumulation of TCC and TCS in species that can be found in receiving waters connected to WWTPs. Specific Aim I: Determine the toxicity and effects of TCC on the

Ceriodaphnia dubia , water flea, using a 7 day static renewal toxicology assay. Specific

Aim II: Using microcosms and fresh water species representative of those found in

streams receiving WWTP effluent determine bioaccumulation of TCC and TCS during a

10 day study.

This research includes a seven day toxicity test using the fresh water daphnid,

Ceriodaphnia dubia, to determine the median lethal concentration ( LC50). Secondly, a bioaccumulation and partitioning study using aquatic microcosms with organisms representative of those found in north american freshwater streams and rivers. The organisms used were fathead minnows (Pimephales promelas), fresh water clams

(Corbicula fluminea) and the plant Anacharis sp. were used to determine the bioaccumulation and partitioning of TCC and TCS. The concentrations of TCC and TCS used in the microcosm experiments are based on a dilution of known TCS and TCC concentrations observed in WWTP effluents and representative of concentrations found in receiving water streams (Table 10).

Ceriodaphnia dubia

Ceriodaphnia dubia (C. dubia) is a microcrustacean or water flea and related organisms can be found in many freshwater environments throughout the world. C. dubia is a filter feeder, using interior and exterior appendages to filter water. The invertebrate uses quick, rapid movements that allow them not only to swim but capture algae and any microorganisms upon which they feed. Use of C. dubia in toxicology studies started in the mid 1980s and continues today. There are multiple advantages of this type of testing with the water flea (Gallagher et al., 2005; Kosmala et al., 1999; Tatarazako et al., 2004).

One advantage of using C. dubia is the short lifetime and reproduction cycle, which typically starts within days of being born. The USEPA uses tests with C. dubia because

they have been found to be one of the most sensitive acute and chronic toxicity indicators

(USEPA, 1989, 2009). The toxicology assays are also designed to determine toxicity and aide in regulations on toxic materials in freshwater environments and WWTPs effluent

(Tatarazako et al., 2004).

The time frames for studies using C. dubia generally span 7-10 days. The assay starts when a neonate is within the third brood of a female and less than 24 hours old.

Within a week the C. dubia neonate will grow and generate three broods of young.

C. dubia typically start having neonates within three days of birth and continue to reproduce every day to every other day after the first brood .The first brood under normal conditions will have 3-5 neonates. The second brood will be approximately 9-12 neonates and the third brood will be in the range of 13-16 neonates (TenEyck and Markee., 2007;

Tatarazako et al., 2004). Having a relatively quick reproduction and life cycle makes

C. dubia an exceptional candidate for both acute and chronic toxicity tests. The toxicity of the aqueous environment is dependent on the most toxic chemical present (Gallegher et al., 2005). As a result, C. dubia is commonly used to determine median effective concentrations, (EC50) and median lethal concentrations (LC50) for suspected contaminants particularly chemicals that can be found in WWTP effluent (TenEyck and

Markee, 2007; Loureiro et al., 2010). For example in a study by TenEyck and Markee

(2007) C. dubia was used to determine the toxicity of chemicals that are found in WWTP effluent and find their way into receiving streams. The chemicals tested were nonylphenol (NP), nonylphenol monoethoxylate (NP1EO), and nonylphenol diethoxylate (NP2EO) and multiple combinations of the chemicals similar to what can be found in WWTPs receiving streams. Using a 48 hour static renewal test the LC50 for NP,

NP1EO, and NP2EO were determined 92.4, 328, 716 µg/L, respectively (TenEyck and

Markee, 2007). A static renewal test with C. dubia was used to determine the toxicity of

TCC in this study.

Table 10: Scientific literature TCS and TCC concentrations found in WWTP influent/ effluent, or freshwater ways (such as rivers or streams connected to WWTP effluent) Author Location / Type of Method TCC ( ng/L) TCS ( ng/L) Sample Klein et al. , 4 Northwest Ohio SBSE-LD- 48 ± 2, 2010 WWTP / WWTP LC/MS/MSa 120 ± 20, 170 ± 30, effluent 244 ± 6, 330 ± 30

Coogan et al., Denton, Texas / Pecan GC-MSb <0.015 - 0.19 <0.01- 0.12 2007, 2008 Creek Kantiani et al, Northeast Spain/ IA, SPE-GC- 83 – 1283 2008 Influent and effluent MSc WWTP, rivers Barber et al., Phoenix, AZ / Tres SPMDd – 86 - 92 2006 Rios Demonstration GC-MS Constructed Wetlands Guidice and 60% of US streams Multiple 109, 213 Young, 2010 (Median, mean respectively) Kumar et al., 4 Savannah, Georgia LC-MS/MSe 25978(I), 2292(E), 18850(I), 1036(E), 2009, 2010a, WWTPs / Influent (I) 9325(I), 2356(E), 13703(I), 180(E), 2010b and effluent (E) from 17357(I), 3045(E), 86161(I), 5370(E), WWTPs 3505(I), 36221(I), 5213(I), 32639(I), 281(E) 274(E) Chalew and Multiple data sources / Multiple 2-50000(I), 35-50000(I), Halden, 2009 WWTP influent(I) and 1.5-3500(E) 1.5-4000(E) effluent(E) (I) - Influent , (E) - effluent aSBSE-LD-LC/MS/MS – Stir bar sorptive extraction and liquid desorption-liquid chromatorgraphy-tandem mass spectrometry, bGC-MS – Gas Chromotography-Mass Spectrometry, cIA-Magnetic particle-based Immunoassay, SPE-GC-MS-Solid Phase Extraction Gas Chromotography-Mass Spectrometry dSPMD-Semipermeable membrane devices eLC-MS/MS-Liquid Chromatography-Mass Spectrometry

Fathead Minnow (Pimephales promelas)

The fathead minnow (FHM), Pimephales promelas, is a fish commonly used in laboratory and field studies. The Pimephales can be found from the south of the US to the northern part of Canada, indicating it is a good representative species to use when doing work in north american freshwater ecosystems (Scott and Crossman, 1973; Kidd et al., 2007 Pimephales are small, easy to maintain, widely distributed, and represent a large family of fish, Cyprinidae (Kidd et al., 2007). Their biology and life history are well characterized making them a perfect candidate for toxicology studies (Kidd et al., 2007).

Because of these factors, the results of laboratory based experiments utilizing Pimephales can generally be extended to natural aquatic systems (TenEyck and Markee, 2007;

Kovacs et al., 1992; Kidd et al., 2007).

There are a number of published studies that have used Pimephales to measure bioaccumulation potential of contaminants from WWTPs and runoff from surface areas

(Arthur et al., 1973; Brungs et al., 1974; Kidd et al., 2007). For example, a pioneering study utilizing Pimephales was performed by Arthur et al. in 1973 to determine the toxicity of sodium nitrolotriacetate, NTA in aquatic environments. In the study

Pimephales were exposed to a similar quantity of NTA as was measured in Lake Superior using a laboratory microcosm. They were able to conclude that there was not an observable change in survival, spawning activity or egg hatchability under chronic exposure to NTA (Arthur et al., 1973). Kovacs and Voss (1992) used Pimephales quantify the effects of newsprint and specialty mill effluent by exposing Pimephales in sub-chronic and chronic tests.

Pimephales used in this research were chosen to represent a commonly occurring freshwater fish species for an accurate comparison to a natural environment in a laboratory microcosm. Fish were used because many studies have been published regarding the toxicity of TCS and TCC in acute and chronic toxicity, however, no studies were found using a bioaccumulation model such as what was used for this thesis (Miller et al., 2008; Coogan et al., 2008; Kantiani et al., 2008; Orvos et al., 2002). For example, a toxicity test using freshwater fish and 150 ng/L TCS showed changes in the thyroid hormone receptor gene expression leading to a reduction in body weight and decrease in swimming (Miller et al., 2008). Triclosan has been shown to cause androgenic changes in

Pimephales specifically in fin lengths and sex ratio; these changes were found in the early stages in fish embryos at concentrations between 0.07 – 0.29 mg/L (Kantiani et al.,

2008). Another test using Pimephales and TCS showed a statistical difference in a 35 day post hatch survival analysis between the fish in TCS fortified water (1000 µg/L ) and the control water (Orvos et al., 2002). Another effect of TCS found in the Pimephales was a delayed swim behavior subsequently to the exposure of TCS at a concentration of 71.3

µg/L (Orvos et al., 2002).

Asian Clams (Corbicula fluminea)

Corbicula fluminea is a freshwater bivalve that is considered an invasive species that was accidentally introduced in North America and fouls freshwater intake systems

(Bidwell et al., 1995). Through their diet, the Corbicula fluminea bioaccumulate toxins by filtering waterborne organic matter and plankton in the freshwater systems they reside

(Vidal et al., 2002a; Jou et al., 2006). Corbicula fluminea have been used in

bioaccumulation studies such as the kinetic uptake of uranium during a 42 day semi- chronic exposure (Simons et al., 2005). The accumulation of uranium is dependent on the amount of sorption-absorption storage and loss by excretion from the organism. It was concluded that there was potential in using this organism for toxicology and bioaccumulation studies using sub-chronic exposure methods (Simons et al., 2005).

Corbicula fluminea is also a good candidate for investigations in freshwater ecosystem environments because they can potentially to bioaccumlate xenobiotic organic pollutants that are present in trace quantities (Labrot et al., 1996; Basack et al., 1998; Vidal et al.,

2001; Vidal et al., 2002b). C. fluminea have also displayed abnormal development and reduction in larval survival at TCC concentrations as low as 30 µg/L (Miller et al., 2008).

In this research they represented a hardy freshwater bivalve mollusk that is a filter feeder.

Anacharis / Elodea

Anacharis also called Elodea or Philotria is a freshwater higher plant often known as the north american or canadian water weed. It is easily grown in aquariums throughout the US, except where it is outlawed for being a competitive species.

Anacharis grows fairly easy using fish waste as its main source of nutrients and it recycles oxygen into the water. Anacharis is similar to vegetation found in most freshwater areas thoughout the US. Studies have used this vegetation to determine the uptake of heavy metals in freshwater systems (Mal et al., 2002; Flora, et al., 1994). The genotoxicity of mercury was found in a short term experiment using Anacharis and mercury containing compounds (Flora et al., 1994). In another short term in vivo exposure Mal et al. (2002) were able to demonstrate the bioaccumulation of copper in

Anacharis. In another experiment completed by the same lab, the exposure of iron was analyzed in Anacharis that was found in drainage ditches (Mal et al., 2002).

In aquatic environments the algae and protozoans that rely on bacteria-like type II fatty acid pathway have been shown to be sensitive to low (µg/L) concentrations of TCS

(Miller et al., 2008). Another study has shown that plant life such as algae are the most sensitive to TCS in low parts per trillion, ppt, concentrations (Chalew and Halden, 2009).

Another study using algae has determined that TCS is toxic at a concentration of 3.4

µg/L (Kantiani et al., 2008). In this study the Anacharis (Egeria densa or Elodea canadensis) that is found in freshwater environments (i.e. microcosms) will be analyzed for TCS and TCC uptake. Anacharis was used as a representative of an aquatic plant species similar to those that are found in natural aquatic environments.

Materials and Methods : Chemicals and Materials used in Research

TCC and TCS (Sigma Aldrich, St Louis MI) were analytical grade, 98% pure

13 13 isotope labeled TCC and TCS ( C13 -TCC and C12 –TCS) (Wellington Laboratories

Ontario, Canada). De-ionized (DI) water was filtered through a MilliQ system for ultra pure (18.1 MΩ) water. MilliQ water is created when laboratory DI water is further treated by a Millipore Simplicity 185. Within the MillQ system the DI water is subjected to UV light to kill all bacteria as well as multiple filters to remove impurities. Ceriodaphnia dubia and FHM along with their food source that were used were purchased from

Aquatic Biosystems, Fort Collins, CO. Anacharis was purchased from Pueblo Pet Store,

Pueblo CO. Oasis HLB 20 cc (1 g) LP Solid Phase Extraction (SPE) Cartridges (Waters

Corp.) was used to filter organics from microcosm water. Acetonitrile (Honeywell

International, Muskegon,MI) , triethylene glycol (Acros Organics, Newark NJ), triclocarban (Sigma Aldrich, St Louis MI), and triclosan (Sigma Aldrich, St Louis MI) were used as received (Table 11).

Toxicology- Effects of Triclocarban on C. dubia

Ceriodaphnia dubia were used to determine the effects of TCC on survival and reproduction. Ten specimens were used for each concentration of TCC and two controls

(RECON and solvent controls). The concentration of TCC used were based on the amounts that have been documented as the no observable effect concentration, or NOEC, of 1.46 µg/L (USEPA., 2009) . The concentrations was based on a logarithmic scale 1, 3,

10, 30, and 100 ng/L. The highest concentration, 100 ng/L is similar to what can be found in streams receiving WTTP effluent (Table 10). The water used in these toxicological studies was reconstituted water, or RECON standard water, prepared in accordance to the

EPA guidelines for moderately hard water (USEPA, 1989). RECON water is MilliQ water that contains the necessary basic salts that could be found in moderately hard fresh water systems. To create RECON water (moderately hard water) the salts (Table 12) are added to the Milli-Q (Millipore Corporation, model Simplicity 185) water and mixed for

24 hours with air, to allow reagent-salts to dissolve. The final conductivity for a majority of this research remained within EPA standards of 249-252 µS/cm, and the water was maintained at room temperature.

Table 11. Chemicals, Specimens and Materials used in Toxicology and Bioaccumulation Investigations. Compound, CAS or product number, Common name, distributor, type, product Abbreviation description, or other product / chemical information

Triclocarban, TCC CAS 101-20-2; 3,4,4‟-trichlorocarbanilide, Sigma Aldrich, St. Louis

Triclosan, TCS CAS 3380-34-5; 5-chloro-2-(2,4-dichlorophenoxy)phenol

13 Triclocarban, C13 –TCC In solution that was 1.0 ppm with methanol solvent, Wellington Laboratories, Ontario Canada, purchased as 50 µg/mL solution with 98% purity in methanol

13 Triclosan, C12 - TCS In solution that was 1.0 ppm with methanol solvent, Wellington Laboratories, Ontario Canada, purchased as 50 µg/mL solution with 98% purity in methanol

Triethylene Glycol, TEG CAS 112-27-6; AC 139590010, Triethylenglykol, 99%, Acros Organics Code New Jersey USA

RECON Water Created in lab with MilliQ water and salts, mixed with water pump, conductivity in EPA standards between 242-254 µS as found in moderately hard water

Acetonitrile, ACN CAS 75-05-8; Burdick and Jackson LC/MS High Purity solvent, Honeywell International, Muskegon, MI 49442

RAPIDVAP N2(gas) NI300; Nitrogen gas tank attached to Rapidvap instrument

YTC Aqueous solution that contains Pines International wheat grass, salmon starter and Fleischmanns yeast, solids are 1780 mg/L total solution. Supplied by Aquatic Biosystems, Fort Collins, CO

Algae Selenastrum capricornutum with a cell cound of 3.0 X 107 cells/mL. Supplied by Aquatic Biosystems, Fort Collins, CO

Salmon Starter Supplied by Aquatic Biosystems, Fort Collins, CO

Ceriodaphnia dubia, Water flea, variable aged, fed YTC and Selenastrum capriconutum immediately, water flea live in controlled environment 22-25°C. Supplied by Aquatic Biosystems, Fort Collins, CO

Anacharis Purchased from pet store located in Pueblo, CO

Corbicula Collected from fresh water source on Arkansas River

Pimephales Supplied by Aquatic Biosystems, Fort Collins, CO ~5 month old species

Beverage Ware Product 028572103745, 1 oz. condiment cups from Plastics Inc. AH HPI Company Coon Rapids, MN 55433

Silicone Rubber Tubing Product 12589; Rubber tubing from Bio-Sil Silicone Rubber .125”ID .250”ODX25‟

Sil-Med Corp. 700 Warner Blvd. Taunton, MA 02780

Within the MillQ system the DI water is subjected to UV light to kill all bacteria as well as multiple filters to remove impurities. MilliQ water is commonly used base water to create soft, medium, or hard water conditions for aquatic specimens. The conditions for this research was to make a moderately-hard water condition specific for toxicity studies using C.,dubia, the water flea, which has become widely used in toxicology labs (USEPA, 1989; US.ASTM, 1993).

Table 12. Reagents added to MilliQ Water to create 1 L EPA moderately hard water –RECON water. Reagent Amount needed for 1 L Amount used to make 15 L NaHCO3 96 mg 1440 mg CaSO4 · H2O 60 mg 900 mg MgSO4 60 mg 900 mg KCl 4 mg 60 mg

Six solutions were made using the RECON water and the stock TCC solutions and all were maintained at room temperature. The concentrations selected to be used in this study were based on a logarithmic scale and related to the concentrations of TCC that can be found the environment. Multiple scientific resources have documented concentrations present in rivers and streams ranging from 15 – 5600 ng/L; these values were taken into consideration when determining the concentrations of TCC solutions to test (Klein et al., 2010; Guidice and Young, 2009; Kumar et al., 2010a; Coogan et al.,

2007). Initially the stock solution was made using methanol as a solvent/ carrier of TCC for six concentrations, 1.000, 10.00, 30.00, 100.0, 300.0 and 1000 ng/L. However, it was determined after an initial test that the methanol was toxic so a new solvent was chosen, triethylene glycol (TEG). It was also determined during the initial test that all the

concentrations used were acutely toxic to the C. dubia, regardless of carrier effect. A new stock solution was made with 0.6010 g of TCC and 100mL of triethylene glycol, TEG.

The initial concentrations tested with TEG as the solvent were 0.1000, 0.3000, 1.000,

3.000, 10.00, 30.00, and 100.0 ng/L TCC (0.3171, 0.9514, 3.171, 9.514, and 31.71 µM

TCC respectively). There was no indication of a carrier effect; however the concentrations used remained too toxic for a successful study. The final concentrations of TCC that were tested were 0.01000, 0.03000, 0.1000, 0.3000, and 1.000 ng/L

(0.0003171, 0.0009514, 0.003171, 0.09514, and 0.3171 µM TCC respectively). The TEG control consisted of the maximum amount of TEG that was present in the highest concentration TCC solution, 20 mL per 2 L in RECON water (0.07325 M).

A system of color coding was used to identify each solution of TCC, rather than concentrations, to protect against bias when conducting the experiment. An 8 X 10 grid was prepared on a Styrofoam board for the study (Figure 17). There were two boards that were set up exactly the same so samples could be transferred from one board to another on a daily basis. The cup used for each sample was a plastic 29.57 mL cup

(Plastica Inc, Coon Rapids, MN, 55433) filled with 13-16 mL of solution.

Figure 17. Grid with color coded solutions for static renewal toxicological study

Ceriodaphnia dubia were purchased from Aquatic BioSystems, Inc. (Fort

Collins, CO). Upon arrival, adult females were transferred evenly to four 500 mL beaker and were fed ~10-15 mL, YTC and algae ad libitum. YTC is a compilation of wheat grass, salmon starter and yeast that gives specimen, such as Ceriodaphnia dubia, the nutrients that are needed on a daily basis for survival. The method used to create the

YTC, the liquid nutrient source, was EPA/505/8-89-002a it was purchased from Aquatic

Bio Systems, Fort Collins CO.

The Ceriodaphnia dubia were fed the YTC and algae daily. After 1-2 days of being in an incubator at 26 °C the most active and largest females were separated using a stereomicroscope at 10X magnification and the individual specimen were placed in their own cup. After 24 hours the neonates that were less than 24 hours old from each female were counted and removed from the cup using a stereomicroscope and placed into a clean dry cup. The females that had 10-18 neonates were in their third brood and the robust

young from the females were kept for use in the toxicology study. Robustness of the young is based on the quantity and quality of motion as an indication of specimen health.

Each cup was filled half way (13-16 mL) with one of the 7 solutions of differing

TCC concentrations, a TEG control, or RECON water control and a single neonate was placed in each cup. In the first cup of each concentration two neonates were placed in the cup for the first two days. This ensured to have an extra organism just in case there are any troubles or losses of a single neonate within the first 48 hours. After 48 hours if the neonate was not needed it was discarded. When all neonates were transferred to the cups they were covered with Shamrock plastic wrap and placed in an incubator at 26 °C for 24 hours (12 hours of light, 12 hours of dark). Randomization was changing the position of the board in the incubator by rotating the board 90º daily.

Each Ceriodaphnia dubia was transferred into a new plastic cup containing the same concentration of solution using a polytheylene pipette assisted by a Bausch and

Lomb stereomicroscope. All the Ceriodaphnia dubia were accounted for on a daily basis along with any other documentable mannerism, such as reproduction, survival, and locomotion using the stereomicroscope. However, if the neonate was swimming and reproducing there was no addition information documented on the daily data sheet. An example of the first few rows of a hypothetical data collection sheet is shown in Figure

18. The sheet got a check mark if the Ceriodaphnia dubia adult female was alive and if alive and with young the checkmark was separated with a slash and the amount of neonates written below the slash. If the concentration was too toxic for the Ceriodaphnia dubia and it did not survive the previous 24 hours the sheet received an X for that specimen. If the Ceriodaphnia dubia was alive, noted by a beating heart, but not moving

and moribund this would be noted in the box under the checkmark, as would any other abnormal behavior that was witnessed. Any observation made for the entire concentration was noted on the margins of the sheet. Appendix II contains all data collected for toxicology study.

After seven (7) days the number of specimens remaining and the total sum of 3 broods of young that each adult produced was averaged for each concentration. The results of the average and standard deviation of young per adult were graphed to determine if there was any statistical difference for a given toxicological endpoint between the RECON water control / TEG control and those in the different TCC concentrations. The data were analyzed using standard t-test in biostatistics software,

Minitab version 5.6.

___Month/Year______Ceriodaphnia dubia______Name (First and Last)___

Date Species Investigator(s)

Pink Yellow Orange Green Blue Red White Orange/Green

 X      X

5 3

       X

5 2 4

 X   X   

Figure 18. Sample data sheet and data used for toxicology analysis of n = 10 C. dubia in TCC.

The data that were compared was the value of young per female for each concentration compared to number of young per female in the control solutions. The endpoints that were used in the analysis were the reproduction of three broods from each female after 7 days of static renewal of TCC fortified solutions and control solutions. The documentation of the mortality, mobility, and mannerisms of the organisms were useful observations for determination of the median lethal concentration of the specimen (LC50) and median effect concentration of the specimen (EC50) for the C. dubia in TCC affected environment. On days 1, 2 and 4 the dissolved oxygen, pH, conductivity, and daily temperature were measured for all the solutions. The dissolved oxygen level was between

6.7 – 7.1; pH was between 7.9-8.0. The conductivity of the solutions was between 265-

279. The temperatures of the solutions when removed from the incubator were measured between 25.7 - 26.0 ºC.

Aquatic Microcosm: Bioaccumulation and Partitioning of Triclosan (TCS) and

Triclocarban (TCC)

Microcosms were used to determine the partitioning and the potential bioaccumulation of TCC and TCS in aquatic life in a model fresh water aquatic system.

The four 4 L fresh water microcosms were prepared and included a RECON water control, solvent (TEG) control, a microcosm with 100 ng/L each of TCC and TCS, and a microcosm with 250 ng/L each of TCC and TCS. The concentrations of TCC and TCS used in the two test microcosms were based on a 10% and 25% dilution of the TCC and

TCS in representative WWTPs effluents (Table 10) into a receiving water (Coogan et al.,

2007; Klein et al., 2010, Kumar et al., 2010a; Kantiani et al., 2008).

The organisms chosen for the study are representative of and commonly found in freshwater environments in North America. The first specimen, Corbicula were collected from the Arkansas River basin directly west of the Pueblo hatchery water drain located at N 38°16.006‟ W 104°42.746‟. These specimens were maintained in water from the collection site in an incubator at 17 °C for six (6) days before they were introduced into the microcosm. Anacharis also known as Elodea, was purchased from the Pueblo Pet Store, Pueblo, CO. Pimephales (5 months old) were purchased from

Aquatic Bio Systems, Fort Collins CO.

Microcosms were established in 4 L glass jars. Each jar was cleaned with 99% ethanol and dried overnight. Once the jar was dry 270 g of clean sand was measured and placed on the bottom of each clean jar (Figure 19).

Figure 19. Microcosm jar with sand on the bottom.

After a liter of one of the four treatments water was added to each jar along with approximately 12 g of the plant Anacharis. The Anacharis was allowed to be free floating within the microcosm above the sand. The jars were then filled with a total of three liters of treatment water. Each jar was marked at the three liters mark. The jars were placed in 63

an incubator programmed to remain at 17°C with compressed air aeration hose attached to a ceramic aerator placed in each microcosm, and each equilibrated for 24 hours (Figure

20).

After 24 hours the completion of microcosm construction started with the addition of seven Corbicula to each microcosm. A glass funnel with a mesh filter attached to rubber tubing (Bio-Sil Corp Taunton, MA, 02780) was added to each microcosm. Seven

Pimephales were added and fed 800-900 mg of salmon starter (Aquatic Biosystems, Fort

Collins, CO). On a daily basis 1 L of treatment water in each microcosm was exchanged with freshly prepared treatment water. After exchanging the water ~800 mg of salmon starter was added to each microcosm. The study was conducted for 10 days and at its conclusion each specimen was weighed, frozen individually, and stored frozen (-20 ºC) until analysis.

At the start of the specimen collection it was noted that the Anacharis had a lot of debris attached to the leaves (Figure 21). In order to just collect the Anacharis, the plants were submerged in DI water and gently rinsed to remove the debris from the plant. After removing the debris from the Anacharis the mass of the plant was measured and the specimen was wrapped in foil and frozen until analysis.

Figure 20. Microcosm set up in incubator upon initial onset of experiment

Debris on the surface of the Anacharis in microcosm.

Figure 21. Anacharis with debris on its surface being removed from one of the microcosms.

When the Pimephales were removed they were placed in ice as to shock the specimen, according to IACUC guidelines and allowed for the blotted wet mass to be measured. After the minnow was removed from the ice they were patted dry with a low- lint wipe. The Pimephales were then weighed, wrapped in aluminum foil, and frozen until analysis. Following collection the Corbicula were padded dry with a low-lint wipe.

They were individually weighed, wrapped in foil, and frozen until analysis. Each set of specimens from a given treatment were placed together in zip top plastic bag. After all the specimens were collected, labeled and placed in the freezer the water was drained into

2 L bottles for analysis of TCC and TCS and as much as possible separated from the sand/ sediment. The sand/sediment was then transferred into trace clean amber glass jars and frozen until analysis of TCC and TCS.

Solid and Semisolid Sample Preparation

Triclosan and Triclocarban were extracted from the sediment, Anacharis,

Pimephales, and Corbicula samples by pressurized liquid extraction (PLE) on a Dionex

ASE 100 (Dionex, Bannockburn, IL, 60015, Figure 22). Pressurized liquid extraction is a common technique used to extract organic compounds from complex matrices such as sediments, soils, and biological samples (Burkhardt et al., 2005; Kinney et al., 2008,

Herklotz et al., 2010; Schultz et al., 2010).

A glass fiber filter is placed at the bottom of the high pressure extraction cell to which about 0.64 cm of ashed (4 hr at 400 ºC) Ottawa sand was added. The specimen is added to the sample cell and any void volume is filled with ashed sand to maintain consistent extraction volumes. The samples were extracted using a 70:30 acetonitrile:

H2O solvent mixture, at 130°C for 3 static cycles taking 7 minutes each at a pressure of

10342 kPa generating a final extracted volume of 22-25 mL. Two milliliters of the extract were filtered through a 13 mm 0.2 µm PTFE syringe filter (Supelco, Bellfonte,

PA 16823, USA) and the acetonitrile was evaporated off to approximately 600µL using a

Labconco N2 RAPIDVAP instrument (Labconco Corporation, Kansas City, MI, 64132).

The extract was reconstituted to approximately 1 mL using 300 µL of a phosphate buffer

13 13 (pH 2.0) and 50 µL (2 mg/L) each of C13-TCC and C12-TCS internal standard solutions. The samples were stored in refrigerator (4 ºC) until LC/MS analysis.

Figure 22. Dionex ASE-100 instrument

Microcosm Water Sample Preparation

The water samples from the microcosm were prepared using a vacuum pump and an Oasis HLB 20 cc (1 g) LP Solid Phase Extraction (SPE) Cartridge using a method similar to that described by Cahill et al. (2004). The SPE was attached to the vacuum manifold and the SPE were conditioned with HPLC grade methanol. The water samples were loaded onto the SPE cartridges under vacuum at a rate of about 20 mL/min. The

TCC and TCS was eluted with 40 mL of HPLC grade methanol. The 40 mL were placed into a LABCONCO vial and the methanol was evaporated to 600 µL using the N2

Evaporator System. The 600 µL was reconstituted to 1000 µL using 300 µL of a

13 13 phosphate buffer (pH 2.0) and 50 µL each of C13-TCC and C12-TCS internal standard solutions. The samples were stored in refrigerator (4 ºC) until LC/MS analysis.

Liquid Chromatography / Mass Spectroscopy ( LC/MS)

One LC/MS instrument that was used was HP 1100 MSD (Hewlett Packard,

California, 94304) employing electrospray ionization with a Synergy hydro-RP 80A 50 X

4.60 mm 4 µ column (Phenomex, CA, 90501) to detect the concentrations of TCC and

TCS in 25 µL‟s of the sample. A second instrument used for the baseline samples was a

Shimadzu LCMS- 2010A (Kyoto, Japan) with the same column along with an electrospray ion probe. The samples on both instruments were analyzed using a binary gradient (Table 13) of 10 mM formate buffer and acetonitrile at a flow rate of 0.400 mL/min. The mass spectrometer was operated in the single-ion monitoring mode to improve sensitivity. Three ions were monitored for each analyte and internal standard

(Table 14). The method detection limit (MDL) for TCS and TCC in soil, sediment, and

biosolid was previously determined to be 2.8 and 5.5 ng/g, respectively. The MDL for

TCS and TCC in plant tissue was 5.7 and 7.4 ng/g, respectively.

Table 13. LC/MS pump flow for data generation Time (minutes) 10 mM Formate Buffer Acetonitrile

0.00 85 15 3.00 65 35 6.00 45 55 9.00 15 85 12.00 0 100 14.00 0 100 14.10 85 15 19.00 STOP STOP

Table 14. Quantifying and Characteristic Ions Monitored for Each Analyte and Internal Standard Compound Quantifying Characteristic Characteristic

Ion Ion Ion

TCS 287 289 291 13 C12-TCS 299 301 303 TCC 313 315 317 13 C13-TCC 326 328 330

Results and Discussion

Toxicology – Triclocarban

The toxicity of TCC and effects on C. dubia were analyzed through a seven-day acute/chronic static- renewal test (USEPA, 2003; TenEyck and Markee, 2007). The test was comprised of multiple TCC concentrations and two solvent controls. Initially methanol was used as a solvent to prepare the TCC stock solution because of the low water solubility of TCC. However, the presence of methanol resulted in a carrier effect on the C. dubia, which interfered with the ability to determine the effects of TCC alone.

C. dubia in the solvent control (85.13 mM methanol) reproduced at a significantly reduced rate compared to the RECON control, 10.7 ± 3.0 and 22.6 ± 5.0 neonates per adult, respectively (p-value = 0.252, two sample t-test).

Triethylene glycol (TEG), which has hydroxyl groups much like methanol was chosen to replace methanol as the solvent for the TCC stock solution per ASTM

Standards (US.ASTM, 1993). Use of TEG as the solvent did not result in a significant difference between the average young per adult compared to the RECON control (p-value

= 0.907, two sample t-test ). The TEG control was created in RECON water in a concentration (0.073 M TEG) equivalent to the quantity of TEG in the most concentrated

TCC solution (US.ASTM, 1993). The concentrations (approximately 1.0, 3.0, 10, 30,

100, 300, and 1000 ng/L) of TCC were created through serial dilutions from a 19.00 mM

TCC stock solution. However, TCC at all these concentrations were lethal to C. dubia.

Therefore the concentrations of TCC tested were reduced to approximately 0.1, 0.3, 1.0,

3.0, 10 ng/L. These concentrations were also too high for C. dubia survival. Nonetheless, the C. dubia in the RECON and TEG standard survived and reproduced comparably in

both of these two initial tests. These two initial tests confirmed that the TEG did not have a significant effect on the C. dubia in regards to survival and reproduction of the species

(RECON vs TEG p-value 0.952, US.ASTM, 1993; USEPA, 2003). The concentrations of the test TCC solutions were further lowered to approximately 0.01, 0.03, 0.1, 0.3, and

1.0 ng/L for the remainder of the study.

The data generated from two separate seven day static renewal studies were combined for final analysis and interpretation (Appendix II). The average young per adult were graphed. (Figure 23). The chart shows that there is a greater difference between the controls and the 0.01 ng/L TCC solution with regards to the number of young the

C. dubia produced. The greatest number of young per adult for the controls minimally was 10 young per adult, whereas the amount of young in the 0.01 ng/L TCC solution did not rise above 6 young per adult. The first incline of the data signifies the first brood of young during days 3-4 which is normal for the C. dubia. The data also shows a paucity of young per adult on day 5 for each control and the TCC solution, this indicating that the females were in their second brood for each solution. The second curvature signifies the third brood that continues through day 7. All the qualities of this initial analysis have shown there is a difference in the amount of young for adult C. dubia that are imperiled in the TCC water during the 7 day static renewal test. The vertical bars represent plus or minus standard deviation.

Unfortunately, in the first study ephippia were identified after more than half of the experiment had been completed indicating the daphnids were stressed and produced males. The stress can be numerous factors, an example of a stress inducing environment is crowding in the culture (US.ASTM, 1993). The males then mate with the females and

together they produce ephippia (US.ASTM, 1993). Instead of disposing all specimens and restarting the study it was decided to complete the analysis. Having ephippia was unfortunate because that meant that the purchased female specimen culture that had been stressed to the point of producing males (US.ASTM, 1993). Males were found in all test concentrations including the RECON and TEG controls, however, only the females were compared between the first and second studies. Using Minitab‟s two sample t-test comparison there were no significant differences between either control group used in either study (RECON1 versus RECON2 p-value = 0.905, TEG1 versus TEG2 p-value =

0.974, TEG1 versus RECON1 p- value = 0.905, TEG2 versus RECON2 p-value = 0.934, two sample t-test, Table 15).

C. dubia Daily Total Average of Young per Adult per Treatment

40 35 30 25 20 RECON water 15 TEG standard 10 0.01 ng/L 5 0 0 2 4 6 8 Total Average ofYoung per Adult Day

Figure 23. C. dubia daily total average of young per adult per treatment on a daily basis in controls and TCC concentrated solutions. Only solutions in which reproduction can be compared, therefore, only the RECON control, TEG control and 0.01 ng/L solution were compared.

Table 15. Daily average of young per adult of C. dubia for RECON and TEG controls Young per Adult Day RECON1 TEG1 RECON2 TEG2 0 0 0 0 0 1 0 0 0 0 2 0.25 0 0 0 3 2.00 4.20 1.30 0.30 4 6.50 5.40 3.80 5.00 5 9.75 9.80 4.60 4.30 6 6.50 0.20 12.30 11.50 7 7.25 10.60 11.40 9.70 Total 32.25 30.20 33.40 30.80

All C. dubia in the 0.1, 0.3, and 1.0 ng/L TCC died within 24 hr of the onset of the test; leaving only those alive in TCC concentrations of 0.01 and 0.03 ng/L. In both tests C. dubia in 0.01 and 0. 03 ng/L TCC displayed a hindrance in their development compared to the organisms in the controls. Males and females in the 0.01 and 0.03 ng/L

TCC did not grow as large or move around as well as those in the controls. The females in the controls were extremely active, healthy, and produced healthy neonates.

In the first study, only two of the C. dubia in the 0.01 ng/L TCC concentration were females that having three broods of neonates. One of the females had only 8 neonates in the third brood. In the second female the number of neonates did not exceed

12 whereas in the controls the amount of neonates in the third brood exceeded 15 for at least one female. For the other females in the controls the amount of neonates in the third brood was at least 11 per female. Although the females were reproducing and the amount of young in the controls and the TCC solution overlapped, however, the quality of the neonates produced in the TCC solution appeared to be less “healthy” than the young in the controls. This indicated that there could be endocrine disruption within this

concentration that affected the healthy development of the adult, and the number and health of the young that were produced.

In the second study, all C. dubia in the 0.01 ng/L TCC concentration were females and only 7 of the 10 survived 7 days. On multiple occasions, I observed that 50% of the neonates were dead; they were counted but not compared with the data in figures.

As it appeared, either the young were born dead at birth or they were too weak to swim and filter water for food. This is an area that could be further reviewed in future work.

This could possibly be the result of TCC causing endocrine disruption in C. dubia similar to what has been observed in other aquatic life (Giudice and Young, 2010). Determining if TCC is an endocrine disruptor and its mode of action in C. dubia is a potential area for future research.

Another example of the interference in the development of C. dubia that was observed in the the 0.03 ng/L TCC solution that by the fifth day of the first study the

C. dubia in 0.03 ng/L were dead without reproducing yet they should have reproduced by day 3 or 4. During the second study, with only females, 7 females were still alive on the fifth day, but and only one reproduced (3 neonates). There was an indication the females that died in the 0.03 ng/L at days 5 and 6 were attempting to give birth when they died. Two of the 10 females appeared to be giving birth to their first brood of neonates but it died in the process. This was 2-3 days later than what is usual for female C. dubia to have the first brood. Since none of the young were alive or detached from the female it is postulated that the female birthed underdeveloped neonates, perhaps from exposure to

TCC (Figure 24). At 0.03 ng/L TCC at least half the C. dubia died within the study

period making this concentration the resulting in a median lethal concentration of a LC50 for TCC (LC50 = 0.03 ng/L).

The C. dubia that died in the 0.1 and 0.3 ng/L TCC solutions were intact with their entire exoskeleton and appendages still as one piece. However, the deceased

C. dubia in the 1 ng/L concentration appeared to be inside out with the outer exoskeleton detached from the specimen (Figure 25). The appearance of the C. dubia in 1 ng/L was similar to the incomplete molting separation (IMS) that was determined for Dimilin, a mosquito control chemical used since the 1970s (Hall et al., 1986).

Neonates on outside of deceased female C. dubia

Figure 24. C. dubia from 0.03 ng/L giving birth 2-3 days late and dying in process, no live young to report

A. B.

Figure 25 A. C. dubia in RECON and TEG controls that were healthy, B. C. dubia in 1.0 ng/L TCC concentration after 24 hours

The findings from this study have shown that TCC has an acute and chronic toxicity to C. dubia in very low concentrations. The lethal concentrations and the concentrations observed to significantly affect reproduction are well below the TCC concentrations commonly measured in WWTP‟s effluent and effluent impacted surface waters (Table 10). Adverse effects on the C. dubia were observed in TCC concentration of 0.01 ng/L resulting in a median effect concentration of 0.01 ng/L. This concentration differs from the USEPA‟s 2009 Interim report which indicates an EC50 at approximately

3.1 µg/L after a 48 hour static test for C. dubia. These results for C. dubia warrant further investigation of effects of TCC on aquatic life. The concentrations of TCC that were observed to impact C. dubia in this study are well below the no observable effect concentration (NOEC) reported by the EPA for C. dubia and TCC, 1.46 µg/L (USEPA,

2003).

Aquatic Microcosm: Bioaccumulation and Partitioning of Triclosan and

Triclocarban

All samples, Pimephales, Corbicula, Anacharis, and sediment from the control and TCC and TCS fortified microcosms were prepared and analyzed in triplicate to obtain an experimental error. Two microcosms had water fortified with TCC and TCS to a final concentration of 100 and 250 ng/L each. The concentration of TCC and TCS in the samples was determined by LC/MS analysis (Tables 16 and 17).

Table 16. TCS in microcosm samples in ng/kg dry weight (mean ± s); n = 3 Sample Baseline RECON control TEG control 100 ng/L TCS 250 ng/L TCS

Anacharis 95.64 ± 70.37 9.77 ± 1.72 17.60 ± 1.37 274.70 ± 35.27 708.21 ± 110.61

Corbicula 175.23 ± 112.92 11.57 ± 1.67 8.64 ± 1.51 13.98 ± 0.47 32.54 ± 4.41

Pimephales 8.28 ± 1.98 11.12 ± 4.32 12.63 ± 3.21 62.92 ± 39.81 213.39 ± 61.77

Sediment *ND 5.94 ± 2.22 4.33 ± 0.11 63.90 ± 8.16 189.88 ± 22.21

*ND = Not detected

Table 17. TCC in microcosm samples in ng/kg dry weight (mean ± s); n= 3 Sample Baseline RECON control TEG control 100 ng/L TCC 250 ng/L TCC

Anacharis 24.90 ± 7.14 63.98 ± 8.36 100.96 ± 13.42 266.82 ± 46.51 760.45 ± 180.15

Corbicula 350.43 ± 115.59 51.04 ± 15.85 17.05 ± 9.00 790.94 ± 31.38 1596.30 ± 173.98

Pimephales 173.32 ± 52.95 27.90 ± 11.95 19.95 ± 4.52 34.32 ± 4.97 190.84 ± 22.04

Sediment *ND 6.50 ± 2.50 2.73 ± 0.47 76.59 ± 14.57 252.58 ± 39.76

*ND = Not detected

A mass balance of TCC and TCS within the microcosms was used to account for the partitioning of each compound within the system. First, the amount of TCC and TCS found in the baseline specimens were added to the totals of each compound added to each fortified microcosm. The 100 ng/L microcosm had an initial 300 ng of each compound the renewal of the water over 10 days another 1000 ng added totaling 1300 ng for the entire study plus the amount of each compound per baseline (approximately, TCC =

724.33 ng/kg and TCS = 464.42 ng/kg) that could partition into other compartments within the microcosm, photo degrade or be removed with the daily renewal.

The 250 ng/L microcosm there was a total of 3250 ng added to the microcosm along with what was in the baseline specimens. A mass balance for TCC and TCS was determined and placed in a table (Table 18 and 19, respectively). From the data it was

determined that in the 100 ng/L microcosm 49.99% of TCC and 60.62% of TCS was removed from the microcosm through the daily water renewal. The data for the 250 ng/L microcosm shows there was 39.66% of TCC and 55.12% of TCS that was removed during the daily water renewal. The amount of TCS that is unaccounted for can indicate the possibility that the TCS did not get removed with the water, but it could have undergone photo degradation (Klamerth et al., 2011). This type of degradation has been shown to reduce the amount of TCS in a water column removing this emerging contaminant from river water samples, the glass microcosm could thereby imitate a natural environment with sunlight degradation (Klamerth et al., 2011; Morrall et al.,

2004; Sanchez-Prado et al., 2006).

Table 18. Mass balance accounting of TCC within the microcosm Microcosm 100 ng/L 250 ng/L Sample Mass balance % TCC Mass balance % TCC Sediment 34.45 45.73 Anacharis 6.492 7.530 Pimephales 0.2954 0.6684 Corbicula 8.707 6.367 Water 11.51 11.51

Table 19. Mass balance accounting of TCS within the microcosm Microcosm 100 ng/L 250 ng/L Sample Mass balance % TCS Mass balance % TCS Sediment 28.75 34.39 Anacharis 6.685 7.013 Pimephales 0.5418 0.7475 Corbicula 0.1538 0.1298 Water 65.01 65.01

The water from the TEG and 100 ng/L were tested for the quantity of TCC and

TCS. The water from the RECON and the 250 ng/L microcosms were lost when the containers containing the water were placed in a refrigerator without temperature control.

The bottles became very cold and it is assumed ice began to form resulting in the

RECON and 250 ng/L water containers burst leaving only the water for the TEG and 100 ng/L microcosms. This circumstance could have been avoided if the bottles were placed in a secondary reservoir. Nonetheless, having at least one control and one fortified water sample was beneficial for the data on the microcosms.

The amount of TCC in both water samples tested was minimal. However TCS was prevalent in both treatments (Figure 26). The vertical error bars are representative of the standard deviation from these data. These data demonstrated that the water contained

TCS whereas TCC partitioned into specimens and sediment in the environment. It was determined that the Corbicula, Anacharis and the Pimephales contained some TCC and

TCS before the onset of the study (Table 16 and 17). Despite the attempt to have „clean‟ starting material for the analysis, this proved difficult. The presence of TCC and TCS in the baseline specimens demonstrates the pervasiveness of both chemicals in the environment. Collection of the Corbicula from the Arkansas Riverbed and the presence of TCC and TCS could have come from the flow of the Pueblo Fish Hatchery drainage pipe cleaning agents.

The hatchery may use TCC and TCS containing substances in the facility that may inadvertently have been released into the river without any treatment and the

Corbicula strategically positioned under the drainage pipe accumulated the organics. The

concentration can also be in the Arkansas River water because the compounds have been found in similar rivers and waterways (Coogan et al., 2007; Klein et al., 2010; Kantiani et al., 2008; Barber et al., 2006).

The Pimephales were raised in captivity in dechlorinated tap water filtered through biofilters. The type of water that was used for the cleaning of the tanks the fathead minnow were raised in is unknown. If tap water is used there have been indications that tap water can contain a small percentage of TCC and TCS (Kantiani et al., 2008). Also if the water is from a local river or stream, as stated above, could introduce the compounds into biota through bioaccumulation (Reiss et al., 2009).

Another possibility is the compounds may be present in the food for the Pimephales. The

Anachris was also cultured in captivity and the concern regarding the type of water of the

Pimephales’ water is also true for the Anacharis. It is also a concern that the Anacharis may have unintentionally exposed to detergent or cleaning substances that contained

TCC and or TCS.

TCC and TCS in Water 80

70 g/L n 60

40 TCC 30 TCS 20

Amountof TCC in TCS and 10

0 TEG control 100 µg/L Microcosm

Figure 26. TCC and TCS in the water samples for TEG control and 100 ng/L microcosms. The error bars a re ± standard deviation.

Compared to the baseline samples, with the exception of the sediment samples, the quantity of TCC and TCS in the organisms from the RECON and TEG control microcosms decreased over the course of the 10 day experiment but there was an increase in TCC and TCS in the sediments and Anacharis (Table 16 and 17; Figure 27 and 28 respectively). The area the TCC and TCS bioaccumulated was not fully unexpected because there is evidence that TCC is more susceptible to bioaccumulate within the sediment and algae or other plant life in a natural environment (Miller et al., 2008;

Chalew and Halden, 2009). The concentrations of TCS in the Pimephales and Corbicula in the control microcosms decreased, however, it bioaccumulated within the sediment of the microcosms. This type of bioaccumulation is found when organisms excrete or release organics or toxins as waste in aquatic environments. Aquatic organisms have been

shown to contain more TCC than TCS which was also true for the microcosms (Chalew and Halden, 2009; Miller et al., 2008; Kumar et al., 2008).

TCC and TCS in Sediment

350

g/kg 300 n 250 200 150 TCC 100 TCS 50

0 Amountof TCC in TCS and Baseline RECON TEG control 100 µg/L 250 µg/L control Microcosm

Figure 27. Amount (ng/kg dry weight) of TCC and TCS in Sediment for each microcosm. The error bars are ± standard deviation.

The concentration of TCC and TCS in the Anacharis in the fortified water microcosms increased during the 10-day period relative to the control microcosms

(Figure 26). This finding substantiated that Anacharis, much like has been reported for algae, exhibited the capability of bioaccumulating both TCC and TCS (Rogers et al.,

2009, Coogan et al., 2007). Orvos et al. (2002) has reported that TCS can impair growth of aquatic plants but does not kill the plants at concentrations between 0.97- 19.1 ng/L.

However, this was not evident in the TCS fortified microcosms where the Anacharis had new growth and in some cases the mass of the plants doubled over the 10-day study. In

the 100 ng/L microcosm both chemicals were >260 ng/kg in the Anacharis. In the 250 ng/L the concentrations of both compounds were >700 ng/kg in the Anacharis.

TCC and TCS in Anacharis 1000

900 g/kg

n 800 700 600 500 400 TCC 300 TCS 200

Amountof TCC in TCS and 100 0 Baseline RECON TEG control 100 µg/L 250 µg/L control Microcosm

Figure 28. Amount (ng/kg dry weight) of TCC and TCS in Anacharis for each microcosm. The error bars are ± standard deviation.

The Pimephales accumulated TCS to a greater extent than TCC (Figure 29) The amount of TCC that was found in the baseline fish sample specimen and the 250 ng/L microcosm were similar. The Pimephales quite possibly could have reached an environmental equilibrium in the highest fortified microcosm; if the test was to have extended to allow reproduction there may have been evidence of the endocrine disruption effects of the TCC in this organism (Kantiani et al., 2008; Miller et al., 2008). The 100 ng/L microcosm may be an environment that allows the Pimephales to excrete the TCC as waste, as seen in bioaccumulation models, even though TCC is renewed daily. The

TCS detected in the baseline and control Pimephales was significantly lower than either of the fortified microcosms.

The greatest accumulation of TCC in the 10 day study was observed in the

Corbicula (Figure 28). Triclosan in the Corbicula decreased in the microcosms compared to TCS in the baseline specimen, suggesting that the Corbicula were exposed to higher concentrations of TCS than what was used in this study. The higher than expected concentrations of TCC and TCS in the baseline Corbicula may be the result of substantial in stream exposure or TCC and TCS contaminated discharge from the Pueblo

Reservoir Hatchery where they were collected. This indicates that Corbicula may represent an ideal sentinel organism for monitoring TCC and possibly TCS contamination in freshwater aquatic environments.

TCC and TCS in Pimephales

300

g/kg 250 n

200

150 TCC 100 TCS 50

0 Amountof TCC in TCS and Baseline RECON TEG control 100 µg/L 250 µg/L control Microcosm

Figure 29. Amount (ng/kg dry weight) of TCC and TCS in Pimephales per micrcosm. The error bars are ± standard deviation.

TCC and TCS in Corbicula

1800

1600 g/kg n 1400 1200 1000 800 TCC 600 400 TCS 200

0 Amountof TCC in TCS and Baseline RECON TEG control 100 µg/L 250 µg/L control Microcosm

Figure 30. Amount of TCC and TCS (ng/kg) in specimen Corbicula per microcosm. The error bars are ± standard deviation.

. The data suggest that the aquatic organisms, similar to those found in freshwater environments, can accumulate TCC and TCS from water in a short period of time.

Recommendation for future work on this project includes monitoring the reproductive cycle of the specimens used. Future researchers may consider an experimental design that includes at least 2 g wet weight of each organism in triplicate per day, so that the rate of accumulation could be measured. Nonetheless, this 10-day study did show that each specimen had the ability to accumulate TCC and TCS. The specimens used in this study were subjected to a quantity of TCC and TCS that is equivalent to an exposure that is 10 and 25% WWTP effluent. I found that TCC and TCS it in the controls the exposure time was sufficient to see a reduction in the concentrations TCC and TCS. The results of this study suggest that further studies are warranted to determine the long term effects and partitioning of TCC and TCS on aquatic organisms.

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APPENDICES

104

Appendix I: NMR spectra of sugar molecules

Mannose Spectra

A-i

A-ii

A-iii

A-iv

A-v

A-vi

A-vii

A-viii

N-acetyl-D-glucosamine NMR Spectra

A-ix

A-x

A-xi

A-xii

A-xiii

A-xiv

A-xv

A-xvi

A-xvii

A-xviii

A-xix

A-xx

A-xxi

D-galactosamine Hydrochloride NMR Spectra

A-xxii

A-xxiii

A-xxiv

A-xxv

A-xxvi

A-xxvii

A-xxviii

A-xxix

A-xxx

A-xxxi

A-xxxii

A-xxxiii

Ceriodaphnia dubia data Test 1

A-xxxiv

A-xxxv

A-xxxvi

A-xxxvii

A-xxxviii

A-xxxix

A-xl

A-xli

Ceriodaphnia dubia data Test 2

A-xlii

A-xliii

A-xliv

A-xlv

A-xlvi

A-xlvii

A-xlviii

Ceriodaphnia dubia data combined

A-xlix

A-l