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Nicotine metabolism in the cabbage looper trichoplusia ni (Hübner) Saremba, Brett 2018

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Nicotine Metabolism in the Cabbage Looper Trichoplusia ni (Hübner)  by  Brett Saremba  B.Sc., The University of British Columbia, 2015  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE  in  THE COLLEGE OF GRADUATE STUDIES  (Biochemistry and Molecular Biology)   THE UNIVERSITY OF BRITISH COLUMBIA  (Okanagan)  April 2018   © Brett Saremba, 2018 iiThe following individuals certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis/dissertation entitled:  Nicotine metabolism in the cabbage looper trichoplusia ni (hübner)   submitted by Brett Saremba               in partial fulfillment of the requirements of   the degree of  Master of Science.   Mark Rheault, Irving K. Barber School of Arts and Sciences Supervisor Susan Murch, Irving K. Barber School of Arts and Sciences Supervisory Committee Member Karen Perry, Irving K. Barber School of Arts and Sciences Supervisory Committee Member Jeff Curtis, Irving K. Barber School of Arts and Sciences University Examiner    iii Abstract  Cabbage looper (Trichoplusia ni) larvae are generalist herbivores that feed on numerous cultivated plants. Consuming plant material from diverse plant species exposes these larvae to a wide variety of plant secondary metabolites involved in chemical defense against herbivory. The ability of the cabbage looper larvae to detoxify plant secondary metabolites, such as nicotine, has been attributed to the rapid excretion via the Malpighian (renal) tubules. However, the role of metabolism prior to excretion in the detoxification of nicotine in cabbage looper larvae has not been studied. The first objective of this thesis was to develop an accurate, precise, and sensitive method for the detection and quantification of nicotine and its major metabolites in tissues, blood and feces from insects. The second objective was to determine the appropriate nicotine dose to study the sub-lethal effects of nicotine in cabbage looper larvae. The third and final objective was to determine the metabolic fate of dietary nicotine in cabbage looper larvae. Previous studies have concluded that the cabbage looper does not metabolize nicotine.  This thesis showed that the 4th instar larvae of the cabbage looper metabolized dietary nicotine into three distinct metabolites.  In addition, the time course for nicotine metabolism and excretion was found to be more rapid than previously reported for other Lepidopteran insects. Taken together these data demonstrate that cabbage looper larvae are capable of efficiently metabolizing nicotine and excreting nicotine and these derived metabolites as a part of their xenobiotic coping mechanism.   ivLay Summary  Agricultural pest insects are a significant economic burden as, they reduce the yield of economically significant crop plants. Insect pests are exposed to many different plants that can contain compounds that are toxic to insects. The most common form of insect pest control is the application of synthetic pesticides. Synthetic pesticides have many unintended off-target effects. Thus, there is a need to find alternative pest control measures. One possible alternative is the application of plant compounds that are toxic to insects. Before implementing these compounds, I must understand how insects cope with toxic plant compounds, to better exploit them. I studied how a caterpillar copes with the toxic plant compound nicotine.   vPreface  Chapter 2 is based on work completed in the Laboratory for Insect Physiology and Biochemistry at UBC Okanagan under the supervision of Dr. Mark Rheault and in Secondary Metabolites Laboratory at UBC Okanagan in collaboration with Dr. Susan Murch. Method development was completed by Brett M. Saremba with assistance by Mark R. Rheault, Susan J. Murch, and Fiona JM Tymm. All experimental work and data analysis was completed by BMS.  Chapter 3 is based on work completed in the Laboratory for Insect Physiology and Biochemistry at UBC Okanagan under the supervision of Dr. Mark Rheault and in Plant Secondary Metabolite Analytical Research Team (PlantSMART) Laboratory at UBC Okanagan in collaboration with Dr. Susan Murch. BMS completed all experimental work and data analysis.  A version of chapter 3 has been submitted for publication as the following manuscript: Saremba BM, Tymm FJM, Murch SJ, Rheault MR (2018) Nicotine metabolism in the cabbage looper (Trichoplusia ni Hübner). Journal of Insect Physiology (Submitted 01/2018).      viTable of Contents  Abstract ................................................................................................................................... iii  Lay Summary ......................................................................................................................... iv  Preface ...................................................................................................................................... v  Table of Contents ................................................................................................................... vi  List of Tables .......................................................................................................................... ix  List of Figures .......................................................................................................................... x  List of Abbreviations ........................................................................................................... xiii Acknowledgements ............................................................................................................... xv Dedication ............................................................................................................................. xvi  Chapter 1 Introduction and Literature Review ................................................................... 1 1.1 Agricultural Pest Insects ....................................................................................................... 1 1.1.1 Trichoplusia ni (Hübner) .................................................................................................. 2 1.2 Synthetic Pesticides............................................................................................................... 3 1.3 Plant Secondary Metabolites ................................................................................................. 4 1.4 Effect of Nicotine in Insects .................................................................................................. 5 1.4.1 Mode-of-Action of Nicotine ............................................................................................. 6 1.4.2 Nicotine Metabolism ........................................................................................................ 7 1.4.3 Effect of Nicotine in Multiple Insect Species ................................................................... 9 1.4.3.1 Gypsy moth (Lymantria dispar) .............................................................................. 9 1.4.3.2 Honey Bee (Apis mellifera)...................................................................................... 9 1.4.3.3 Parasitic Wasp (Cotesia congregata and Hyposoter annulipes) ............................ 10 1.4.3.4 Southern Armyworm (Spodoptera eridania) ......................................................... 10 1.4.3.5 Tobacco Hornworm (Manduca sexta) ................................................................... 11 1.4.4 Effect of Nicotine in the Cabbage Looper (Trichoplusia ni) .......................................... 11 1.5 Nicotine resistance in insects .............................................................................................. 12 1.5.1 Excretion of Nicotine in Multiple Species ...................................................................... 12 1.5.2 Oxidative Detoxification of Nicotine in Multiple Species ............................................. 14 1.5.2.1 Tobacco Whitefly (Bemisia tabaci) ....................................................................... 15  vii1.5.2.2 Common Fruit Fly (Drosophila melanogaster) ..................................................... 15 1.5.2.3 Tobacco Hornworm (Manduca sexta) ................................................................... 16 1.5.2.4 Honey Bee (Apis mellifera).................................................................................... 16 1.5.3 Other Resistance Strategies ............................................................................................ 17 1.6 Excretion and Oxidative Detoxification of Nicotine in T. ni .............................................. 18 1.6.1 Excretion of Nicotine in T.ni .......................................................................................... 18 1.6.2 Oxidative Detoxification of Nicotine in T. ni ................................................................. 18 1.7 Quantifying Oxidative Detoxification................................................................................. 19 1.7.1 UPLC-MS/MS Analysis of Nicotine and Related Metabolites ...................................... 20 1.8 Hypothesis and Objectives .................................................................................................. 22 Chapter 2 Accurate Method for the Detection and Quantification of Nicotine and its Metabolites............................................................................................................................. 23  2.1 Background ......................................................................................................................... 23 2.1.1 Alkaloid Analysis ........................................................................................................... 23 2.1.1.1 Chromatographic Variations .................................................................................. 26 2.1.1.2 Spectroscopic Variations........................................................................................ 27 2.1.2 Hydrophilic Interaction Liquid Chromatography ........................................................... 28 2.2 Materials and Methods ........................................................................................................ 29 2.2.1 Chemical Sources ........................................................................................................... 29 2.2.2 Preparation of Nicotine, Cotinine, Nicotine-N-oxide, Cotinine-N-oxide, and 4-Hydroxy-4-(3-pyridyl) butanoic acid Standards ......................................................................................... 30 2.2.3 Nicotine, Cotinine, Nicotine-N-Oxide, Cotinine-N-Oxide, and 4-Hydroxy-4-(3-pyridyl) Butanoic Acid Standard Curves ................................................................................................... 30 2.2.4 Sample Preparation ......................................................................................................... 31 2.2.4.1 Preparation of Tissue and Fecal Samples............................................................... 31 2.2.4.2 Preparation of Haemolymph Samples .................................................................... 31 2.2.5 Spiked Samples for Estimation of Nicotine, Cotinine, Nicotine-N-Oxide, Cotinine-N-Oxide, 4-Hydroxy-4-(3-pyridyl) Butanoic Acid Recovery ......................................................... 34 2.2.6 Hydrophilic Interaction Liquid Chromatography ........................................................... 34 2.2.7 Tandem Mass Spectrometry ........................................................................................... 35 2.2.8 Data Analysis .................................................................................................................. 37 2.2.8.1 Processing of Acquired Raw Data ......................................................................... 37 2.2.8.2 Quantification of Processed Raw Data .................................................................. 38  viii 2.2.8.3 Method Detection Limit ......................................................................................... 38 2.2.8.4 Method Limit of Quantification ............................................................................. 38 2.2.8.5 Statistical Analysis ................................................................................................. 41 2.3 Results ................................................................................................................................. 41 2.3.1 Chromatographic Separation of Nicotine and its Metabolites ........................................ 41 2.3.2 Optimization of Mass Spectrometry ............................................................................... 43 2.3.3 Method Detection Limit and Limit of Quantification .................................................... 53 Chapter 3 Nicotine Metabolism in 4th Instar Cabbage Looper Larvae ........................... 54 3.1 Background ......................................................................................................................... 54 3.2 Materials and Methods ........................................................................................................ 57 3.2.1 Chemicals ....................................................................................................................... 57 3.2.2 Insect Rearing ................................................................................................................. 58 3.2.3 Dietary Nicotine Exposure and Sample Collection ........................................................ 58 3.2.4 UPLC-MS/MS Detection and Quantification of Nicotine and its Metabolites .............. 59 3.3 Results ................................................................................................................................. 61 3.3.1 Dietary Nicotine Exposure ............................................................................................. 61 3.3.2 Distribution of Nicotine and Metabolites in Haemolymph, Tissues, and Frass ............. 63 3.3.3 Metabolic Fate of Dietary Nicotine ................................................................................ 65 3.3.4 Distribution Kinetics of Nicotine Metabolism ................................................................ 69 3.4 Discussion ........................................................................................................................... 72 Chapter 4 Conclusion ........................................................................................................... 80  4.1 Objective 1: Accurate Method for the Detection and Quantification of Nicotine and its Metabolites ....................................................................................................................................... 80  4.2 Objective 2: Metabolic Fate of Dietary Nicotine in 4th Instar Trichoplusia ni Larvae ....... 81 4.2.1 Dietary Nicotine Exposure ............................................................................................. 81 4.2.2 Metabolic Fate of Dietary Nicotine ................................................................................ 82 4.2.3    Time Course of Nicotine Metabolism and Excretion...........................................86 Chapter 5 Bibliography.........................................................................................................89    ixList of Tables  Table 2.1 Gradient used in the chromatographic separation of nicotine, cotinine, nicotine N-oxide, cotinine N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid.....................34  Table 2.2 Cone voltages, collision energies, and MRM transitions for nicotine and derived metabolites..............................................................................................................34  Table 3.1 The modified McMorran Grisdale insect diet.........................................................58  xList of Figures  Figure 1.1 Nicotine, a common plant alkaloid produced by plants of the Solanaceae family. Created in Chemdraw Direct (Perkin Elmer)...........................................................6  Figure 2.1 The chemical structure of A) the pyridine-derivative alkaloid, nicotine and B) the tropane-derivative alkaloid, atropine.....................................................................23  Figure 2.2 The chemical structure of the highly toxic indol alkaloid, strychnine...................23  Figure 2.3: Schematic diagram of sample preparation for analysis of nicotine and its metabolites in tissue samples (A) and fluid samples (B)......................................31  Figure 2.4 Graphical output from segmented regression of the log[nicotine] plotted vs the log(peak area) in R-studio, using the Segmented package. Dashed line represents the LOQ................................................................................................................38  Figure 2.5 Chromatographic separation of nicotine and cotinine (A) and chromatographic separation of NNO, CNO, and HYB (B). A and B represent the respective total ion count (TIC) MRM channels...........................................................................40  Figure 2.6 Optimization of capillary voltage (kV) for nicotine.............................................42  Figure 2.7 Optimization of capillary voltage (kV) for cotinine.............................................43  Figure 2.8 Optimization of capillary voltage (kV) for nicotine-N-oxide..............................44  Figure 2.9 Optimization of cone energy (V) for the three traces of nicotine: 163 > 132 (A), 163 > 130 (B), and 163 > 106 (C).......................................................................45   xiFigure 2.10 Optimization of cone energy (V) for the three traces of cotinine: 177 > 80 (A), 177 > 98 (B), and 177 > 146 (C)............................................................................46  Figure 2.11 Optimization of cone energy (V) for the three traces of NNO: 179 > 132 (A), 179 > 130 (B), and 179 > 148 (C).................................................................................47  Figure 2.12 Optimization of collision voltage (V) for the three traces of nicotine: 163 > 132 (A), 163 > 130 (B), and 163 > 106 (C)..................................................................48  Figure 2.13 Optimization of collision voltage (V) for the three traces of cotinine: 177 > 80 (A), 177 > 98 (B), and 177 > 146 (C)....................................................................49  Figure 2.14 Optimization of collision voltage (V) for the three traces of NNO: 179 > 132 (A), 179 > 130 (B), and 179 > 148 (C).........................................................................50  Figure 3.1 Kaplan-Meier curve representing Trichoplusia ni survival on diets containing various doses of nicotine.......................................................................................60  Figure 3.2 The distribution of nicotine and derived metabolites over the entire 24 h after nicotine exposure......................................................................................................................62  Figure 3.3 Nicotine and derived metabolites detected in T. ni tissue, haemolymph, and frass samples 24 h after being moved to a nicotine-free diet.........................................64  Figure 3.4 Nicotine and derived metabolites detected in T. ni the various hemolymph, tissue, and frass samples 24 h after being moved to a nicotine-free diet..........................66  Figure 3.5 Time course of nicotine in the cabbage looper......................................................68  Figure 3.6 The distribution of nicotine and derived metabolites in isolated tissues at the sampling points after nicotine exposure................................................................69  xii Figure 3.7 Three major biochemical pathways for the enzymatic modification of nicotine to its common metabolites: cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid....................................................................76  Figure 3.8 Representative UPLC-MS (MRM) chromatograms accompanied by the proposed fragmentation for nicotine and the 4 metabolites scanned………........................77  Figure 4.1 Generalized model for metabolism and excretion in T. ni....................................83  xiii List of Abbreviations   MT – Malpighian tubules CYP – The cytochrome P450 enzymes FMO – Flavin-containing monooxidase enzymes cyp – Cytochrome P450 genes fmo - Flavin-containing monooxidase genes GABA – gamma-Aminobutyric acid IPM – Integrated pest management ED50 – Effective dose for 50 % of a population LD50 – Lethal dose for 50 % of a population USD – US dollars ACh – Acetylcholine nAcHR – Nicotinic acetylcholine receptor mAcHR – Muscarinic acetylcholine receptor  MDL – Method detection limit LOQ – Limit of quantification NPLC – Normal phase liquid chromatography RPLC – Reverse phase liquid chromatography HILIC – Hydrophilic interaction liquid chromatography MS – Mass spectrometry MRM – Multiple reaction monitoring MS/MS – Tandem mass spectrometry HPLC – High performance liquid chromatography UPLC – Ultra high performance liquid chromatography TLC – Thin-layer chromatography m/z – Mass-to-charge ratio ppt – parts per trillion UV-vis – Ultraviolet-visible light spectroscopy NMR – Nuclear magnetic resonance TOF – Time-of-flight mass spectrometry  xivFT – Fourier transfer mass spectrometry QqQ – Triple quadrupole mass spectrometry Q1 – Mass analyzer 1 Q2 – Mass analyzer 2 q – Non-mass analyzer CID – Collision induced dissociation  qPCR – Quantitative real-time polymerase chain reaction NNO – Nicotine-N-oxide CNO – Cotinine-N-oxide HYB – 4-hydroxy-4-(3-pyridyl) butanoic acid S2 – Variance SD – Standard deviation RSD - % relative standard deviation RT – Retention time ESI – Electrospray ionization  FW – % Wet food weight ABC - ATP Binding Cassette    xv Acknowledgements  Thank you to The University of British Columbia and the Natural Sciences and Engineering Council of Canada for financial support of this research.  I offer my enduring gratitude to my supervisor Dr. Mark Rheault for his stellar advice, both professional and personal, and for all his help with this project. In addition, I’d like to thank Dr. Susan Murch who has been a terrific mentor throughout my time at UBCO and has provided me many fantastic opportunities. Lastly, many thanks to Dr. Karen Perry, who has been an excellent teacher and mentor since my time as an undergraduate. Thank you to Fiona Tymm for the invaluable technical support and training she provided. I appreciate all the friends I made during my time at UBCO, you are all beauties. Thank you to all my lab mates in the Facility for Insect Physiology and Biochemistry and the Plant Secondary Metabolites Laboratory. Thanks to Nadia Ighaninazhad Matanagh for help with rearing T. ni.  My friends and family have provided an immense amount of aid and encouragement, whether they realized it or not. Cheers to the boys for their assistance in many matters. I could not have done this without the succor of my girlfriend Nicole, who is an endless supply of inspiration. Special thanks are owed to my parents, who have supported to the utmost always.   xviDedication  This thesis is dedicated to my friends, family, and the caterpillars that died gloriously for this project.   1Chapter 1 Introduction and Literature Review  Insects are found in nearly every ecosystem on the planet. Current estimates of the number of insect species on earth fall between 2.75 – 10 million (Gaston, 1991; May, 1988). Over 900 thousand insect species have been classified, representing roughly 80 % of all identified species on the planet. This abundance means that insects have a large effect on the ecosystem they inhabit. Society benefits from this effect in cases where insects provide us with beneficial services such as, pollination, decomposition, and predation of unwanted organisms. In contrast, herbivorous insects can cause enormous losses in economically significant crop yield. In this chapter I will review the literature regarding agricultural pest insects, with an emphasis on Trichoplusia ni, and some of the chemical control strategies for these insects. I will focus on literature surrounding nicotine tolerance in agricultural pest insects.  1.1 Agricultural Pest Insects  Agricultural pest insects are those insects that damage agriculturally significant crop plants. Two major ways that insects damage plants are pathogen transmission and herbivory. Many insects act as plant-pathogen-vectors, facilitating the transmission of pathogens from infected to uninfected plants. For example, the Dutch elm beetle (Scolytus multistriatus) carries fungal spores that cause Dutch elm disease on its legs hairs, from infected to healthy elms trees, transmitting the disease through elm populations (Lee and Seybold, 2009). Pathogen-vector insects affect agriculture by reducing the overall health of a plant population. Herbivorous insects consume and inflict direct physical damage to plants which may lead secondarily to further pathogen infection.  Herbivorous, or phytophagous, insects can be differentiated by their feeding strategy; examples include leaf chewers, sap suckers, stem borers, root pruners, gall makers, leaf miners, and pollen or nectar collectors. Agricultural pest insects are those whose feeding strategy causes significant damage to a plant tissue, such as leaf chewers. For example, the leaf chewing larvae of the diamondback moth, a major pest of canola, incurs a worldwide  2cost between $1.3 – 2.3 billion USD annually. These yearly costs include both crop yield loss and associated pest control measures (Zalucki et al., 2012).  1.1.1 Trichoplusia ni (Hübner)  The cabbage looper Trichoplusia ni is a leaf eating agricultural pest insect. T. ni has a worldwide distribution, found any place that crucifers (Cruciferae) are cultivated. Having no diapause, this insect cannot overwinter where cold temperatures persist for long periods of time. During winter months T. ni can be found in places such as the Southern US and South-East Asia, where mean daily temperatures are more moderate. Development from egg to adult takes between 18 and 25 days when exposed to temperatures between 32 and 21 °C, respectively (Toba et al., 1973). Temperatures below 10 °C and above 40 °C can be lethal. Cabbage looper eggs are hemispherical in shape, yellowish white in color, measure 0.6 mm in diameter and 0.4 mm in height and are deposited one at a time on the upper or lower surface of a leaf. Eggs take two, three, and five days to hatch at 32, 27, and 20°C, respectively (Jackson et al., 1968). Cabbage looper larvae have 5 instars, young larvae are dusky white but become greener the older they get. In addition, at the later instars the larvae gain a white stripe along both of their sides. Larval bodies are thicker at the posterior end and narrow considerably at the anterior end. Once mature, the body measures between 3 and 4 cm long. Cabbage loopers reared on cabbage at 23 and 32 °C took 19.9 and 20.8 days, respectively, to complete larval development (Shorey et al., 1962). The larval form of the cabbage looper is the leaf eating life stage; therefore, the larvae are considered an agricultural pest insect and are the focus of most studies in this field.   Cabbage looper larvae feed primarily on crucifers, but can survive on several field crops, weeds, and flower crops (Soo et al., 1984; Vail et al., 1991). Larvae are capable of eating three times their own body weight in plant material daily (McEwen and Hervey, 1960). Their voracious appetite is what makes T. ni larvae such an effective agricultural pest insect. In addition, cabbage looper populations are capable of building resistance to common pest control strategies. There is evidence that commercial greenhouse populations of T. ni can rapidly develop resistance to the microbial insecticide Bacillus thuringiensis, leading to  3increased operating costs (Janmaat and Myers, 2003). In addition, populations of cabbage loopers were found to be resistant to the alkaloid defenses of the invasive Hemlock species Conium maculatum, a chemical defense that is highly toxic to vertebrates and pharmacologically similar to nicotine (Castells and Berenbaum, 2008). Resistance to commonly used pest control measures is one of the foremost issues in agriculture, entomology, and related fields. In subsequent sections I will highlight literature reviewing current and future directions of the chemical component of pest control. Furthermore, I will discuss the resistance to these chemical components in insects and how understanding this resistance can help make decisions regarding future pest control strategies.  1.2 Synthetic Pesticides  Synthetic pesticides are the most common chemicals in current pest control programs. There are many classes of synthetic insecticides used, including organophosphates, carbamates, organochlorines, and pyrethroids, and neonicotinoids. Neonicotinoids are the most recently developed and currently the most commonly used of these insecticide classes. Imidacloprid, a neonicotinoid, is the most widely used pesticide worldwide and has been since the early 1990s (Yamamoto, 1999). Neonicotinoids like imidacloprid have insecticidal activity because, like nicotine, they can mimic acetylcholine, acting as agonists at insect nicotinic acetylcholine receptors resulting in hyper-excitability of neurons. These compounds exhibit acute toxicity in many insect species, effectively removing large portions of an insect pest population. However, many synthetic pesticides have negative off-target effects and insect populations exposed to synthetic pesticides for prolonged periods become increasingly resistant to pesticide treatments (reviewed in Ihara et al., 2017). Off target effects occur when pest control measures negatively effect organisms other than the intended target.  Pesticides cannot be contained to agricultural areas, they can be aerosolized, solubilized, or ingested, allowing them to disperse through the atmosphere, hydrosphere, and cascade through trophic levels of the biosphere, respectively. These compounds often have low selectivity, negatively affecting neighboring ecosystems and accumulating throughout trophic levels (Damalas and Eleftherohorinos, 2011).  Due to their unintended off-target  4effects related socio-economic costs are incurred when pesticides accrue in food and water, especially in developing countries (Lopes et al., 2009). The selectivity of a synthetic pesticides often increases with the cost of the substance (Ecobichon, 2001). Agriculture in developing countries depends on less expensive synthetic pesticides, further health and economic issues (Ecobichon, 2001). There is a demand for alternatives to synthetic pesticides (reviewed in Chowański et al., 2014). In the context of agriculture, pesticides that are effective, cheap, and environmentally safe are urgently needed.  1.3 Plant Secondary Metabolites  Plants synthesize many compounds that are not required for their normal development, growth, or reproduction. These compounds are plant secondary metabolites, they serve the plant in a variety of ways. For example, plants of the Solanaceae family produce nicotine that can protect them from insect herbivory (Steppuhn et al., 2004). Plant secondary metabolites are a feasible successor to synthetic pesticides (Amoabeng et al., 2014). Alkaloids, such as nicotine, are a group of nitrogen-containing plant metabolites that exhibit many insecticidal properties. These compounds are produced in many plants and are ubiquitous in almost all plant tissues. The effects of many plant alkaloids on insects have been catalogued (reviewed in Chowański et al., 2016). These plant alkaloids often lower the attractiveness of the plant to insects. Plants can increase the concentrations of these compounds temporally and spatially in response to damage by insect pests (Wu and Baldwin, 2010). Using insecticidal plant compounds as pest control measures is a strategy that, while potentially effective, must be assessed on the molecular level before implementation.   Attributes that make plant secondary metabolites attractive pest control agents include: a range of physiological effects making it more difficult to develop resistance, relatively high selectivity, and a short half-life in most natural ecosystems (Buss and Brown, 2014). The mode-of-action of most plant secondary metabolites disrupts regular cellular and physiological function in an insect, thus disturbing homeostasis of the organism, leading to eventual death. Non-lethal changes to key structures and tissues are another way plant secondary metabolites undermine insect populations. Unlike synthetic pesticides, not all plant  5secondary metabolites result in acute mortality; sub-lethal effects can reduce organisms fitness, fecundity, and can impose a high energetic cost related to detoxification (Kliot and Ghanim, 2012). Using a compound that exhibits sub-lethal effects in pest control can be advantageous, because it reduces the risk of the pest population gaining resistance. Plant secondary metabolites represent a possible constituent of an integrated pest management program (IPM). For example, the successful implementation of plant-derived essential oils, from five different plant species including Piper callosum, Adenocalymma alliaceum, Pelargoium graveolens, and Plectranthus neochlius, are used to control a common tomato pest the silverleaf whitefly (Fanela et al., 2016). The goal of IPM is to limit the use of toxic synthetic pesticides by first using other strategies.   Biological control agents are a common IPM strategy. For example, natural predators, physical damage to pest insects, crop rotation, or insecticidal plant derived material. Biological control agents such as natural predators, physical damage to pest insects, crop rotation, or insecticidal plant derived material are common IPM strategies. For example, biological control measures are more cost efficient than synthetic pesticides; for every dollar invested in a biological control agent the investor receives $30 - $100 dollars back, compared to $4 for each dollar invested in synthetic pesticides (Pimentel, 2005).   Integrating plant secondary metabolites into IPM strategies requires a detailed understanding of the biology and molecular genetics involved in insects’ resistance to insecticides (Perry et al., 2011). Thus, in the following sections I will review literature relating to the effects of a model plant alkaloid, nicotine, on various insects. This will be followed by a review of the mechanisms by which insects garner resistance to nicotine.  1.4 Effect of Nicotine in Insects  Nicotine (Fig. 1.1) is an alkaloid produced by plants of the Solanaceae family that protects plants from insect herbivory and is toxic to higher vertebrates (Castells and Berenbaum, 2008; Krischik et al., 1991). Nicotine makes the plant less attractive to most potential insect herbivores. Furthermore, nicotine disrupts regular neurotransmission by mimicking  6acetylcholine in animals, leading to paralysis and death (Millar and Denholm, 2007). Nicotine has become a significant environmental pollutant, as environmental residue from tobacco processing contains a high level of nicotine (Briški et al., 2003). Nicotine has garnered much attention, due to the number of tobacco related human fatalities, and is well studied, making it a prime model alkaloid. Nicotine has been demonstrated to be toxic and/or repellant to insect herbivores. Dietary nicotine was shown to be toxic to the cabbage looper Trichoplusia ni (Krischik et al., 1991). Nicotine has a lower relative toxicity in insects than in vertebrates, the dose required to induce 50 % mortality (LD50) of nicotine in the common house fly is roughly five times higher than in mice (Hukkanen et al., 2005). Due to this increased resistance to nicotine toxicity, insects represent ideal models for the study of nicotine resistance and metabolism.    Figure 1.1 Nicotine, a common plant alkaloid produced by plants of the Solanaceae family. Created in Chemdraw Direct (Perkin Elmer).   1.4.1 Mode-of-Action of Nicotine  Acetylcholine is the primary excitatory neurotransmitter and agonist for rapid neurotransmission in insect central nervous systems (Matsuda et al., 2009; Thany, 2010a). After release from the presynaptic membrane acetylcholine interacts with the agonist binding site of the acetylcholine receptor ion channel complex (Thany, 2010b). The interaction results in a conformational change in the receptor and opening of the ion channel, followed by an influx of extracellular Na+ and efflux of intracellular K+. Nicotine mimics  7acetylcholine at nicotinic acetylcholine receptors (nAChRs), acting as an agonist. The low toxicity of nicotine in insects relative to mammals is largely due to small structural difference in sub-types of nicotinic acetylcholine receptors (Tomizawa et al., 2000; Tomizawa and Casida, 2009, 2005, 2003, 1999). The mechanism of acute toxicity is paralysis of the nerve centers. Chronic exposure may also cause delayed growth and lowered fertility (Gordon, 1961). The metabolism of nicotine to substrates that are not agonist of nAChRs can mitigate the neurotoxicity of nicotine. The subsequent sections will outline general nicotine metabolism and review the effects of nicotine in various insects.  1.4.2 Nicotine Metabolism  Most nicotine metabolism pathways are facilitated by Cytochrome P450 (CYP) enzymes. While nicotine metabolism has some similarities between diverse animal classes, there are distinct differences. Common nicotine metabolites found in most animals include cotinine, and nicotine-N-oxide. Common nicotine metabolites in vertebrates also include nicotine glucuronide, nornicotine, and 2’ hydroxynicotine (Hukkanen et al., 2005). In insects, additional metabolites include, oxidation products of cotinine, including cotinine-N-oxide, norcotinine, 3’-hydroxycotinine,  and 4-hydroxy-4-(3-pyridyl) butanoic acid (Du Rand et al., 2017, 2015; Kumar et al., 2014a; Self et al., 1964).  Nicotine metabolism can be mediated by multiple independent metabolic pathways. These include the 5’ C-oxidation of nicotine to cotinine and its subsequent metabolites, N-oxidation to nicotine-N-oxide, and 2’ C-oxidation to 4-hydroxy-4-(3-pyridyl) butanoic acid (Fig 3.7). Metabolism of nicotine to cotinine, nornicotine, and 2`hydroxynicotine are mediated by CYPs (Hukkanen et al., 2005). Flavin-containing monooxygenases (FMOs) mediate the transformation of nicotine to nicotine-N-oxide (ibid). The enzymes responsible for the production of 4-hydroxy-4-(3-pyridyl) butanoic acid remain to be determined (Du Rand et al., 2017).  Between insects there are further subtle differences in nicotine metabolism. For example, the major nicotine metabolite in M. sexta was cotinine-N-oxide (Kumar et al., 2014a) however, the major metabolite in A. mellifera was 4-hydroxy-4-(3-pyridyl) butanoic acid (Du Rand et  8al., 2017). Variation in nicotine metabolism in insects translates to different effects and energetic costs in different species.   91.4.3 Effect of Nicotine in Multiple Insect Species  1.4.3.1 Gypsy moth (Lymantria dispar)  A previous study (Shaw and Waranch, 2008), exposed Gypsy moth larvae to nine alkaloids, including nicotine, to determine the effects of the plant metabolites on the insects’ feeding. Diet-choice bioassays showed that nicotine had a significant deterrent effect on this generalist herbivore’s feeding behaviour. The effective dose (ED50) that decreased feeding in 50% of the larvae for the nicotine treatment was 28.3 mM. Nicotine was estimated to be the third most potent antifeedant studied, after berberine and aristolochic acid, although there was greater variance in the nicotine treatment. While L. dispar has some capacity to detoxify nicotine this insect was highly sensitive to low levels of nicotine and tended to avoid plant material containing nicotine.  1.4.3.2  Honey Bee (Apis mellifera)  Honey bees encounter trace amounts of nicotine naturally in the nectar of some plant species. Foraging bees exposed to >50ppm nectar-nicotine for a period of 15d had decreased flight habits and decreased feeding. Hatching success was not significantly affected by 50ppm nectar nicotine treatment. However, larval survival was reduced by approximately 30% in the 50 ppm nicotine treatment (Distl et al., 2006). Natural concentrations (3 – 31 µM) of nectar-nicotine had no adverse effect, but higher concentrations increased larval mortality (Köhler et al., 2012). Larvae are sensitive to ingested nicotine due to their high nutritional requirements suggesting there is a threshold for nicotine detoxification in A. mellifera (ibid).  However, it has been shown that low concentrations of nectar-nicotine increase feeding preference, suggesting that psychoactive properties of these alkaloids may be a mutualistic reward for pollinators, encouraging return to the plant (Singaravelan et al., 2005).  101.4.3.3 Parasitic Wasp (Cotesia congregata and Hyposoter annulipes)  C. congregata is a parasitic wasp whose larvae are laid on and are hosted by the larvae of the tobacco hornworm Manduca sexta. The tobacco hornworm is a specialist insect herbivore which feed on Nicotiana species containing high amounts of nicotine. Larval emergence of the parasitoid C. congregata was compared in populations of M. sexta reared on diets containing varying doses of nicotine. In populations of M. sexta reared on nicotine diets larval emergence of the parasitoid was reduced significantly, relative to the nicotine-free diet condition. Furthermore, there was a significant decrease in the number of cocoons formed by the parasitoid in the nicotine exposed population (Barbosa et al., 1986; Harvey et al., 2007).  Hyposter annulipes, is another parasitic wasp that parasitizes the larvae of the fall armyworm Spodoptera frugiperda. The effects of nicotine-fed host insects on the fitness of this parasitoid were investigated. Parasitoid mortality increased by approximately 30% in the nicotine-fed host condition (Barbosa et al., 1986). Nicotine ingested by common pest insects negatively affecting their natural parasitoids suggests that some insects repurpose insecticidal compounds for their own benefit (See section 1.5.3 Other resistance strategies).  1.4.3.4  Southern Armyworm (Spodoptera eridania)  Exposure of the southern armyworm Spodoptera eridania to a diet containing 0.5% nicotine by fresh food weight significantly decreased the amount of food ingested, reduced weight gain, and decreased relative growth rate of larvae suggesting that detoxification of insecticidal compounds imposes a significant energetic cost. In contrast  dietary exposure to α-(+)-pinene, a plant monoterpene, used by insects as a chemical communication signal and substrate of the same detoxification system as nicotine, had no significant effect on the health of the insects (Cresswell et al., 1992).      111.4.3.5  Tobacco Hornworm (Manduca sexta)  The tobacco hornworm, a specialist insect herbivore that feeds on plants of the Solanaceae family, primarily tobacco plants is the most nicotine tolerant insect (Wink and Theile, 2002). The developmental period of M. sexta was not affected by dietary nicotine (Harvey et al., 2007).  As a result, M. sexta is commonly used to study the physiological effects of nicotine. Field caught M. sexta exposed to an artificial diet containing 0.1% nicotine showed no difference in mortality, compared to control. However, a decrease in larval mass was observed (Kumar et al., 2014b). The lack of any effect on mortality shows that M. sexta larvae are indeed capable of detoxifying and excreting nicotine effectively. Decreasing larval mass suggests that there is a significant energetic cost imposed by the detoxification and excretion.  1.4.4 Effect of Nicotine in the Cabbage Looper (Trichoplusia ni)  The effect of nicotine in cabbage looper larvae remains poorly understood, relative to other insects reviewed in the preceding sections (1.4.3). T. ni larvae reared on an artificial diet containing more than 0.125 % nicotine by wet weight did not survive (Krischik et al., 1991), indicating that T. ni is more sensitive to dietary nicotine than other Lepidoptera.  However, the length of nicotine exposure was not reported for this assay. Therefore, the acute toxicity of dietary nicotine in cabbage looper larvae remains to be determined. Sub-lethal chronic exposure to dietary nicotine levels less than 0.125 % increased the number of days to pupation and decreased pupal weight (Krischik et al., 1991). While high levels of nicotine result in mortality in the cabbage looper, there is a paucity of information regarding the mechanisms by which they can survive sub-lethal chronic dietary nicotine exposure.   121.5 Nicotine resistance in insects  1.5.1 Excretion of Nicotine in Multiple Species  Excretion is a common strategy employed by many insects to remove xenobiotic compounds, such as nicotine, thus reducing the amount of xenobiotic that can reach its target receptor. Excretion is facilitated by the Malpighian tubule of an insect and membrane-bound multidrug, or ABC, transporters. Malpighian tubules are the renal organ in insects, analogous to the mammalian kidney. ABC transporters are non-specific transporters that transport a wide range of structurally diverse xenobiotic compounds and have been implicated in insecticide resistance (reviewed in Merzendorfer, 2014). These proteins are thought to play a significant role in generalist insects that are exposed to a wide variety of xenobiotics (Bretschneider et al., 2016a; Dermauw and Van Leeuwen, 2014; Labbé et al., 2011). Excretion results in rapid removal of unmetabolized xenobiotic and, potentially, its metabolites. Chemical analysis of insect excretions allows determination of the degree to which a xenobiotic is excreted in its unmetabolized or metabolized form.  Unmetabolized nicotine was detected in the feces of  M. sexta after direct injection into the haemolymph (Maddrell and Gardiner, 1975). 14C-labeled nicotine added to the bathing medium of a Ramsay preparation is taken up by the isolated MTs of M. sexta and secreted into the primary urine. The 14C-nicotine in the secreted fluid was much more concentrated than in the bathing medium, indicating that M. sexta could actively transport nicotine from the haemolymph for subsequent excretion. The ability to excrete plant alkaloids via MTs and associated organs has been demonstrated in numerous insect orders, at both larval and adult life stages, in both phytophagous and hematophagous insects. The following insect species all demonstrated a capacity to transport nicotine: Oncopeltus. fasciatus, Rhodnius prolixus, Drsophila melanogaster, Aedes aegyptii, Acheta domesticus, Locusta migratoria, Tenebrio molitor, Periplaneta Americana, and Trichoplusia ni (Rheault et al., 2006). The apparent conservation of this mechanism for removal of nicotine highlights the importance of excretion when considering how insects cope with plant secondary metabolites and in the implementation of plant secondary metabolites as pesticides.   13 Excretion in insects is key to the normal physiological state of the organism. For example, excretion facilitates osmoregulation, ion regulation, and removal of toxins. Understanding how plant derived compounds affect excretion in insects could lead to novel insect control measures (Ruiz-sanchez and O'Donnell, 2015). For instance, compounds that inhibit ATP-binding cassette transporters involved in excretion may increase the efficacy of insecticidal compounds (Merzendorfer, 2014).   141.5.2 Oxidative Detoxification of Nicotine in Multiple Species  Insects have an array of enzymes that are capable of detoxifying xenobiotics, including insecticidal compounds, to less toxic or more easily excreted derivatives or compounds (Li et al., 2007; Xu et al., 2005). Cytochrome P450 proteins (CYP) are the most studied Phase I detoxification enzymes, in this regard. In general, CYPs oxidize toxic compounds, decreasing toxicity and increasing their polarity, thus facilitating excretion or allowing the oxide to act as substrates for further Phase II detoxification reactions (Kumar et al., 2014b). Phase II reactions add charged functional groups to the substrates further increasing their hydrophilicity, increasing their water solubility and thus facilitating excretion. Phase II reactions are mediated by enzymes such as glutathione-S-transferases (GST). Phase III detoxification in insects involves an array of membrane bound transporters that transport Phase I or Phase II products from cells and across epithelia for subsequent excretion. This generic detoxification is well understood; however, hundreds of detoxification genes have been recently identified (Chung et al., 2009; You et al., 2013) and in most insects, the specific Phase I, II, and III proteins involved in detoxification in vivo are unknown.  The oxidation of nicotine in most insects is mediated by CYPs and is an effective detoxification mechanism (reviewed in Li et al., 2007). The product of 5’ C-oxidation of nicotine, cotinine and the further oxidization product cotinine N-oxide, exhibit ≤ 3% toxicity relative to nicotine when applied to the cuticle of two fly species, Luciliu caesar and Culliphoru vomitoria (Riah et al., 1997). Oxidative detoxification incurs a significant energetic cost on most species of insects. For example, A. mellifera induces Phase I detoxification enzymes to confer nicotine resistance (Du Rand et al., 2015). This has been associated with an increased high energetic cost, as indicated by the upregulation and increased expression of genes, proteins and substrates involved in the glycolytic cycle (ibid).  151.5.2.1 Tobacco Whitefly (Bemisia tabaci)  Overexpression of CYP6CM1 in the tobacco whitefly Bemisia tabaci is associated with imidacloprid resistance in two different populations (Karunker et al., 2008). Imidacloprid is a neonicotinoid and is one of the most widely used agricultural insecticides. Nicotine resistant and susceptible populations of the tobacco whitefly had significantly higher expression of the cyp6cm1 gene post nicotine exposure (Kliot et al., 2014). Taken together, these findings suggest that exposure to natural plant secondary metabolites can promote insect resistance to commercial pesticides. B. tabaci populations that have higher expression of cytochrome P450 genes (cyp) are found to be more tolerant to a wide array of insecticides; in fact, the populations with higher expression of cyp genes have become the primary population in affected agricultural systems (Guo et al., 2014). The capacity of an insect pest to induce the CYP detoxification system seems to be related to the herbivore’s success in the field.  1.5.2.2 Common Fruit Fly (Drosophila melanogaster)  Recently, the increased expression of ornithine aminotransferase transcripts was strongly associated with nicotine resistance in populations of the common fruit fly D. melanogaster. Increased expression of ornithine aminotransferase likely inhibits the accumulation of glutamate and leads to a higher gamma-Aminobutyric acid (GABA): glutamate ratio. GABAergic signaling on dopaminergic neurons is thought to decrease the activity of nicotine on acetylcholine (ACh) receptors (Passador-gurgel et al., 2007). In both nicotine-susceptible and resistant populations CYPs were associated with nicotine resistance. Marriage et al., (2014) attempted to map the loci of nicotine resistance in D. melanogaster. Their results supported the hypothesis that fruit fly larvae exposed to nicotine metabolize the xenobiotic to less toxic, excretable compounds.   161.5.2.3 Tobacco Hornworm (Manduca sexta)  The tobacco hornworm has been studied extensively, in the context of nicotine detoxification. In the past the CYP detoxification system was identified as a major route by which nicotine is metabolized (Self et al., 1964; Snyder et al., 1995, 1994, 1993). Recently, some of the specific genes and enzymes responsible for the detoxification of nicotine have been identified, including enzymes from the CYP6 and CYP9 subfamilies, such as CYP6B46 (Kumar et al., 2014a). However, there is evidence that other mechanisms, such as the excretion of unmetabolized nicotine by Malpighian tubules may play a role in M. sexta’s nicotine tolerance (Section 1.5.1).  1.5.2.4 Honey Bee (Apis mellifera)  Nectar nicotine is extensively metabolized in the honey bee A. mellifera (Du Rand et al., 2017). The primary nicotine metabolite detected in this study was 4-hydroxy-4-(3-pyridyl) butanoic acid, a product of 2`C-oxidation of nicotine (ibid). The 2`C-oxidation pathway is also employed by the aphid M. persicae to confer nicotine resistance. In the aphid, nicotine resistance has been associated with the upregulation of CYP6CY3 (Bass et al., 2013; Wink and Theile, 2002). The specific detoxification enzymes in adult honeybees have not been identified. 2`C-oxidation is not thought to play a significant role in nicotine metabolism in other insect classes.  In contrast, it is the 5’C-oxidation pathway of nicotine metabolism to cotinine and its subsequent derivatives that has been shown to be the primary pathway in Lepidoptera.  CYP6B46 upregulation has been associated with the presence of 5’C-oxidation metabolites in M. sexta (Kumar et al., 2014a).   171.5.3 Other Resistance Strategies  Insect herbivores have many specialized strategies for coping with xenobiotic compounds that are neither excretion, nor detoxification. These strategies arise from many centuries of co-evolution with their host plants (reviewed in Aniszewski, 2007). The nicotine-tolerant M. sexta is able to repurpose ingested nicotine for defense against its natural predator the wolf spider (Kumar et al., 2014b). Unmetabolized nicotine present in the haemolymph is expunged from spiracles in the outer cuticle to the headspace surrounding the caterpillar. M. sexta larvae fed nicotine are less susceptible to predation than those fed a nicotine-free diet. Nicotine halitosis is an interesting example of an insect repurposing an insecticidal compound to increase its overall fitness.  Investigating insect gut microbiota`s role in the detoxification of xenobiotics is a growing field of research. Gut microbes play a role in xenobiotic detoxification in several insect species (Hammer and Bowers, 2015). In a major pest of coffee, the coffee berry borer Hypothenemus hampei , caffeine detoxification is mediated by the insects’ gut microbiota (Ceja-Navarro et al., 2015). The implication of these findings remain unclear regarding Lepidopteran insects. Caterpillars’ guts tend to have high pH, simple structure, and rapid transit time, making them a poor environment for common insect gut microbiota. A study investigating gut microbiota in many caterpillar species found that these insects lacked host-specific, resident gut microbes; the majority of microbe species identified were transient plant bacteria (Hammer et al., 2017). Thus, it is unlikely that the gut microbiota of Lepidopteran insects, such as the cabbage looper, contribute significantly to the detoxification of nicotine or other plant secondary metabolites.   181.6 Excretion and Oxidative Detoxification of Nicotine in T. ni  1.6.1 Excretion of Nicotine in T.ni  It has long been recognized that Lepidopteran insects can excrete xenobiotic compounds present in haemolymph via their Malpighian tubules (Maddrell and Gardiner, 1975). Previous research by Self et al., (1964) has established that cabbage looper larvae are capable of excreting dietary nicotine, as evidenced by the presence of  nicotine in the feces. Rheault et al., (2006) examined the capacity of cabbage looper Malpighian tubules to secrete nicotine from haemolymph for subsequent excretion. Rapid induction of nicotine flux was observed in vitro. Of eleven species studied from nine different insect orders, the Malpighian tubules from 4th instar T. ni had the second highest nicotine secretion rate (~48 pmol-1 h-1 mm-1) (ibid). Excretion is used to cope with xenobiotic exposure; however, due to the lack of information regarding oxidative detoxification the relative contributions of these strategies remains unclear in T. ni.  1.6.2 Oxidative Detoxification of Nicotine in T. ni  There has never been a study explicitly investigating oxidative detoxification of nicotine in T. ni larvae. Data from two previous studies investigating excretion indicated that cabbage looper larvae were not capable of detoxifying nicotine and relied completely on rapid excretion of unmetabolized nicotine (Rheault et al., 2006; Self et al., 1964). However, given the growing body of evidence that insects, including ones closely related to T. ni, have many enzymes capable of detoxifying xenobiotics (reviewed in section 1.5.2 and Li et al., 2007) it is probable that T. ni is capable of metabolizing nicotine. In the next section I will discuss how oxidative detoxification in insects has been quantified in the past and how, with new technology, I can readdress the question of whether T. ni larvae metabolize nicotine.   191.7 Quantifying Oxidative Detoxification  The most common method of quantifying xenobiotic metabolism in insects has been chemical analysis of haemolymph, tissue, and excretions to detect and measure a compound and its putative metabolites (Du Rand et al., 2017, 2015; Kumar et al., 2014a; Rheault et al., 2006; Self et al., 1964). Excretions represent the end-point of both metabolism and excretion of a xenobiotic. Thus, by quantifying the amount of a xenobiotic and its metabolites one can infer the relative contribution of different metabolic pathways.   Early studies attempting to quantify nicotine metabolism in T. ni larvae relied on thin-layer chromatography (TLC) to separate nicotine and its potential metabolites and UV-spectroscopy to measure them (Rheault et al., 2006; Self et al., 1964). TLC has low resolution relative to more recent techniques, such as high and ultra high-performance liquid chromatography (HPLC; UPLC). Low resolution can lead to overlapping bands and spots, potentially reducing separation of structurally similar compounds. In comparison, UPLC can have higher resolution separation, as operational parameters such as flow rate, column oven temperature, mobile phase buffer composition, etc can be tightly controlled.   Modern detection techniques have become extremely sensitive. For example, tandem mass spectrometers (MS/MS) can detect analytes in the low parts per trillion (ppt) range (Svahn et al., 2017). It is likely that previous studies concluded nicotine was not metabolized in cabbage looper larvae because the separation of nicotine and its metabolites was difficult to detect, due to the low resolution associated with previous TLC technology. Thus, nicotine metabolism in T. ni larvae should be revaluated with higher resolution, more sensitive, modern analytical instrumentation.   201.7.1 UPLC-MS/MS Analysis of Nicotine and Related Metabolites  Ultra high-performance liquid chromatography coupled to mass spectrometry is a powerful analytical technique used for chemical analysis that separates plant chemicals by chromatography and detects individual compounds by ionization and fragmentation.  Previous methods have been developed for nicotine and its metabolites (Chang, 2013; Du Rand et al., 2017; Vlase et al., 2005; Wang et al., 2000).   Due to the oxidative nature of nicotine metabolism, its resulting metabolites are polar hydrophilic molecules. Polar and hydrophilic molecules can be difficult to separate using the popular reversed-phase liquid chromatography (RPLC). RPLC columns have a hydrophobic stationary phase; therefore, polar compounds experience little or no retention under standard mobile phase conditions. In the past, normal-phase liquid chromatography (NPLC) has been used when separating polar or hydrophilic compounds; however, NPLC depends on non-aqueous, environmentally unfriendly solvents that tend not to dissolve polar or hydrophilic compounds. Hydrophilic interaction liquid chromatography (HILIC) is an emerging type of chromatography that uses hydrophilic stationary phases. HILIC chromatography separates metabolites by complex electrochemical or vanderwall interactions and solvent partitioning between a polar stationary phase, a less-polar boundary layer coating the stationary beads and a gradient of elutents from hydrophobic to hydrophilic. Hydrophilic stationary phases allow for adequate retention of polar compounds while using semi-aqueous mobile phases that are better for the environment and thoroughly solubilize polar compounds and  HILIC eluents are more compatible with mass spectrometry than NPLC, increasing the sensitivity of UPLC-MS coupled systems, respectively (Appelblad et al., 2008).   I propose that a HILIC-MS/MS system is ideal for detecting and quantifying nicotine and its metabolites in T. ni tissue, haemolymph, and excretion samples to reassess whether this insect metabolizes nicotine. Chemicals eluted from the chromatography column are ionized by electrospray ionization and detected based on their mass-to-charge ratio (m/z). Selected ions (precursor or parent) are then exposed to high voltage under argon gas to split into fragments known as product or daughter ions. The pattern of fragmentation is characteristic  21of the metabolite and is used to identify the compounds. The most recent investigation of insect nicotine metabolism was conducted using UPLC coupled to MS/MS, highlighting the relevance of my proposed method (Du Rand et al., 2017).  221.8 Hypothesis and Objectives  The increasing volume of evidence for CYP mediated nicotine metabolism in several insect species has led me to propose the following hypothesis:   The 4th instar larvae of the cabbage looper Trichoplusia ni will metabolize dietary nicotine to one or more of its common metabolites, cotinine, cotinine-N-oxide, nicotine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid, as evidenced by the presence of these compounds in the tissue, haemolymph, or excretions of the insects.  To test this hypothesis, I propose the following three objectives:  Objective 1) To develop an LC-MS/MS method for the accurate detection and quantification of nicotine, cotinine, nicotine-n-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid in T. ni tissues, haemolymph and frass.  Objective 2)  To determine the appropriate dietary nicotine dose to study the sub-lethal effects of nicotine on xenobiotic metabolism in 4th instar T. ni larvae   Objective 3)  To determine the metabolic fate of dietary nicotine in 4th instar T ni. larvae using our newly developed LC-MS/MS method (objective 1)   Chapter 2 of this thesis addresses Objective 1 and describes the development of a novel LC-MS/MS method for the detection and quantification of nicotine and its metabolites in insects. Chapter 3 of this thesis addresses both Objectives 2 and 3 outlined above and describes the metabolic fate of dietary nicotine exposure in the larvae of the cabbage looper Trichoplusia ni.  23Chapter 2 Accurate Method for the Detection and Quantification of Nicotine and its Metabolites  2.1  Background  2.1.1  Alkaloid Analysis  Alkaloids can be considered any organic base that contains a nitrogen group. This background information will introduce the reader to the alkaloid chemical class, using well known compounds as examples. Furthermore, the author will describe how these compounds have been analyzed in the past while explaining some of the different instrumentation used for analysis. Although, the distinction between alkaloids and other nitrogen-containing compounds is not well defined in the literature, within the alkaloids class there is large variation in chemical structure and pharmacological activity. For example, nicotine (Fig 2.1A) and atropine (Fig 2.1B) are both alkaloids, nicotine is a pyridine-derivative alkaloid and atropine is a tropane-derivative. Consequently, these two compounds have markedly different pharmacological effects. Nicotine is a parasympathomimetic stimulant that acts as an agonist at most nicotinic acetylcholine receptors (nAChRs). Atropine, however, is an anticholinergic drug that acts as a competitive antagonist at most muscarinic acetylcholine receptors (mAChRs). Nicotine and atropine are both found in plants of the Solanaceae family.  Plants are the largest source of naturally occurring alkaloids, it has been estimated that between 10 and 25% of higher plants have some alkaloid content (Aniszewski, 2007.) Additionally, alkaloids have been found in some types of fungi, in marine organisms, and in animals. When initially isolated from their original sources it is common for several alkaloids to be present in the isolate. Considering many alkaloids are drugs and poisons it is vital to be able to identify which compounds are present in the isolate and have the capability of extracting each compound into separate isolations.    24Analysis and extraction of alkaloids offer economic benefit when applied to a proprietary compound. Nicotine is a good example of a proprietary compound that can be extracted from the original plant material and must be analyzed before it can be sold in the form of various tobacco products. The ability to successfully analyze plant material or extractions of such can benefit personal and public health. Consider the indol alkaloid strychnine (Fig 2.2), a compound present in the seeds of the fruits of the strychnine-tree (Strychnos nux-vomica L.) that, while highly toxic to mammals, is often used in herbal remedies in India and South-East Asia. Methods have been developed to screen strychnine containing products to protect and educate consumers. The method includes a simple ultra high-performance liquid chromatography separation coupled to ultra-violet visible spectroscopy (UV-Vis) analysis (Han et al., 2008). Analysis methods range from simple thin-layered chromatography (TLC) experiments to convoluted systems containing multiple instruments coupled to one another. These often include a modern chromatography technique coupled to a high-throughput analytical technique. Such as, nuclear magnetic resonance (NMR) and mass spectrometry (MS). However, there are variations in the broad techniques here mentioned, so I will focus on insect studies that used different variations of chromatography and mass spectrometry to analyze alkaloids.       Figure 2.1 The chemical structure of A) the pyridine-derivative alkaloid, nicotine and B) the tropane-derivative alkaloid, atropine.  25         Figure 2.2 The chemical structure of the highly toxic indol alkaloid, strychnine.  262.1.1.1 Chromatographic Variations  Early research investigating the excretion of alkaloids in various insects most often employed TLC for the detection and identification of analytes (Maddrell and Gardiner, 1975; Rheault et al., 2006). TLC is a chromatographic technique that depends on separation of analytes present in the mobile phase based on their mass as they ascend the solid phase (plate) via capillary action. For example, Self et al., (1964) studied the metabolism of nicotine in tobacco-feeding insects by exposing these insects to dietary nicotine and subsequently analyzing the frass, or feces, excreted by the insects. Nicotine and related metabolites were extracted from the frass via an acid-base extraction before being chromatographed on a TLC plate, the plate was further analyzed using UV-Vis. Contemporary research that requires the analysis of alkaloids generally exploits UPLC as the chromatographic aspect of its method. UPLC is based on the same underlying principles as in TLC, the analytes in the mobile phase will interact uniquely with the stationary phase based on some physio-chemical property. However, HPLC pumps are used to force the mobile phase (solvent and analyte mixture) into a column filled with the solid phase (solid adsorbent material) with which the analyte(s) interact. Different analytes will interact differently with the solid phase causing each compound to flow through the column at a different rate and thus become separated from one another. There are several types of HPLC that are currently used, I will outline a few examples relevant to the analysis of alkaloids in insect systems.  Normal-phase HPLC (NPLC) was one of the first types of HPLC developed and is still used in modern analyses. NPLC employs a polar stationary phase and depends on analytes affinity for said polar stationary phase to separate the compounds within the column. Non-polar, organic mobile phases are used; therefore, NPLC is best suited for analysis of compounds that are soluble in solvents such as chloroform. Tendency of an analyte to adsorb to the NPLC stationary-phase is affected by the analyte’s polarity and certain steric factors. Because steric hindrance can affect an analyte’s affinity for the stationary phase one can use NPLC when elucidating structural isomers. NPLC has been used in a rapid liquid-chromatography coupled to tandem mass spectrometry (MS/MS) method for determination  27of nicotine and cotinine in serum and saliva samples. Furthermore, the researchers observed their method had increased accuracy, precision, and sample throughput, relative to the previously used radioimmunoassay (Byrd et al., 2005).   Recent research investigating nicotine-mediated antipredator defense in the Tobacco Hornworm, Manduca sexta used reversed-phase UPLC to detect and quantify nicotine and derived metabolites during the defense response of this herbivorous insect (Kumar et al., 2014a). Reversed-phase chromatography (RPLC) is considered any chromatographic method where the solid-phase is hydrophobic. Hydrophobicity is achieved by covalently bonding alkyl chains to a standard solid phase. A hydrophobic stationary phase means that hydrophobic analytes will tend to adsorb to the column, while hydrophilic analytes will flow through and elute first. Elution of hydrophobic compounds can be accomplished by increasing the concentration of a non-polar organic solvent in the mobile phase, thus lowering the pH and decreasing hydrophobic interactions between the column and the analyte(s) (Horváth, 1976).  2.1.1.2 Spectroscopic Variations  TLC has been used in previous insect studies, to separate nicotine and the potentially present nicotine derived metabolites. UV-Vis was then used to estimate the relative quantities of nicotine and derived metabolites present in the mobile phase (Self et al., 1964). Whereas, chromatography serves to separate, detect, and identify analytes, spectroscopy and spectrometry techniques allow the researchers to quantify analytes. Clearly, the most powerful analysis methods would incorporate two or more techniques of both chromatographic and spectrometric origin. UV-Vis can be considered absorption or reflectance spectroscopy in the visible and directly adjacent spectrums. According to the Beer-Lambert Law the absorbance of a solution is directly proportional to the path length and the concentration of absorbing chemical species; thus, UV-Vis can be used as the detector in a chromatography system. In fact, analytical systems using UV-Vis as a detector have been widely used in the past and are still used to some degree presently (Yal et al., 2014).   28Recently, mass spectrometry (MS) has been used in tandem with chromatographic separation techniques as the detector. Generally, MS is a technique that ionizes analytes, then sorts the ions based on their mass-to-charge ratios and can be applied to single compound solutions or complex matrices. The basic MS procedure includes: ionization of the analyte(s), separation of ions based on mass-to-charge ratio by acceleration and application of an electro-magnetic field, detection, and displaying data as relative abundances of ion as a function of mass-to-charge ratio (Sparkman, 2001). The nicotine-mediated defense response of M. sexta has been studied by analyzing the alkaloids produced in this nicotine-feeding insect during its defense response. For analysis, nicotine and its metabolites were separated using normal-phase UPLC, this separation was done in tandem with a time-of-flight (TOF) MS analysis. TOF-MS was employed by the researches as the detector for their analysis, allowing them to quantify the identified alkaloids (Kumar et al., 2014a). The mass-to-charge ratio of an ion in a TOF-MS experiment is determined based on the amount of time it takes for the ion to be accelerated by an electric field of known strength, across the range of known length. Increasing mass decreases the velocity of the ion, while increasing the charge increases the velocity. Several types of MS are often employed in the analysis of alkaloids, including: tandem MS (MS/MS) (Vlase et al., 2005; Wang et al., 2000),  Fourier transfer (FT) MS (Dudley et al., 2010), and linear quadrupole ion trap (LTQ) MS (Du Rand et al., 2015), to name a few.  2.1.2 Hydrophilic Interaction Liquid Chromatography  The instrumentation I used to develop and run the method in the following chapter includes UPLC, specifically hydrophilic-interaction liquid chromatography (HILIC), coupled to a triple quadrupole tandem mass spectrometer. HILIC is an iteration of normal-phased chromatography that uses a hydrophilic stationary-phase and utilizes solvents also used in reverse-phased chromatography. HILIC can be considered a liquid-liquid partition chromatography; in a HILIC column, analytes elute in order of increasing polarity. HILIC has become more popular because it provides separation that is ideal for complex mixtures containing multiple charged and uncharged polar compounds; further, the solvents used usually have a high portion of acetonitrile (ACN), facilitating electrospray ionization  29coupling to MS (Irgum, 2006).  Past research has shown that HILIC can be used in the analysis of alkaloids, specifically nicotine, with high sensitivity, accuracy, and precision. Moreover, peak tailing was reduced by a factor of 1.13 when compared to identical analyses using different types of HPLC (Wang et al., 2000).  Triple-quadrupole mass spectrometry (QqQ) is a type of tandem mass spectrometry that has two mass-analyzer quadrupoles (Q1, Q3) set-up in tandem with a non-mass-analyzing quadrupole (q2) in between the other two. Q1 and Q3 are used to resolve the mass-to-charge ratios of the different ions present; whereas, q2 provides a site for collision-induced dissociation (CID). CID is a technique used to cause fragmentation of the molecular or precursor ions (e.g. nicotine) being analyzed. Fragmentation occurs when the gaseous precursor ions collide with a neutral gas (nitrogen, helium, etc). Conversion of kinetic energy results in the breakage of chemical bonds and the formation of product ions. The product ions are then analyzed by MS/MS, providing information that can facilitate structure elucidation and increase method sensitivity (Sleno and Volmer, 2004).  2.2 Materials and Methods  2.2.1 Chemical Sources  (-)-Nicotine, (-)-cotinine were purchased from Sigma-Aldrich (St. Louis, MO) and Toronto research Chemicals (Toronto, ON). Cotinine-N-oxide (CNO), nicotine-N-oxide (NNO), and 4-hydroxy-4-(3-pyridyl) butanoic acid (HYB) were purchased from Toronto Research Chemicals (Toronto, ON). MS-grade methanol and MS-grade acetonitrile were purchased from Fisher Scientific (Mississauga, ON). Centrifugal filters were purchased from VWR (Radnor, PA). QuEChERS AOAC tubes, ceramic homogenizers, buffering salt packets, and dispersive SPE-cleanup tubes were purchased from Agilent Technologies (Santa Clara, CA).    302.2.2 Preparation of Nicotine, Cotinine, Nicotine-N-oxide, Cotinine-N-oxide, and 4-Hydroxy-4-(3-pyridyl) butanoic acid Standards  Approximately 5 and 15 mg of cotinine and nicotine were weighed out, respectively. 1mL of acetonitrile was added to create a stock solution containing 5 mg/mL and 15 mg/mL cotinine and nicotine, respectively. Approximately 3 mg of NNO, CNO, and HYB were weighed out. 1 mL of E-pure H2O was added to create a stock solution containing 3 mg/mL of NNO, CNO, and HYB. 100 μL of each stock was combined and then diluted with 900 μl of acetonitrile or E-pure H2O. The resulting solutions were diluted ten-fold three times. Further dilutions were four-fold and were repeated four times to produce a seven-point calibration curve approximately ranging from 0.65 ng/μL to 3.0x10-4 ng/μL.  2.2.3 Nicotine, Cotinine, Nicotine-N-Oxide, Cotinine-N-Oxide, and 4-Hydroxy-4-(3-pyridyl) Butanoic Acid Standard Curves  Calibration standards were prepared for every run, as outlined in 2.2.3. These curves contained seven points and were used to quantify experimental data. Initial standard curves for each analyte included fifteen points. Experimental standard curves included seven points for logistical reasons, method validation curves required fifteen points.   312.2.4 Sample Preparation 2.2.4.1 Preparation of Tissue and Fecal Samples  Whole body and isolated tissue samples (midgut, Malpighian tubules…) were processed with the Agilent QuEChERS AOAC extraction kit (p/n 5982-5755CH) and the Agilent QuEChERS dispersive SPE kit (p/n 5982-5022). Workflow was adapted from an Agilent Technologies application note (Chang, n.d.). Approximately 300 mg of frozen tissue, from nicotine exposed and control insects, was weighed out into a 50 mL QuEChERS AOAC extraction tube. 15 mL of milli-Q water was added to the 50 mL extraction tube and then the tube was vortexed for 5 seconds. This solution was adjusted to pH 11 with sodium hydroxide. An AOAC extraction salt package, 15 mL acetonitrile, and two ceramic homogenizers were added to the 50 mL AOAC extraction tube. The tube was sealed and then shaken rapidly for one minute, by hand, then centrifuged at 5000 rpm for 5 minutes. A 1 mL aliquot was taken from the top acetonitrile layer of the AOAC extraction tube, transferred to a dispersive SPE tube, and vortexed for 10 seconds. The SPE tube was centrifuged at 10 000 rpm for 5 minutes. 600 μL of supernatant from the SPE tube was transferred to a 1.5 mL centrifugal filter tube. This was centrifuged at 5000 rpm for 5 minutes. A 100 μL aliquot was transferred to glass conical insert in an autosampler vial (Fig 2.3A).  2.2.4.2 Preparation of Haemolymph Samples  Haemolymph and secreted fluid samples were prepared using an extraction method adapted from Du Rand et al., (2017) and the Agilent QuEChERS dispersive SPE kit (p/n 5982-5022). A known volume of haemolymph (20 μL) was added to a 1.5 mL tube containing 500 μL of acetonitrile and 500 μL milli-Q H2O and then vortexed for 10 seconds. This was centrifuged at 10 000 rpm for 5 minutes. A 500 μL aliquot of the supernatant was transferred to a 1.5 mL centrifugal filter tube and centrifuged at 5000 rpm for 5 minutes. This solution was transferred to an AOAC dispersive SPE cleanup tube and vortexed for 10 seconds. 450 μL of supernatant was transferred to a centrifugal filter tube and centrifuged at 5000 rpm for 5  32minutes. 100 μL of the solution was transferred to a glass conical insert in an autosampler vial (Fig 2.3B).  33  Figure 2.3: Schematic diagram of sample preparation for analysis of nicotine and its metabolites in tissue samples (A) and fluid samples (B). Weigh ~300 mg tissue and add to 50 mL AOAC extraction tube with 15 mL H2O. Vortex for 5 seconds. Adjust pH to ~11 with NaOH. Add 15 mL ACN, 2 ceramic homogenizers, and 1 extraction salt Shake rapidly by hand for 1 minute. Centrifuge @ 5000 rpm for Transfer 1 mL of top ACN layer to SPE tube. Vortex 10 sec. Centrifuge at 10 000 rpm for 5 minutes. Transfer 600 μL of supernatant to 1.5mL centrifugal filter tube. Centrifuge at 5000rpm for 5 Transfer 100 μL aliquot to glass conical insert in autosampler vial. Inject on LC/MS/MS for analysis using a HILIC column. Add known volume of hemolymph/secreted fluid to 1.5 mL tube containing 500 μL ACN and 500 μL H2O. Vortex for 10 seconds. Centrifuge at 5000 rpm for 5 minutes. Transfer 800 μL supernatant to SPE tube. Vortex for 10 seconds.  Centrifuge SPE tube at 10 000 rpm for 5 minutes.  A B  342.2.5 Spiked Samples for Estimation of Nicotine, Cotinine, Nicotine-N-Oxide, Cotinine-N-Oxide, 4-Hydroxy-4-(3-pyridyl) Butanoic Acid Recovery  One of the most common techniques for determining the accuracy of a new analytical method is spike recovery (Betz et al., 2011). The spike recovery involves adding a known amount of the analyte(s) of interest to a matrix at the beginning of the sample preparation workflow. Subsequently, the matrix is analyzed, and the amount detected is compared to the amount added to give the researcher an estimate of their method’s accuracy. Spike recovery can be done in parallel with experimental samples to account for the possible presence of endogenous analyte(s). Recovery is often concentration dependent. Thus, the FDA suggests validation be done by spiking samples at 80%, 100%, and 120% the expected analyte(s) concentration (CDER, 1994). The metric used to compare expected value and measured value was detector response per mass unit of analyte. Therefore, I directly compared the response of a known amount of analyte in the standard to the amount of analyte detected in the spike recovery samples. Samples were spiked in the first step of the sample preparation workflow, after adding 15 mL of milli-Q H2O (Fig 2.3). Spiked samples contained all analytes at concentration between 0.8 ng/µL and 1.4 ng/µL. The calculated percent recoveries of nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid were 41 %, 84 %, 83 %, 37 %, and 53 %, respectively.  2.2.6 Hydrophilic Interaction Liquid Chromatography  For each sample, a 10 μL aliquot was injected on to a HILIC column (100 x 2.10 mm, 2.6 μm HILIC, Kintex (p/n: 00A-4461-AN) in a Waters I-class UPLC. Flow rate and column temperature and flow rate were set to 0.7 mL/min and 30 °C, respectively. Mobile Phase A was 10mM ammonium formate (adjusted to pH 3 with glacial acetic acid) in milli-Q H2O. Mobile phase B was 100% acetonitrile. The following gradient was used: 0.0 min = 90.0% B, 4.0 min = 70.0% B curve 6, 4.5 min = 70% B curve 6, 4.6 min = 90% B curve 6, 6.0 min = 90% B curve 6 (Table 2.1).   352.2.7 Tandem Mass Spectrometry  QqQ MS/MS data were collected with a Xevo TQ-XS Triple Quadrupole Mass Spectrometer (Waters, ON) (p/n: 720005650EN, l/n: LITR134891424). Data were collected in multiple reaction monitoring (MRM) mode for all analyses. There was a 0.75 amu resolution across all quadrupoles, with an applied span of 0.5 amu. Each MRM transition had a dwell time set to 15 ms and every channel was monitored from 0 to 6.0 minutes. Data acquisition was completed with the MRM transitions listed in Table 2.2. This instrument was always run in ES+ mode. Cone voltage, capillary voltage, and source offset were set to 20 V for nicotine and 25 V for cotinine, 1.5 kV, and 40 V, respectively. Desolvation temperature was set to 300 °C, the gas flow and cone gas flow were set to 300 L/h and 150 L/h, respectively. The collision gas used in all runs was ultra-pure nitrogen, regulated to 7 psi.                  36Table 2.1 Gradient used in the chromatographic separation of nicotine, cotinine, nicotine N-oxide, cotinine N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid. Mobile phase A and B are 10 mM ammonium formate in milli-Q H2O (pH = 3) and acetonitrile, respectively. Time Point (minutes) % Mobile Phase A % Mobile Phase B Curve 0.0 10 90 6 4.0 30 70 6 4.5 30 70 6 4.6 10 90 6 6.0 10 90 6    Table 2.2 Cone voltages, collision energies, and MRM transitions for nicotine and derived metabolites.  Compound Cone  Voltage (V) Collision  Energy (V) ES+ MS  transition (m/z)+ Nicotine 20 12 163.0 → 132.1 Nicotine 20 6 163.0 → 130.1 Nicotine 20 8 163.0 →106.0 Cotinine 25 28 177.0 → 80.0 Cotinine 25 28 177.0 → 146.0 Cotinine 25 14 177.0 → 98.0 Nicotine N-oxide 25 22 179.3 → 132.1 Cotinine N-oxide 25 15 193.2 → 162.0 4-hydroxy-4-(3-pyridyl)butanoic acid 25 18 182.2 → 164.0    372.2.8 Data Analysis  2.2.8.1 Processing of Acquired Raw Data  Processing acquired data was done with Mass Lynx V4.1 (Waters). Chromatogram smoothing was done using a 3x2 mean smooth, i.e. I applied a mean smooth with a window of 3 data points and did a total of 2 scans.  A mean smooth is achieved by applying a moving average to the raw data during integrations, this serves to increase the signal to noise ratio and clarify trends in the data. Mean smoothing is easily applied to spectrometric data and is sufficient when signal to noise ratios are relatively low. However, as baseline becomes convoluted and data approaches the MDL a polynomial or method of least squares smoothing technique, also known as the Savitzky-Golay smooth, becomes more effective and appropriate (Savitzky and Golay, 1964).  382.2.8.2 Quantification of Processed Raw Data  Raw detector responses that represent peak area are exported from Mass Lynx V4.1 (Water) to Microsoft Excel (Microsoft Office 365, version 1706) for further processing. Initially, the peak areas from the standard curve data were log transformed, the triplicate values were then averaged, and plotted against the log of the corresponding analyte concentrations. The equation of this line was used for subsequent quantification. Experimental data was considered next. Peak area was log transformed and then substituted into the equation of the line created in the previous step. The anti-log of this value gave the concentration of the analyte in the auto-sampler vial, dilution was then accounted for, before averaging the triplicate analyte concentration for all analytes in each sample.  2.2.8.3 Method Detection Limit  Method detection limit (MDL) is the lowest possible analyte concentration detectable with 99% confidence in a matrix containing said analyte. Calculation of the MDL is outlined by the EPA in the 40 CFR Appendix B to Part 136, where they give a complete workflow of the calculation and highlight possible complications (EPA, 2013). To summarize, the researcher estimates the MDL for a given compound, creates a blank sample spiked with the compound at or close to the estimated MDL, and analyses n aliquots (minimum seven) of the sample, calculates variance (S2) and standard deviation (SD). The MDL is equal to the product of the student t-test value, appropriate for 99% confidence with n replicates, and the calculated standard deviation. Furthermore, there is an optional iterative workflow to verify the previous calculations.  2.2.8.4 Method Limit of Quantification  The limit of quantification is the amount at which a compound can dependably be appointed a quantitative value (Betz et al., 2011). Practically, this is all concentrations of an analyte that  39fall within the linear range of the associated standard curve used to quantify. Isolating the linear range in our method was done using segmented regression. Segmented regression is a type of regression analysis where the independent variable, in our case analyte concentration, can be separated into distinct segments and have a line fit separately to each segment. This separation is valuable because, it allows us to determine where the linearity of our standard curve deteriorates, i.e. the LOQ. Segmented regression was done in R-studio (version 3.4.0 (2017-04-21)) using the Segmented package (version: 0.5-2.1; published: 2017-06-14; source: segmented_0.5-2.1.tar.gz). Standard curve data was plotted, and a linear regression model fit to it. The coefficients from the model were extracted and used to add a single line to the graph. A segmented regression model was created and fitted to the graph. The new segmented line was added to the graph and a dashed line was added to illustrate the LOQ (Fig 2.4), the numerical value of which is output by the breakpoint estimator function in the Segmented package.   40 Figure 2.4 Graphical output from segmented regression of the log[nicotine] plotted vs the log(peak area) in R-studio, using the Segmented package. Dashed line represents the LOQ.  412.2.8.5 Statistical Analysis  Statistical analysis of experimental data was completed in GraphPad Prism (version 5.03 (2017-12-10)) including calculation of the mean of experimental samples and the corresponding standard error about the mean (SEM). Regression analyses were completed in R-studio as reported in Section 2.2.8.4.  2.3 Results  2.3.1 Chromatographic Separation of Nicotine and its Metabolites  Two chromatographic separations were developed (Fig 2.4) based on an application note from Agilent (Chang, n.d.), first for nicotine and its metabolite cotinine and second for the three nicotine metabolites NNO, CNO, and HYB. Both displayed a minimum baseline separation that is achieved in under six minutes. Separation is preserved across several insect tissue types, including: whole body, frass, hemolymph, midgut, hindgut, and salivary glands. The results show that complete separation on our HILIC column was only possible when using an organic mobile phase of appropriate polarity. Misidentification of peaks based on retention time variability was not observed; percent relative standard deviation (RSD) for retention times (RT) of nicotine cotinine, NNO, CNO, and HYB were 1.8%, 6.5%, 0.8%, 1.4%, and 0.9%, respectively.   42   Figure 2.5 Chromatographic separation of nicotine and cotinine (A) and chromatographic separation of NNO, CNO, and HYB (B). A and B represent the respective total ion count (TIC) MRM channels. The subsequent chromatograms represent the quantification ion MRM channel for each individual compound. Nicotine (163.124 > 132.082), cotinine (177.102 > 80.109), CNO (193.2 > 162.000), HYB (182.2 > 164.000), and NNO (179.3 > 130.000).  A B  432.3.2 Optimization of Mass Spectrometry  Source parameters were optimized in the mass spectrometer for nicotine and cotinine. Parameters for CNO and HYB were taken from du Rand et al., (2017) who conducted an analysis of the same compounds with identical instrumentation under the same matrix conditions. The observed parameter maximums for nicotine, cotinine, and NNO were close or identical to those found by Du Rand et al., (2017); thus, coupled with identical instrumentation, I deemed it appropriate to use the reported values to conserve time and expense.  For nicotine, cotinine, and NNO the parameters tested included: capillary voltage, cone voltage, and collision energy (Figures 2.5 – 2.13). Collision gas flow was taken from Chang, (n.d.). Source offset and dwell time showed no significant effect on optimization. Capillary voltage was found to have a maximum at 1.50 kV for the nicotine and cotinine MS scans. For the NNO, CNO, HYB MS scan this value was 3.95 kV. Optimized source parameters and their respective MRM channels can be found in Table 2.2.  44  Figure 2.6 Optimization of capillary voltage (kV) for nicotine. Optimized for the 163.124 > 132.082 trace and found to be representative of all other nicotine traces.   45  Figure 2.7 Optimization of capillary voltage (kV) for cotinine. Optimized for the 177.102 > 80.109 trace and found to be representative of all other cotinine traces.  46  Figure 2.8 Optimization of capillary voltage (kV) for nicotine-N-oxide. Optimized for the 179.3 > 130.000 trace and found to be representative of all other NNO traces.  47  Figure 2.9 Optimization of cone energy (V) for the three traces of nicotine: 163 > 132 (A), 163 > 130 (B), and 163 > 106 (C).  48  Figure 2.10 Optimization of cone energy (V) for the three traces of cotinine: 177 > 80 (A), 177 > 98 (B), and 177 > 146 (C).  49  Figure 2.11 Optimization of cone energy (V) for the three traces of NNO: 179 > 132 (A), 179 > 130 (B), and 179 > 148 (C).  50  Figure 2.12 Optimization of collision voltage (V) for the three traces of nicotine: 163 > 132 (A), 163 > 130 (B), and 163 > 106 (C).  51  Figure 2.13 Optimization of collision voltage (V) for the three traces of cotinine: 177 > 80 (A), 177 > 98 (B), and 177 > 146 (C).  52  Figure 2.14 Optimization of collision voltage (V) for the three traces of NNO: 179 > 132 (A), 179 > 130 (B), and 179 > 148 (C).  532.3.3 Method Detection Limit and Limit of Quantification  The calculated MDL values for nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-3-(4-pyridyl) butanoic acid were: 1.25x10-4, 1.69x10-7, 9.32x10-5, 8.90x10-5, and 7.83x10-6 ng/μL, respectively. The resulting LOQ for nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid were calculated to be: 4.31 x 10-4, 2.78 x 10-4, 1.05 x 10-4, 1.51 x 10-4, and 1.23 x 10-5 ng/µL, respectively. Nicotine had the highest MDL and LOQ of all five analytes, and nicotine is the least polar of the compounds analyzed (PubChem: ChemSpider).   54Chapter 3 Nicotine Metabolism in 4th Instar Cabbage Looper Larvae  3.1 Background  The cabbage looper (Trichoplusia ni) is a generalist, leaf-eating herbivore that feeds on a wide range of plant species. Cabbage looper larvae are most often found feeding on crucifers, but can succeed on many other weed, flower, and field crops (Soo et al., 1984; Vail et al., 1991). Polyphagous insect herbivores are exposed to a wide variety of plant secondary metabolites produced in plant chemical defense against herbivory. Secondary metabolites are found in the vegetative tissues of many cabbage looper host plant species (Edward et al., 2013). Secondary metabolites, including alkaloids, non-protein amino acids, and phenolics, often have a bitter taste and toxic pharmacological effects.  Secondary metabolites positively and negatively affect insects exposed to them (Stevenson et al., 2017). For example, nectar containing secondary metabolites has been shown to increase foraging efficiency and memory of reward in honey bees (Wright et al., 2013). Secondary metabolites have been shown to reduce parasitic infections (Baracchi et al., 2015; Simone-Finstrom and Spivak, 2012). However, most plant defense metabolites are toxic to insects at high doses and repellant at low doses (Chowański et al., 2016). In addition to increasing mortality, plant secondary metabolites can reduce growth, development, and reproductive performance in insects (Cresswell et al., 1992; Fragoyiannis et al., 1998; Mekhlif, 2017). Nicotine is an example of a toxic plant secondary metabolite the cabbage looper larvae can encounter in the field. Found mainly in plants of the Solanaceae family, nicotine is a pyridine alkaloid that has been used in organic farming pest control programs as tobacco tea (reviewed in Isman, 2006). Nicotine affects the central nervous system of insects, acting as an agonist of the post-synaptic nicotinic acetylcholine receptors, akin to the mode of action of neonicotinoid insecticides (Casida and Durkin, 2013).   Nicotine is present in several common cabbage looper host plants. In one study eggplant samples contained 0.1 ppm nicotine (Castro and Monji, 1986).  Another study showed that cabbage looper larvae were able to tolerate an artificial diet containing up to 0.064% wet  55weight nicotine. Above this dose mortality and days to pupation increased, larval weight decreased (Krischik et al., 1991). Cabbage looper larvae have some mechanism that enables them to tolerate ecologically relevant (and higher) dietary doses of the toxic plant secondary metabolite, nicotine.  Excretion, or the physical removal of a substance that disrupts an organism’s metabolism, is a strategy often employed by insects to deal with toxic compounds such as nicotine. Lepidopteran insects have been shown to use excretion extensively when exposed to xenobiotic compounds (Maddrell and Gardiner, 1975). Non-specific alkaloid pumps allow the Malpighian tubules to remove alkaloids from insects haemolymph at very high rates (Kumar et al., 2014a; Snyder et al., 1994; Wink and Theile, 2002). These pumps can also be found at the insect blood-brain barrier, where they disrupt delivery of nicotine to the nicotinic acetylcholine receptors (Murray et al., 1994). Of eleven insect species from nine different orders studied, the Malpighian tubules of 4th instar cabbage looper larvae had the second highest measured nicotine secretion rate (~48 pmol h-1 mm-1) (Rheault et al., 2006). Excretion is a tolerance strategy used by both polyphagous alkaloid-intolerant and monophagous alkaloid-tolerant Lepidopterans; in addition, the monophagous alkaloid-tolerant Manduca sexta has been shown to use detoxification, or oxidative-metabolism, as an alkaloid tolerance strategy. However, detoxification is not well understood in polyphagous alkaloid-intolerant insects, such as the cabbage looper.   In addition to direct  excretion, enzymatic modification, or detoxification,  of toxic compounds making them less toxic or more easily excreted is another strategy used by insects to deal with xenobiotic compounds (reviewed in Li et al., 2007). The three most common enzyme super families involved in enzymatic modification are the cytochrome P450 monooxygenases (CYPs), glutathione transferases (GSTs), and carboxylesterases (Li et al., 2007). Nicotine tolerance in insects is often associated with the overexpression of one or several enzymes from CYP, GST, or carboxylesterase families. For example, in nicotine-tolerant lines of B. tabaci and M. persicae overexpression of CYP6CM1 and CYP6CY3 is associated, respectively, with the insects’ nicotine-tolerance (Bass et al., 2013; Kliot et al., 2014).  56A third strategy employed by insects to cope with exposure to xenobiotic compounds, especially plant secondary metabolites, is sequestration (reviewed in Nishida, 2002). Lepidopteran insects have been shown to accumulate toxic plant secondary metabolites in various tissues; thus, removing the xenobiotic from their haemolymph and making the insect less palatable for natural predators (Dixon et al., 1978; Frick and Wink, 1995; Seiber et al., 1980). Several species of Arctiid moths have been shown to extensively sequester pyrrolizidine alkaloids and their N-oxides, highlighting a synergy between the two strategies of oxidative detoxification and sequestration, for coping with xenobiotics (Nickisch-Rosenegk et al., 1990; Wink and Nickisch-Rosenegk, 1993). However, while sequestration is thought to play a predominant role in sequestration-specialist insects, such as some Arctiid moths,  it is not believed to  significantly contribute to xenobiotic detoxification in generalist insects (Nishida, 2002).  Nicotine-tolerance in the monophagous Lepidopteran M. sexta has been attributed to the overexpression of CYP6B46 (Kumar et al., 2014a). Evidence of oxidative detoxification as a dietary nicotine coping strategy in M. sexta is indicated by the detection of common nicotine metabolites in the excretions of the insects (Kumar et al., 2014a; Snyder et al., 1994; Wink and Theile, 2002). In addition to excretion in frass it has been demonstrated  that M. sexta is able to exude unmetabolised nicotine from its cuticular spiracles, rendering the insect less palatable and  thus less susceptible to natural predation (Kumar et al., 2014).   Metabolism as a defense strategy to xenobiotic exposure in polyphagous nicotine-intolerant insects has not been well studied. Two studies have provided limited information regarding the role of metabolism in coping with dietary nicotine in cabbage looper larvae. Both studies  concluded that cabbage looper larvae only used excretion, but not metabolism, to cope with dietary nicotine (Rheault et al., 2006; Self et al., 1964). However, the methods used to detect nicotine metabolites in these studies had low sensitivity and resolution relative to modern analytical techniques.  These previous studies examined nicotine metabolism in cabbage looper larva excretions only. Alkaloid detection was done with paper chromatography in combination with UV- 57spectroscopy (Self et al., 1964) and thin-layer chromatography  (Rheault et al., 2006). TLC has lower resolution than, for example, HPLC. Low resolution can lead to overlapping bands and spots, making differentiation between compounds difficult. In comparison, separation is often much clearer in HPLC columns as parameters such as flow rate, column oven temperature, mobile phase buffer composition, and elution gradient can be controlled. In addition, HPLC provides many more detection methods than TLC. Modern detection techniques, such as MS/MS make analyte detection possible in the low ppt range (Svahn et al., 2017). It is possible that past studies investigating nicotine metabolism in cabbage looper larvae were confounded by poor separation due to the low resolution of TLC techniques, leading to the false conclusion that nicotine was not metabolized in this insect.  Thus, nicotine metabolism in cabbage looper larvae needs to be reassessed using analytical techniques with higher resolution and sensitivity.  Nicotine tolerance in cabbage looper larvae has been associated with rapid removal from the hemolymph by the Malpighian tubules and rapid excretion. In the tobacco hornworm nicotine tolerance has been linked to both rapid excretion and oxidative detoxification by cytochrome P450 enzymes. However, the role oxidative detoxification plays in the nicotine tolerance of the cabbage looper is neither well understood, nor well studied. The aim of the present study is to understand the metabolic fate of dietary nicotine in 4th instar cabbage looper larvae using UPLC-MS/MS analysis to confirm the presence or absence of common nicotine metabolites as well as determine the time course of potential nicotine metabolism.  3.2 Materials and Methods  3.2.1 Chemicals  All chemicals used were analytical grade unless otherwise stated. Acetonitrile Optima® used for UPLC was purchased from Fisher Scientific (Hampton, NH, US). (-)-Nicotine and (-)-cotinine were purchased from Sigma-Aldrich (St. Louis, MO, USA). (1’S, 2’S)-Nicotine 1’Oxide, (S)-cotinine N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid were purchased from Toronto Research Chemicals (Toronto, ON, CA). Extractions and solutions were made  58up with E-pure water from a Milli-Q® Synthesis water purification system made by Millipore Sigma (Etobicoke, ON, CA) unless otherwise expressed.  3.2.2 Insect Rearing  Trichoplusia ni eggs were obtained from Insect Production Services (Sault Ste. Marie, ON, CA). Eggs were received in 22 mL plastic rearing cups with cardboard lids containing 20 - 30 eggs and approximately 5 mL of insect diet. Upon receipt the cups were placed upside down (food side up, lid side down) in an incubator (CU-36L4, Percival®; Perry, IA, US) at 27 °C, 60 % RH, and 18:6 l:d. Seven days after receiving the eggs insects were removed from the original cups and placed in new cups containing approximately 5 mL of a revised McMorran diet (Hervet et al., 2016) at a density of 3 insects per cups, to avoid cannibalism.  3.2.3 Dietary Nicotine Exposure and Sample Collection  4th instar larvae were placed in 24-well culture plate (VWR, Radnor, PA). Each well contained 1.5 mL of nicotine-containing (1% FW) revised McMorran diet (Table 3.1) at a density of 1 insect per well. The larvae stayed on the nicotine-containing diet for 24 hours then were switched to a nicotine-free diet for an additional 24 hours. Hemolymph (45 µL), midgut, hindgut (ileum, Malpighian tubules, and rectum), remaining carcass, and frass (excretions) samples were collected from 4 randomly selected larvae from both the control and nicotine exposed plates. Samples were collected 0, 3, 6, and 24 hours after being switched back to the nicotine-free diet. The Malpighian tubules, ileum, and rectum were collected together as the hindgut sample. Hemolymph was collected by making a small incision on the ventral side of each larva, pressuring both ends of the insect, and collecting the resulting droplet with a micropipette. The control diet used was the revised McMorran diet. Control insects changed plates following the same schedule as nicotine-exposed insects.  59Table 3.1 The modified McMorran Grisdale insect diet. Ingredient Volume (mL) Mass (g) E-pure H2O 840  Agar  17.36 Alphacel  5 Casein  35 Wesson’s Salt Mix  10 Sugar  35 Toasted wheat germ  30.69 Choline chloride  1 Ascorbic acid  4 Formalin (37 % formaldehyde) 0.5  Methyl paraben  1.5 Aureomycin  2.1 Raw linseed oil 5  Vitamin solution 10  Anti-fugal spray - - Potassium hydroxide 5    3.2.4 UPLC-MS/MS Detection and Quantification of Nicotine and its Metabolites  The extraction method was adapted from a QuEChERS for HILIC LC/MS/MS method (AOAC 2007.01) from Agilent Inc. ® (Chang, 2013). Nicotine and its metabolites were homogenized and extracted in a buffered acetonitrile-water extraction then centrifuged for 5 minutes at 5000 rpm. The supernatant from the extracts was collected and vortexed on an SPE-cleanup column (AOAC 2007.01) to remove pigments, lipids, and proteins, then centrifuged for 5 minutes at 10 000 rpm. Supernatant from the clean extracts was collected and filtered through 0.2 µm centrifugal filters to remove particulates.   60Experimental extracts were analyzed on a Waters® I-Class ACUITY UPLC™ (Milford, MA, US) coupled to a Waters® Xevo™ TQ-S tandem quadrupole mass spectrometer (Milford, MA, US) using MassLynx™ software. Chromatographic separation of nicotine and its metabolites were achieved using a Kintex™ HILIC column (100 x 2.10 mm, 2.6 μm, 100 Å; p/n: 00A-4461-AN) purchased from Phenomenex®. Flow rate was set at 0.7 mL/min and column temperature kept at 30 °C. Mobile phase A was 10 mM ammonium formate in E-pure water (adjusted to pH 3 with acetic acid) and mobile phase B was acetonitrile. The following gradient was used: 0.0 min = 90.0% B, 4.0 min = 70.0% B curve 6, 4.5 min = 70% B curve 6, 4.6 min = 90% B curve 6, 6.0 min = 90% B curve 6. 10 µL of each experimental sample was injected on to the column. The mass spectrometer was operated in electrospray positive mode, data collection was done in multiple reaction monitoring (MRM) mode. Two scans were completed for each sample, one separating and detecting nicotine and cotinine; the other nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid. The subsequent electrospray ionization (ESI) conditions were held constant for the nicotine and cotinine scan: capillary voltage 1.50 kV; source offset 40 V; cone gas (nitrogen) 788 L/h; cone gas (nitrogen) 51 L/h; and collision gas flow rate 0.2 mL/min. For nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid scan capillary voltage was 3.95 kV and the rest of the parameters were the same as the nicotine-cotinine scan. Analyte specific cone voltages, collision energies, and MRM transitions are provided in Table 2.2.   613.3 Results  3.3.1 Dietary Nicotine Exposure  Trichoplusia ni survivorship on a nicotine-containing diet was assessed by exposing insects to increasing doses of dietary nicotine ranging from 0 - 1.0 % nicotine by wet food weight (FW) (Fig. 3.1). Insects fed the highest dose (1.0 % FW) had the highest mortality at the end of the 120 h exposure period, with 93 % dead. Control insects fed the nicotine-free diet had the lowest mortality, with 7 % dead at after 120 h of exposure. After 24 h of exposure to nicotine there were no significant differences in mortality, regardless of nicotine dose.  Thus, I determined that exposing insects to a dietary dose of 1.0 % FW nicotine for 24 h was appropriate to study the sub-lethal effects of nicotine on metabolism in the cabbage looper, denoted by the vertical dashed line on Fig 3.1.   62 0501000 % nicotine0.25 % nicotine1.0 % nicotine24 48 72 96 120HoursPercent survival Figure 3.1 Kaplan-Meier curve representing Trichoplusia ni survival on diets containing various doses of nicotine, ranging from 0 % FW to 1.0% FW. Data represents one experiment conducted over 120 h (n = 15).  633.3.2 Distribution of Nicotine and Metabolites in Haemolymph, Tissues, and Frass  The concentrations of nicotine and its metabolites were measured in extracts of isolated tissues, haemolymph and frass of T. ni which had been exposed to 1.0 % dietary nicotine by food weight for 24 h.  The isolated tissues studied included salivary glands, midgut, hindgut (ileum and Malpighian tubules), and carcass, in addition to haemolymph and frass samples. Common Phase I nicotine metabolites identified in insects (Lepidoptera and Hymenoptera) are cotinine, cotinine-N-oxide, nicotine-N-oxide, and 4-hydroxy-4-(3-pyridyl)-butanoic acid (Du Rand et al., 2017; Kumar et al., 2013).  Nicotine and nicotine metabolites were detected in all sample types analyzed over the 24 h sampling period (Fig. 3.2). The compounds detected included: nicotine, cotinine, nicotine-N-oxide, and cotinine-N-oxide. Frass samples contained 98 % of the total nicotine and derived metabolites detected over the entire 24 h sampling period. Haemolymph samples contained the smallest portion of nicotine and derived metabolites (0.1 %). Evidently, the two sample types that encompass the digestive tract, midgut (0.6 %) and hindgut (0.3 %), contained the second and third largest proportions of nicotine and derived metabolites (Fig. 3.2).   64HaemolymphSalivary glandsMidgutHindgutCarcassFrass0120406080100% of total nicotine and derived metabolites Figure 3.2 The distribution of nicotine and derived metabolites over the entire 24 h after nicotine exposure. Nicotine, cotinine, and cotinine-N-oxide were present in all sample types. Nicotine-N-oxide was only detected in frass samples. No 4-hydroxy-4-(3-pyridyl)-butanoic acid was detected in any sample types. Data represents the mean of four experiments ± SE and are expressed as % of total nicotine and derived metabolites detected. (n = 4; pooled samples of 3 randomly selected insects).   653.3.3 Metabolic Fate of Dietary Nicotine  Cotinine, the product of the 5’ C oxidation of nicotine, was the most abundant metabolite detected over the entire 24 h sampling period in all sample types, comprising 84% of the total nicotine and its derived metabolites (Fig. 3.3). Unmetabolized nicotine was the second most abundant compound at 16% of total nicotine and its derived metabolites. Nicotine-n-oxide and cotinine-N-oxide were detected in trace amounts. No 4-hydroxy-4-(3-pyridyl)-butanoic acid was detected in any samples tested.   66 NicotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.0000.0250.05020406080100% of total nicotine and derived metabolites Figure 3.3 Nicotine and derived metabolites detected in T. ni tissue, haemolymph, and frass samples 24 h after being moved to a nicotine-free diet. Data represents the mean of four experiments ± SEM and are expressed as % of total nicotine and derived metabolites detected. (n = 4; pooled samples of 3 randomly selected insects). HyPyBut: 4-hydroxy-4-(3-pyridyl)-butanoic acid. Where error bars are not visible they are smaller than the lines used. Where no bars are indicated there was no detection of the compound of interest and is 0 percent.  67I also examined the percent of nicotine and its metabolites found in isolated haemolymph, tissue, and frass samples (Fig 3.4). In the midgut, I detected nicotine (3 %), cotinine (97 %), and cotinine-N-oxide (0.004 %), indicating that nicotine is absorbed by the midgut epithelium and further metabolized in the midgut epithelium (Fig. 3.4A). Similar relative percent distributions of nicotine, cotinine, and cotinine-N-oxide were measured in hindgut (Fig. 3.4C) and salivary gland (Fig. 3.4D). In all remaining tissues, represented by the carcass sample, nicotine (16%), cotinine (84%) and trace amounts of cotinine-N-oxide (Fig 3.4E) were detected (Fig 3.4E). Haemolymph samples had a relatively higher proportion of un-metabolized nicotine at 30%, while the major nicotine metabolite cotinine constituted 70% of total nicotine derived metabolites detected. Haemolymph had trace amounts of cotinine-N-oxide as well (Fig 3.4B). Frass samples, representing final excretion and metabolism of nicotine, contained un-metabolized nicotine (16%), cotinine (84%), and cotinine-N-oxide (0.04%) (Fig. 3.4F). In contrast to all other samples, nicotine-N-oxide; a product of N-oxidation of nicotine, was detected in trace amounts (0.001%) in the frass as well (Fig. 3.4F).  No 4-hydroxy-4-(3-pyridyl)-butanoic acid, the product of the 2’ C-oxidation of nicotine, was detected in any samples tested (Fig 3.4A-F).  68 NicotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.0000.00520406080100120Midgut% of total nicotine and derived metabolitesNicotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.000.0520406080100Haemolymph% of total nicotine and derived metabolitesNicotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.0000.00820406080100Hindgut% of total nicotine and derived metabolitesNicotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.0000.01520406080100120Salivary glands% of total nicotine and derived metabolitesNicotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.000.0120406080100Carcass% of total nicotine and derived metabolitesNictotineCotinineNicotine-N-oxideCotinine-N-oxideHyPyBut0.000.020.040.062060100Frass% of total nicotine and derived metabolitesA BCE FD Figure 3.4 Nicotine and derived metabolites detected in T. ni the various hemolymph, tissue, and frass samples 24 h after being moved to a nicotine-free diet. Data represents the mean of four experiments ± SE and are expressed as % of total nicotine and derived metabolites detected. (n = 4; pooled samples of 3 randomly selected insects). HYB: 4-hydroxy-4-(3-pyridyl)-butanoic acid.  69In midgut samples the absolute amounts of nicotine, cotinine, and cotinine-N-oxide detected were 2.8 x 10-2 ± 6.7 x 10-3, 0.94 ± 0.23, and 2.9 x 10-5 ± 6.5 x 10-6 μmol/g of tissue, respectively. The same compounds were found in haemolymph samples at the following concentrations 59.3 ± 18.5, 140.0 ± 38.2, and 3.5 x 10-2 ± 2.2 x 10-2 μmol/L, respectively (Supplementary Table 1).  3.3.4 Distribution Kinetics of Nicotine Metabolism  During the 24 h sampling period the frass always contained the highest amount of unmetabolized nicotine and derived metabolites. Nicotine and derived metabolites all decreased markedly over the 24 h sampling period (Fig. 3.5). In fact, a 99 % decrease in nicotine and its metabolite content was observed 6 h after being switched to a nicotine-free diet. Notably, there was an increase in detected nicotine and derived metabolites from the 3 h (1.6 %) to the 6 h (8.4 %) time point in carcass samples suggesting some degree of xenobiotic sequestration in the insect fat body.  Figure 3.6 illustrates the kinetic distribution of nicotine and its metabolites in isolated haemolymph, tissue, and frass samples. Frass contained the largest portion of nicotine and metabolites at all time points. The 0 h timepoint represents the nicotine and metabolite content immediately after being removed from the nicotine-containing diet.     70 012204060801000 3 6 24Sampling timepoint (hrs post nic exposure)% of total nicotine and derived metabolites Figure 3.5 Time course of nicotine in the cabbage looper. Nicotine and derived metabolites detected and quantified during the 24 h sampling period. Cotinine was the major metabolite (84 %). Nicotine-N-oxide and cotinine-N-oxide were detected at trace levels. 4-hydroxy-4-(3-pyridyl)-butanoic acid was not detected in any samples. Data represents the mean of four experiments ± SE and are expressed as % of total nicotine and derived metabolites detected. (n = 4; pooled samples of 3 randomly selected insects).    71    Midgut0 3 6 240.00.20.40.60.8Time (h)% of total nicotine and derived metabolitesHaemolymph0 3 6 240.00.20.40.60.8Time (h)% of total nicotine and derived metabolitesHindgut0 3 6 240.00.20.40.60.8Time (h)% of total nicotine and derived metabolitesSalivary glands0 3 6 240.00.20.40.60.8Time (h)% of total nicotine and derived metabolitesCarcass0 3 6 240.00.20.42060100Time (h)% of total nicotine and derived metabolitesFrass0 3 6 24020406080100Time (h)% of total nicotine and derived metabolitesA BCE FD Figure 3.6 The distribution of nicotine and derived metabolites in isolated tissues at the sampling points after nicotine exposure. Nicotine, cotinine, and cotinine-N-oxide were present in all sample types. Nicotine-N-oxide was only detected in frass samples. No 4-hydroxy-4-(3-pyridyl)-butanoic acid was detected in any sample types. Data represents the mean of four experiments ± SE and are expressed as % of total nicotine and derived metabolites detected. (n = 4; pooled samples of 3 randomly selected insects).  723.4 Discussion  Oxidative detoxification of nicotine was observed in 4th instar cabbage looper larvae. Cotinine represented over 80 % of the total nicotine and derived metabolites detected during the 24 h sampling period post nicotine exposure (Fig. 3.2). Haemolymph, tissue, and frass samples all contained nicotine metabolites; cotinine was the major metabolite in all sample types. 99 % of nicotine and its metabolites were cleared 6 h after being placed on the nicotine-free diet (Fig. 3.5). In contrast, M. sexta larvae fed a comparable dietary dose of nicotine still had detectable levels of nicotine and its metabolites sequestered in tissues 48 h after being switched to a nicotine-free diet (Snyder et al., 1994). The time course of nicotine and derived metabolite clearance in cabbage looper larvae was faster than expected. Over 98 % of the total nicotine and metabolites detected over the 24 h sampling period were found in frass samples. Cotinine was the major metabolite found in frass samples (84 %), low levels of nicotine (16 %) were also detected, and trace levels of nicotine-N-oxide (0.001 %) and cotinine-N-oxide (0.04 %). However, the product of 2’C oxidation of nicotine, 4-hydroxy-4-(3-pyridyl)-butanoic acid (Fig. 3.7), was not detected in any samples types, whereas it was the major metabolite found in honey bees fed nectar nicotine (Du Rand et al., 2017).   Several possible routes for the elimination of nicotine exist in cabbage looper larvae.  Ingested nicotine first reaches the midgut, at the beginning of the digestive tract. Nicotine detoxification in M. sexta is thought to occur in the midgut (Snyder et al., 1995, 1993; Stevens et al., 2000). I found evidence of nicotine detoxification in the midgut of 4th instar cabbage looper larvae. 97 % of the total nicotine and metabolites detected in midgut samples over the 24 h sampling period was cotinine, the product of 5’C oxidation of nicotine. Trace levels of cotinine-N-oxide (0.004 %) were detected in midgut samples. Nicotine detoxification is facilitated by multidrug or ABC transporters. Studies have shown that insect midgut tissue contains high levels of ABC transporters (Bretschneider et al., 2016a; Dermauw and Van Leeuwen, 2014; Hawthorne and Dively, 2011; Labbé et al., 2011; Lanning et al., 1996; Liu, 2011; Merzendorfer, 2014). These transporters facilitate the movement of organic compounds, like nicotine, into midgut epithelial cells, where oxidative metabolism can occur. Nicotine and its metabolites can then be transported back, across the  73apical membrane to the midgut, or further across the basal membrane to the haemocoel. The presence of nicotine metabolites in midgut samples provides support for this pathway. Nicotine and its metabolites did not accumulate to a large degree in midgut samples. The level of nicotine and its metabolites in midgut samples decreased steadily over the 24 h sampling period (Fig. 3.6A), suggesting transport to the haemocoel or rapid transit through the digestive tract and subsequent excretion.   Transcellular diffusion of neutral nicotine is a likely path for nicotine moving from midgut to haemocoel. Diffusion is possible because insect midgut epithelium lacks a cuticle, making it permeable to small organic compounds (Huang et al., 2015; Klein et al., 1996; Sacchi and Wolfersberger, 1996; Turunen and Crailsheim, 1996). In addition, nicotine can take the paracellular route across the epithelium (Huang et al., 2015).  I detected nicotine in haemolymph samples, providing support for transepithelial diffusion to the haemocoel. Nicotine metabolites could travel from midgut to haemocoel via the same diffusion pathway, however, because they are more polar than nicotine the diffusion would likely be slower. Of all sample types measured, the highest nicotine level (30 %) was found in haemolymph samples (Fig. 3.4B). Cotinine (69 %) and cotinine-N-oxide (0.02 %) were also present in haemolymph samples, suggesting that nicotine and its metabolites are transported across the apical and basal membranes of the midgut epithelium.  Many Lepidopteran insects can remove xenobiotics from their haemolymph and store the compounds in various tissues, such as fat-body, a strategy known as sequestration (Dixon et al., 1978; Frick and Wink, 1995; Nishida, 2002; Seiber et al., 1980). In fact, studies have shown that moth larvae are capable of sequestering plant secondary metabolites, like the pyrrolizidine alkaloids, reducing xenobiotic content in their haemolymph and making them less palatable for natural predators (Nickisch-Rosenegk et al., 1990; Wink and Nickisch-Rosenegk, 1993). 0 h after nicotine exposure carcass samples contained less than 0.2 % of the total nicotine and derived metabolites; however, 6 h post nicotine exposure the carcass samples contained 27 % of the total (Fig. 3.6E). The increasing relative proportion of nicotine and derived metabolites in carcass samples from the 0 h to the 6 h sampling point suggests that cabbage looper larvae can sequester nicotine and its metabolites to some  74degree. There is evidence for carrier-mediated sequestration in Lepidopteran insects (Frick and Wink, 1995). Nicotine and its metabolites were not detected in haemolymph samples 6 h post nicotine exposure (Fig. 3.6B). Rapid clearance from the haemolymph coupled with an increase of nicotine and its metabolites in the carcass at 6 h suggests that nicotine and its metabolites move passively and by some carrier-mechanism from the haemolymph to the fat-body, or some other tissue in the carcass sample.   Cabbage looper larvae are able to remove nicotine from their haemolymph by inducing rapid excretion via their Malpighian tubules (Rheault et al., 2006). The removal of organic compounds, like nicotine, is facilitated by the same multidrug transporters that facilitate xenobiotic detoxification in the midgut. Multidrug transporters are abundant in the Malpighian tubules of most insects (Dermauw and Van Leeuwen, 2014). After uptake by the Malpighian tubules it is possible that nicotine returns to the digestive tract, entering the ileum, then the rectum, before being excreted in the frass. Nicotine (5 %), cotinine (95 %), and cotinine-N-oxide (0.007 %) were detected in hindgut samples (Fig. 3.4C), suggesting that nicotine does return to the digestive tract after being removed from the haemolymph. The relative proportion of cotinine is higher in hindgut samples than in haemolymph samples, suggesting cotinine is removed from the haemolymph more rapidly than nicotine. Once nicotine and its metabolites have returned to the digestive tract they can be excreted, permanently removing them from the organism.   Frass samples contained the three compounds found in all other samples types: nicotine (16 %), cotinine (84 %), and cotinine-N-oxide (0.04 %). In addition, trace levels of nicotine-N-oxide (0.001 %) were detected. Nicotine-N-oxide was only present at detectable levels in frass samples. It is possible that nicotine-N-oxide’s transit through the digestive tract is much more rapid than the other compounds, however I found no additional evidence for this. There is evidence that the Malpighian tubules are the prominent site of xenobiotic detoxification in D. melanogaster (Yang et al., 2007). It is possible that N-oxidation of nicotine, resulting in nicotine-N-oxide, only occurs in the cells of the cabbage looper Malpighian tubules. Lastly, nicotine-N-oxides are thermally unstable (Jacob et al., 1986). It is possible that inside the organism some portion of the nicotine-N-oxide produced degraded before the samples were  75taken. Representative UPLC-MS (MRM) chromatograms of nicotine, cotinine, cotinine-N-oxide, nicotine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid from frass samples are illustrated in Figure 3.8; notably, there was no discernable signal detected in the MRM trace of 4-hydroxy-4-(3-pyridyl) butanoic acid.  Cotinine was the most abundant compound detected, illustrating the importance of the 5’ C-oxidation pathway in cabbage looper.  The 5’C-oxidation of nicotine is the primary metabolic pathway used to detoxify nicotine in the monophagous nicotine-tolerant caterpillar M. sexta (Snyder et al., 1994; Wink and Theile, 2002). In the honey bee the major pathway is the 2’C-oxidation of nicotine (Fig. 3.7), producing 4-hydroxy-4-(3-pyridyl)-butanoic acid (Du Rand et al., 2017). I hypothesized that nicotine detoxification would occur in 4th instar cabbage looper larvae and that it would closely resemble the detoxification observed in M. sexta. I found the cabbage looper depends on the same metabolic pathway as M. sexta to detoxify nicotine, 5’ C-oxidation (Fig. 3.7). However, the cabbage looper did not employ the 2’ C-oxidation, used by honey bees, to any degree. Nicotine detoxification in the cabbage looper was similar to some mammals, for example humans employ the same major metabolic pathway (5’ C-oxidation) oxidizing 75 % of the nicotine dose to cotinine and excreting 15 % of the nicotine metabolized (Hukkanen et al., 2005).   In most nicotine tolerant insects, their tolerance is due to the activity of CYPs. Nicotine tolerance in the tobacco hornworm has been attributed, in part, to the enzyme CYP6B46 (Kumar et al., 2014a). Nicotine tolerant lines of B. tabaci and M. persicae gain their tolerance from the overexpression of CYP6CM1 and CYP6CY3, respectively (Bass et al., 2013; Kliot et al., 2014). The cabbage looper transcriptome has recently been released (preprint in Yu et al., 2017). This will facilitate the discovery of the enzymes responsible for nicotine detoxification in cabbage looper larvae.   The degree of transformation from nicotine to its major metabolite in M. sexta was lower than I observed in the cabbage looper (Wink and Theile, 2002), but similar to the degree of transformation in another nicotine-intolerant insect, S. ocellatus (Wink and Theile, 2002). In addition, the observed time course for removal of nicotine in the cabbage looper was similar to M. sexta and roughly 24 h faster than in S. ocellatus (Wink and Theile, 2002). In caged  76honey bees that do not defecate, transformation of nicotine to 4-(3-pyridyl)-butanoic acid had a similar time course to nicotine removal in cabbage looper larvae.   This study shows that polyphagous nicotine-intolerant insects, that were thought to only use rapid excretion when dealing with xenobiotics, extensively employ the 5` C-oxidation pathway, transforming nicotine to cotinine. The cabbage looper did not use the 2`C-oxidation pathway (Fig. 3.7) at all, although this was the major pathway used in honey bees (Du Rand et al., 2017). The product of the 2`C-oxidation pathway is 4-(3-pyridyl)-butanoic acid, which has the lowest LogP-value (Chemspider, Pubchem) of the nicotine metabolites investigated in this study. A lower LogP-value corresponds to slower passive transport through membranes. Conversion of nicotine to a compound with poor membrane clearance confines the xenobiotic to the digestive tract in honey bees (Du Rand et al., 2017). In comparison, cabbage looper larvae mainly transform nicotine to a compound with higher membrane clearance; the cabbage looper can very rapidly remove xenobiotics from the haemolymph, thus a faster passive diffusion of the xenobiotic into the haemocoel may result in faster overall clearance from the organism.  This was the most extensive study of phase 1 detoxification of nicotine in the cabbage looper. It is currently unknown whether or not nicotine induces phase 1 enzymes commonly associated with nicotine-tolerance, in insects and mammals, and in cabbage looper larvae (Bass et al., 2013; Hukkanen et al., 2005; Kliot et al., 2014; Kumar et al., 2014a). I have shown that phase 1 metabolites are present in the cabbage looper after dietary nicotine exposure. Thus, it would be prudent to investigate the presence of inducible or constitutively expressed detoxification enzymes. Detoxification enzymes in insects are often members of the CYP6 and CYP9 families (Li et al., 2007). Expression and functional characterization of these enzymes will be facilitated by the recently released cabbage looper transcriptome (preprint in Yu et al., 2017).   Gut microbiota have been implicated in the detoxification of xenobiotics in some insects (Hammer and Bowers, 2015). In coffee berry borers, the major pest of coffee, a study found that detoxification of caffeine was mediated by the insects’ gut microbiota (Ceja-Navarro et  77al., 2015). Caterpillar guts generally have high pH, simple structure, and fast transit times making them inhospitable for common insect gut microbiota. In fact, a study found that many caterpillar species lacked host-specific, resident gut microbes (Hammer et al., 2017). Therefore, it is unlikely that gut microbiota contributes to nicotine detoxification in the cabbage looper larvae.  In conclusion, I have shown that the polyphagous nicotine-intolerant Lepidoptera Trichoplusia ni extensively metabolizes nicotine to cotinine, the product of 5’ C-oxidation of nicotine. In addition, I found that T. ni did not rely on the 2’ C-oxidation of nicotine to 4-(3-pyridyl)-butanoic acid, like in the honey bee (Du Rand et al., 2017). Understanding in vivo metabolic pathways for the detoxification of plant secondary metabolites in this significant agricultural pest will help shape future decisions regarding insecticide development and application.      78                   Figure 3.7 Three major biochemical pathways for the enzymatic modification of nicotine to its common metabolites: cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid. Relative amounts (%) of nicotine and derived metabolites are shown in parentheses. These values represent the relative percentage of the total nicotine and its metabolites detected in all sample types over the entire 24 h sampling period. The major pathway in honey bees is the 2’ C-oxidation (green). In mammals and tobacco hornworm the major pathway is 5’ C-oxidation (blue). N-oxidation of nicotine is a minor pathway in many organisms (purple). Figure adapted from (Hukkanen et al., 2005).  2 3 4 5 Nicotine (16 %) Pseudooxynicotine 4-Oxo-4-(3-pyridyl) butanoic acid 4-Hydroxy-4-(3-pyridyl) butanoic acid (0 %) Nicotine-N-oxide (0.001 %) Cotinine (84 %) Cotinine-N-oxide (0.04 %) 2’ C-oxidation 5’ C-oxidation Oxidation N-oxidation  79                      Figure 3.8 Representative UPLC separation- Multiple Reaction Monitoring (MRM) chromatograms accompanied by the proposed fragmentation for nicotine and the 4 metabolites I scanned for. Nicotine-N-oxide fragmentation involves a ring expansion and is not well represented by a dashed line, and thus is not shown. Chromatograms are from frass samples, representing the product of nicotine metabolism. Cotinine was the most abundant compound (84 %), nicotine (16 %), nicotine-N-oxide (0.001%), and cotinine-N-oxide (0.04 %) were also detected. 4-hydroxy-4-(3-pyridyl) butanoic acid was not detected. Time (min) MRM of 6 Channels ES+ 163.1 > 132.1 (nicotine) 4.13e6 MRM of 6 Channels ES+ 177.1 > 80.1 (cotinine) 1.06e7 MRM of 6 Channels ES+ 193.2 > 162.0 (cotinine-N-oxide) 2.63e6 MRM of 6 Channels ES+ 179.3 > 130.0 (nicotine-N-oxide) 1.04e6 MRM of 6 Channels ES+ 182.2 > 164.0 (HyPyBut) 5.75e3  80Chapter 4 Conclusion  4.1 Objective 1: Accurate Method for the Detection and Quantification of Nicotine and its Metabolites  In this thesis I successfully developed and validated a HILIC-MS/MS method for the detection and quantification of nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-4-(3-pyridyl) butanoic acid. The method was validated according to current EPA guidelines for MDL and LOQ calculation (Betz et al., 2011).  To account for possible effects on analyses due to daily variable environmental conditions (e.g room temperature, humidity) I analyzed spiked samples in parallel with experimental samples. This allowed us to clearly identify peaks of interest in all chromatograms analyzed.   In addition, the %RSD for each compound was calculated and was used to determine the degree of retention time drift.  Minimal retention variability was observed in this study (< 2 %RSD), significantly decreasing the possibility of misidentification of analyte peaks; %RSD for RTs of nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-3-(4-pyridyl) butanoic acid were 1.8%, 1.5%, 0.8%, 1.4%, and 0.9%, respectively. In addition, percent recoveries of those same five compounds were calculated for this novel method and were determined to be 41 %, 84 %, 83 %, 37 %, and 53 % for nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-3-(4-pyridyl) butanoic acid, respectively. MDL values for nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-3-(4-pyridyl) butanoic acid were in ng/μL: 1.25x10-4, 1.69x10-7, 9.32x10-5, 8.90x10-5, and 7.83x10-6 respectively. LOQs for nicotine, cotinine, nicotine-N-oxide, cotinine-N-oxide, and 4-hydroxy-3-(4-pyridyl) butanoic acid were calculated to be in ng/µL: 4.31 x 10-4, 2.78 x 10-4, 1.05 x 10-4, 1.51 x 10-4, and 1.23 x 10-5, respectively. Taken together the respective %RSD, recovery percentage, MDL and LOQ values indicated a consistent trend in increasing analyte performance throughout the method developed in this study, with increasing polarity of the analytes themselves.   In general, the more polar an analyte, the better it performed in my HILIC-MS/MS method. For example, nicotine, the least polar analyte (PubChem: ChemSpider), had the highest  81%RSD in retention times, one of the lowest recoveries, the highest MDL, and the highest LOQ. The degree of retention of an analyte to a HILIC column is determined by the hydrophilic interaction between the column’s stationary phase and the analyte. Thus, because nicotine was the least polar, or hydrophilic, analyte it likely had the weakest interaction with the column’s stationary phase, resulting in poor performance, as evidenced by the relatively high MDL, LOQ, and %RSD, and low recovery. As would be expected, I have confirmed that HILIC-MS/MS is an appropriate and robust method for the detection and quantification of the polar hydrophilic analytes produced as a result of nicotine oxidative metabolism.  4.2 Objective 2: Metabolic Fate of Dietary Nicotine in 4th Instar Trichoplusia ni Larvae  4.2.1 Dietary Nicotine Exposure  The first step in completing objective 2 was determining the appropriate dietary dose of nicotine to use for the study of nicotine’s effect on xenobiotic metabolism in 4th instar T. ni larvae. Past studies that exposed cabbage looper larvae used dietary nicotine levels up to 0.125 % FW (Krischik et al., 1991; Self et al., 1964). However, in these studies the length of exposure not reported. In this study, I exposed 4th instar cabbage looper larvae to increasing amounts of nicotine (0 % FW – 1.0 % FW) for 120 h to determine the ideal dose and length for exposure. 4th instar larvae were used as they are the largest feeding larval stage and thus would represent the stage which would consume the highest level of plant secondary metabolites and are the stage which is responsible for the greatest amount of agricultural damage and economic loss.  I showed that after 24 h of dietary nicotine exposure there was not an observable difference in mortality, between the insects fed 1.0 % and 0 % nicotine FW. However, as the length of exposure increased the percent mortality increased with increasing dose of dietary nicotine. Thus, while higher doses of nicotine do induce mortality in 4th instar cabbage looper larvae, it takes > 24 h to have a significant effect on mortality. It is important to contrast the results of this thesis in context of the previous literature. Krischik et al. (1991) showed that cabbage looper larvae mortality increased, as determined by reduced larval emergence and decreased body weight above a 0.064 % nicotine FW dose.  82They found no survivorship above a 0.125 % nicotine FW dose.  Our study showed that T. ni larvae can survive much higher doses of dietary nicotine exposure, up to 1% nicotine FW dose. The differences observed between previous studies and our study may reflect the differences in larval life stages exposed to nicotine. Krischik et al. (1991) reared their insects from egg to emergence on diets containing nicotine, whereas our study only exposed larva to diets containing nicotine once they had reached the 4th instar larval stage. It is important to note that toxic effects are influenced by allometry (Bliss, 1935; Newman and Heagler, 1991), with the general principle that larger size reflects greater resistance to toxicant effects. I suggest it is most appropriate to study toxic and metabolic effects on the life stage which has the greatest environmental impact and is the largest in body size, and thus should according to these principles be the most resistant. In the case of T. ni this would be represented by the 4th instar larval stage and thus I believe our study represents the most appropriate life stage to evaluate the metabolism of nicotine and other plant secondary defense compounds in this insect. The mortality studies and subsequent metabolic fate results of this study suggest that T. ni have a much greater capacity for nicotine detoxification than has been previously reported.   4.2.2 Metabolic Fate of Dietary Nicotine   A small number of previous studies have reported that cabbage looper larvae are not capable of metabolizing dietary nicotine, and simply detoxified nicotine through rapid excretion of unmetabolized nicotine (Rheault et al., 2006; Self et al., 1964).  The results in Chapter 3 of this thesis clearly support our overall hypothesis that dietary nicotine is indeed metabolized by the cabbage looper Trichoplusia ni.  Furthermore, this thesis clearly shows that the major metabolite of nicotine metabolism is cotinine with minor trace amounts of cotinine-N-oxide and nicotine-N-oxide metabolites produced as well. This is the first time that the metabolic fate of nicotine in this generalist agricultural pest has been reported.   The results of this thesis support a generalized model describing the metabolism and excretion of nicotine and its derived metabolites in the cabbage looper (Fig 4.1). Briefly, ingested nicotine is transported across the midgut epithelium as unmetabolized nicotine into  83the haemolymph, or is alternatively metabolized by the midgut epithelium into cotinine, cotinine-N-oxide, and nicotine-N-oxide, and subsequently transported into the haemolymph. I suggest that nicotine and its metabolites are further metabolized and secreted from the haemolymph to the lumen of the Malpighian tubule and subsequently excreted downstream in the frass. It is possible that the unmetabolized nicotine detected in frass samples represents nicotine that was not absorbed in the alimentary canal and simply excreted. However, I have shown that unmetabolized nicotine was detected in haemolymph and other tissue samples strongly indicating that ingested nicotine is absorbed unmetabolized across the midgut epithelium. The presence of unmetabolized nicotine in the excretions of the 4th instar cabbage looper larvae suggests that some portion of the ingested nicotine was excreted unmetabolized. In fact, it has been well established that the isolated Malpighian tubules of T. ni are capable of actively secreting unmetabolized nicotine from the haemolymph to the lumen of the Malpighian tubule and hindgut for subsequent excretion (Maddrell and Gardiner, 1975; Rheault et al., 2006; Self et al., 1964). Transport  of alkaloids such as nicotine across arthropod epithelia is facilitated by the multidrug resistance transporters of  ATP Binding Cassette (ABC) transporter Superfamily (Bretschneider et al., 2016b; Dermauw and Van Leeuwen, 2014; Labbé et al., 2011). In addition, studies on vertebrate cell lines have demonstrated that nicotine can be at least partially transported by the Organic cation transporter 2 (OCT2) as evidenced by the partial blockage of nicotine by the OCT substrate cimetidine (Uwai et al., 1998; Zevin et al., 1998).   The transformation of nicotine to cotinine and cotinine derivatives, such as cotinine-N-oxide, in many animals, including mammals and insects, is highly conserved and is mediated by CYPs (Hukkanen et al., 2005). In M. sexta the CYPs associated with nicotine resistance include CYP6CM1 and CYP6B46 (Kumar et al., 2014a; Stevens et al., 2000). This detoxification of nicotine utilizes the Phase I, 5’ C-oxidation pathway (see Fig 3.8). Nicotine-N-oxide is the result of an N-oxidation of nicotine. Detection of trace nicotine-N-oxide in the excretions of the insects suggests that the larvae can utilize multiple metabolic pathways to detoxify dietary nicotine. The N-oxidation of nicotine is mediated by FMOs (Hukkanen et al., 2005). Our data suggest that T. ni either can induce expression of CYPs and FMO when faced with dietary nicotine exposure, or have certain constitutively expressed CYPs and  84FMO. Currently, none of the enzymes responsible for nicotine detoxification in the cabbage looper have been identified or characterized. The recent release of  T. ni transcriptome (Yu et al., 2017) may facilitate further detection and characterization of the enzymes responsible for nicotine detoxification in the cabbage looper. I propose that using a similar experimental design regarding nicotine dietary exposure and quantification of subsequent changes in putative cyp and fmo gene expression may identify the CYP and FMO enzymes responsible for the 5’C oxidation and N-oxidation pathways described in this thesis.    While there appears to be considerable conservation in the mechanisms and pathways used for nicotine metabolism amongst various animals as demonstrated in the previous paragraph there are also studies that show marked differences in nicotine metabolism within insects. For example, in a study using a similar method to the one used in this thesis 4-hydroxy-4-(3-pyridyl) butanoic acid was the most abundant metabolite of nicotine detoxification in the honey bee A. mellifera (Du Rand et al., 2017). 4-hydroxy-4-(3-pyridyl) butanoic acid is the result of 2’ C-oxidation of nicotine (Hukkanen et al., 2005), however, the particular enzymes responsible for mediating this biochemical reaction in insects remain to be elucidated (Du Rand et al., 2017). I did not detect any 4-hydroxy-4-(3-pyridyl) butanoic acid in our study, indicating that T. ni do not rely on the 2’ C-oxidation pathway for nicotine detoxification, to any measurable degree. These data highlight the differences in xenobiotic detoxification between different orders of insects.  A greater understanding of the detoxification pathways behind insect resistance to plant secondary compounds, such as nicotine, may facilitate the development and implementation of more environmentally friendly and cost-effective alternatives to synthetic pesticides for the chemical control of insect agricultural pests.   85                  Figure 4.1 Generalized model for metabolism and excretion in T. ni. The model shows the three basic cavities of the insect body: the gut, the haemocoel, and the Malpighian tubules. The colored acronyms are compounds: nicotine (NIC), nicotine N-oxide (NNO), cotinine (COT), and cotinine N-oxide (CNO). The yellow circle represents cytochrome P450 enzymes (CYP). The blue circle represents the multi-drug resistant transporter (MDR). Dashed lines represent passive diffusion and dotted lines represent metabolism.   864.2.3 Time Course of Nicotine Metabolism and Excretion   Previous studies on the time course of nicotine metabolism and elimination in the nicotine tolerant specialist M. sexta have shown that nicotine is still detected in haemolymph, tissue and frass up to 48 h post exposure (Kumar et al. 2014). In contrast I showed that  T. ni cleared over 99 % of the total nicotine and derived metabolites detected by 6 h after removal from a nicotine exposure. 24 h after removal only frass samples had detectable levels of analytes. Thus, it appears that T. ni larvae are very efficient at clearing nicotine and its metabolites post exposure, thus making T. ni a putative model insect for the study of nicotine detoxification and excretion. Notably, M. sexta employs nicotine halitosis as a novel anti-predation defense strategy. There is no reported evidence for this in generalist herbivorous insects such as T. ni. Thus, it is possible that the delayed time course for the clearance of nicotine observed in M. sexta may, in fact be beneficial due to the use of nicotine as a chemical deterrent against predation.    874.3 Future Directions   Although several questions and future lines of investigation have been raised in the preceding chapters and paragraphs of this thesis there are several additional questions regarding the mechanisms responsible for the detoxification and excretion of nicotine metabolites that could be addressed using information described in this thesis.   1) Does dietary exposure to nicotine result in the increased expression of Cytochrome P450s and Flavin-containing monooxygenases?  The specific enzymes responsible for nicotine metabolism in cabbage looper larvae have not been identified. There remains a gap in the research surrounding phase I metabolism enzymes in T. ni. However, I have demonstrated that phase I metabolites of nicotine are present in the tissue, haemolymph, and frass of cabbage looper larvae fed dietary nicotine. Therefore, it is likely that expression of phase I enzymes should be altered upon nicotine exposure in T. ni.  A major stumbling block to these proposed studies is the lack of genetic information on T. ni. However, a very recent paper (Yu et al., 2017) describing the transcriptome of T. ni should allow for design of primers which will facilitate the study of mRNA expression levels in T. ni exposed to dietary nicotine. Previous studies on M. sexta have shown that mRNA expression of cyp transcripts are increased upon dietary nicotine consumption (Kumar et al., 2014). I propose that expression levels of putative cyp and fmo transcripts could be evaluated through targeted gene expression studies using quantitative polymerase chain reaction (qPCR) or using high-throughput transcriptomic analysis using the experimental feeding design used in this thesis.   2) Do the Malpighian tubules of T. ni actively secrete the oxidative metabolites of nicotine?  A previous study by Rheault et al. (2006) have demonstrated that the Malpighian tubules of a number of species from diverse orders of insects are capable of secreting nicotine. As demonstrated in chapter 3 of this thesis I now know that the oxidative metabolites of nicotine  88are excreted in the frass of T ni.  Taken together with previous studies this suggests that the Malpighian tubules of T. ni have the capacity to secrete both nicotine and its metabolites. I propose that the ability of the Malpighian tubules to transport nicotine, cotinine, cotinine-N-oxide, and nicotine-N-oxide can be measured using a combination of traditional Ramsay assay techniques (Rheault et al. 2006) and the HILIC-MS/MS method developed in chapter 2 of this thesis. Briefly, isolated Malpighian tubules can be stimulated to secrete in vitro, the secreted fluid collected and analyzed to measure secretion rates and fluxes of transepithelial metabolite transport. In addition to answering the question of whether the Malpighian tubules are capable of transporting the oxidative metabolites of nicotine, the method developed in this thesis can also be used to determine if the Malpighian tubules themselves also are capable of the oxidative metabolism of nicotine. In this case Malpighian tubules can be exposed in vitro to nicotine and their secretions can be assayed for the relative abundance of individual metabolites.   In summary, this thesis has demonstrated that the larvae of the generalist phytophagous insect Trichoplusia ni is capable of oxidative metabolism of nicotine and has the potential to be used as a model insect for future studies of plant secondary metabolite detoxification. This thesis provides a robust method that can be used to further characterize plant secondary metabolite detoxification in insect tissues, body fluids and excretions.  A greater understanding of the processes and mechanisms underlying insect detoxification and excretion of xenobiotics has the potential to contribute critical knowledge to the future evaluation and development of novel pesticides and insect control measures.   89Bibliography   Allelochemicals, P., 1986. Plant Allelochemicals and Insect Parasitoids Effects of Nicotine on Cotesia congregata (Hymenoptera: Braconidae) and Hyposoter annulipes 12, 1319–1328. Amoabeng, B.W., Gurr, G.M., Gitau, C.W., Stevenson, P.C., 2014. 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