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The physical and behavioral effects of embryonic ethanol exposure in Caenorhabitis elegans Lin, Conny 2008

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THE PHYSICAL AND BEHAVIORAL EFFECTS OF EMBRYONIC ETHANOL EXPOSURE IN CAENORHABDITIS ELEGANS  by Conny Lin B.Sc., The University of British Columbia, 2004    A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE  in  THE FACULTY OF GRADUATE STUDIES  (Neuroscience)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December 2008 © Conny Lin, 2008 ii  ABSTRACT In this thesis I used Caenorhabitis elegans as a model of Fetal Alcohol Spectrum Disorder (FASD) to study the physical and behavioral effects of ethanol exposure during embryonic development. Davis et al. (2008) found that ethanol exposure during larval development in C. elegans produced physical/developmental and behavioral effects; however, whether exposure during embryonic development might produce similar outcomes remained to be elucidated. Because the type and degree of effects caused by developmental ethanol exposure was dependent on the pattern of ethanol treatment, in the first part of the thesis I investigated the physical/developmental effects of embryonic exposure to various ethanol doses, exposure durations, onsets and frequencies. I found that exposure to >30% ethanol for an hour during embryonic development was necessary to lower hatch rate, delay reproductive onset, and reduce body size in C. elegans. Furthermore, exposure during early embryonic development caused a larger effect than exposure during later stages, and multiple exposures produced a worse outcome than a single exposure for a comparable duration. In the second part of the thesis, I investigated  locomotory activities and habituation of adult C. elegans exposed to various patterns of embryonic ethanol treatment. I found that the rate of locomotion was altered differently by chronic and acute embryonic ethanol exposure, but I did not find any effect in short- or long-term habituation. In summary, I have characterized the pattern of embryonic ethanol exposure necessary to produce physical/developmental effects in C. elegans, and identified the types of exposure conditions that would cause worse outcomes than others; in addition, I have found that embryonic ethanol exposure affects the rate of locomotion in C. elegans. In this thesis, I have established a foundation for the future investigation into the physical and motor defects caused by embryonic ethanol exposure in C. elegans. iii  TABLE OF CONTENTS Abstract......................................................................................................................................................................... ii Table of Contents......................................................................................................................................................... iii List of Tables ................................................................................................................................................................. v List of Figures ...............................................................................................................................................................vi Acknowledgments ..................................................................................................................................................... viii Chapter 1. Introduction .................................................................................................................................................. 1 Fetal Alcohol Spectrum Disorder (FASD) ................................................................................................................ 1 Many factors determine the outcome of alcohol teratology ...................................................................................... 3 Dose (Concentration) ........................................................................................................................................... 4 Timing & pattern of exposure .............................................................................................................................. 5 Mandate for high-throughput animal models ....................................................................................................... 6 Caenorhabditis elegans the model organism ............................................................................................................ 7 Thesis objectives ..................................................................................................................................................... 11 Chapter 2. Physical/Developmental Abnormalities Induced by Embryonic Ethanol Exposure in C. elegans ............ 14 Introduction ............................................................................................................................................................. 14 Embryonic ethanol exposure in C. elegans ........................................................................................................ 14 Chapter objective and hypothesis ....................................................................................................................... 18 Materials & Methods .............................................................................................................................................. 19 Animals maintenance ......................................................................................................................................... 19 Embryo (egg) collection ..................................................................................................................................... 19 Ethanol exposure procedures ............................................................................................................................. 20 Hatch rate ........................................................................................................................................................... 22 Reproductive onset ............................................................................................................................................. 22 Egg-laying (reproductive) pattern & brood size ................................................................................................. 23 Body size ............................................................................................................................................................ 23 Lifespan.............................................................................................................................................................. 24 Statistical analysis .............................................................................................................................................. 24 Results..................................................................................................................................................................... 27 Experiment A-1: Acute 20% ethanol exposure at different developmental stages induced different developmental abnormalities ............................................................................................................................. 27 Experiment A-2: Ethanol dose and duration needed to surpass a certain threshold to induce physical/developmental deficits ......................................................................................................................... 33 Experiment A-3 Early and the repeated exposure induced the worst outcome .................................................. 55 Discussions ............................................................................................................................................................. 66 Treatment severity needs to surpass a threshold to stabilize physical/developmental deficits ........................... 66 Early development is the most vulnerable stage ................................................................................................ 67 The result of repeated exposure is a product of compounded effects................................................................. 69 Potential for this data set is enormous ................................................................................................................ 70 Chapter 3. Behavioral Effects of Embryonic Ethanol Exposure in C. elegans ............................................................ 72 Introduction ............................................................................................................................................................. 72 Materials & Methods .............................................................................................................................................. 76 Exposure protocol .............................................................................................................................................. 76 Behavioral recording & scoring ......................................................................................................................... 76 Procedures for body bends and spontaneous reversals....................................................................................... 77 Habituation procedures ...................................................................................................................................... 78 Statistical analysis .............................................................................................................................................. 79 Results..................................................................................................................................................................... 81 Experiment B-1: Exposure of 30% ethanol for an hour during middle development increased the rate of body bends .................................................................................................................................................................. 81 Experiment B-2: Chronic exposure to 10% ethanol during embryonic development resulted in a decreased rate of body bends and an increased rate of spontaneous reversals ........................................................................... 85 Experiment B-3: Exposure during early development produced hyper-responsiveness to tap .......................... 87 Experiment B-4: Habituation to taps at 10s or 60s ISI was not affected in worms chronically exposed to ethanol during embryonic development ............................................................................................................. 90 iv  Experiment B-5: Long-term memory of habituation training was intact in animals chronically exposed to ethanol during embryonic development ............................................................................................................. 94 Discussions ............................................................................................................................................................. 97 Alteration in the number of body bends and spontaneous reversals in the 10% chronic animals may be either a result of motor dysfunction or developmental delay .......................................................................................... 97 Body bend was increased by acute exposure but decreased by chronic exposure ............................................. 98 A more difficult task may be required to see learning and memory deficits in exposed worms ...................... 101 Chapter 4. General Discussions and Conclusions ...................................................................................................... 103 General Discussions .............................................................................................................................................. 105 Phenotypic variability caused by developmental ethanol exposure occurs in an isogenic population ............. 105 The ethanol concentration used was high, but the serum concentration in the egg was unknown ................... 109 Limitations and Future Directions ........................................................................................................................ 111 References ................................................................................................................................................................. 114 Appendices ................................................................................................................................................................ 122 Appendix A: Hatch rate, Egg and Larva Counting ............................................................................................... 122 Appendix B: iMovie, QuickTime, and Image J Operation for the Measurement of Body Size in C. elegans ...... 123 Appendix C: Water Effect: The Timing of Water Control Treatment .................................................................. 125 Appendix D: Water vs. M9 ................................................................................................................................... 126   v  LIST OF TABLES Table 2.1 Summary of findings from ethanol exposure groups in Experiment A-2. ................... 54 Table 2.2. Descriptive statistics of hatch rate, body size and lifespan of worms exposed to 30% ethanol for 20 or 60min singly or repeatedly at early, middle or late exposure. .................. 59  vi  LIST OF FIGURES Figure 1.1 C. elegans anatomy ....................................................................................................... 7 Figure 1.2 C. elegans Life cycle ..................................................................................................... 9 Figure 2.1 Ex-utero development of C. elegans. .......................................................................... 14 Figure 2.2 Apparatus for filming worm in iMovie ....................................................................... 23 Figure 2.3 Ethanol exposure timing in Experiment A-1 ............................................................... 27 Figure 2.4 Acute exposure of 20% ethanol at different times during embryonic development delayed reproductive onset in the late exposure group but did not altered the reproductive patterns or brood sizes. ......................................................................................................... 29 Figure 2.5 1hr exposure to 20% ethanol at different times during embryonic development did not alter the growth pattern of body sizes (µm2)  or body sizes within particular days. ............ 30 Figure 2.6 Lifespan and adult survival rate was not altered by acute exposure of 20% ethanol during early (2-3h EUE), middle (5-6h EUE), and late (8-9h EUE) embryogenesis. .......... 31 Figure 2.7 Treatment conditions in Experiment A-2. Groups exposed to chronic exposure for 8.5hr was treated from 0.5-9hr ex-utero development. ......................................................... 35 Figure 2.8 Chronic exposure reduced hatch rate .......................................................................... 37 Figure 2.9 Hatch rate dropped when ethanol treatment severity reached a threshold, and this threshold was unique to each developmental stage and treatment frequency. ..................... 38 Figure 2.10 Reproductive onsets were delayed by chronic exposure, and more effectively by the repeated exposure than by single exposures. ........................................................................ 41 Figure 2.11 Body sizes were smaller in worms exposed to 10% ethanol for 8.5hr throughout embryonic development. ....................................................................................................... 44 Figure 2.12 The body sizes of 2-d-old worms exposed to 5-10% ethanol for 8hr throughout embryonic development were drastically smaller. ................................................................ 45 Figure 2.13  For exposure during early embryonic development, the growth of body sizes were reduced in worms exposed to 30% ethanol for 60min, and exposure for shorter durations did not have large impact on the body sizes. .............................................................................. 47 Figure 2.14  For exposure during middle embryonic development, the growth of body sizes were reduced in worms exposed to 40% ethanol for 60min on Day 4 only. ................................. 49 Figure 2.15 Exposure during late embryonic development did not alter the growth of body. ..... 50 Figure 2.16 Repeated exposure for a total of 60min (20min per episodes) reduced the body sizes................................................................................................................................................ 52 Figure 2.17 Models for the magnitude of physical deficits observed in repeated exposure. ........ 56 Figure 2.18 Exposure timing in Experiment A-3.......................................................................... 59 Figure 2.19 Reproductive onset was delayed in early and repeated exposure group at 30% ethanol concentration in Experiment A-3. ............................................................................ 60 Figure 2.20 The magnitude of reproductive delay in repeated exposure group more closely resembles the sum of three 60min exposure at three different stages. Sample size was 45 per group. .................................................................................................................................... 61 Figure 2.21. Reproductive onset organized in categories reveled that repeated exposure and early exposure for 60mins had larger proportions animals with delayed reproductive onset. ...... 62 Figure 2.22 Survival rate was not altered by 30% ethanol exposure during embryonic development. ......................................................................................................................... 64 Figure 2.23 Early exposure produced more severe phenotype. .................................................... 68 Figure 2.24 Repeated exposure produced the most severe effect ................................................. 70 vii  Figure 2.25 An example of questions you can ask from the systematic evaluation in Experiment 1.2.......................................................................................................................................... 71 Figure 3.1 Body bend punctuated by spontaneous reversal was not counted. .............................. 73 Figure 3.2 Apparatus for behavioral recording ............................................................................. 77 Figure 3.3 Protocol for body bend and spontaneous reversal experiments. ................................. 78 Figure 3.4 Training protocol for long term habituation ................................................................ 79 Figure 3.5 Worm handling timeline .............................................................................................. 81 Figure 3.6 Body bends (BB) before and after stimulus of worms exposed to 30% ethanol for 20 or 60min singly during either early, middle or late development, or repeatedly at each of the three developmental stages. .................................................................................................. 82 Figure 3.7 The activity of body bends across the full 6min behavioral recording. ...................... 84 Figure 3.8 Spontaneous reversals of worms exposed to 30% ethanol for 20 or 60min singly during early, middle or late embryonic development or repeated during all 3 stages. ......... 85 Figure 3.9 locomotory behavioral recording for Experiment B-2 ................................................ 86 Figure 3.10 Body bends and spontaneous reversal of chronic ethanol exposure. ........................ 87 Figure 3.11 The reversal response to tap stimuli of worms exposed to 20% ethanol for 60min during different times of embryonic development. ............................................................... 88 Figure 3.12 Habituation curves of worms exposed to 20% ethanol for 60min during different times of embryonic development. ......................................................................................... 89 Figure 3.13 The response to initial and last 3 stimuli of chronically exposed worms at a 10s ISI................................................................................................................................................ 91 Figure 3.14 Habituation to tap stimuli delivered at a 10s ISI of worms exposed chronically for 8hrs to ethanol during embryonic development.................................................................... 92 Figure 3.15 The response to initial and last 3 stimuli of chronically exposed worms at a 60s ISI................................................................................................................................................ 93 Figure 3.16 Habituation to tap stimuli delivered at a 60s ISI of worms exposed chronically for 8hrs to ethanol during embryonic development.................................................................... 94 Figure 3.17 Long term memory of habituation training in worms exposed chronically for 8hr during embryonic development. ........................................................................................... 95 Figure 3.18 The difference in the protocol to obtain behavioral recordings in Experiment B-1 (A) and B-2 (B) ........................................................................................................................... 99 Figure 4.1 Models to explain the variable phenotypes observed in different ethanol treatment severity. ............................................................................................................................... 107 Figure 4.2 More severe ethanol exposure during embryonic development produced more worms with smaller body sizes, and less severe ethanol exposure produced more worms with normal body sizes. .............................................................................................................. 108  viii   ACKNOWLEDGMENTS   First I would thank my supervisor, Catharine Rankin, for all her patient support, both academically and mentally. Her positive and persistent encouragements made the journey to the completion of this thesis enjoyable and rewarding. I would also like to thank her for giving me a right amount of freedom to learn from mistakes. She was there to hold me up when I had lost my hope and confidence. I have not only learned how to do science from her, but also learned how to be a better person. I have grown academically and mentally during these years, thanks to her.  I would like to thank my parents and my grandfather who valued academics more than anything else in life. With that I could pursue my late Master’s degree without worrying too much about the pressure of reality. My veteran grandfather, who passed away a few days before I submit this thesis, said to me, “Study as much as you can. Everything else in life you can lose, but no one can take away your education because it stays in you.” My mother was the same way. She had tolerated me through my worst rebellious times and supported me for whichever many interests and dreams I might have. At this point, I also need to thank John Wang for he forced me to finished high school, and without that, I would not be here today. I would like to specially thank my father for his support and understanding for my rocky journey that finally take me to finish my undergraduate degree and now my Master’s thesis (“late bloomer”, he called me).  I also like to thank all the lab members, especially Yun Li (data collection of Experiment B-1 was completely done by her; I did the analysis), who had been through rough nights of experiments with me, and sometimes my rough temper. My interactions with every lab members have taught me tremendously (didn’t matter ones who stayed or left). Thanks for letting me learn from you and helping me through my degree. Love for all of you. 1  CHAPTER 1. INTRODUCTION Fetal Alcohol Spectrum Disorder (FASD) Fetal alcohol syndrome (FAS) was first recognized as a consistent pattern of birth defects that (craniofacial dysmorphology, growth retardation, and brain damage) occurred in children of alcoholic mothers (Jones and Smith, 1973). Curiously, only 3-4% of children from alcoholic mothers qualify for the diagnostic criteria for FAS (Sampson et al., 1997; Koren et al., 2003). Later it became clear that children of alcoholic mothers often manifested neurodevelopmental, behavioral, and/or cognitive problems (Koren et al., 2003) not originally covered in the criteria for FAS. Affected children who did not clearly fall into the diagnostic criteria of FAS were then placed into diagnostic categories such as partial FAS, alcohol-related birth defects, and alcohol-related neurodevelopment disorder (Burd et al., 2003). Finally, Barr and Streissguth (2001) united the full range of adverse effects induced by prenatal alcohol exposure under the umbrella term “Fetal Alcohol Spectrum Disorder (FASD)”.  The incidence of FASD has been estimated as 1 in every 100 live births in the United States, causing a financial burden of more than $1.6 billion a year (Rice et al., 1991; Chudley et al., 2005). Unfortunately, FASD incidence has remained unchanged for the last decade despite increasing awareness of FASD (Cudd, 2005). In North America, 10% of women persisted in drinking alcohol during pregnancy (Wattendorf and Muenke, 2005). The problem is worse in other countries. Recent surveys suggested over half of the Australian, 52% of French, and 40% of Brazilian (Elizabeth J. Elliott, 2008) (Moraes and Reichenheim, 2007; de Chazeron et al., 2008) women drank alcohol during pregnancy. Therefore, in addition to advocating abstinence, research into FASD treatments must be explored. 2  FASD is the leading cause of mental retardation in North America, even more so than the Down’s syndrome and cerebral palsy (Nash et al., 2008). The intelligence scores of children with FAS often fall more than 2 standard deviations below the mean, and approximately half of the children with full-blown FAS are mentally retarded (Streissguth et al., 1996). For less severe cases, a wide range of alterations in brain structures, behavioral and psychological functioning may be present in individuals affected with FASD (Riley and McGee, 2005). Human studies using magnetic resonance imaging (MRI) revealed that children with FASD exhibited alterations in several brain regions, especially in the cerebellum, corpus callosum, and basal ganglia (Riley and McGee, 2005). Other neurobehavioral impairments, such as motor dysfunction, learning and memory difficulties, and problems in attention, impulse control, and judgment, were also common consequences of prenatal alcohol exposure (Chudley et al., 2005). For example, the type of motor dysfunctions may include delayed motor development (Kalberg et al., 2006), poor motor coordination (Mattson et al., 1998), and slowed motor selection speed (Simmons et al., 2006). The degree of motor dysfunction and motor developmental delay correlated also with the degree of cognitive delays (Osborn et al., 1993). A range of learning and memory problems, such as deficits in habituation, spatial navigation, working memory, and associative learning were commonly found in children affected with FASD (Kodituwakku, 2007).  These neurobehavioral problems sacrificed the quality of life in affected individuals, and in many cases, limited their ability to live independently (Spohr et al., 2007).  A vast amount of effort has been poured into finding the mechanistic basis for the neurobehavioral problems in FASD, but so far the situation has been very complicated and difficult to comprehend. Using learning deficiencies as an example, many molecular targets have been identified in rodent models: AMPA-type Glutamate receptors (Vaglenova et al., 2008; 3  Wijayawardhane et al., 2008), NMDA-type Glutamate receptors (Toso et al., 2006b; Samudio-Ruiz et al., 2007), GABA receptors (Toso et al., 2006b; Toso et al., 2006a), BDNF (Puglia et al., 2007; Caldwell et al., 2008), and many others. Unfortunately, our understanding on how the molecular targets interact to produce deficits in learning is vague at best. To complicate the matter even further, ethanol can affect each brain region and cell type differently. For example, the same dose of prenatal ethanol exposure reduced the number of Purkinje cell and Mitral cells, but increased the number of CA3 cells (Maier et al., 1999). The same dose of ethanol increased the volume of dentate gyrus, but decreased the volume of cerebellum and olfactory bulb (Maier et al., 1999). Consequently, our current understanding of learning deficits in FASD is that it involves many biological processes. However, we do not understand how each process comes together to produce the learning problems seen in FASD.  Many factors determine the outcome of alcohol teratology The outcome of fetal alcohol exposure depends on many factors, including maternal environment, fetal genetic background, and severity of alcohol exposure. Many maternal factors such as maternal genes, age, and diet can play important role. For example, some maternal alcohol dehydrogenase (ADH) variants were shown to decrease FASD vulnerability (Warren and Li, 2005). We know that FASD-affected children born to older mothers (30 years or older) have a wider range of cognitive deficits than affected children born to younger mothers (Jacobson et al., 2004). The severity of FASD was associated with maternal diet deficiency in folic acid, zinc, and thiamine (Ba et al., 1999; Summers et al., 2008; Xu et al., 2008; Yanaguita et al., 2008). On the other side of the story, minor fetal genetic differences can affect the outcome of prenatal alcohol exposure. Dizygotic (fraternal) twin studies in humans repeatedly find one twin to be more severely affected by prenatal alcohol exposure than the other twin (Christoffel and 4  Salafsky, 1975; Chasnoff, 1985; Streissguth and Dehaene, 1993; Riikonen, 1994; Mathelier and Karachorlu, 1999). Animal studies have also demonstrated the importance in fetal genetic differences. An experiment exposing different strains of rats to the same amount of prenatal alcohol showed that one strain exhibited lower birth weights and higher mortality rates than other strains (Abel, 1989). In another rat study, the susceptibility for facial dysmorphology was shown to differ between two strains (Parri Wentzel, 2008). In a chicken model, Su et al. (2001) and Cavieres & Smith (2000) concluded that strain was predictive of the type and severity of facial dysmorphology and cardiac defects produced by fetal ethanol exposure. These differences in maternal environment and fetal genetics ultimately interact with different patterns of maternal alcohol consumption to create a unique outcome in each individual.  Dose (Concentration) Many studies have found a unidirectional relationship between ethanol concentration and the severity of deficits, but such a relationship may not be true under all circumstances. A linear and unidirectional relationship between dose and the severity of symptoms was seen in the viability of cerebellar cell culture, myelin protein expression level of oligodendrocyte cell line, protein kinase C expression level in chicken, seizure threshold in rats, and the delay in axon outgrowth, thymidine uptake and DNA metabolism in cell culture  (Snyder et al., 1992b; Snyder et al., 1992a; Pantazis et al., 1993; McIntyre et al., 1999; Bichenkov and Ellingson, 2001; Bonthius et al., 2001; Bonthius et al., 2002; Lindsley et al., 2003). It seems that the higher the ethanol concentration, the worse the condition, but not all responses to ethanol are unidirectional. The response to ethanol can be bi-directional. For example, lower doses of prenatal ethanol exposure decreased amino acid uptake, but higher doses increased amino acid uptake (Snyder et al., 1992b). The neuron number in the dentate gyrus increased with lower maternal blood alcohol 5  concentration, but decreased in higher blood alcohol concentration (Miller, 1995). Some other effects only occur if ethanol concentration goes over a threshold level. For example, only the highest ethanol dose tested decreased the number of mitral cells in the rat olfactory bulb (Maier and West, 2001a). Even more interestingly, some effects only occur within a specific range of ethanol concentrations. For example, the number of CA3 cells increased in the medium dose, but did not change in response to either lower or higher doses (Maier and West, 2001a). Because responses sometimes do not follow a simple unidirectional relationship with changing ethanol concentration, the dose-response relationship of a new target must be experimentally determined.  Note that the ways to express ethanol doses used in different animal systems were different from each other depending on the administrative method. The same dose may be severe for one animal model but not the other model, partly due to the different rates of alcohol metabolism in different animal systems. Therefore, the “low”, “moderate”, or “high” dose of ethanol exposures described in FASD studies often referred to the range of doses tested in a particular study relative to a particular animal system.  Timing & pattern of exposure The process of development is highly environment- and temporal- dependent. Cell fate and cell migration often are guided by specific combinations of molecules at specific concentrations in their environment (extracellular space), and certain events must occur before the subsequent event can occur normally. Introduction of a foreign chemical (ethanol) can disturb the dynamics of these delicately positioned molecules in the extracellular space (Martinez Arias and Stewart, 2002). The consequences of this disturbance can be detrimental at certain times for one group of cells, but not for other cells. In addition, every cell can potentially respond to this disturbance differently at a given time.  6  Ethanol exposure occurring during a “critical period” produces the most severe damage. One study had found that acute ethanol exposure during postnatal day (PD) 5 induced a reduction in Purkinje cell number in rats, but exposure at a later time during development (PD10) did not induce a reduction (Pauli et al., 1995). Later the same group found that Purkinje cells were especially vulnerable during PD5-9 (Miki et al., 1999).  Similarly, Hsiao et al (2004) discovered a critical period for GABA current density. They found 2 days of exposure during PD8-9 in rats was sufficient to induce the same magnitude of reduction in GABA current density as exposure for 6 days during PD 4-9. These findings demonstrated that ethanol can produce the most detrimental outcome when introduced during a specific period in development. The timing of the critical period can be different for each target. In addition, longer periods of exposure do not necessarily result in worse outcomes.  Mandate for high-throughput animal models The severity of ethanol exposure is determined by many factors. One treatment condition may result in some, but not all symptoms. In order to understand the effect of prenatal alcohol exposure on a specific target, we must explore a range of exposure conditions. However, this is not practical with the current rodent or primate models because of the time and the sheer number of animals required.   For example, to investigate the expression level of a neurotransmitter receptor for 5 different ethanol concentrations, 3 exposure durations at 3 different times during development, would require (5 doses x 3 exposure duration x 3 timings x 2 controls) 90 groups. For a sample size of 10, that comes to 900 animals. 2700 animals will be needed to do 3 replications. This number is neither practical nor economical in mammal studies. This large combination of ethanol exposure conditions is one of the most challenging problems in FASD research. 7  Imagine an animal that can produce 70 progeny during its peak reproductive day. In a day, 13 individuals of that species can produce enough progeny to do one replication of the hypothetical experiment described above. The essential baseline studies considering different exposure patterns seem more approachable in an animal model of this kind. The nematode Caenorhabditis elegans offers such a model system. Caenorhabditis elegans the model organism C. elegans was first proposed as a model organism for the study of the genetic control of development by Sydney Brenner (Brenner, 1974). Soon it became a very popular genetic model due to its rapid generation time and large progeny pool. This animal is a 1mm long, soil-dwelling nematode (round worm) found in temperate regions (Figure 1.1). The main form of C. elegans is a hermaphrodite, capable of self-fertilizing to produce 300 isogenic progeny during its lifespan.   Figure 1.1 C. elegans anatomy  A) Normaski image of an adult C. elegans, and B) cartoon representation of C. elegans anatomy. (adapted from www.wormbase.org) This isogenic population is a convenient way to eliminate the contribution from genetic variation bewteen individuals. As a pilot project for the Human Genome Project, C. elegans was the first multi-cellular animal to have its genome fully sequenced (Hodgkin et al., 1998). Its genome 8  contains approximately 20,000 genes. The continually expanding mutant library is massive (~6000 genes coverage to date), which is as big as the recently completed yeast Genome Deletion Project. Best of all, these strains are inexpensive to obtain and maintain. Strains can be requested via email through the C. elegans Genetic Center, safely delivered through FedEx, and conveniently stored in -80oC freezers. The body of this animal is transparent, which allows in vivo observation of proteins tagged with green fluorescent protein (GFP). C. elegans expressing these GFP-tagged proteins are relatively easy to make because C. elegans can duplicate plasmids. Therefore, plasmids containing genes coded for these GFP-tagged proteins can be cloned and injected into C. elegans gonads. The resulting progeny will express these transgenic proteins.  A number of features of C. elegans make it an excellent model in which to study ethanol exposure during development. Its entire cell lineage has been mapped (Sulston et al., 1983), and the timing of important developmental events has been determined. Therefore, ethanol can be administered during a chosen developmental event or even at the time when a chosen neuron is born. Developmental abnormalities following ethanol exposure can be found by observing cell divisions that deviate from the cell lineage map. In addition, C. elegans embryos develop into adults in only 2.5 days at 20oC. This rapid development is particularly advantageous for longitudinal studies. C. elegans is the only multicellular animal with its neuron wiring diagram completely mapped out. The hermaphrodite nervous system consists of only 302 neurons with fewer than 10,000 connections. In contrast to billions of neurons and trillions of connections in the mammalian nervous system, the exploration of ethanol teratology on a neuron-to-neuron basis is actually practical in C. elegans. With an established battery of behavior assays, this model is 9  equipped with all the tools we need to explore the underlying mechanisms for behavioral deficits in FASD. Embryonic development in C. elegans occurs first inside the hermaphrodite for approximately 2.5hr and then continued after the egg was laid outside of the hermaphrodite for approximately 11.5hr. Fertilization of the oocyte and the following 2.5hr of cell proliferation  Figure 1.2 C. elegans Life cycle  (adapted from www.wormbase.org) occur inside the hermaphrodite. Soon after gastrulation initiates (approximately 30-cell stage), the hermaphrodite lays the egg, and the ex-utero embryonic development starts. Gastrulation is completed 3hr into ex-utero embryonic development. Gastrulation is followed by organogenesis and morphogenesis, during which the embryo develops from comma to 3 fold stages. For the 8.5hr organogenesis and morphogenesis, cell differentiation occurs without further cell 10  proliferations. Matured embryos hatch into larval stage 1 (L1) at the end of the 11.5hr ex-utero embryonic development.  Postembryonic (larval) development in C. elegans consists of 4 stages (L1-4) (Figure 1.2). L1 develops to L2, L3, L4, with each stage completed by a shedding (molting) of the old cuticle. (the cuticle composition is specific for each larval stage). L1 development is marked by the birth of more neurons and differentiation of the reproductive system, which takes approximately 12hr. L2 development takes 8hr during which 2 more neurons are produced, and the gonad starts to elongate. Gonadogenesis continues in L3 stage (8hr), and completes at L4 stage (10hr). Finally, L4 matures into young adult which then takes approximately 8hrs before laying its first egg as an adult. Davis et al. (2008) found that exposure to ethanol from the time of hatching until the end of  larval development led to several physical/developmental effects in worms. Worms exposed to 0.4M ethanol, which resulted a tissue alcohol concentration equivalent to a 0.1% blood alcohol concentration, exhibited smaller body length, delayed reproductive onset, smaller brood size, and shorter lifespan than control animals. These physical/developmental abnormalities were not as prominent in worms exposed to lower doses (0.1M or 0.2M).  Interestingly, some of these physical/developmental deficits seem to be caused by separate mechanisms. For worms exposed to ethanol for their entire lives, their reproductive onset was delayed and body widths were thinner than control worms of the same age. The vulva of a 4-day-old worm exposed chronically to ethanol was as immature as the vulva of a 2-days-old in control group. If ethanol exposure acted through a common mechanism to produce both a delay in reproductive maturity and body width, 4-d-old ethanol exposed worms should have the same body width as the 2-d-old larva in the control group. However, that was not the case. The 11  body width of 4-d-old worms exposed to ethanol was smaller than the untreated L4 larva. This finding suggests that ethanol exposure impedes reproductive development and body width growth at different rates. Ethanol must act through at least two different mechanisms, one responsible for reproduction and the other responsible for body width, in order to impede two traits at different rates. Davis et al (2008) conducted the first experiment to explore the effect of ethanol exposure at different times during embryonic development in C. elegans (Davis et al., 2008). Because of the chitinous egg shell and a poorly permeable vitelline membrane protecting the embryo (Schierenberg and Junkersdorf, 1992), embryonic exposure requires a much higher concentration of ethanol than larval exposure in order to produce effects; 60min exposure to 20% ethanol, which was approximately 10X as strong as 0.4M ethanol, reduced hatch rate, but exposure to 10% ethanol did not reduce hatch rate. In addition, Davis et al. (2008) showed that the timing of embryonic exposure was important: embryonic exposure early in development reduced hatch rate more than exposure later in development. These results suggested that early embryonic development was most vulnerable to ethanol toxicology. However, whether embryonic ethanol exposure could produce similar physical effects as larval ethanol exposure could and whether different timing of embryonic exposure could produce different outcomes remained to be elucidated. Thesis objectives The experiments described in this thesis were designed to investigate the effects of ethanol exposure during embryonic development in C. elegans. The ultimate goal of this thesis was to develop a model in C. elegans in which to investigate the underlying mechanisms responsible for behavioral problems produced by embryonic exposure to ethanol. Starting with 12  the knowledge that exposure at different times during embryonic development produced different hatch rates (Davis et al. 2008); I hypothesized that worms exposed during early embryonic development would exhibit more physical/developmental and behavioral deficits than worms exposed during later development. My first objective was to describe physical/developmental effects of exposure to different doses of ethanol for different periods of time during embryonic development. First, I examined the effects of exposure to 20% ethanol for 60min at 3 different times during embryonic development: early, middle and late embryonic development. Because there were no previous data available on the possible physical effects ethanol may have during embryonic development in C. elegans, a range of physical/developmental parameters were explored to establish what sorts of physical effects might be induced. In a second experiment, based on insights gained from the 20% ethanol study, I explored a wider range of exposure patterns. This second study addressed factors important in ethanol exposure, such as dose, duration, timing, and frequency. In the third study, I investigated whether there were differences between single and multiple exposures to ethanol during embryonic development.  In these 3 studies, I found the severity of the alterations in development were dependent on the ethanol dose, exposure duration, and time.  The second objective was to study a number of behaviors in order to identify measurable behavioral abnormalities in adult animals exposed to ethanol during embryonic development. I hypothesized that C. elegans exposed to ethanol during embryonic development would exhibit altered locomotory behavior, learning, or memory. In the first and second experiments, I investigated differences in basal activity and mechanosensory responses between exposed and non-exposed animals. In the third and fourth experiment, I looked for abnormalities in short term learning in the form of habituation to mechanical stimuli (tap) in exposed animals and for 13  deficits in long term memory formation in exposed animals. I found that basal locomotory activity was affected in C. elegans exposed to chronic ethanol during embryonic development, but that learning and memory were not.  14  CHAPTER 2. PHYSICAL/DEVELOPMENTAL ABNORMALITIES INDUCED BY EMBRYONIC ETHANOL EXPOSURE IN C. ELEGANS Introduction Embryonic ethanol exposure in C. elegans The in-utero development of a C. elegans embryo begins at the time of fertilization (Figure 2.1A). The total duration of in-utero development is approximately 2.5 hr in 20oC, at which time the egg is laid outside of the hermaphrodite as a 30-cell gastrula (Figure 2.1B), which then continues to develop ex-uterally (Altun and Hall, 2005). The end of the gastrulation is marked by ventral cleft closure 3 hours into ex-utero development in 20oC, and is followed by elongation. Embryonic movement initiates soon after 1.5 fold (tadpole) stage at around 4.2 hours into ex-utero development (Figure 2.1C). After approximately 6.7 hours of ex-utero development, the 3-fold (Pretzel) stage embryo already looks like a worm (Figure 2.1D). More complex behaviors such as head movements are observable at the Pretzel stage.  Figure 0.1 Ex-utero development of C. elegans.  The important developmental events were adapted from a figure in WormAltas in www.wormatlas.org. The timing was converted from 25oC to 20oC, and adjusted to the duration of in-utero development to express timing of events occurred ex-uterally. 15  Davis et al. (2008) found that 1hr of 20% ethanol exposure during ex-utero embryonic development had timing-dependent effects on hatch rate (the number of larva hatched from eggs). Hatch rates differed for embryos exposed for an hour to 20% ethanol at different times during ex-utero development. Hatch rate in embryos exposed during early ex-utero development (1-2, 2-3, or 3-4 hrs) was approximately 10-20%, during middle ex-utero development (4-5, 5-6, or 6-7 hrs) hatch rate was approximately 50-75%, but during late ex-utero development (7-8 or 8-9 hrs) hatch rate was the same as the control embryos. Early embryonic development, 0-4hrs into ex-utero development, consists primarily of gastrulation (Altun and Hall, 2005), the purpose of which is to position 3 germ layers through cell migration and division (Wilt and Hake, 2004). Correct gastrulation is critical as it lays the foundation for organogenesis that follows. Middle embryonic development (4-7hrs ex-utero development) is mainly composed of organogenesis (Altun and Hall, 2005), the process in which 3 germ layers develop into rudimental tissues and organs (Wilt and Hake, 2004). During this time the embryo is transformed from a ball of cells into the shape of a worm (Mango, 2007). The late embryonic development is mainly composed of morphogenesis, the process in which tissues and organs take shape and specialized cells are positioned (Wilt and Hake, 2004). At this stage, deposition of the larval cuticle occurs, and the neuronal network matures to become fully functional (Sulston et al., 1983; Altun and Hall, 2005). Based on hatch rate results from Davis et al. (2008), ethanol exposure during gastrulation, organogenesis or morphogenesis produced different survival probabilities with the lowest survival rate if exposed during gastrulation and the highest if exposed during late embryonic development (3 fold stage).  In humans, ethanol exposure during development can kill the fetus. Heavy maternal alcohol consumption in humans often results in spontaneous abortion and stillbirth (1985; 16  Windham et al., 1992; Windham et al., 1997; Kesmodel et al., 2002). Similar results were found in animal models such as monkeys at 5g ethanol per kg body weight per day (5g/kg/day) (Scott and Fradkin, 1984), rats at as low as 1g/kg/day (Vaglenova and Petkov, 1998), mice at 30% ethanol-derived calories (Middaugh and Boggan, 1995), zebrafish at 2.9% ethanol (Bilotta et al., 2004), and chickens at 30% ethanol (Satiroglu-Tufan and Tufan, 2004). In C. elegans, exposure of eggs to 20% ethanol for an hour decreased the hatching probability of embryos, especially if eggs were exposed during early embryonic development (i.e. gastrulation) (Davis et al., 2008). However, the effects on hatch rate from other patterns of embryonic ethanol exposure remained to be established in C. elegans.  Growth retardation, both prenatally and postnatally, is the most commonly found characteristic in human FAS (Abel, 1981). In rat models, growth retardation was prominent in animals exposed to prenatal ethanol (4-6g/kg/day throughout gestation) (Abel and Dintcheff, 1978), but this growth retardation may or may not last into adulthood. The body weights of rats exposed to 4g/kg/day ethanol treatment eventually caught up in growth of the non-exposed rats, but rats exposed to  higher dose of ethanol (6g/kg/day) remained smaller well into adulthood (5months-old) (Abel and Dintcheff, 1978). Similarly, Davis et al. (2008) found that C. elegans exposed to a high dose (0.4M or 2% v/v) of ethanol during larval development were smaller than non-exposed worms throughout their life, but C. elegans larva treated with lower doses (0.1-0.2M or 0.5-1% v/v) reached normal size as they aged (Davis et al., 2008). In contrast, Abel (1981) found that a low dose of ethanol (1-2g/kg/day) did not produce any postnatal growth retardation. This dose-dependent characteristic in growth retardation can be very useful to judge the severity of ethanol exposure in C. elegans. 17  Reproductive abnormalities have been demonstrated in some studies of rat models of FASD. A few studies have found that the onset of puberty was delayed (Boggan et al., 1979; McGivern et al., 1992; McGivern and Yellon, 1992), and the cessation of menstrual cycle was earlier (Wilson et al., 1995). Together these two effects resulted in a shorter time window for reproduction (Wilson et al., 1995). If such reproductive abnormalities occur in C. elegans, the brood size (total number of eggs laid) should be reduced. Davis et al. (2008) found smaller brood sizes in worms that experienced larval exposure of 0.1, 0.2 and 0.4M ethanol. In addition, reproductive onset was delayed in the worms exposed to the highest dose (0.4M), though not in the worms treated with lower doses. The current study will investigate whether reproductive abnormalities also occur in animals exposed to ethanol during embryonic development.    Health problems caused by ethanol teratology can induce a shorter lifespan. Studies on the effects of FASD on human life expectancy are scarce, which may be due to difficulties in tracking individuals and in controlling numerous factors involved in lifespan expectancy. A North Dakota study suggested that FASD was associated with higher mortality rate (Burd et al., 2008). This study suggested that FASD may be associated with shorter life spans due to health problems that were already present at birth, or problems that normally occurred at an older age, for example, death due to cerebral palsy at the age of 22, respiratory disease at the age of 11, and cardiac abnormalities at the age of 28. More intriguing was that the siblings of the subjects all died at infancy due to birth defects. A study using a rat model of FASD also determined that exposed animals lived shorter lives (Abel et al., 1987). In C. elegans, worms exposed to a high dose of ethanol during larval development had a shorter lifespan than non-exposed controls (Davis et al., 2008). Thus, lifespan is an interesting variable to examine in any model of FASD.   18  Chapter objective and hypothesis The main objective of Chapter 1 was to establish an ethanol embryonic exposure protocol that produces consistent physical/developmental phenotypes that may mirror abnormalities found in FASD. Since Davis et al. (2008) found that exposure to 20% ethanol for an hour at earlier stages during embryonic development produced a lower hatch rate than exposures at later stages, I hypothesized that physical abnormalities produced by exposure to 20% ethanol during early embryonic development would be more severe than by exposure during middle or late embryonic development. In Experiment A-1, I investigated changes in hatch rate, body size, reproductive onset, reproductive pattern, brood size, and lifespan in C. elegans exposed to 20% ethanol for an hour during early, middle or late embryonic development. I found that 20% ethanol produced different sets of physical/developmental abnormalities in animals exposed during different times, but abnormalities in animals exposed during early development were not worse than those of animals exposed later. For Experiment A-2, I hypothesized that the ethanol dose we used in Experiment A-1 wasn’t strong enough to produce consistent deficits. Bilotta and colleagues (2004), using a zebrafish model of FASD, demonstrated that only with exposure to a near-lethal dose of ethanol could you find a 100% frequency of physical abnormalities which eventually causes death in the affected fish; when the ethanol dose was severe enough to reduce the survival rate by 15%, only 60% of the subjects exhibited physical dysmorphology. Therefore, in Experiment A-2, I explored exposures to a broad range of ethanol concentration that lasted for specific lengths of time at specific periods of embryonic development. In addition, since pilot findings from our lab suggested that multiple exposures at different times during embryonic development caused lower hatch rate than a single exposure (unpublished result), the effects of three exposure episodes at 19  early, middle and late development (repeated exposure) were also investigated. I found that worms exposed during early development and worms that experienced repeated embryonic exposures to 30% ethanol had higher levels of growth retardation than ones experienced only a single exposure episode during middle and late development. Moreover, worms that experienced repeated exposure to 40% ethanol for a total of 15min exhibited more signs of ethanol teratology than worms that experienced a single 15min exposure episode to 40% ethanol during early development. Therefore, I hypothesized that repeated exposure would produce worse physical deficits than a single exposure. In Experiment A-3, I investigated the effects of repeated exposures to 30% ethanol for 60min, and compared that to the effects of single exposures for 20 or 60min during early, middle or late development. I found that repeated exposure indeed produced worse outcomes. Materials & Methods Animals maintenance Wildtype (Bristol N2) C. elegans obtained from the C. elegans Genetic Center were used in all experiments. Animals were cultivated on Nematode Growth Medium (NGM) at 20oC seeded with Escherichia coli (OP50) as described in Brenner (1974).  Embryo (egg) collection To collect age-synchronized eggs (embryo), 30 or more 4-days-old adult worms were transferred onto a 5-cm NGM plate seeded with E Coli to lay eggs for 15-30 minutes (for all experiments except for experiments involving 8hr of chronic exposure, for which adult worms were allowed to lay eggs for 60min in order to collect more eggs; adult worms transferred to a new NGM plate often did not lay eggs for the first 10-15min, thus a longer egg laying time was allowed to collect more eggs). Hour zero was defined as the time when the first adult animals 20  were placed onto the plate. For example, 2-hour-old eggs were defined as 120 minutes after placing the first adult animals onto the plate for egg laying. All eggs were laid at the gastrula stage (determined by visual inspection); however, the age of the eggs on a plate may not be precisely identical. The only way to precisely synchronize the age of the eggs was to pick individual eggs that were at 2- or 4-cell-stage, and that was not a feasible method for large scale studies such as those described in this thesis. As a result, the ages of the eggs collected by the method described in this thesis were not precisely synchronized, and our conclusions have taken this age variation into consideration. Ethanol exposure procedures Two alternative ethanol exposure procedures were used in this study. The Centrifuge procedure was adapted from Davis et al. (2008), and was used in Experiment A-1. Later, the Centrifuge procedure was modified into the Agar-Bath procedure to address issues such as contamination, reliability of egg counting (Appendix A), and effect of spinning on embryos. All exposure procedures were conducted in a 20oC temperature controlled room. A. Centrifuge procedure: 10-15 minutes before the onset of exposure, eggs were washed off plates with ddH2O into 1.5ml Eppendorph tubes. This egg solution was spun for 5 minutes at 6600rpm in Fisher Scientific Mini Centrifuge, and the supernatant was first removed by aspiration with pipette, and a small part of the leftover supernatant was absorbed by a piece of Kimwipe® so that the subsequent ethanol solution would not be diluted by the leftover supernatant. At the onset of exposure, 1ml of 20% ethanol or ddH2O was added to the 1.5ml tube containing eggs. Eggs were re-suspended in this 1ml treatment solution, and incubated in it for the duration of treatment.  To remove the ethanol a multistep procedure was: 5 minutes before the end of exposure, the treatment solution was spun at 6600rpm for 5minutes and the 21  supernatant was removed by aspiration. The pellet was washed twice with ddH2O and re-collected by centrifugation. Any remaining liquid was absorbed away from the pellet by a small piece of Kimpwipe. Pellets were resuspended in 100-200µl ddH2O and dispensed onto a fresh NGM plate. The eggs then were allowed to air-dry. B. Agar-bath procedure: 5 minutes before the onset of exposure, a piece of agar with eggs laid on top of it was cut out from an agar filled 5cm Petri plate using a sterilized scalpel and placed into an empty 5-cm Petri plate. At the onset of exposure, eggs were immersed in either ddH2O or freshly prepared ethanol solution. Because the discovery that water exposure at different times during development may have different effects (detail see Appendix C), experiments using the Agar-Bath procedure implemented ddH2O (water) controls for every treatment combination. The “untreated group” was used to control for the effects caused by water immersion; the eggs in the untreated group were left alone on the plate without liquid immersion of any kind. The difference between the untreated and water control was assumed to be induced by water, and the difference between the ethanol exposure group and its corresponding water control was assumed to be induced by ethanol alone. At the end of exposure, the solutions were poured away. The piece of agar was washed once briefly with ddH2O to eliminate any residual ethanol solution, and then incubated 5 minutes in ddH2O to allow diffusion of any residual ethanol inside the egg. Eggs were dried briefly, and transferred to a fresh NGM plate using a clump of E. Coli as sticking agent. Eggs in the untreated group were also transferred in the same manner to control for possible mechanical damage resulted from egg transferring.    22  Hatch rate Hatch rate was defined as the number of larva divided by the number of eggs collected. In the Centrifuge Procedure, egg counting was only possible after the suspension liquid was completely dried, which was about 30 to 120 min after the exposure. In the Agar-Bath Procedure, eggs were counted immediately after transfer onto a fresh NGM plate. In each procedure, the number of hatched larva was counted 24 hour after the commencement of egg collection.  Reproductive onset  In this experiment, reproductive onset was defined as the age in hours when an animal laid its first egg. Individual 48-hr-old animals were transferred onto their own isolate plates (NGM with a drop of E. coli). Isolate plates were checked hourly for signs of eggs starting from 65hr of age until 80hr of age. If an animal had no eggs on its plate at 71hr but had eggs at 72hr, the reproductive onset for this animal was 72. In Experiment A-1, worms that did not lay eggs before 80hr were excluded from the analysis. In subsequent studies (1.2 & 1.3), animals that failed to lay eggs before 80hr were given an estimated reproductive onset according to the number of eggs present the next day at hour 96. Since undisturbed C. elegans lay about 4-10 eggs per hour, the number of eggs were divided by 8 in order to estimate the reproductive onset. For example, if 49 eggs were present at hour 96, the first egg laid was estimated to have been laid 6hr ago, and hour 90 was recorded as the reproductive onset for this animal. Animals that failed to lay eggs after hour 96 were recorded as “Never Laid”. Any worms that died before laying their first egg were omitted from the numerical analysis but were included in the categorical analysis of reproductive onset.  23  Egg-laying (reproductive) pattern & brood size The number of progeny (eggs and hatched larva) from an individual was counted after the worm was transferred to a fresh NGM plate seeded with a drop of E. Coli; the transferring occurred every 24hr starting at 72hr of age until the worm stopped laying eggs; this transferring was necessary to avoid losing the original hermaphrodite among its large number of fast-growing progeny. Eggs lacking distinct oval shapes or with darkened interiors which often laid by older hermaphrodites were not counted because those were thought to be unfertilized eggs. Egg-laying patterns were represented as the number of progeny produced on each reproductive day. Brood size was defined as the total number of progeny an animal produced in its lifetime.  Body size Body size was defined as the total surface of a worm on a 2-D image. Animals were filmed with iMovie using the Mac OS-X operating program through a Leica ICA digital camera attached to a Leica WILD M3Z dissecting microscope at 60X (Figure 2.2). JEPG images of each worm were clipped from iMovie, and worm sizes were digitally measured with Image J software. A detailed procedure on how measure worm size in Image J program is described in Appendix B, which also lists macro programs written to automate part of the scoring process.   Figure 0.2 Apparatus for filming worm in iMovie 24  Lifespan Lifespan was defined as the age in days when the animal ceased to react to mechanical stimuli. Animals were checked daily for locomotory activity and any non-active worms were touched once gently on the head with a platinum wire since old worms were often inactive but would respond to touch with slight twitching. If that first touch did not initiate any movement, three consecutive touches were further administered. If the animal failed to respond to those stimuli, the animal was considered dead.  Statistical analysis In Experiment A-1, statistical differences among groups in the reproductive onset, brood size, and lifespan were analyzed with Analysis of Variance (ANOVA) between treatment condition (water, early, middle & late) using SPSS 17.0; a one-way ANOVA was used; the post-hoc Tukey (HSD) method was used to compare between two groups (Glass and Hopkins, 1995); for the pattern of reproduction and body growth, the differences among groups were analyzed with repeated measures two-way ANOVA across ages (Days) between treatment solutions and onsets; the reproductive patterns between two groups were analyzed with post-hoc HSD as described above, and the number of eggs laid within a day was analyzed with post-hoc Dunn (Bonferroni) method of multiple planned comparison (Glass and Hopkins, 1995) with α = 0.05 using QuickCalcs Online Calculators for Scientists (GraphPad Software Inc.) which used the following formula and procedure: 1) t statistics was calculated for each comparison as  where MSresidual was the mean square for the residuals calculated in ANOVA and compared to 25  t* which was the value of the t ratio that correspond to a two-tailed p value of 0.05 divided by the number of comparisons made within the dataset taking into account the number of degrees of freedom (df) used in the ANOVA, and 2) if the t statistics was greater than t* than the comparison was significant at p = 0.05; for the survival curves, the Gehan Wilcoxon test, designed to compare whether a treatment changed survival rate, were used to analyze for the survival time during adulthood as described in Murakami and Johnson (1996), and the difference between two survival curves were analyzed with post hoc pair-wise comparison for Wilcoxon survival tests using SPSS.  Due to the large number (55) of the treatment conditions in Experiment A-2, data were analyzed in groups: 1) chronic exposure of 8.5hr, 2) chronic exposure of 8hr, acute exposure during 3) early, 4) middle, and 5) late development, and 6) repeated exposure; attempt to analyze all 55 groups with three-way multiple ANOVA revealed significance of all 3 factors (dose, duration, and onset), but post-hoc tests that followed needed to compare 1485 pairs of groups; for post-hoc statistical tests that took the number of comparison into account, the power became so low that none except for a few pair of groups were significantly different. The hatch rate was analyzed using Chi-Square test because only one dataset was collected with a sample size of 50-150; the number of hatched egg was designated as one category and the number of un-hatched eggs was designated as the second category within a treatment onset; the frequency of hatched and un-hatched eggs of the ethanol exposure groups were compared with the untreated and water controls; only a hatch rate that showed a significant difference from both the untreated and water controls was considered as significantly altered by ethanol exposure. The reproductive onset was analyzed within groups described above with a two-way ANOVA between treatment solution and duration. The growths of body sizes from Day 1-4 were analyzed within the same treatment 26  onset and duration with one-way multiple ANOVA using ethanol dose as a between subject variable and age (Day1-4) as a multiple variable; the body sizes were not analyzed with repeated measures ANOVA because the data were randomly sampled from a group of worms. All pair-wise comparisons were analyzed using Tukey’s method as described above instead of planned comparisons because every possible pair was analyzed for statistical significance.  In Experiment A-3, a total of 6 replications were collected for each experimental group. Thus the statistical differences in the hatch rate, reproductive onset, lifespan, and body size were analyzed by multiple ANOVAs for treatment solution, onset, and duration, and differences between two groups were analyzed by post-hoc planned comparison described above; the survival curve during adulthood was analyzed in the same manner as in Experiment A-1. In Experiment A-2 & 3, strict criteria were used to designate significant changes caused by ethanol exposure: a measure from the ethanol group was considered as showing a significant difference only if the measure was significantly different from both the corresponding water control and untreated control because 1) the difference between the ethanol and water groups were assumed to be caused by ethanol alone, and 2) the measurement collected from untreated control group, which did not go through any kind of liquid immersion, represented the norm; in addition, the direction of change must be the same; for example, if the hatch rate of the ethanol group was 85%, water control was 90%, and untreated control was 80%, it would be illogical to conclude that ethanol treatment reduced hatch rate compared to water control but increased hatch rate compared to norm (untreated control). 27   Results  Experiment A-1: Acute 20% ethanol exposure at different developmental stages induced different developmental abnormalities  In Experiment A-1, I characterized physical/developmental deficits resulting from 1 hour of 20% ethanol exposure at 3 different times during embryonic development in C. elegans (Figure 2.3). The exposure method used in this experiment was described in Davis et al. (2008). The eggs in the early exposure group was treated with 20% ethanol in ddH2O (v/v) during 2-3hr ex-utero development, the middle exposure group during 5-6hr ex-utero development, and the late   Figure 0.3 Ethanol exposure timing in Experiment A-1 Early exposure started from 2hr ex-utero development, middle exposure from 5hr ex-utero development, and late exposure from 8hr ex-utero development.  exposure group during 8-9hr ex-utero development; the water control group was treated with 0% ethanol (ddH2O) during 2-3hr ex-utero development. One problem with the Centrifuge procedure was that it prohibited reliable egg counting (details described in Appendix B); for example, the egg count would be 50 but the larva count would be 60, which resulted in an unrealistic 120% hatch rate; this unreliability was due to the fiber debris and residual washing liquid obscuring the experimenter’s view and the finding that the eggs often clumped together which made accurate 28  counting extremely difficult; therefore, the hatch rate results were excluded from the report of Experiment A-1 and efforts were made in the subsequent experiments (A-2 & 3) to resolve this problem.  Reproductive onset, Brood Size, and Reproductive Pattern The study of reproduction in Experiment A-1, in which worms were exposed to 20% ethanol during early, middle, and late embryonic development, included the measures of reproductive onset, brood size, and reproductive patterns (the number of eggs laid across the age of worms). Reproductive onset, which was the time when a worm laid its first egg, was significantly different among ethanol exposure groups and the water control (F(3, 124) = 4.598, p = 0.004) (Figure 2.4A). The post hoc HSD analysis showed that the reproductive onset of the late exposure group was significantly delayed compared to the water control group (p = 0.003, HSD) and the early exposure group (p = 0.028, HSD). However, the brood size, the total number of eggs laid during a worm’s lifetime, did not show any differences among treatment groups according to the ANOVA (Figure 2.4B). The reproductive pattern was illustrated as the number of eggs laid across age (days), and a repeated measures ANOVA revealed no difference between groups from Day 2-9 (Figure 2.4C), but within groups, age (F(7) = 174.066, p < 0.0001) had a significant effect on the number of eggs laid, which made sense because worms laid the most of their eggs during Day 3-4, the number gradually diminished from Day 5 and approached zero after Day 9. Post-hoc HSD comparison also suggested the reproductive patterns weren’t different between any two groups. Planned comparison using Dunn (Bonferroni) method revealed no difference in the number of eggs laid within a particular day between any two treatments conditions that were hypothesized to be different (Day 2, 3, and 7). This lack of difference was 29  likely due to the large number of planned comparisons executed. Overall, only the late exposure group exhibited any reproductive abnormalities.  Figure 0.4 Acute exposure of 20% ethanol at different times during embryonic development delayed reproductive onset in the late exposure group but did not altered the reproductive patterns or brood sizes.  (A) The late exposure group began to lay their first egg later than the water control and early exposure group. The brood sizes (B) and reproductive patterns (C) were not altered by exposure to 20% ethanol for an hour during embryonic development. The number within the bar in A) and B) are sample sizes. *, p < 0.05, **, p < 0.01  Body Size In mammals, the smaller birth weights and slower body weight gain are common effects caused by heavy prenatal ethanol exposure (Abel, 1982); because the body weight of a single C. elegans was technically difficult to obtain, the measurement of body sizes from a 2-D image were used in place of body weight measurement. A repeated measures ANOVA indicated that 30  the pattern of body size did not show difference between groups throughout the age for Day 1-8, 9-10, 11-12, 13-14, and 15-21 (Figure 2.5); age had a significant effect (F(11) = 153.505, p < 0.0001) on body size, as expected, as worms grew more than 10 times in size from Day 1 to Day 4. The comparisons of the growth patterns of body sizes did not show any difference between the ethanol and water groups, and post-hoc comparison within Day 1, 3, and 15-21 between the ethanol and water groups did not show any difference. Thus body size wasn’t affected by exposure to 20% ethanol for an hour during embryonic development.    Figure 0.5 1hr exposure to 20% ethanol at different times during embryonic development did not alter the growth pattern of body sizes (µm2)  or body sizes within particular days.  The mean body sizes of  the late exposure group on Day 3 appeared to be smaller but post-hoc analysis did not reveal significant difference.  The decreasing body sizes after Day 13 was a normal  aging process in C. elegans.  Lifespan Ethanol exposure during larval development reduced lifespan (Davis et al., 2008) so we hypothesized that ethanol exposure during embryonic development would also do the same. However, lifespan was not affected by exposure to 20% ethanol for an hour at early, middle or late stages of embryonic development (Figure 2.6A). Although the mean lifespan of the early  31   Figure 0.6 Lifespan and adult survival rate was not altered by acute exposure of 20% ethanol during early (2-3h EUE), middle (5-6h EUE), and late (8-9h EUE) embryogenesis.  A) Lifespan was not different among and between groups, and that was partially because B) lifespan was a highly variable measure. C) Survival rate plot among all 3 exposure groups and water control were not different. D) Early, E) Middle, nor F) Late exposure group showed difference in survival rate.  exposure group was 2 days (12 days) shorter than the water control (14 days), however, because lifespan was another highly variable measure (Figure 2.6B), no significant differences were found between the two groups. The survival curves during adulthood displayed in Figure 2.6 C-F shows the percentage of individuals that remained alive starting from Day 4 to 20, a method to 32  assess survival probability adapted from Murakami and Johnson (1996); the larval survival curves were not compared because very few worms died during larval stages (Day 1-3). Although the survival curve of the early exposure group appeared to be lower than that of the water control, the two curves weren’t significantly different (p = 0.078, Wilcoxon pair-wise), and the survival curves of the rest of the ethanol exposure groups weren’t different from that of the water control either. Thus life expectancy didn’t appear to be affected by one hour of 20% ethanol exposure during embryonic development.  Summary of Experiment A-1 Based on the findings described by Davis et al (2008) that the exposure of 20% ethanol for an hour during early stages of embryonic development reduced hatch rate more than did exposure during middle or late stages, I hypothesized that the same dose of ethanol would produce more physical/developmental effects in worms exposed during early embryonic development. However, that did not appear to be the case. The only physical/developmental effect I found was a slight delay of reproductive onset in the worms exposed during late embryonic development; the reproductive onset of the late exposure group was 72.4hr±0.6 and the water group was 69.1hr±0.5. Considering that some worms in the water control group also laid eggs at 73hr, the delay found in the late exposure group was small. In addition, the reproductive onset of the worms exposed to 0.4M (~2%) ethanol during larval development was delayed by approximately 70hrs, which is almost 3 days (Davis et al., 2008); the 3.3hr delay found in the worms exposed during late embryonic development was very small compared to the delay caused by larval exposure. The body sizes, on the other hand, appeared to be larger in the ethanol exposure groups than water groups after Day 13 (Figure 2.5). Interestingly, Davis et al. (2008) also showed a 33  similar finding of larger body size in older ethanol exposed worms compared to water control. Davis et al. (2008) showed that a bigger mean body size during old age in the ethanol exposure group was due to smaller worms dying first. In summary, Experiment A-1 did not support the hypothesis that the worms exposed to 20% ethanol for an hour at early embryonic development would exhibit more physical effects; in fact, the worms exposed during late embryonic development were the only group that showed any consistent physical effect. Experiment A-2: Ethanol dose and duration needed to surpass a certain threshold to induce physical/developmental deficits   The unexpected results from Experiment A-1 lead to the hypothesis that the ethanol dose used in the previous experiment wasn’t strong enough to produce considerable physical/developmental effects. In Experiments A-2, I investigated a range of ethanol exposure patterns with different doses, durations, and frequency during embryonic development. This experiment had four main goals:  1) to establish the ethanol concentration and 2) exposure duration required to induce physical effects at particular times during embryonic development, 3) to explore whether exposure during early development would result in more severe physical/developmental effects at a higher ethanol dose, and 4) to investigate whether multiple exposures would produce more severe physical/developmental effects than a single exposure.   The exposure protocol (Centrifuge method) used in Experiment A-1 created several problems. One problem was that after centrifugation, the complete removal of water, which was used to wash off eggs from the agar plate, required both aspiration using pipette and absorption using a Kimwipe®; the fiber debris from the Kimwipe® increased the chance of contamination and obscure the visibility of the eggs, which created problems with the egg counting for hatch 34  rates. In addition, the centrifuge introduced a centrifugal force at 6600rpm, which might stress the developing embryos, and also caused the eggs to clump together after the treatment, which further obscured the egg counting. Moreover, the time required to air-dry eggs varied substantially (1-3hr), which introduced variable durations of osmotic stress to different replications.  The yield of this egg collection method was also very unpredictable; sometimes no eggs were left at the final collection stage even though hundreds of eggs were laid on the starting plate; for example, if 2 different experimental plates each had 100 eggs, after several washings and centrifugations, there might be 95 eggs left for one plate but zero eggs for another plate. This unreliable egg collection had created severe problems for the experimental design as one replication of an experiment might successfully collect eggs from only 2 treatment groups but other replications successfully collected from all 4 groups .To address these problems, an Agar-Bath method was established and implemented in Experiment A-2: the eggs were left on the agar instead of being washed off; the pieces of agar containing eggs were dissected off and incubated in the treatment solutions; since no liquid was required to transfer eggs onto a new growth medium, the drying time after the treatment was only 5-15min; 1-10 eggs were transferred each time onto the new growth medium and eggs were carefully separated from each other so that egg counting became highly accurate. This new method not only improved many aspects of the protocol used in Experiment A-1, but also made short and long exposure durations possible. The centrifuge method required 5min of centrifugation and approximately 30 sec to remove ethanol solution; the Agar-Bath method required 2-5 sec to pour away ethanol solution and wash away residual ethanol solution with ddH2O. Therefore, in Experiment A-2, exposure duration as short as 5min could be performed with reasonable speed of handling. The centrifuge method was also limited by the number of slots available within the centrifuge equipment, which was 6 in our 35  portable centrifuge, so that only a restricted number of treatment conditions could be done within a given time period. There was no such limitation with the Agar-Bath method; as a result, 16 different treatments could begin at the same time in Experiment A-2. Additional consideration was given to the possible effects exerted by osmotic stress induced by exposure to distilled water at different time for different duration and frequencies (detail see Appendix C). Therefore, every ethanol exposure pattern was controlled with a corresponding water control. Theoretically, M9 (the physiological saline for C. elegans) would be the best vehicle for the ethanol treatment, but the salt crystals left behind after M9 evaporated caused tremendous difficulties with egg counting (Appendix D). In addition, the salt crystals created a local hypotonic environment for hatched larva, which would introduce even more confounds into our study; therefore, we used distilled water as the vehicle. Finally, an untreated group in which no liquid immersion of any kind was experienced by the embryos was included to control for effects induced by water control treatment.  Figure 0.7 Treatment conditions in Experiment A-2. Groups exposed to chronic exposure for 8.5hr was treated from 0.5-9hr ex-utero development.  Groups exposed to chronic exposure for 8hr was treated from 1-9hr ex-utero development. Groups exposed during early exposure initiate their treatment at 2hr ex-utero development, middle exposure at 5hr ex-utero development and late exposure at 8hr ex-utero development. Groups exposed to repeated exposure condition were treated for 3 episodes each began at 2hr, 5hr, or 8hr ex-utero development. 36   There were 55 conditions in this experiment. I explored 5 different exposure durations (8.5hr, 8hr, 1hr, 30 min, and 15 min), 3 developmental periods (early, middle, and late), 5 doses for an 8.5hr exposure (5, 10, 20, 30, and 40%), 6 doses for an 8hr exposure (5-10%), 3 doses for acute exposure (20, 30, 40%), and 2 exposure frequencies (single and triple). Figure 2.7 is a schematic diagram showing when exposures occurred in relation to ex-utero embryonic development. For the 8.5hr chronic exposure, embryos were exposed during 0.5-9hr ex-utero development; for the 8hr chronic exposure, embryos were exposed during 1-9hr ex-utero development. For acute exposures, the treatment of early exposure groups began at 2hr ex-utero development, middle exposure groups at 5hr, and late exposure groups at 8hr. The repeated exposure group was treated for 3 times which began at 2, 5, or 8hr ex-utero development; each exposure episode lasted for a third of the total duration specified. The results from this experiment are organized into “chronic exposure”, “single exposure”, and “repeated exposure” for each variable: hatch rate, reproductive onset, and body size. Hatch Rate Chi-square analysis was used to compare the hatch rate frequency (the number of hatched eggs vs. the number of un-hatched eggs) of the ethanol exposure groups with the untreated and water controls. A significant change of hatch rate was determined to have occurred when the hatch rate of the ethanol exposure group was different from both the untreated and water controls.  Chronic Exposure: Hatch rates varied significantly between groups of embryos exposed to treatments chronically for 8.5hr from 0.5hr to 9hr ex-utero development (X2=454.428, df = 5, p < 0.0001) (Figure 2.8A). Hatch rate of embryos exposed to a 5% concentration were not reduced but in 10% decreased significantly (X2=130.325, df = 1, p < 0.0001). No embryo 37  exposed to 10% ethanol or higher for 8.5hr chronic survived. Inspired by the dramatic difference in hatch rate between the 5% and 10% chronic exposures, a total of 8hr chronic exposure starting from 1hr-9hr ex-utero development were explored for concentrations between 5-10%; the 8hr duration was chosen this time to allow for a more realistic time frame for egg preparation, as   Figure 0.8 Chronic exposure reduced hatch rate  A) Chronic exposure for 8.5hrs reduced hatch rate to nearly zero at 10%. No embryos hatched at 20% or higher ethanol. B) Chronic exposure for 8hrs reduced hatch rate at 8% or higher ethanol. “Unexposed”  or “NR” represented no treatment groups, in which no liquid immersion was applied to embryos.  30min was a too brief of a window to obtain, prepare and treat enough eggs for all of the required groups. Chronic exposure for 8hr induced significantly different hatch rates between groups (X2= 71.182, df = 7, p < 0.0001) (Figure 2.8B); hatch rates began to decrease significantly at 8% ethanol or higher (8hr, X2= 4.728, df = 1, p = 0.03, 9hr, X2= 9.878, df = 1, p = 0.002, 10%, X2=13.872, df = 1, p < 0.0001). Interestingly, the chronic exposure for 8hr began only 30min 38  later than the chronic exposure for 8.5hr (1hr vs. 0.5hr ex-utero development, respectively), but this 30min delay produced drastic differences in the resulting hatch rate: the chronic exposure for 8.5hr to 10% ethanol reduced hatch rate to nearly zero while the chronic exposure for 8hr to the same dose only decreased hatch rate to 0.71. This observation suggested that the first 30min of ex-utero development was particularly vulnerable to the effect of ethanol.  Single Exposures: The description of the results from single exposures was partitioned into early, middle and late exposure conditions. Firstly, analysis of early exposure groups found that 30% ethanol exposure for 60min during early development decreased hatch rate compared to the water control (X2= 134.993, df = 1, p < 0.0001) and untreated group (X2= 89.086, df = 1, p < 0.0001); no difference in hatch rate were found in any dose treated for 30 and 15min during early   Figure 0.9 Hatch rate dropped when ethanol treatment severity reached a threshold, and this threshold was unique to each developmental stage and treatment frequency.  A) Hatch rate reduced when ethanol treatment severity reached 30% or higher for an hour in early exposure groups. B) Hatch rate was reduced in exposure for an hour at 40% ethanol during middle development. C) Hatch rate was reduced when ethanol exposure reached 40% in late exposure groups. D) Hatch rate was reduced when ethanol exposure reached 30% or higher for an hour or 40% for  30 and15mins in repeated exposure group. The dotted line represents hatch rate in No Treatment group.  39  exposures (Figure 2.9A). For exposures that occurred during middle development, only exposure to 40% ethanol for 60min decreased hatch rate (vs. water, X2= 14.587, df = 1, p < 0.0001, vs. untreated, X2= 8.632, df = 1, p = 0.003) (Figure 2.9B), and no other treatment conditions that occurred during middle development made a significant difference in hatch rate. For exposures during late development, exposure to 40% ethanol for 60min induced decreased hatch rates compared to the water control group (40% 60min vs. water, X2= 96.730, df = 1, p < 0.0001, vs. untreated, X2= 96.122, df = 1, p < 0.0001); no other treatment conditions during late development altered hatch rate.  In summary, to reduce embryo survival rate (hatch rate), treatment strength stronger than 30% ethanol for 60min was required for exposure during early development, and 40% ethanol for 60min was required for exposure during middle and late development. The ethanol dose required to reduce survival rate of early embryos appeared to be less than that of middle and late embryos. This result supported the hypothesis that early embryos were more easily killed by ethanol exposure than middle and late embryo, which was similar to the hatch rate result reported in Davis et al. (2008). Repeated Exposure: In repeated exposure conditions, the total specified duration was divided across three developmental time points; for example, worms exposed in the 15min repeated exposure condition were actually exposed to 5min each at early, middle and late embryonic development. For the 60min repeated exposure condition (20min each at early, middle and late), hatch rates were significantly reduced by exposure to 30% and 40% ethanol (30%, X2= 57.089, df = 1, p < 0.0001, 40%, X2= 146.039, df = 1, p < 0.0001). (Figure 2.9D). For both 30 and 15min repeated exposure to 40% ethanol significantly decreased hatch rate (30min, X2= 18.573, df = 1, p < 0.0001, 15min, X2= 24.525, df = 1, p < 0.0001). Repeated exposure for a 40  total of 30 min or 15min to 40% ethanol reduced hatch rate; however, the same strength of ethanol treatment in early, middle or late single exposure groups did not. Considering the 30 and 15min repeated exposure protocol lasted merely 10 and 5min at each stage, the effect of multiple exposures produced much greater decline in hatch rate. Generally, this result supported the hypothesis that repeated exposure produced a more severe outcome in hatch rate than single exposures.  Reproductive Onset Chronic Exposure: Reproductive onset for the worms exposed to ethanol chronically for 8hrs starting from 0.5hr ex-utero development were different among groups (F(2) = 9.249, p = 0.001) (Figure 2.10A). Since only the 5% ethanol group had a high enough number of hatched worms to analyze statistically, other ethanol doses were excluded in the study of reproductive onset. The reproductive onset of worms treated with 5% ethanol chronically was delayed compared to the water control (p = 0.002, HSD) and untreated control (p = 0.03, HSD).  Single Exposure: The reproductive onsets of worms exposed to a single exposure to 0, 20, 30 or 40% ethanol for 60, 30 or 15min during early, middle or late embryonic development were described here. For exposure during early embryonic development, dose (F(3) = 4.458, p = 0.006) and duration (F(2) = 90.708, p < 0.0001) each had significant effect on the reproductive onsets, but a two-way ANOVA did not show a significant interaction between dose and duration (Figure 2.10B); comparisons between groups showed that the reproductive onset of worms exposed to 30% ethanol for 60min was delayed compared to the untreated control (p = 0.021, HSD), but not to the water control; exposure to 40% ethanol for 30min was delayed compared to both untreated (p < 0.0001, HSD) and water controls (p = 0.016, HSD); the rest of the exposure  conditions in the early exposure groups did not produce significant change in the reproductive 41    Figure 0.10 Reproductive onsets were delayed by chronic exposure, and more effectively by the repeated exposure than by single exposures.   A) Chronic exposure to 5% ethanol for 8.5hrs caused delay in reproductive onset. B) Only early exposure to 40% for 30min delayed reproductive onset. No treatment conditions tested for the middle (C) or late (D) exposure groups resulted in reproductive delay. E) Repeated exposure produced delay in worms exposed to 30% ethanol for a total of 60min (20min per exposure episode), 20-40% ethanol for 30min (10min per exposure episode), and 40% ethanol for 15min (5min per exposure episdoe. ***, p < 0.0001, **, p < 0.001, *, p < 0.05, for comparison to both water and untreated controls. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition. Solid horizontal line in B-E) represents the mean reproductive onset of untreated control. X, no embryo survived the treatment. 42  onsets. For worms treated during middle development, a two-way ANOVA showed that duration (F(2) = 4.400, p = 0.015), but not dose, had a significant effect on the reproductive onsets, and there was no significant interaction between dose and duration (Figure 2.10C); middle exposure for 60min to all 3 doses showed a significant delay compared to the untreated control (20%, p =0.032, 30%, p = 0.025, 40%, p = 0.010, HSD), but all 3 were not significantly different from the water control; the rest of the exposure conditions in the late exposure groups did not produce significant change in the reproductive onsets. The exposures during late development revealed that only duration had a significant effect to reproductive onset (F(2) = 11.082, p < 0.0001) (Figure 2.10D), and no ethanol group had a significantly different reproductive onset from its corresponding water control. In summary, single exposures did not produce dramatic effects on reproductive onsets, and the only exposure condition that produced any significant delay was the early exposure to 40% ethanol for 60min. Repeated Exposure: This section describes the reproductive onsets of worms that were exposed 3 times during early, middle or late embryonic development; each exposure episodes lasted for one third of the total duration specified. A two-way ANOVA showed that both dose (F(3) = 29.257, p < 0.0001) and duration (F(2) = 16.574, p < 0.0001) alone had significant roles to reproductive onset (Figure 2.10E). In addition, the interaction between dose and duration also had significant effect on reproductive onset (F(5) = 8.701, p < 0.0001). The repeated exposure to 30% ethanol for a total of 60min, which was 20min at each developmental stage, produced significantly delayed reproductive onset compared to that produced by the water (p <  0.0001, HSD) or untreated controls (p < 0.0001, HSD); the 40% ethanol wasn’t tested because no worms were hatched from this exposure condition. All three doses tested for the 30min repeated exposure condition produced significantly delayed reproductive onsets compared to both water 43  (20%, p = 0.033, 30%, p = 0.018, 40%, p = 0.002, HSD), and untreated controls (20%, p = 0.004, 30%, p = 0.002, 40%, p < 0.0001, HSD). Exposure for a shorter period of time, 15min in total or 5min for 3 episodes each at early, middle or late development, only produced a reproductive delay in worms exposed to 40% ethanol (40% vs. water, p = 0.002, vs. untreated, p = 0.004, HSD). In summary, repeated exposure produced much more severe effects on reproductive onset than did single exposure for comparable durations; 40% ethanol exposure for only 5min for 3 times during embryonic development produced marked delay in the reproductive onset. These results supported the hypothesis that repeated exposure to the same dose of ethanol produced a worse outcome than a single exposure for the same duration. Body size  Growth retardation  has commonly been found in animal models of FASD and in humans exposed to heavy doses of alcohol during development (Abel, 1985). Heavy ethanol exposure during C. elegans larval development was also found to reduce worm size (Davis et al., 2008). Therefore, reduced body sizes were predicted to be found in worms exposed to ethanol during embryonic development. Body size was measured as the body area of 2D images of worms. Chronic Exposure: For worms exposed to 8.5hr of treatment throughout embryonic development, body sizes across Day1-4 were compared between untreated, 0% (water) and 5% group and repeated measure ANOVA did not demonstrate that dose had an effect on body sizes (Figure 2.11). Interestingly, although no embryos exposed to 10% ethanol hatched on Day1, a few hatched on Day 2. However, due to lack of body size data on Day 1 for this group, repeated measures ANOVA that analyzed difference across Day1-4 would not include this group. Therefore, a repeated measure ANOVA was done on difference across Day 2-4 in order to include the 10% ethanol group; in this case, dose had significant effect on body size (F(3) =  44   Figure 0.11 Body sizes were smaller in worms exposed to 10% ethanol for 8.5hr throughout embryonic development.  The body sizes of worms treated with 5% ethanol chronically for 8.5hrs during embryonic development were not different from the untreated or water controls, but the body sizes of 10% exposed worms were much smaller than all other 3 groups. The lack of Day 1 body size data for 10% group was because only 1 worm was hatched in that group so no statistics could be done. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition 16.242, p < 0.0001), and the body sizes of the 10% ethanol group across Day 2-4 was significantly slower compared to the 5% ethanol group (p = 0.001), water control (p < 0.0001, HSD), and untreated controls (p < 0.0001, HSD). Figure 2.12A shows pictures of 2-d-old worms treated with 8hr of ethanol from 1-9hr during ex-utero development. Note that body sizes of the water control groups were indistinguishable from the unexposed group, suggesting that water exposure did not change body sizes. The body sizes for the 5% ethanol exposure group are visibly smaller than the untreated and water controls, and as ethanol concentration increased, the body sizes visibly decreased. The analysis of body sizes of 2-d-old animals demonstrated that dose had significant effect on body size (F(7) = 40.964, p < 0.0001, one-way ANOVA) (Figure 2.12B). The body sizes of all ethanol groups were substantially smaller than both water and untreated controls with p-values less than 0.0001 (HSD) for all pairs compared. The body sizes of the 10% ethanol exposure group were 45    Figure 0.12 The body sizes of 2-d-old worms exposed to 5-10% ethanol for 8hr throughout embryonic development were drastically smaller.  A) Pictures 2-d-old worms from each treatment. Worms in the untreated group looked identical to water (0%) control. A clear dose-dependent reduction in body sizes was seen in 5% to 10% ethanol exposure. The ultra-small worms found in ethanol exposure groups were pointed by arrows. B) The body size measurements showed significant reduction in worms treated with 5-10% chronic ethanol exposure comparing to both untreated and water controls. The box plot (C) revealed showed outliers in black dots; the dataset used in (C) was the same as (B). ***, p < 0.0001. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition  46  smaller than that of the 5 or 6% groups (10% vs. 5%, p = 0.001, 6%, p = 0.004, HSD), sizes of the 9% group were smaller than that of 5 and 6% groups (9% vs. 5%, p = 0.005, vs. 6%, p = 0.010, HSD), but sizes of 7 and 8% group were not smaller than that of the other ethanol groups, and sizes of 5 and 6% groups weren’t different from each other. These result demonstrated that 9-10% ethanol exposure for 8hr throughout embryonic development produced marked reduction in body size of 2-d-old animals, and this reduction was worse than that produced by 5-6% ethanol exposure. It is possible that these decreased body sizes were the result of delayed development.  Future studies will need to separate the effects of delayed development and specific effects of growth. An interesting observation from Figure 2.12A was the presence of ultra-small worms in the pictures of 7% and 10% ethanol exposure groups, as pointed by arrows. These ultra-small worms might also be present in the groups treated with other doses but were absent in the pictures displayed in Figure 2.12A.Therefore, a box plot was constructed in order to reveal the ultra-small worms, displayed as outliers below the mean in black dots (Figure 2.12C). Interestingly, these outliers (ultra-small worms) did not occur in the untreated or water controls. Even more curiously, one normal sized worm appeared in the 10% ethanol group (the dot above the mean). Note that these large phenotypic variations were commonly observed in worms exposed to ethanol, but not a unique finding from this particular experiment. Taking into the consideration that C. elegans culture was genetically homogeneous, this large variation between individuals within the same treatment condition would unlikely be caused by genetic differences, but more so by some unknown environmental, epigenetic, or maternal factors. Single Exposure: The growth in body size from Day 1-4 were analyzed within the same treatment onset and duration with one-way multiple ANOVAs using ethanol dose as the between 47  subject variable and age as the multiple variable. For the early exposures shown in Figure 2.13, dose had significant effect on the growth of body size of the 60min exposure duration (F=3.558, p = 0.001, Pillai’s Trace, multivariate tests of multiple ANOVA), 30min (F = 1.822, p = 0.040),   Figure 0.13  For exposure during early embryonic development, the growth of body sizes were reduced in worms exposed to 30% ethanol for 60min, and exposure for shorter durations did not have large impact on the body sizes.  Early exposure for 60min to 30% ethanol reduced body size on Day 2-3 (A), but not on Day 1 (B). Early exposure for 30min to 30% ethanol reduced body sizes on Day 1 (C-D), but not on Day2-4 (C). Exposure for 15min did not produce reduction in body sizes from Day1-4 (E) or on Day 1(F). *** p < 0.0001, *, p < 0.05. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition 48  but had no effect on the 15min exposure duration. Analysis of the effect of dose between-subjects in the 60min exposure duration showed that dose had significant effects on the body sizes on all 4 Days (early 60min Day 1, F(3) = 3.772, p = 0.027, Day 2, F(3) = 16.202, p < 0.0001, Day 3, F(3) = 6.191, p = 0.004, Day 4, F(3) = 5.516, p = 0.008); in the 30min exposure  duration, dose had significant effects on Day 1 and 4 (early 30min Day 1, F(4) = 5.567, p = 0.003, Day 4, F(4) = 2.959, p = 0.041); and in the 15min exposure duration dose did not have significant effects on any Day; these result suggested that ethanol had more severe effect on body sizes if embryos were exposed for longer durations. Tukey post hoc pair-wise comparisons on the body sizes showed that worms exposed to 30% ethanol for 60min during early development were smaller on Day 2-3 (Day 2 vs. untreated, p < 0.0001, vs. water, p = 0.014, Day 3 vs. untreated, p = 0.016, vs. water, p = 0.005, Day 4 vs. untreated, p = 0.008, vs. water, p = 0.046); for 30min exposure duration, only worms exposed to 30% ethanol were smaller on Day 1 (30% vs. untreated, p = 0.001, vs. water, p = 0.049); for 15min exposure duration, no dose induced a reduction in body size on any Day. For the middle exposures shown in Figure 2.14, dose had significant effect on the growth of body size for the 60min exposure duration (F=3.691, p < 0.0001, Pillai’s Trace, multivariate tests of multiple ANOVA), 15min (F = 2.137, p = 0.009), but had no effect on the 30min exposure duration. Analysis of the effect of dose between-subjects in the 60min exposure duration showed that dose had significant effects on the body sizes on Day 1, 3, and 4 (middle 60min Day 1, F(4) = 5.166, p = 0.002, Day 3, F(4) = 3.070, p = 0.029, Day 4, F(4) = 11.434, p < 0.0001); in the 30min exposure duration, dose had significant effects on only Day 1 (middle 30min Day 1, F(4) = 4.243, p = 0.008); in the 15min exposure duration, dose had significant effects on Day 1 and 4 (middle 15min Day 1, F(4) = 3.413, p = 0.018, Day 4, F(4) = 3.056, p =  49   Figure 0.14  For exposure during middle embryonic development, the growth of body sizes were reduced in worms exposed to 40% ethanol for 60min on Day 4 only.  Middle exposure for 60min to 40% ethanol reduced body size on Day 4 (A), but not on Day 1 (B). Early exposure for 30min did not produce reduction in body sizes cross Day1-4(C-D). Exposure for 15min did not produce reduction in body sizes from Day1-4 (E) or on Day 1(F). *, p < 0.05. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition 0.028). Tukey post hoc pair-wise comparisons showed that worms exposed to 40% ethanol for 60min during middle development were smaller on Day 4 (Day 4 vs. untreated, p = 0.043, vs. water, p = 0.012; for 30 and 15min exposure duration, no dose induced a reduction in body size on any Day. 50  For the late exposures, dose had significant effect on body sizes across Day1-4 in the 60min exposure duration (F=1.940, p = 0. 043, Pillai’s Trace, multivariate tests of multiple ANOVA), and 30min (F=2.872, p = 0. 001), but had no effect on the 15min exposure duration (Figure 12.15). Analysis of the effect of dose between-subjects in the 60min exposure duration showed that dose had significant effects on the body sizes on Day 4 (late 60min Day 4, F(4) = 3.350, p = 0.034); in the 30min exposure duration, dose had significant effects on only Day 2-3   Figure 0.15 Exposure during late embryonic development did not alter the growth of body.  Late exposure for 60min (A-B), 30min (C-D), and 15min (E-F) did not alter body sizes on Day 1-4. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition 51   (late 30min, Day 2, F(4) = 4.541, p = 0.005, Day 3, F(4) = 7.599, p < 0.0001); in the 15min  exposure duration, dose had significant effects on Day 2 (late 15min Day 2, F(4) = 2.721, p =0.046). Tukey post hoc pair-wise comparison showed that no ethanol groups of the middle exposure groups had smaller mean body size on any Day compared to the untreated and water controls. Repeated Exposure: Repeated exposures consisted of 3 exposure episodes each at early, middle and late development, and each episode lasted for a third of the total duration specified, for example, 60min repeated exposure group had 3 episodes each lasted for 20min. In Repeated exposures, dose had significant effect on the growth of body sizes of the 60min exposure duration (F=2.856, p = 0. 004, Pillai’s Trace, multivariate tests of multiple ANOVA), 15min (F=2.191, p = 0. 008), but had no effect on the 30min exposure duration (Figure 2.16). Analysis of the effect of dose between-subjects in the 60min exposure duration showed that dose had significant effects on the body sizes on Day 1-4 (repeated 60min Day 1, F(3) = 8.649, p = 0.001, Day 2, F(3) = 15.620, p < 0.0001, Day 3, F(3) = 9.878, p < 0.0001, Day 4, F(3) = 15.575, p < 0.0001); in the 30min exposure duration, dose had significant effects on only Day 4 (repeated 30min, Day 4, F(4) = 2.826, p = 0.040); in the 15min exposure duration, dose had significant effects on Day 2 and 4 (repeated 15min Day 2, F(4) = 2.951, p = 0.033, Day 4, F(4) = 4.283, p = 0.006). Tukey post hoc pair-wise comparisons on the body sizes showed that worms exposed to 30% ethanol for 60min repeatedly were smaller on Day 2-4 (Day 2 vs. untreated, p = 0.008, vs. water, p = 0.008, Day 3 vs. untreated, p = 0.019, vs. water, p = 0.022, Day 4 vs. untreated, p = 0.003, vs. water, p = 0.001); none of the dose for 30 and 15min exposure duration induced a reduction in body size on any Day.  52   Figure 0.16 Repeated exposure for a total of 60min (20min per episodes) reduced the body sizes.  Repeated exposure for 60min to 30% ethanol (A-B) produced body size reduction on Day 2-3 (A), but not on Day1 (B); the lack of statistical significance was due to low sample size. Repeated exposure for 30min (C-D), and 15min (E-F) did not alter body sizes on Day 1-4. ** p < 0.001, *, p < 0.05. Sample size was 10-15 except for 40% 60min early, and 40% 60min repeated condition  53  Summary of Experiment A-2 The vast amount information from Experiment A-2 is difficult to digest; therefore I arranged all the results obtained from the ethanol exposure groups into Table 2.1, in which the black arrows indicated statistically significant increase (▲) or decrease (▼), crosses () indicated no worms survived for measurements to be taken, blank cells indicated no data was collected, and hyphens (-) indicated statistically non-significant findings. As described earlier, a measurement of the ethanol exposure groups was considered to be altered when the measurement significantly differed from that of both untreated control and water control. Chronic exposures for 8.5hr decreased hatch rate when the ethanol dose was 8% or more. Chronic exposure for 8.5hr also produced lasting growth retardation in body sizes. In Chronic exposure for 8hr, growth retardation could be seen in 2-d-old worms exposed to 5% or higher ethanol concentration. This result led to the conclusion that growth retardation could be induced by ethanol exposure during embryonic development if the ethanol dose was high enough. Within single exposure conditions (early, middle and late exposures), early development was the most vulnerable stage. As in exposure to 30% ethanol for 60min during early development condition, smaller body sizes were seen from Day 2-4; however, exposure to the same dose during middle and late development did not produce any physical effects. Similarly, exposure to 40% ethanol for 60min during early development killed all embryos, but the same dose did not induce any physical abnormalities if exposed during middle and late development. These results supported the hypothesis that ethanol exposure during early development would produce more physical effects, and somewhat replicated the findings reported in Davis et al. (2008) that ethanol exposure during early development killed more embryos than exposure of the same dose during middle and late development. 54   Table 0.1 Summary of findings from ethanol exposure groups in Experiment A-2.  Black arrows = significant and expected increase (▲) or decrease (▼) comparing to both water and unexposed controls. Open arrows = unexpected but significant increase (). Hyphend bar (-) indicated non-significant result. Cross () indicated no data was collected due to death of all animals in the group.  Empty cells indicated that no data was collected. H.R. = hatch rate. R.O. = reproductive onset. Timing Dose (%) Duration H.R. R.O. Body size Day 1 Day 2 Day 3 Day 4 Chronic 5 8.5h - ▲ - - - - 8h -   ▼   6 8h -   ▼   7 8h -   ▼   8 8h ▼   ▼   9 8h ▼   ▼   10 8.5h ▼   ▼ ▼ ▼ 8h ▼   ▼   Early 20 60min - - - - - - 30min - - - - - - 15min - - - - - - 30 1h ▼ - - ▼ ▼ ▼ 30min - ▼ ▼ - - - 15min - - - - - - 40 60min ▼      30min - ▲ - - - - 15min - - - - - - Middle 20 60min - - - - -  30min - - - - - - 15min - - - - - - 30 60min - - - - - - 30min - - - - - - 15min - - - - - - 40 60min ▼ - - - - ▼ 30min - - - - - - 15min - - - - - - Late 20 60min - - - - - - 30min - - - - - - 15min - - - - - - 30 60min - - - - - - 30min - - - - - - 15min - - - - - - 40 60min ▼ -  - - - 30min - - - - - - 15min - - - - - - Repeated 20 60min - - - - - - 30min - ▲ - - - - 15min - - - - - - 30 60min ▼ ▲ - ▼ ▼ ▼ 30min - ▲ - - - - 15min - - - - - - 40 60min ▼      30min ▼ ▲ - - - - 15min ▼ ▲ - - - - 55   Another point illustrated in this table was that repeated exposure often produced worse outcomes than a single exposure of the same duration. When 30% ethanol was applied to the early exposure group for 60min, the hatch rate and body sizes from Day 2-4 were affected. However, when the same dose was used in the repeated exposure condition, in which 30% ethanol was applied for 20min during early, 20min during middle, and 20min during late development, not only hatch rate and body sizes from Day 2-4 were affected, but reproductive onset was also affected. When 40% ethanol was applied for a total of 30min repeatedly or 10min at each stage, hatch rate was decreased and reproductive onset was delayed; however, the same dose only induced reproductive delay in the early exposure condition; in addition, the same dose did not cause a change in hatch rate or reproductive onset in the middle or late exposure condition. Similarly, repeated exposure to 40% ethanol for a total of 15min decreased hatch rate and delayed reproductive onset, but the same dose did not cause any physical changes in the other three single exposure conditions. These results demonstrated that repeated exposure produced more severe physical effects than a single exposure to the same concentration and duration, even though each exposure episode of the repeated exposure condition was two-third shorter at each developmental stage, Experiment A-3 Early and the repeated exposure induced the worst outcome  Based on the findings in the previous experiment, Experiment A-3 was an attempt to assess how repeated exposure might produce worse outcome than a single exposure. The physical effects induced by the repeated ethanol exposure could simply be accumulated from each single exposure, or, the magnitude of effect could be the result of a complex interaction between the three exposure episodes (Figure 2.17). In the Additive Model, the final physical 56  phenotype resulting from the repeated exposure is the sum of the physical effects caused by each exposure. In the Compounded model, the damage caused by previous exposures is exacerbated or modified by the following exposure episode; for example, the damage from the early exposure is compounded by the middle exposure, and the compounded physical damage after the middle exposure was further exaggerated by the late exposure. In other words, the final product depends on the sequential developmental events occurring during and between the exposure episodes.  Figure 0.17 Models for the magnitude of physical deficits observed in repeated exposure.  There are two possible ways to produce the magnitude of physical deficits observed in the repeated exposure groups. The damage may be additive (Additive Model). The damage incurred during early development resulted in a certain portion of physical damage, and the damage during middle and late development resulted in each another portion. In compounded model, the final magnitude of physical deficits produced is dependent on a sequence of events. Damage incurred during early exposure can be amplified or modified by exposure during middle development, and so on. The final physical deficits observed in repeated exposure group are a product of all three exposures. In Compounded Model, previous exposure can modify the potential of the second exposure to have an effect on a physical deficit. For example, a single exposure at a certain developmental timing normally may not produce a physical deficit, but this same exposure became harmful if it was proceeded with a previous exposure episode. The Compounded Model is more likely the case because developmental processes are highly dependent on previous events and experiences. However, to experimentally prove the Compounded model, one needs to find a measurable and comparable phenotype across different time points during embryonic stages, which may be technically difficult to doto test the .  Alternatively, I designed an experiment to test the Additive model. If the Additive model was true, the degree of the effect created by 3 episodes of repeated exposure should be the sum of the amount of the same effect produced by 3 single exposures. For example, the amount of a specific 57  effect produced by three 20min repeated exposures at early, middle and late development should be the same as the sum of the same effects produced by three 20min single exposures at early, middle or late development. Similarity, if the Additive Model was true, the 60min repeated exposure condition used in Experiment A-2 should produce a magnitude of a specific effect somewhere in between the sum of three 15min single exposures (a total of 45min) and three 30min single exposures (a total of 90min). According to the results found in Experiment A-2, as 60, 30, or 15min exposure to 30% ethanol during middle and late development did not produce any reproductive delay, the sum of reproductive delay produced by the three single exposures should be the same as the amount of delay produced by early exposure. However, the 60min repeated exposure produced a reproductive delay more similar to the 60min early exposure than to the 30min early exposure, which was not as predicted by Additive Model.   Figure 0.18 Exposure timing in Experiment A-3.  Single exposure groups were treated for either 60 or 20min at early, middle or late development. Repeated exposure groups were treated for a total of 60min, of which three 20min individual episodes began at early, middle and late development. Treatment at early exposure began at 2hr ex-utero development, middle at 5hr, and late at 8hr. 58  In Experiment A-3, the physical effects produced by 20 or 60min single exposures during early, middle or late development were compared with the effects produced by the 60min repeated exposure (Figure 2.18). A 30% ethanol exposure was chosen for this study because this treatment condition did not kill all of the embryos at any exposure treatment so that the subsequent events such as reproductive onset, body sizes and lifespan could be measured. Experiment A-3 contained 6 replicates per experimental group, and the experimenter recording the data was blind to the condition of the worms. Hatch Rate The hatch rates were combined from 6 replications of each experimental group as described above (Table 2.2).  A three-way ANOVA using ethanol dose, treatment duration and exposure onset as between-subjects variables showed that dose alone had an effect on hatch rates (F(5.231, p = 0.025), but duration and onset both did not have effects on hatch rates. Post-hoc pair-wise comparisons did not show significant reduction in the hatch rates of the ethanol exposure groups compared to the untreated and water controls. Reproductive Onset In Experiment A-3, the reproductive onset data was analyzed in two ways: numerical or categorical. Worms that laid eggs before 96hr of age were included in the numerical analysis, but worms that failed to lay eggs before 96hr or died before laying any egg were excluding from the numerical analysis, and instead, were assigned as “Not Laid” and “Dead”, respectively, and included into categorical analysis. The categorical analysis method was incorporated in this experiment because excluding the “Not Laid” and “Dead” worms may undermine the effect of ethanol by only looking at worms that were healthy enough to lay eggs before the cut-off time of reproductive onset study. 59  Table 0.2. Descriptive statistics of hatch rate, body size and lifespan of worms exposed to 30% ethanol for 20 or 60min singly or repeatedly at early, middle or late exposure.  No results from the ethanol exposure groups were significantly different from the untreated and the corresponding water controls.    Hatch Rate  Body size on Day 4  Lifespan Onset Duration Dose N Mean ± SE N Mean ± SE N Mean ± SE Untreated 6 .93 ± .02  35 69.99 ± 2.36  35 11.37 ± .68 Early 60min water 6 .95 ± .02  42 63.14 ± 2.04  42 11.17 ± .69 30% EtOH 6 .96 ± .01  36 66.74 ± 1.30  36 9.19 ± .58 20min water 6 .97 ± .02  43 73.81 ± 1.98  43 12.07 ± .58 30% EtOH 6 .92 ± .02  37 68.27 ± 2.18  37 12.68 ± .71 Middle 60min water 6 .97 ± .01  35 71.62 ± 1.84  35 11.74 ± .69 30% EtOH 6 .94 ± .01  32 68.57 ± 2.73  32 11.28 ± .85 20min water 6 .99 ± .01  39 68.60 ± 2.19  39 10.05 ± .55  30% EtOH 6 .96 ± .02  40 68.66 ± 2.31  40 11.68 ± .71 Late 60min water 6 .92 ± .05  42 70.38 ± 1.86  42 11.90 ± .58 30% EtOH 6 .93 ± .02  35 74.96 ± 1.23  35 12.17 ± .73 20min water 6 .99 ± .03  39 67.48 ± 1.91  39 11.08 ± .60 30% EtOH 6 .97 ± .01  41 72.34 ± 1.84  41 11.44 ± .71 Repeated 60min water 6 .98 ± .02  40 66.39 ± 2.78  40 11.20 ± .59 30% EtOH 6 .91 ± .04  43 66.71 ± 1.56  43 11.09 ± .65  Reproductive onset analyzed numerically: A three-way ANOVA using dose (untreated, water & 30%), onset, and duration as between-subjects variables revealed significant interaction between all three factors (F(2) = 3.574, p = 0.029), between duration and timing (F(2) = 13.992, p < 0.0001) and timing and dose (F(3) = 8.994, p < 0.0001) (Figure 2.19); dose, onset and duration alone all had significant effect on the reproductive onset (dose, F(1) = 10.306, p = 0.001, onset, F(3) = 14.496, p < 0.0001, duration, F(1) = 42.517, p < 0.0001). Post hoc pair-wise comparisons showed that the reproductive onsets of worms exposed during early ethanol exposures for 60min (vs. untreated, p < 0.0001, vs. water, p = 0.008), and that of worms exposed repeatedly (vs. untreated, p < 0.0001, vs. water, p = 0.037) were delayed. In addition, the reproductive onset of the repeated exposure groups was significantly more delayed than all other ethanol exposure groups except for 60min early exposure group (repeated exposure vs. 60min 60  middle, p = 0.001, late, p < 0.0001, vs. 20min early, p < 0.0001, middle, p < 0.0001, late, p < 0.0001). Similar to Experiment     Figure 0.19 Reproductive onset was delayed in early and repeated exposure group at 30% ethanol concentration in Experiment A-3.   A) Early exposure and repeated exposure for 60min at 30% delayed  reproductive onset. reproductive onset was more delayed in repeated exposure group than middle and late exposure groups. B) Early exposure for 20min delayed reproductive onset. The repeated exposure group shown in B) was identical as in A).  Repeated exposure was more delayed in reproductive onset than all three 20min-exposed groups. Sample size was 45 per group. A-2, only the early and repeated exposure groups induced delay in reproductive onset but the middle and late exposure did not. Curiously, the magnitude of delay produced by the repeated exposure group was similar to the magnitude of delay produced by the 60min early exposure group. If the additive model for the repeated exposure were true (Figure 2.17), the magnitude of reproductive onset delay should be more similar to the sum of delay from all three 20min exposure groups, rather than more similar to the magnitude of delay from all three 60min exposure groups; in fact, the reverse was true: the magnitude of reproductive delay produced by the 60min repeated exposure was more similar to the sum of all 60min exposure groups for a total of 180min exposures, and was much higher than the sum of all 20min exposure groups for a total of 60min exposure (Figure 2.20). Thus the numerical study of reproductive onset suggested 61  that the Compounded Model more precisely describes the magnitude of developmental effects produced by the repeated exposure paradigm.   Figure 0.20 The magnitude of reproductive delay in repeated exposure group more closely resembles the sum of three 60min exposure at three different stages. Sample size was 45 per group. Reproductive onset analyzed by categories: Reproductive onset was also analyzed categorically (Figure 2.21) by Chi-square analysis. The “Normal” category included worms that laid eggs between 70-75hr of age, “Delayed” included worms that laid eggs between 76-80hr of age, “Severely delayed, Not Laid, and Dead” included worms that laid eggs between 80-96hr of age, did not lay eggs after 96hr of age, and were dead before laying any egg.  The categorical reproductive onset were different than the untreated and water control for worms exposed for 60min during early development (vs. untreated, X2(2) = 27.495, p < 0.0001, vs. water, X2(2) = 10.654, p = 0.005), for 60min during late development (vs. untreated, X2(2) = 6.276, p = 0.043, vs. water, X2(2) = 10.758, p = 0.005), for 60min repeatedly (vs. untreated, X2(2) = 29.404, p < 0.0001, vs. water, X2(2) = 7.793, p = 0.020). None of the categorical reproductive onsets of the 20min ethanol exposure groups were significantly different. As demonstrated in Figure 2.21, the  62   Figure 0.21. Reproductive onset organized in categories reveled that repeated exposure and early exposure for 60mins had larger proportions animals with delayed reproductive onset.  A) Animals exposed to 60mins of 30% ethanol exposure during early and late development showed difference in the range of time they laid their first egg. Repeated exposure produced the worst and similar outcome in reproductive onset as in early exposure for 60mins. B) No reproductive onset categories were different in animals treated for 20mins. Repeated exposure group in B) was the same group in A. Repeated ethanol exposure was different from all other ethanol treated groups in 20mins exposure duration. Sample size was 45 per group.  63  categorical reproductive onset of the repeated exposure group closely resembled that of the 60min early exposure group. Life expectancy Life expectancy was analyzed in two ways. Firstly, the mean Day of death (lifespan) was compared between groups with three-way ANOVA using dose, duration and onset as between-subject variables. Secondly, survival curves of adulthood were analyzed using Wilcoxon survival test in order to determine whether the rates of survival were lower in the ethanol exposure groups. The analysis of lifespan showed a significant interaction between duration and onset (F(2) = 6.516, p = 0.002, three-way ANOVA), and duration and dose (F(1) = 4.287, p = 0.039), but dose, duration, and onset alone did not have significant effects on the lifespan. Post-hoc pair-wise analyses did not show any differences; in other words, lifespan was not altered by exposure to 30% ethanol for 60min or 20min during embryonic development (Table 2.2).  Analysis of survival rate using the Wilcoxon survival test showed significant differences between groups (Wilcoxon statistics = 25.433, df = 14, p = 0.031), but post-hoc pair-wise comparison showed no difference for ethanol treated groups compared to their corresponding water controls (Figure 2.22). The survival curve for 60min early exposure using 30% ethanol began to deviate from the water control, but this deviation was not significant according to Wilcoxon statistics (Wilcoxon statistic = 3.291, df = 1, p = 0.070) (Figure 2.22); a larger sample size in this case may be required to find significance with this statistical test. In summary, life expectancy was not altered in worms exposed to 30% ethanol for 60min or less during embryonic development.   64    Figure 0.22 Survival rate was not altered by 30% ethanol exposure during embryonic development.    65  Body sizes Analysis of body size of 4-d-old adults using a three-way ANOVA showed a significant interaction between treatment onset and duration (F(2) = 5.511, p = 0.004), and treatment onset alone had an effect on body size (F(3) = 2.665, p = 0.047). Post-hoc pair-wise comparisons did not show any change in body sizes of the ethanol exposure groups. The descriptive data for body sizes was listed in Table 2.2. In summary, 30% ethanol exposure for 60min or less during embryogenesis did not alter body size of 4-d-old worms. Exposure for 60min at early (A), middle (B), or late (C) development did not reduce survival rate. Exposure for 20min at early (D), middle (E), or late (F) development did not reduce survival rate. Repeated exposure (G) did not reduce survival rate. Sample size was 45 per group.  Summary of Experiment A-3 Although no hatch rate, life expectancy, or body sizes changes were found in Experiment A-3, the study of reproductive onset showed several interesting findings. Firstly, early development was again found to be more vulnerable than middle and late development in that reproductive delay was only found in the single exposure occurred during early embryonic development. Secondly, the repeated exposure was found to produce reproductive delays as large as 60min early exposure. In addition, the magnitude of reproductive delay produced by the 60min repeated exposure was much larger than the sum of reproductive delay produced by three 20min single episodes, and was more similar to the sum of reproductive delay produced by three 60min single exposures. This finding suggested that the Additive Model did not predict the magnitude of reproductive delay produced by the repeated exposure. Instead, the Compounded model was more likely to explain the magnitude of physical effects caused by repeated exposures. 66  Discussions  This chapter describes the first FASD study that incorporates all 4 variables of ethanol exposure patterns into one comprehensive experiment. The examination of these variables has been a major methodological constraint in FASD research. Many studies in the past attempted to address one or two variables at a time. A study in zebrafish model looked at 6 different treatment conditions involving different timing, duration and dose (Bilotta et al., 2004), but this study did not examine the contribution of these 3 factors systematically. In Experiment A-2, I looked at a total of 55 treatment conditions, and derived information from 550 individual animals. This study demonstrated that C. elegans is a powerful tool to investigate the interaction between dose, duration, timing, and frequency of ethanol exposure.  Three major findings were drawn from this study. First of all, the severity of ethanol exposure needed to surpass a threshold that was unique to each developmental stage in order to produce lasting physical defects. Secondly, early development was the most vulnerable stage for the effect of ethanol. Thirdly, the magnitude of the damages produced by each repeated exposure episodes were was by subsequent exposures.  Treatment severity needs to surpass a threshold to stabilize physical/developmental deficits  An idea presented in this chapter was that ethanol exposure must surpass a certain threshold in order to induce physical/developmental effects in exposed animals, and that this threshold is dependent on the timing of exposure. I found that exposure during early development required exposure to 30% ethanol or more for at least 60min in order to produce physical/developmental effects. Middle and late development, on the other hand, required at least 60min of 40% ethanol to generate very minor effects. A speculation is that 40% ethanol for 60min was still too low a dose for middle and late development. A higher dose of ethanol should 67  be tested in the future to see whether physical/developmental effects can be induced by ethanol exposure at any developmental stages as long as the dose is strong enough.  I also predict that the threshold for inducing physical effects is likely to be close to the lethal dose. In a chicken model of FASD, only at the ethanol dose that eventually killed 100% of the chicken embryos 7 days post-fertilization could you find 100% of the chickens exhibiting physical abnormalities; when the ethanol treatment was only severe enough to reduce survival rate to approximately 80%, only 55-60% exhibited physical dysmorphology (Bilotta et al., 2004). Therefore, it is reasonable to predict that the dose required to induce consistent physical effects in C. elegans may come close to one that produces 100% lethality.  Early development is the most vulnerable stage A common goal of Experiments A-1 and A-3 was to determine a critical period for ethanol-induced physical/developmental abnormalities. The susceptibility of the early embryonic period was first supported in Figure 2.8 where chronic exposure initiated 30min earlier produced nearly zero hatch rate but initiated 30min later produced less than 50% reduction in hatch rate. Table 2.1 also clearly illustrated that the same ethanol treatment administered during early development produced more physical damages than during the other two stages. To illustrate this idea further, I compared the growth in body size and the reproductive onset of worms exposed to 30% ethanol for 60min (Figure 2.23). The early exposure group, illustrated as a solid line in Figure 1.23A, demonstrated more severe growth retardation than the middle and late exposure groups. Similarly, the reproductive delay of the early exposure group was significantly larger than that of the middle and late exposure groups (F(2) = 4.009, p = 0.039, ANOVA; early vs. middle, p = 0.027, early vs. late, p = 0.012, HSD). It is clear that, worms treated during early 68  A)*B) Figure 0.23 Early exposure produced more severe phenotype.  A) Body size and B) reproductive onset comparison between animals exposed to ethanol for an hour at 30% ethanol during different developmental stages. Data extracted from Experiment A-2. development were more affected than ones treated during middle and late development given the same treatment does and duration. It is important to emphasize that this conclusion was not intended to generalize to other animal models or to other ethanol-related symptoms. Some FASD symptoms, such as the reduction in hippocampal CA1 neuron number, were better induced by ethanol exposure during later developmental stages, such as synaptogenesis (Tran and Kelly, 2003). However, in regards to physical abnormalities, studies of other animal models have led to conclusions similar to my findings. Human studies found that the critical period of craniofacial abnormalities was from around the time of conception (Ernhart et al., 1987), overlapping to the time that gastrulation occurred in the human fetus. A study on mice also determined that gastrulation was the critical stage for the establishment of physical malformations (Sulik, 1984). However, another study using mice as an FASD model suggested that later (gestation 12-17) rather than earlier (gestation 5-10) ethanol exposure more readily induced growth retardation (Middaugh and Boggan, 1991). In contrast, early ethanol exposure more readily induced growth retardation in C. elegans.  69  The result of repeated exposure is a product of compounded effects In humans, the symptoms of FASD are most likely a product of multiple exposures. The reason may be that each additional exposure can worsen the damaged incurred by the previous exposure. However, to experimentally differentiate how each exposure episode interacts with others can be very methodologically challenging. In this study, I hypothesized that if multiple exposures would produce a worse outcome, then the magnitude of effects generated by multiple exposure should be larger than the sum of effects generated by single exposures equivalent to the individual episodes from multiple exposure. In Experiment A-3, I found that the repeated exposure could induce worse outcomes than the sum of 3 single exposures for the same total duration. Additional evidence was be found in Experiment A-2 in which Figure 2.24A showed the mean reproductive onset of worms treated to 40% ethanol for 15min. The repeated exposure delayed reproductive onset for longer than the late exposure. Moreover, the effect of the repeated exposure often resembled the outcome of the early exposure for the same duration. Figure 2.24B showed the growth in body size over Day 1 to 4 of worms treated for 60min with 30% ethanol. The growth curve of the repeated exposure group was quite similar to the growth curve of the early exposure group. This resemblance was most clearly illustrated in Experiment A-3. Both the numerical and categorical analysis of reproductive onset demonstrated that the effect produced by repeated exposure was very similar to that produced by 60min early exposure. These findings suggested that repeated exposure can at least produce physical/developmental abnormalities as severe as early exposure of the same duration.  I have been unable to find any previous study that systematically compared the effects of multiple exposures with the effects of single exposures. This study may be the first to examine the effect of multiple exposure episodes quantitatively and has the potential to tease apart the 70  underlying mechanisms that differ between the exposure methods. If repeated exposure indeed produces symptoms following the Compounded model, the mechanisms involved in the repeated exposure may be very different from those involved in chronic or acute exposure. In that case, this robust FASD model in C. elegans may become a starting point for the investigation into the differences between repeated and single exposure conditions.  Figure 0.24 Repeated exposure produced the most severe effect  A) Mean reproductive onsets in animals exposed to 40% ethanol for15min during early, middle or late development or repeatedly (5mins at each developmental stage). Repeated exposure produced more delay in reproductive onset than late exposure group. B) Growth curve in body sizes in animals exposed for 1hr at 30% ethanol during early, middle or late development, or repeated for 20mins at each stage. Growth curve in repeated exposure closely resembles growth curve in early exposure group. Potential for this data set is enormous The strength of this thesis is that it takes into account of the 4 factors in ethanol exposure patterns. A systematic approach to this study can answer many interesting questions. For example, is drinking a small amount repeatedly better than binge drinking only once?  To simulate the situation, I used the repeated exposure for an hour (20min at each stage) at the lowest 2 doses, and compared that to a single episode exposure for 30min at a higher dose (Figure 1.25). The resulting graph, translated into plain language, suggests that the chance of 71  inducing growth retardation is higher: 1) if you binge drink one time during early pregnancy than repeatedly drink 50% less concentrated alcohol, or 2) if you drink only 25% less alcohol repeatedly than binge drink once at any time during early pregnancy. A variety of potential questions can be addressed with this dataset, but the analysis of all those possibilities deserves a separate chapter. To keep this thesis more concise, I’ll provide only this one hypothetical example to illustrate how this data set can be further analyzed to address questions about the interactions between dose, duration, timing, and frequency of ethanol exposure.  Figure 0.25 An example of questions you can ask from the systematic evaluation in Experiment 1.2.  72  CHAPTER 3. BEHAVIORAL EFFECTS OF EMBRYONIC ETHANOL EXPOSURE IN C. ELEGANS Introduction In humans, prenatal alcohol exposure may produce neurobehavioral and cognitive deficits such as problems in learning and memory, motor function, attention, executive functioning, and social skills (Riley and McGee, 2005). These problems can impair an individual’s ability to maintain a job, or even to live independently (Spohr et al., 2007). The mechanisms responsible for these nervous system problems in individuals with FASD are not well understood, however, the study of a model system such as C. elegans with a small, tractable nervous system may uncover possible mechanisms by which alcohol affects the development of nervous systems. To generate this data, first we need to examine the types of behavioral deficits that may be induced by ethanol exposure in this model system. The objective of this chapter therefore was to find behavioral, learning, and memory deficits in C. elegans exposed to ethanol during embryonic development.  In humans, numerous types of motor dysfunctions such as delayed motor development and fine-motor function, tremors, motor speed and precision, weak hand grasp, and poor hand-eye coordination have been shown to occur in individuals suffering from FASD (Mattson and Riley, 1998; Riley and McGee, 2005). The source of these motor dysfunctions has been associated with functional damage to the cerebellum (West, 1986). Animal models of FASD have also demonstrated delays in reflex development, gait disturbances, and poor balance (Mattson and Riley, 1998). Recent report comparing young children from 1 to 6 years old exposed to prenatal alcohol confirmed that gross and fine motor development was more delayed 73  than children who were not exposed (Kalberg et al., 2006). The severity of the developmental motor delays correlated with cognitive delays (Osborn et al., 1993). It may be possible that ethanol exposure in C. elegans may produce similar damage to the neuronal circuit responsible for motor functions, and manifested as abnormalities in locomotion. The normal locomotion in worms consists of forward movement with a sinusoidal motion punctuated by occasional backward movement, or “spontaneous reversals” (Figure 3.1). The rate   Figure 0.1 Body bend punctuated by spontaneous reversal was not counted.  (1) The worm moved backward (spontaneous reversal) in (2) long enough distance to form a complete reversal body bend, but (3) moved forward before forming another reversal body bend. The first reversal body bend was not counted because the worm did not end up moving to another location (1) & (3).  of worm locomotion can be measured by counting the number of body bends produced by the sinusoidal motion and the number of spontaneous reversals within a given time period (Hart, 2006). Several problems can be identified during this type of locomotory assay: visible motor abnormalities in terms of un-coordinated movements, hyper-activity in terms of higher rate of locomotion, hyper-sensitivity in terms of slower return to the baseline locomotory rate after stimulation, and problems in the neuronal circuit responsible for the balance of forward and reversal movements. Davis (2006) found that larval ethanol exposure in C. elegans  reduced the rate of body bends and the magnitude of the reversal response to tap (Davis, 2006). These findings suggested there were some motor dysfunction in the worms exposed to ethanol during 74  larval development. In Experiment B-1, I investigated the locomotory behavior in worms exposed to 30% ethanol during embryonic development. In Experiment B-2, I investigated locomotory behavior in worms chronically exposed to ethanol during embryonic development. Prenatal alcohol exposure can produce several types of learning and memory deficits (Mattson and Riley, 1998; Kodituwakku, 2007). Studies in humans suggested that deficiencies in auditory memory, spatial memory, and some aspects of working memory occurred in FASD children (Mattson and Riley, 1998). Memory problems have also been found in animal models of FASD such as rats (Petkov et al., 1991), chickens (Rao and Chaudhuri, 2007), and mice (Summers et al., 2006). Rat models of FASD found that memory of spatial learning (Zimmerberg et al., 1991), response perseveration (Riley et al., 1979b), and alternation test (Zimmerberg et al., 1989) were affected. However, other studies in humans (Fried and Watkinson, 1990; Fried et al., 1992; Jacobson et al., 1993) and rats (Riley et al., 1979a) reported that some types of memory are relatively intact. A well studied learning and memory impairment was spatial navigation (Kodituwakku, 2007). Spatial memory studies using Morris Water Maze in rat model of FASD showed that the rats exposed to prenatal ethanol were more impaired in delayed recall of spatial memory and less impaired in immediate recall (Matthews and Simson, 1998). Children affected with FASD showed impairment in place learning but not cue-navigation in a virtual Morris Water Maze paradigm (Hamilton et al., 2003), suggesting a specific rather than general impairment in spatial learning. Object recognition was another type of learning and memory task found to be impaired in delayed but not in immediate recall in both children with FASD (Uecker and Nadel, 1996) and in rat model of FASD (Popovic et al., 2006). Furthermore, in studies with rats (Cohen et al., 1985) and humans (Streissguth et al., 1981), prenatal alcohol exposure was found to impair habituation.  75  Habituation is a non-associative form of learning defined as a decrease in responses to repeated stimuli (Rose and Rankin, 2001). Nervous systems receive a tremendous amount of sensory input from the environment at any given time, but only a few inputs can be processed at a given time. Habituation acts like a filter to decrease input from unimportant sensory stimuli. Habituation has been found in all levels of organisms including protozoa (Wood, 1970), fruit flies (Duerr and Quinn, 1982), Aplysia (Pinsker et al., 1970), rats (Davis, 1970), and humans (Geer, 1966).. Worms respond to mechanical stimuli delivered as a tap on the side of the Petri dish by moving backwards (Rankin et al., 1990). When no unpleasant consequences are associated with the repeated tap stimuli, worms habituate as they learn to ignore the tap and stop responding to it.  Streissguth et al. (1983) assessed habituation in infants using the Brazelton Neonatal Assessment Scale, and found that defects in habituation were related to the amount of alcohol consumed by the mother. In rat model of FASD, impaired habituation of the heart rate orienting response to an olfactory stimulus was found in neonatal rats exposed to a maternal postnatal ethanol binge (Hunt and Phillips, 2004). It may be that embryonic ethanol exposure will have the same effect in C. elegans. In Experiment B-3, I investigated short-term habituation to taps delivered at 10s inter-stimulus interval (ISI) in worms exposed to 20% ethanol for 60min at 3 different times during embryonic development. In Experiment B-4, I investigated short-term habituation to taps in worms chronically exposed to ethanol for 8hr throughout ex-utero embryonic development at either 10s or 60s ISIs.To see whether memory could be affected by ethanol exposure in a nematode system, in Experiment B-5, I investigated the effect of chronic ethanol exposure on long-term memory for habituation training in C. elegans.  76  Materials & Methods Exposure protocol Worms were exposed to ethanol as described in Chapter 2. In Experiment B-1, embryos were exposed to 30% ethanol singly or repeatedly for 20 or 60min at 3 different times of embryonic development as described in Experiment A-3; the sample size was 30-45. In Experiment B-2, 4 and 5, embryos were chronically exposed to 5 or 10% ethanol for 8hrs throughout embryonic development as the 8hr chronic exposure group described in Experiment A-2; the sample size was 20-30. In Experiment B-3, embryos were exposed to 60min of 20% ethanol during early, middle or late development as described in Experiment A-1, and the sample size was 20-30. Behavioral recording & scoring Worm images were recorded using a Panasonic AG-1960 video cassette recorder connected to a Panasonic D5000 camera on a stereomicroscope (Wild Zeiss Canada) and the movements were observed with a JVC color monitor connected to the cassette recorder (Figure 3.3). The distances of the reversal responses to taps and the lengths of worms were traced onto acetates using a stop-frame VCR. The acetates containing these tracings were scanned (UMAX Astra 2100U) into a Macintosh computer and were digitally measured using NIH Image software. Body bends and spontaneous reversals were counted by reviewing behavior recordings with a stop-frame VCR. For body bends, one count was defined as the time when the part of the worm just behind the pharynx reached a maximum bend in the opposite direction as the bend last counted. In the case a spontaneous reversals occurred, the reversal body bends that retraced the forward bend occurred just before the reversal did not count (Figure 3.2) (Hart, 2006). Similarly, when the spontaneous reversal was followed by forward movement without finishing a complete 77  body bend, the forward body bend that retraced the reversal body bend did not count. One spontaneous reversal count was defined as the time when a worm moved backwards after a period of stationary or forward movement.  Figure 0.2 Apparatus for behavioral recording Procedures for body bends and spontaneous reversals In Experiment B-1, approximately 20 4-d-old worms of the treatments described above were transferred to a 5-cm NGM plate seeded with a drop of E. Coli and then carefully placed into the plate holder (Figure 3.3); after a 5 min rest period, the activities of the worms were taped for 3mins; on the 4th min from the start of the recording, one tap was administered and the resulting activity was recorded for another 3min. In Experiment B-2, 20 single animals was transferred one at a time onto the center of a NGM plate spread evenly with E. Coli and rested for 1 min before being recorded for 3mins. 78   Figure 0.3 Protocol for body bend and spontaneous reversal experiments.  A) Protocol for Experiment B-1. B) Protocol for Experiment B-2. Habituation procedures Mechanical stimuli were delivered using a Grass Instruments (Quincy, MA) S88 stimulator connected to a tapper exerting 1-2 N of force to the side of the Petri plate (Figure 3.2). For both short- and long-term habituation, a Petri plate containing worms was placed into the plate holder 6mins before the first stimulus. For short-term habituation, 10 taps were delivered to a plate containing 8-12 4-d-old worms at either a 10s or a 60s ISI. Recovery taps were administered 30s, 5min, and 10min after the 10th stimuli; in 10s ISI, responses to all 3 recovery taps were scored and analyzed; in 60s ISI, only responses to the 5min and 10min recovery taps were used. For long-term habituation, 12-15 4-d-old worms were transferred from a worm colony that was raised on an anti-vibration shelf to a fresh NGM plate seeded with a drop of E. Coli 1hr before the training. To deliver mechanical stimuli to several plates of worms at the same times, plates containing experimental worms were placed into a plastic box and covered with a piece of foam. The purpose of the covering foam was to avoid bouncing and sliding of the plates inside the box while stimuli were delivered. One mechanical stimulus was delivered by dropping the box 1-inch from a solid surface. Four blocks of 20 drops at a 60s ISI separated by 1hr rest periods were given to experimental worms (Figure 3.4). Plates containing control worms were 79  given only a single drop at the end of the training period. To test for long-term habituation, 22-28 hours after the training 5 test taps were given in the same manner as in short-term habituation, and the responses to these 5 taps were scored and used in data analysis.   Figure 0.4 Training protocol for long-term habituation Statistical analysis In Experiment B-1, the number of body bends and spontaneous reversals recorded 3min before and after the tap stimulus were analyzed separately by three-way ANOVAs using ethanol dose, exposure duration, and onset as between-subjects variables; Tukey (HSD) post-hoc analysis was used to analyze for statistical significance between two groups. In Experiment B-2, the number of body bends and spontaneous reversals of the untreated, water, 5 and 10% groups were analyzed with a one-way ANOVAfor the 4 different treatment conditions, and pair-wise comparisons were made with Tukey post-hoc analysis; in Experiment B-4 and 5 body length was analyzed the same way. In Experiment B-3 to 4, the difference in the response to the first stimulus (initial response) was analyzed by a one-way ANOVA between treatment conditions, and Tukey post-hoc analysis between two groups; evidence of short-term habituation was established when the responses to the last three stimuli were significantly lower than the response to the first stimulus; this difference was analyzed with an ANOVA between the two responses within a treatment group. In Experiment B-5, long-term habituation was considered to 80  have been demonstrated if the response of trained worms was significantly lower than that of the untrained worms; a one-way ANOVA was used to compare between responses from trained and untrained worms within a treatment group.  81  Results Experiment B-1: Exposure of 30% ethanol for an hour during middle development increased the rate of body bends The rate of body bends and spontaneous reversals were observed in 4-d-old adult worms exposed to 30% ethanol singularly or repeatedly for a total of 20 or 60min during early, middle or late development (protocol adapted from Experiment A-3) (Figure 2.18). A group of 12-15 worms were transferred to a new Petri plate seeded with a drop of E. coli (Figure 3.3A & Figure 3.5). The number of body bends and spontaneous reversals were recorded for a period of 3min after a 5min  rested period (rested locomotory behavior), and then for another 3min after a tap stimulus (stimulated locomotory behavior). The level of locomotory behavior right after the tap was considered as a measure of “reactivity” or “sensitivity” after stimulation; worms that responded more than controls after the stimulation was considered as hyper-sensitive.  Figure 0.5 Worm handling timeline For the number of body bends during the first 3min, a three-way ANOVA using ethanol dose (untreated, 0%, 30%) treatment onset (early, middle, late or repeated) and duration (untreated, 20 or 60min) as between-subject variables showed no significant interaction of the 3 variables, or any of the two variables (Figure 3.6); however, dose (F(1) = 4.543, p = 0.034), duration (F(1) = 23.641, p < 0.0001), and onset (F(3) = 6.976, p < 0.0001) alone had effects on the number of body bends in rested worms. Post-hoc pair-wise comparisons showed that only worms exposed to ethanol for 60min during middle development had an elevated number of  82   Figure 0.6 Body bends (BB) before and after stimulus of worms exposed to 30% ethanol for 20 or 60min singly during either early, middle or late development, or repeatedly at each of the three developmental stages.  The number of body bend was increased in worms exposed to ethanol for 60min during middle development compared to the untreated and water controls. The upper solid horizontal line indicated the number of body bends of the untreated control during the first 3min and the lower dotted line indicated body bends during the last 3min. Grey vertical lines were used to divide each pairs of ethanol group and its corresponding water control. body bends of the first 3min (vs. untreated, p < 0.0001, vs. water, p = 0.008, HSD); no other ethanol groups showed significant differences when compared to the untreated and water controls. For the number of body bends during the last 3min, dose and onset showed an interaction (F(3) = 5.035, p = 0.002), and dose (F(1) = 2.672, p = 0.103), duration (F(1) = 15.872,  p < 0.0001), and onset (F(3) = 10.762, p < 0.0001) alone had effects. No ethanol groups showed any 83  difference in the number of body bends after the tap stimulus compared to the untreated and water controls. The analysis of body bends plotted across the entire behavioral recording (6min) demonstrated that treatment duration and onset alone both had significant effects on the activity curves (duration, F(1) = 25.913, p < 0.0001, onset, F(3) = 12.330, p < 0.0001, three-way repeated measures ANOVA), and the interaction between dose and onset had a significant effect (F(3) = 4.383, p = 0.005) (Figure 3.7). Pair-wise comparisons showed that only worms exposed for 60min during middle development had significantly elevated body bend activity compared to the untreated and water controls (vs, untreated, p < 0.0001, vs. water, p = 0.003, HSD). Analysis of the number of spontaneous reversals before the tap stimulus revealed a significant interaction between treatment and duration (F(1) = 13.987, p < 0.0001), and duration alone had an effect (F(1) =10.027, p = 0.002) (Figure 3.8). The rates of spontaneous reversals after the stimulus showed a significant interaction between duration, treatment and onset (F2) = 6.654, p = 0.001). However, no ethanol conditions caused a change in the number of spontaneous reversals either before or after the stimulus. In summary, the body bend activity of a rested worm was increased by ethanol exposure during middle embryonic development, but the rate of spontaneous reversals was not altered. In addition none of the groups showed a significant difference in either body bends or spontaneous reversals in the minute following the tap stimulus. This result suggested that there was no hypersensitivity in response to a mechanical stimulus. 84   Figure 0.7 The activity of body bends across the full 6min behavioral recording.  Tap stimulus occurred in between the 3rd and 4th min. The activity of body bends was higher in worms exposed to ethanol for 60min during middle development (C), but not other exposure conditions: early 60min (A) and 20min (B) middle 20min (D), late 60min (E) and 20min ( F) repeated exposure (G). ***, p < 0.001. No Rx = untreated control. The vertical lines between the 3rd and 4th minutes separate the responses before and after the stimulation, respectively.  85   Figure 0.8 Spontaneous reversals of worms exposed to 30% ethanol for 20 or 60min singly during early, middle or late embryonic development or repeated during all 3 stages.  No ethanol exposure condition caused significant changes in spontaneous reversals before or after the stimulus Experiment B-2: Chronic exposure to 10% ethanol during embryonic development resulted in a decreased rate of body bends and an increased rate of spontaneous reversals This experiment examined the locomotory activity of worms exposed chronically to 5 or 10% ethanol for a total of 8hr throughout embryonic development. Single worms were transferred onto a test plate and rested for 1min before the 3min behavioral recording began (Figure 3.9). Recall that worms exposed to 5 or 10% ethanol were smaller in size (Figure 2.12 in 86  Chapter 2). A pilot study of body lengths of chronically exposed worms showed that body length of 5% exposed worms were not shorter but 10% exposed worms were shorter than controls (data not described in this thesis). Because behavioral scoring involved monitoring and tracing worm responses on a video monitor instead of computer, body lengths were more conveniently obtained from behavioral studies. In this chapter, body lengths were measured instead of body sizes as in Chapter 2; the idea was that if chronic exposure was effective, the body lengths of the 10% exposure group should be shorter than the control groups. As expected, the body lengths of 4-d-old worms in this experiment were different among the untreated (308.56±4.62 pixel), water (291.97±6.30 pixel), 5% (296.82±6.08 pixel) and 10% (253.24±8.18 pixel) ethanol groups (F(3) = 9.761, p < 0.0001). Only the body lengths of the 10% group were significantly shorter than the other three groups (10% vs. No Rx, water, 5%, all p < 0.0001, LSD), which replicated the earlier finding and showed that 10% chronic ethanol exposure produced persistent effects on size.  Figure 0.9 locomotory behavioral recording for Experiment B-2 The number of body bends in 3min were different among the 4 groups (F(3) = 5.626, p = 0.001) (Figure 3.10A). The worms exposed to 10% ethanol exhibited fewer body bends than the other 3 groups (10% vs. untreated, p = 0.008, vs. water, p = 0.001, vs. 5%, p = 0.049, HSD). This result suggested that the 10% exposure group either moved more slowly, or spent less time being active. However, from my observation, the 10% group was not less active than the other groups; 87  during the 3min behavioral recording, no worms from any group stopped moving for a noticeable period of time. Therefore, the reduced number of body bends for the 10% ethanol group should be taken as an indication of a slower rate of locomotion. An analysis of spontaneous reversals revealed a significant difference among the 4 groups (F(3) = 3.910, p = 0.010), with the 10 % group reversing spontaneously at a higher rate than the untreated and water controls (vs. untreated, p = 0.023, vs. water, p = 0.015, HSD) (Figure 3.10B). This result suggested that 10% chronic ethanol exposure during embryonic development altered the neural circuit responsible for the regulation of spontaneous reversal, but 5% ethanol did not.  Figure 0.10 Body bends and spontaneous reversal of chronic ethanol exposure.  Worms exposed to 10% ethanol exhibited A) lower number of body bends and B) more number of spontaneous reversals Experiment B-3: Exposure during early development produced hyper-responsiveness to tap Worms exposed to 20% ethanol for 60min during either early, middle, or late development were given 10 taps at a 10s ISI to investigate for the effect of this treatment on the response to tap and habituation; worms exposed to water for 60min during early embryonic development were used as controls. The exposure protocol was the same as in Experiment A-1 (Figure 2.3). 88  Worms exposed to ethanol early during their embryonic development demonstrated elevated responses to tap (Figure 3.11A). An ANOVA indicated that treatment conditions (water, early, middle & late) had a significant effect on the initial response to tap (F(3) = 4.427, p = 0.007). Post-hoc analysis showed that 20% ethanol exposure during early development significantly increased the length of the reversal response to tap compared to the water control (p = 0.023, HSD), and late exposure group (p = 0.009, HSD).  If worms successfully habituated to the test stimuli, the magnitude of response to the last 3 stimuli (the 8th – 10th tap) should be less than that to the first stimulus. Comparison between the initial response and the last three responses demonstrated that all 4 groups successfully habituated to the test taps (water group, F(1) = 43.441, p < 0.0001, early, F(1) = 18.159, p < 0.0001, middle, F(1) = 18.719, p < 0.0001, late, F(1) = 14.606, p < 0.0001) (Figure 3.13B). Thus all groups showed short-term habituation to 10 taps at a 10s ISI. ****A) B)****** *****050100150200250300Water Early Middle Latemean response magnitudeS1S8-10050100150200250300Water Early Middle Latemean response magnitude  Figure 0.11 The reversal response to tap stimuli of worms exposed to 20% ethanol for 60min during different times of embryonic development.  A) Initial response of worms exposed to 20% ethanol during early development was higher than that of water control and late exposure group. B) All 4 conditions successfully habituated to tap delivered at a 10s ISI as the response to the last 3 stimuli (S8-10) was significantly lower than the response to the first stimuli (S1). ***, p < 0.0001, **, p < 0.01, *, p < 0.5. 89  The response curve to the 10 tap-stimuli delivered at a 10s ISI (habituation curves) showed that the magnitude of the tap response across the 10 stimuli in the early exposure group was generally higher than that of the other 3 groups (Figure 3.12A). The examination of spontaneous recovery (responses to R1-3 in Figure 3.12A) suggested that all groups did show recovery from habituation 30sec, 1min, and 5min after the last test stimuli. In order to compare the pattern of habituation curves when the initial responses were different, the reversal responses were normalized to the initial responses in Figure 3.12B, where the response was expressed as a proportion to the initial response. When response curve of the early exposure group was normalized to initial response, the curve became indistinguishable from the curves of the other three groups. This suggested that although initial response was larger for early exposed worms, habituation to the 10s ISI was not impaired in worms exposed during early development.  A) B) Figure 0.12 Habituation curves of worms exposed to 20% ethanol for 60min during different times of embryonic development.  A) The magnitude of reversal response and B) response normalized to the magnitude of the initial response across 10 taps delivered at a 10s ISI. In summary, worms exposed to 60min of 20% ethanol during embryonic development successfully learned to habituate to taps delivered at 10s ISI. In worms exposed during early development, their reversals in response to tap were generally larger, but the habituation pattern was not altered. 90  Experiment B-4: Habituation to taps at 10s or 60s ISI was not affected in worms chronically exposed to ethanol during embryonic development Since 20% ethanol exposure for 60min did not alter learning in habituation, in Experiment B-4, I examined a stronger exposure condition than that used in Experiment B-3. Worms were exposed to 5 or 10% ethanol chronically for 8hrs throughout embryonic development (Figure 3.10) and then tested for habituation to taps delivered at either a 10s or 60s ISI.   10s ISI In this experiment, the chronic ethanol exposure successfully induced difference in body lengths between the 4 groups (F(3) = 8.238, p < 0.0001, Mean of body lengths: untreated, 238.1±6.2 pixel, water, 251.1±5.1 pixel, 5%, 231.4±8.3 pixel, 10%, 200.2±8.9 pixel). The body length in the 5% exposure group wasn’t different from the water or untreated controls, but the body length in the 10% exposure group was significantly shorter than all other groups (10% vs. water, p < 0.0001, vs. untreated, p < 0.0001, vs. 5%, p < 0.0001, HSD).  The responsiveness to mechanical stimuli was examined using the magnitude of response to the first tap stimuli (Figure 3.13A). Although worms exposed chronically to 10% ethanol appeared to have a lower initial response to tap, this difference was not statistically significant. Both ethanol groups as well as the untreated and water control also appeared to habituate normally to taps delivered at 10s ISI: the magnitude of response to the last 3 taps was significantly lower than that to the first tap (untreated, F(1)= 21.143, p < 0.0001, water, F(1)= 37.162, p < 0.0001, 5%, F(1)= 44.593, p < 0.0001,10%, F(1)= 17.569, p < 0.0001) (Figure 3.15B). The habituation pattern to taps delivered at a 10s ISI is displayed in Figure 3.16A. Worms exposed to 5% chronic ethanol during embryonic development produced a response  91  050100150200250untreated water 5% 10%mean response magnitudeS1S8-10050100150200250untreated water 5% 10%mean response magnitudeA) B)********* *** Figure 0.13 The response to initial and last 3 stimuli of chronically exposed worms at a 10s ISI. The initial response to tap (A) and the difference between the response to the first (S1) stimulus and to the last three stimuli (S8-10) delivered at a 10s ISI of worms treated chronically for 8hr during embryonic development curve very similar to those produced by the water and untreated controls. In contrast, 10% exposed worms appeared to respond slightly less than the other groups, but this lower reversal distance might be associated with their shorter length. Therefore, Figure 3.14B shows the response curve normalized to body length; after reversal distance was corrected for body length, the habituation curve of 10% group apears very similar to the other 3 groups, which suggests that the slightly lower response observed in Figure 3.14A was partially due to a shorter mean body length. When the response was expressed as a proportion of the intitial response,the 4 habituation patterns became indistinguishable from each other (Figure 3.14C). Therefore, visual examination of the pattern of response to tap delivered at a 10s ISI did not suggest any changes made by ethanol exposure during development. Thus, habituation to mechanical stimuli delivered at a 10s frequency was not affected by chronic ethanol exposure during embryonic development in C. elegans. 92  00.10.20.30.40.50.60.70.80.91 2 3 4 5 6 7 8 9 10mean response magnitude normalized to body lengthStimuliuntreatedwater5%10%0204060801001201401601802001 2 3 4 5 6 7 8 9 10Mean response magnitudeStimuliuntreatedwater5%10%00.20.40.60.811.21 2 3 4 5 6 7 8 9 10Mean response magnitude normalized to intial responseStimuliuntreatedwater5%10%A)B) C) Figure 0.14 Habituation to tap stimuli delivered at a 10s ISI of worms exposed chronically for 8hrs to ethanol during embryonic development.  A) raw response. B) response normalized to worm length. C) Response normalized to initial response.   60s ISI The body lengths in worms participated in the 60s ISI study were significantly different among the 4 conditions (F(3) = 27.187, p < 0.0001, ANOVA), the body lengths in the 10% ethanol group (182.59±5.27 pixel) were significantly shorter than the untreated (240.66±6.79 pixel), water (236.25±4.62 pixel) and 5% ethanol groups (233.12±6.21 pixel) (10% vs. untreated, water, and 5%, all p < 0.0001, HSD), but in the 5% ethanol group were not. The initial response to tap, although variable, was not significantly different between groups or between any two groups (Figure 3.15A). Analysis of the difference between the reversal response to the first and  93  050100150200250300350untreated water 5% 10%Initial responseS1S8-10050100150200250300350untreated water 5% 10%Initial responseA) B)***** **** Figure 0.15 The response to initial and last 3 stimuli of chronically exposed worms at a 60s ISI. The initial response to tap (A) and the difference between the response to the first (S1) stimulus and to the last three stimuli (S8-10) delivered at a 60s ISI of worms treated chronically for 8hr during embryonic development last three stimuli suggested that all 4 groups, including ethanol exposed groups, had successfully habituated to taps delivered at a 60s ISI (untreated, F(1) = 9.822, p = 0.002, water, F(1) = 32.068, p < 0.0001, 5%, F(1) = 11.466, p = 0.001, 10%, F(1) = 11.322, p = 0.001) (Figure 3.15B).These findings suggested that habituation at 60s ISI was not impaired in worms treated with 5% or 10% ethanol chronically during embryonic development. Visual examination of the habituation curve suggested that all 4 groups responded more or less the same to the tap stimuli delivered at a 60s frequency (Figure 3.16A). When the responses were normalized to body length, in order to accommodate the shorter body length of 10% ethanol group, the responses of 10% group appeared to be larger proportional to the body length (Figure 3.16B). When the curve was normalized to initial response, the responses produced by untreated, water, 5 or 10% ethanol groups appeared similar to each other (Figure 3.16C). In summary, ethanol exposure to 5 or 10% ethanol did not alter the initial reversal responses and habituation to tap delivered at 10s or 60s ISI. Thus, short-term habituation to mechanical stimuli at these two frequencies was normal in worms exposed to chronic ethanol during embryonic development. 94   Figure 0.16 Habituation to tap stimuli delivered at a 60s ISI of worms exposed chronically for 8hrs to ethanol during embryonic development.  A) raw response. B) response normalized to worm length. C) Response normalized to initial response.   Experiment B-5: Long-term memory of habituation training was intact in animals chronically exposed to ethanol during embryonic development Worms exposed to 8hr of chronic ethanol during embryonic development were assessed for their ability to retain memory of habituation training that occurred a day earlier. The body lengths of the tested worms were different between the 4 treatment conditions (F(3) = 10.685, p < 0.0001), and the body lengths of 10% groups were shorter than that of the 5% ethanol group (p = 0.009, HSD), untreated (p < 0.0001, HSD) and water controls (p < 0.0001, HSD); these demonstrated the effectiveness of the 10% ethanol exposure.  95   Figure 0.17 Long term memory of habituation training in worms exposed chronically for 8hr during embryonic development.  Lower response of trained worm compared to un-trained worm (control) indicated that trained worm retained memory of habituation training. Long-term habituation training was delivered as 4 blocks of 20 taps delivered every 60s and each training block was separated by an hour of resting period (Figure 3.4). If worms remembered the training, they should respond less to the test taps delivered to them 24hr after training. Within the untreated control, trained worms responded significantly less than the un-trained (control) animal (F(1) = 9.380, p = 0.003), which indicated that the training procedure was successful at producing memory (Figure 3.17). Both 5% and 10% ethanol exposure groups also had intact long term memory as the trained worms from each of the two ethanol groups responded significantly less than the untrained worms did (5%, F(1) = 6.682, p = 0.010, 10%, F(1) = 7.874, p = 0.006, ANOVA). Unexpectedly, trained worms from the water exposure control was the only group that failed to demonstrate long term memory (F(1) = 1.262, p = 0.263). This finding was puzzling. Independent analysis of the two replications used in this study suggested that the memory failure in the water control occurred only in the first replication. The water and 10% groups from the second replication displayed the slightly elevated initial responses as seen in Experiment B-4. This difference in initial response of the water and 10% 96  groups observed from the second replications was not likely due to environmental difference between the two replications because the responses from both the train and untrained groups of the untreated control are visually identical between the two replications. This suggested that some unknown factors present in the second replication might be responsible to cause the elevated response found in the water and 10% groups, and the absence of the same factor might be responsible for the failure of long-term habituation in the water control. More replications of this experiment will be required to confirm whether the elevated response found in the water and 10% groups are repeatable phenotypes, or, whether the absence of the memory found in the water group was repeatable. In summary, chronic exposure to ethanol strong enough to induce developmental delay did not impair memory to habituation training in C. elegans.  97  Discussions Alteration in the number of body bends and spontaneous reversals in the 10% chronic animals may be either a result of motor dysfunction or developmental delay  Worms exposed to 10% chronic ethanol during embryonic development moved at a slower rate and reversed spontaneously at a higher rate. Factors that may influence the rate of locomotion include age, the presence of food, and the humidity in the agar and air (Chiba and Rankin, 1990; Zhao et al., 2003). The humidity in the agar and air should not be an issue because those were carefully controlled as our experiment were carried out in a temperature and humidity-controlled facility and the agar plates were stored in 4oC and used within a few days of each other. The amount of food was also controlled carefully. That left us the age effect to consider. Chiba and Rankin (1990) found that the frequencies of spontaneous reversals were quite stable across different ages except that it was higher in young adult worms. Since the shorter body lengths in the 10% group might result from developmental delay, most individuals in the 10% group might still be at the young adult stage. Curiously, I found 1 individual in the 10% group was still at the Larva-4 stage, but most other individuals looked older than L4, and a few of them looked as old as the controls. This observation suggested that the higher spontaneous reversals in the 10% group might simply be a product of delayed development. For the rate of body bends, in contrast, there was no published developmental study to compare with the body bend findings from this study so it is unclear whether the reduced number of body bends found in the 10% group could also be a result of developmental delay.  To investigate whether developmental delay was the sole reason for the alteration of body bends and spontaneous reversals, the developmental ages of the worms must be known. Since 10% chronic ethanol exposure during embryonic development produced a heterogeneously aged 98  population, each worm in the 10% group must be aged individually. If being delayed and in the young adult stage on Day 4 was the reason for the higher spontaneous reversal of the 10% group, only worms staged at young adult should exhibit higher spontaneous reversals. Additionally, the 10% group may be tested when they grow to the developmental stage that matches that of the other groups. If the age-matched 10% worms still have the same spontaneous reversals as controls, then the alteration in spontaneous reversals seen in this study may not be simply caused by developmental delay induced by ethanol exposure. Similarity, the analysis of body bends can be done during at the same time. If it turns out that the alteration in body bends and spontaneous reversals of the 10% group were not simply due to a developmental delay then that suggests the neural circuitry responsible for the locomotory regulation may be altered by ethanol exposure during embryonic development. As neuronal circuitries involved in the locomotory behavior in C. elegans have been mapped out and many neuronal proteins involved were deciphered (Tsalik and Hobert, 2003; Gray et al., 2005; Glodowski et al., 2007), we are equipped to investigate further problems that underlie this locomotory abnormalities found in the 10% exposed worms. Body bend was increased by acute exposure but decreased by chronic exposure A comparison of Experiment B-1 and B-2 should take into account two main variables: the behavioral and the exposure protocols. For behavioral differences, in Experiment B-1 worms were given 5min rest period, but only 1min rest period in Experiment B-2 (Figure 3.20).  Even after the 5min of rest, body bend activity gradually declined during the first 3min of the behavioral recording in Experiment B-1 (Figure 3.7). This decline suggested that 1) the body bend activity had not reached a baseline level after being transferred, which was a stronger source of mechanical stimuli than a tap, and 2) the body bend activity during the 5min of rest period should be even higher than that was observed from the first 3min of behavioral recording 99  in Experiment B-1. According to the Figure 3.18, Experiment B-2 was recording a period of time equivalent to the 2nd-4th min of the rest period in Experiment B-1. If the previous prediction was true, the number of body bends observed in Experiment B-2 should be higher than that in Experiment B-1. That was indeed the case: the untreated worms in Experiment B-2 moved an average of 85.28 body bends within 3min while the untreated worms in Experiment B-1 only did an average of 53.29 body bends within 3min. Different types of locomotory neuronal circuit may predominate in Experiment B-1 and 2. After experiencing a mechanical stimulus, a worm may have one priority: running as far away from the stimulus as possible. During this time, the escape   Figure 0.18 The difference in the protocol to obtain behavioral recordings in Experiment B-1 (A) and B-2 (B) circuit may dominate leading to a period of rapid forward movement aiding the worm to move away from the source of the stimulus (Gray et al., 2005). After a short period of time, the worm gradually slows down from escaping and looking for food and exploring the foreign environment slowly becomes the new priority (Gray et al., 2005).  During this “exploratory period” with the presence of food (as in Experiment B-1 and 2), the worm moves at a slower rate (Sawin et al., 2000), and this activity involves more complicated neuronal circuits as the worm may be taking 100  more sensory information (i.e. chemical, temperature) into consideration. In Experiment B-2, the escape circuit may have dominated the exploratory circuit because the worm just experienced a stimulus 1min ago. However, in the first 3min of behavioral recording in Experiment B-1, the exploratory circuit took more weight than that in B-2 because the worm experienced the stimulus 5mins ago. This difference in the circuit balance between Experiment B-1 and 2 not only offers a possible explanation for why the body bends in B-2 were larger than in B-2, but also suggests that the body bend enhancement seen in B-1 might reflect abnormalities in the exploratory circuit. In Experiment B-2, the decrease of body bends might suggest problems that occurred in the escape circuit. Alternatively, the opposite effects on body bend found in Experiment B-1 and 2 may be simply caused by different exposure protocol. Worms in Experiment B-1 experienced acute ethanol exposure for 60min or less at specific times but worms in Experiment B-2 experienced chronic ethanol exposure for 8hr throughout embryonic development. As described previously, the changes in locomotion of the 10% chronically exposed worms may be due to developmental delay. However, worms exposed to 30% acute ethanol did not exhibit marked delay in development so the locomotory abnormalities found in worms exposed during middle development should not be associated with delay in development. Moreover, the modes of damage by acute and chronic ethanol exposure may be different from each other; however, the difference in the underlying mechanisms responsible for the damage exerted by chronic or acute ethanol exposure during development is still poorly understood. Further investigation into this area may reveal very interesting and useful information to enrich the understanding of the complex interaction between early ethanol exposure and nervous system development.   101  A more difficult task may be required to see learning and memory deficits in exposed worms Although all worms tested at the 10s ISI displayed significant habituation to taps, there were subtle differences in the amount of decrease from the initial to last three responses. The magnitudes of decreased responses were not different in animals exposed to 20% acute ethanol in Experiment B-3, but in chronic exposure, the magnitudes of decreased response were smaller in worms exposed to 10% ethanol compared to the untreated control (p = 0.043, HSD), which suggested that though the 10% group had successfully habituated to 10s ISI mechanical stimuli,  it did not habituate to the same extent as seen in the untreated control. In other words, the smaller decrease in response might suggest that the 10% group did not habituate as quickly as the untreated controls. Whether worms in the 10% exposure group can habituate to the same asymptotic level as controls, or whether the 10% group may take longer to reach the same asymptotic level may be revealed through examinations into the habituation pattern to an increased number of taps (i.e. 30 taps instead of 10 taps). No significant problems were found in the short- or long-term habituation of worms exposed to ethanol during embryonic development. The lack of effect from habituation assays did not replicate the finding described in the rat study (Cohen et al., 1985), however there are several possible explanations for this. In humans, the problem with habituation is evident in infants (Streissguth et al., 1983), but the impairment fades with time, so do many other types of neurodevelopmental problems caused by prenatal alcohol exposure (Nagahara and Handa, 1999). One possibility is that habituation may be a problem in larval C. elegans but the defect disappeared in adult worms. Another possibility is that only a specific type of habituation was affected, for example, the habituation to smell or taste, and the habituation to mechanical stimuli was not affected by ethanol exposure. A third possibility is that habituation was not affected by 102  ethanol exposure in C. elegans. However, this finding does not eliminate the possibility that other types of learning and memory may be affected by ethanol exposure during embryonic development. Kodituwakku (2007) suggested that FASD children were impaired more in complex tasks than in simple tasks. The habituation tasks we used to challenge the worms may be too easy to reveal ethanol-related defects. An example of a more difficult task for C. elegans is salt conditioning, which requires the worm to associate high salt concentration with the presence or absence of food. In addition, long-term habituation may be tested 48hr instead of 24hr later, which requires the worm to retain memory for a longer period of time.  In conclusion, the results from this chapter suggested that worms exposed to 10% ethanol chronically during embryonic development had alterations in locomotory activity. However, none of the exposure protocols tested produced noticeable impairments in short- or long-term habituation to mechanical stimuli. Further investigations using more difficult learning and memory tasks are needed to conclude whether ethanol exposure during embryonic development in C. elegans produces learning and memory deficits.   103  CHAPTER 4. GENERAL DISCUSSIONS AND CONCLUSIONS The goal of this thesis was to assess physical/developmental and behavioral effects of embryonic ethanol exposure in C. elegans. The findings of physical/developmental and behavioral effects were parsed into two chapters. In Chapter 2, I investigated the relationship between embryonic ethanol dose and time of exposure on hatch rate, reproduction onset, lifespan, and body size in C. elegans. I found that early embryonic development was particularly vulnerable to ethanol toxicity, and that multiple exposures induced larger physical effects than did single exposures. In Chapter 3, I examined the relationship between embryonic ethanol dose and time of exposure on locomotory activity and habituation to tap. I found locomotory activity was altered in the worms exposed to acute or chronic ethanol during embryonic development, but I did not find effects on either short- or long-term habituation. In chapter 2, describing physical/developmental effects of ethanol exposure, delayed reproductive onset and smaller body size were characteristics of worms exposed to high dose of ethanol during embryonic development. Interestingly, larval ethanol exposures (Davis et al., 2008) produced strikingly similar physical effects to those seen in embryonic exposures. Delays in reproductive development and retardation in body growth are general physical/developmental phenotypes found in worms exposed to ethanol during both developmental stages: a high dose of ethanol is required to induce these physical effects. These findings are parallel to findings in other animal models of FASD. For example, postnatal growth retardation is prominent and long lasting in rats exposed to a high dose of ethanol (6g/kg/day throughout gestation) (review see Abel, (1985). However, growth retardation is rarely seen in rats exposed to a low dose of ethanol (2g/kg/day throughout gestation); while with a moderate dose (4g/kg/day throughout gestation) the resulting growth retardation is subtle and is short-lived. A similar dose-dependent 104  relationship of growth retardation was found in zebrafish (Bilotta et al., 2004),  chickens (Satiroglu-Tufan and Tufan, 2004) and humans (Little, 1977). The results described in chapter 2 demonstrated that in C. elegans growth retardation was a common consequence of developmental ethanol exposure. Therefore some mechanisms by which ethanol induced growth retardation may be conserved in nematodes. In chapter 3 on behavioral effects of ethanol, locomotory activity was found to be altered by both acute and chronic ethanol exposure conditions. The rate of body bends was increased in worms exposed to 30% ethanol for 60min during middle development, but was decreased in worms exposed to 10% chronic ethanol throughout embryonic development. This opposite finding may due to the difference in the state of worms or the difference in the exposure pattern between the two experiments. Moreover, the rates of spontaneous reversals were increased in worms exposed to ethanol chronically, which might suggest hyperactivity. In humans, hyperactivity is a common symptom of children suffering from FASD (Mattson and Riley, 1998), however rodent models of FASD showed mixed results: some reported hyperactivity (Osborne et al., 1980; Bond, 1981; Meyer and Riley, 1986), but some studies did not (Vorhees, 1989; Downing et al., 2008). Downing et al. (2008) suggested that, in the mouse model of FASD, the presence of hyperactivity was influenced by differences in the age of the animals tested and different patterns of ethanol administration. These results imply that locomotory problems of FASD may be caused by a combination of factors, and the details are not well understood. The observation that locomotion can be altered by embryonic ethanol exposure in C. elegans provides a novel avenue for the analysis of mechanisms underlying the locomotory problems in FASD. 105  Modeling human disease in C. elegans has already led to a number of important discoveries because many biochemical pathways in humans are highly conserved in nematodes. A few examples are the insulin pathway of Type II diabetes (Kimura et al., 1997), the p53 pathway of cancer (Derry et al., 2001), and the SREBP control of obesity (Yang et al., 2006). Metabolism of alcohol is a by-product of many highly conserved biochemical pathways, such as fatty acid synthesis, glycerolipid metabolism, and bile acid biosynthesis (Lehninger et al., 2000): it is essential for organisms to have catabolic pathways to eliminate alcohol. In fact, the amino acid sequence of the zinc-binding alcohol dehydrogenase family, which is responsible for the oxidation of ethanol into acetaldehyde, is conserved from bacteria to humans (Anthony, 2001); the homologs of the zinc-binding alcohol dehydrogenase  have also been found in C. elegans (Glasner et al., 1995). Many pathways responsible for damage resulting from ethanol exposure may also be conserved in C. elegans. This thesis has established a foundation upon which to examine the mechanistic cause of ethanol teratology.  Further research in this area will inevitably lead to novel sights for this complex disorder. General Discussions Phenotypic variability caused by developmental ethanol exposure occurs in an isogenic population The high phenotypic variation resulting from developmental ethanol exposure is a curious but frustrating topic in FASD research. In human and rodent studies the variability is often attribute to individual genetic variation. Because the C. elegans population is isogenic, or genetically identical, the variability observed in this thesis was not likely a result of genetic variability. Thus the observed variation in this model system must be caused by some other, as 106  yet unknown factors such as environmental differences, exact developmental time exposure begins and/or ends, epigenetic background, or even random chance events.    The developmental program is highly specific and sensitive. Any external changes, such as exposure to ethanol, may produce damage or alter the progression of subsequent development. Developmental processes may respond to environmental changes in an adaptive or maladaptive way (Figure 4.1). Cells may repair themselves from damage made by ethanol exposure or may alter themselves to compensate for the presence of ethanol. In this case, the developing embryo successfully adapts to ethanol, and the resulting worm appears normal, at least superficially. On the other hand, the embryo can fail to adapt to ethanol exposure and the consequences may be embryonic arrest (death), growth retardation, physical and or later behavioral problems. However, this is not to say that the adapted embryo does not harbor any changes; instead, the changes made (i.e. repair & compensation) have led to a relatively normal individual compared to ones that fail to make the right changes. Embryos may not adapt to all but only some effects of ethanol exposure; and in that case, the worm becomes a partially “affected worm” and does not display all symptoms observed in the “sick worm”.  This model may also explain why more worms were affected under high ethanol dose, and more appeared normal under low dose (Figure 4.1). When the ethanol treatment is harsh, a greater number of embryos fail to adapt. When the dose is low, more adapt and fewer fail to adapt. Figure 4.2 showed an illustration of this idea using body size data collected from various repeated exposure conditions. Body sizes of worms exposed to 30% ethanol for 60min are all smaller than ones of the water control (0% 60min); the majority of worms in the 20% 60min condition had normal body size, but some have small sizes similar to ones in the 30% 60min group; the 40% 30min group, however, consisted of some normal size, some slightly smaller,  107   Figure 0.1 Models to explain the variable phenotypes observed in different ethanol treatment severity.  108  and one very small worm (40% 60min was not used here because all embryos died, and 30min of 40% ethanol was more severe than 20% 60min, but less severe than 30% 60min [See Table 2.1]). The data presented in Figure 4.2 follow the prediction made by the proposed model: as the dose increased, the proportion of “normal” worms decreased and the “sick” worms increased. However, whether this idea may hold true for other animal models or for other phenotypes in C. elegans requires further investigation. In addition, the testability of this model is questionable as the definition of “normal” requires further refinement. In other words, this model describes an attempt to conceptualize the underlying forces responsible for the vast variability observed from a population that is genetically identical and is in a highly controlled environment.  Figure 0.2 More severe ethanol exposure during embryonic development produced more worms with smaller body sizes, and less severe ethanol exposure produced more worms with normal body sizes.  The data was extracted from repeated exposure conditions described in Experiment A-2.  The body sizes of 3-d-olds exposed to water exposure repeatedly for a total of 60min, 20min each at early, middle and late exposure (0% 60min ), 30% ethanol repeatedly for 60min (30% 60min), 40% ethanol repeatedly for 30min (40% 30min), and 20% ethanol repeatedly for 60min (20% 60min). 30% 60min was considered to be a relatively harsher treatment than 40% 30min because the latter did not produce reduction in body size, but 30% 60min did (see Table 2.1).  109  The ethanol concentration used was high, but the serum concentration in the egg was unknown Data presented in Davis et al. (2008) and in this thesis demonstrated that the embryonic exposure required a much higher dose than larval exposure to induce physical and behavioral effects in C. elegans. While exposure to more than 40% ethanol for an hour killed 100% of embryos, 0.5-0.6M ethanol (approximately equivalent to 2.5-3% ethanol, v/v) infused in agar was enough to kill larvae or adult worms within an hour (Davies and McIntire, 2004; Davis et al., 2008). Since 0.5M ethanol is equivalent to 3% ethanol (v/v), the ethanol dose required to kill embryo was more than 13 times the concentration required to kill worms outside of eggs. This higher lethal dose for embryos is not unique in the C. elegans system. Shell-less chicken embryos required 48hr of 70% ethanol to achieve 100% lethality, and 24-48hr of 40% ethanol to induce physical effects (Giles et al., 2008). Compared to that, the hour of 40% ethanol exposure that was required to achieve 100% lethality in C. elegans was not abnormal. This higher resistance to alcohol of C. elegans eggs compared to that of larva and adult was previously reported by Williamson and colleagues (1991).  In contrast, in rat models of FASD, growth retardation was reported to be more closely related to blood alcohol levels (Abel, 1981) than to the amount of alcohol administered (Maier and West, 2001b). That suggested the actual serum alcohol concentration in the C. elegans embryo would be a critical piece of information. I attempted to measure that using methods described in Davis and McIntire (2004) designed for the measurement of the alcohol content in adult worms. A major problem, aside from collecting enough eggs, was the elimination the egg shell. The C. elegans embryo is enveloped by a poorly permeable vitelline membrane, and then structurally protected by a chitinous egg shell (Schierenberg and Junkersdorf, 1992). The embryo can develop normally without an egg shell but not with a damaged vitelline membrane. 110  Therefore, the measurement of serum alcohol concentration should concern the contents within the vitelline membrane only, and ethanol trapped between the egg shell and vitelline membrane should not be included. However, so far the complete egg shell removal of a large population of eggs (ten of thousands) has been technically challenging.    A plausible way to directly measure the amount of ethanol entered into the embryo would be the use of a radioactive ethanol molecule. The oxygen atom of ethanol (CH3CH2O16H) may be replaced with a radioactive one (CH3CH2O15H). The radioactivity detected within the exposed embryo could be measured and calculated to give precise kinetics of ethanol entry and ethanol metabolism. One limitation of this method is that it is generally considered undesirable due to concerns for safety so other indirect measures are usually explored before choosing the radioactive technology. An indirect way may employ the activity of ethanol-response genes, such as alcohol dehydrogenase and others described in Kwon et al. (2004), as reporters of relative ethanol serum levels. The good candidate would be a gene that is only expressed upon ethanol exposure, or a gene that alters expression level considerably upon ethanol exposure. A reporter gene can be tagged with green fluorescent protein (GFP), and the level of GFP expression following ethanol exposure can be visualized and measured. We coould then compare the activity of such reporter gene with ones in the mammalian models of FASD and humans. One caveat is that the activity level of the reporter gene in C. elegans must be assumed to be the same as the level in the other animal models; another is that this reporter method only provides the relative genetic response to ethanol exposure so the actual serum alcohol concentration still remain unknown.   111  Limitations and Future Directions  FASD research using a simple model such as C. elegans can aid in uncovering the conserved pathways involved in ethanol teratology, but there are some limitations with the generalizability of this research to humans. The route of ethanol entry in C. elegans is not the same as humans. C. elegans eggs were bathed in a stable ethanol concentration while human fetuses are exposed through ethanol content in the blood stream, which may oscillate depending on the rate of maternal alcohol consumption and metabolism. C. elegans also does not have a placenta serving as a selective barrier, or a liver designated to catabolize alcohol. Moreover, as suggested in Abel (1982), the rate of ethanol metabolism in rats was higher than that in humans; as a result, a higher amount of ethanol was required to maintain similar blood alcohol concentration in rats. Similarly, the rate of metabolism in C. elegans may be many magnitudes faster than humans. In fact, adult C. elegans can recover from a nearly lethal dose of ethanol exposure in about an hour (Davies and McIntire, 2004). Consequently, Cudd (2005) speculated that the high alcohol concentration necessary to induce defects in non-mammalian systems creates a problem when using them as models of FSAD because the mechanisms involved in the ethanol toxicology at high concentration might be different than ones involved at low concentration. Therefore, extra consideration should be taken before applying findings from C. elegans to humans. C. elegans is a great model for nervous system disorders because it contains a manageable number of neurons. However, the simplicity may be a limitation when the type of behavior of interest requires higher order brain functions. C. elegans does not have a central nervous system (CNS), the neuronal synapses are formed en passant, and the total neuron numbers are fixed; these characteristics of C. elegans nervous system restrict the use of this 112  model to study problems that involve a highly specialized nervous system. For example, specific types of neuronal cells and some brain regions were affected more than others by prenatal alcohol exposure (Riley et al., 2004). Ethanol exposure was found to shorten the length and number of dendrites (Qiang et al., 2002). Studies of ethanol teratology on brain regions or dendrite morphology are not feasible in C. elegans.  The detailed cell lineage map for C. elegans provides a great advantage to study the effects of ethanol to specific developmental processes; however, nematode and human development have many differences. The specifics in gastrulation, organogenesis, and morphogenesis differ between phyla (i.e. Phylum Nematoda, Arthropoda, and Chordata) (Wilt and Hake, 2004). Neural tube formation, an important process that is affected in FASD does not occur in invertebrates. The brain spurt period during the third trimester of human fetus may not have an equivalent in nematode development. Therefore, ethanol’s effect at some developmental stages in humans may not be investigated in C. elegans.  Although there are a few constraints, using C. elegans as a model for FASD provides many advantages. The 4 factors (concentration, timing, duration, and frequency) involved in ethanol exposure pattern can be analyzed easily in C. elegans due to its short generation time and large progeny size. The effect of embryonic ethanol exposure or the expression of specific proteins can be visualized in vivo with the aid of green fluorescent proteins. 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In the Centrifuge method described in Chapter 2, eggs were washed off of an agar plate, centrifuged into a pellet, and dried with a Kimpwipe before the introduction of the treatment solution; at the end of the treatment, the eggs were centrifuged into a pellet, removed from treatment solution, washed twice with water, centrifuged into a pellet, resuspended with 100-200µl of ddH2O, and then dispensed onto a fresh agar plate. This method set a limitation to the accuracy of egg counting. Eggs are most accurately counted if: a) the eggs are dispersed within a minimal area on the agar, b) the agar surface is free of liquid and debris, and d) the eggs are not clumped together. Any of the above obscures the visibility of eggs thus deestroys the accuracy of egg counting. Unfortunately, dispensing eggs within a restricted area lengthened the time for it took the suspension liquid to dry completely (1-2hr), condensed the density of debris left from Kimwipe, and increased the likelihood of egg clumps. Conversely, dispensing eggs about a larger area exponentially increased the duration of egg counting and the chance of overlooking eggs and especially L1 larva; in addition, doing so did not eliminate the chance of debris obscuring some eggs or larvae because the quantity of debris remained the same independent of the dispensed area. Therefore, the Agar-Bath method was developed in order to accommodate many problems mentioned above. 123   Appendix B: iMovie, QuickTime, and Image J Operation for the measurement of body size in C. elegans Worms were filmed as described in Chapter 2. After clipping JPEG images of worms, the iMovie file was converted into a QuickTime file as follows: under “Share”, choose “QuickTime”, and select “compress movie for CD-ROM” with “15 frames per second, 320x240” option. This conversion process condensed the iMovie file into approximately 1.5 megabytes instead of the original size of 1.5 gigabytes. This conversion procedure was important to maintain the original scale of the iMovie recording in the QuickTime file. To reload a large QuickTime movie back into iMovie, the QuickTime file needed to be split into smaller pieces using QuickTime Pro and each pieces were loaded individually back into iMovie. Note that the newer versions of iMovie does not contain picture clipping functionality. Therefore, if recording is done with the newer version, the file needed to be converted into files readable by the older version for the 2-D images to be extracted from the older iMovie program. To measure worm body size in Image J, first open a JPEG image file, and then open the macros 1-4 described below. Macro 1 was used to set global measurement conditions such as pixel-to-millimeter conversions and the types of measurements desired to be taken (area = body size, perimeter = the length that encircled the body area, feret’s distance = the length bewteen the two points furthest away from each other). Micro 2 was used to convert images into 8-bit format and set a desired threshold. Sometimes threshold needed to be re-adjusted according to the light conditions used during body size recording. However, adjusting threshold introduces inconsistencies between readings. To avoid changing threshold, one should keep the light source as uniform as possible between recordings. Micro 3 was used to convert the image so that the worm body was covered with black. After running Macro 3, check if the worm body was 124  correctly represented by the blackened area. If not, trace the outer edge of the worm manually using the pen function in Image J, and then re-run Macro 3 to refill in the encircled area. Next, Macro-4 was used to measure body size, save the image as TIF file, and then prompt to open the next JPEG file for the next worm. Once a set of measurement had been taken, the result was cut and paste from image J to Excel, and each measurement was labeled accordingly.  Macro 1 run("Set Measurements...", "area perimeter feret's limit redirect=None decimal=9"); run("Set Scale...", "distance=225 known=1 pixel=1 unit=mm global");  Macro 2 run("8-bit"); setThreshold(26, 58);  Macro 3 run("Make Binary", "thresholded remaining black"); run("Fill Holes");  Macro 4 run("Measure"); saveAs("Tiff", ""); close(); open("");  Worm size and length measurements were represented in µm2. Image J was calibrated to automatically convert pixel2 readings into µm2. The number of pixels corresponding to the length of a millimeter was determined by filming a ruler under the same magnification of the body size recording. Since the mark of the ruler was quite thick under the microscope, the pixel distance of between the two millimeter marks including the width of the first mark but excluded the width of the second mark was measured 10 times and the mean was used for conversion. The number used in this thesis was 225pixel/mm.  125   Appendix C: Water effect: The timing of water control treatment A discovery made during this project was that incubating embryos in distilled water somtimes produce physical effects in the hatched worms. In Figure C, water treatment for 60min during middle and late development, or repeatedly for 60min (20min each at early, middle or late development) produced a delay in reproductive onset, but treatment during early development did not. This data suggested that water treatment at different developmental stages produced different physical effect. Therefore, it is important to have a water control for every ethanol exposure patterns. In addition, an untreated control, which was not immersed in liquid of any kind, was used in order to compare for the effect resulting from water treatment.    Figure C  Reproductive onset in worms exposed to 60min of water during early, middle, or late development singly or repeatedly. **, p < 0.001, ***, p  0.0001.      126   Appendix D: Water vs. M9  Figure D. Pictures of the surface of agar after depensing 100-200µl of H2O/EtOH solution or M9/EtOH solution. Note the prominent number of salt crystals on the surface of the M9/EtOH plate  Since bathing embryos in water appeared to produce physical effects, M9 (physiological saline for C. elegans) was thought to be a more logical solvent for ethanol treatments. However, pilot experiments yielded an unexpected result. As shown in Figure D, M9 left many salt crystals on the agar plate after its liquid component evaporated. The appearance of these crystals made the egg and larva counting virtually impossible. In addition to producing low visibility, these salt crystals would also create an extreme hypertonic environment. Ironically, our original thought to use M9 was due to concerns for the hypotonic environment created by ddH2O during treatment of the eggs, but it appeared that the use of M9 would result in hypertonic environment not just for the eggs but also for the newly hatched worms. This undesired high salt environment would introduce an even larger confound into the study. One way to eliminate these crystals would be to wash off the M9 buffer with water. However, the original purpose of the switch to M9 was to avoid possible osmotic pressure exerted by water. If we need to use water to wash off M9, we defeat our original purpose to switch. Therefore, water remained to be the solvent of choice for this thesis.  

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