The Effects of the General Anaesthetic Propofol on Drosophila Larvae Drew Min Su Cylinder Bachelor of Science

The effects of the general anaesthetic propofol on Drosophila larvae Drew Min Su Cylinder Bachelor of Science

A thesis submitted for the degree of Master of Philosophy at The University of Queensland in 2019 Queensland Brain Institute

Abstract

Although general anaesthetics have been in use since the mid-19th century, the mechanism by which these drugs induce reversible loss of consciousness is still poorly understood. Previous research has indicated that general anaesthetics activate endogenous sleep pathways by potentiating GABAA receptors in wake-promoting neurons. However, more recent studies have demonstrated that general anaesthetics also inhibit synaptic release through interactions with the SNARE complex, an integral part of presynaptic neurotransmitter release machinery in all neurons. The presynaptic and postsynaptic mechanisms may thus be linked in a two-step process: at low doses, general anaesthetics activate sleep-promoting circuits, thereby producing unconsciousness, while at the higher doses necessary for surgery, general anaesthetics inhibit presynaptic release machinery brain-wide, thereby causing a total loss of behavioural responsiveness. While this hypothesis remains speculative, it is testable in animal models. This study develops larval Drosophila melanogaster as an animal model to test this hypothesis in the context of a common intravenous GABA-acting general anaesthetic, propofol. Although presynaptic effects of general anaesthetics have been studied in larval neuromuscular junction preparations, there is not much data for how these drugs affect larval behaviour or brain activity. General anaesthesia is easily addressed in animal models because it can be described as a state of decreased responsiveness which can be assessed using diverse behavioural endpoints. In this study, a series of behavioural assays were designed and tested to assess the effect of GABA-acting general anaesthetics and sedative drugs on Drosophila larvae. These assays were then used to explore key parts of the pre- and postsynaptic mechanisms thought to underlie general anaesthesia. It was found that knockdown of the RDL subunit, the main anaesthetic target in the GABA receptor, did not confer resistance to propofol. However, deletion of the N-terminus of the H3 domain in syntaxin1A, a key component of the SNARE complex, resulted in resistance to propofol and isoflurane. These results suggest that propofol and isoflurane act through similar mechanisms and that this mechanism involves a presynaptic component that transcends life stage or brain size. Larval Drosophila thus provide a valuable counterpart to adult flies for investigating presynaptic mechanisms of anaesthesia.

Declaration by author

This thesis is composed of my original work, and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, financial support and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my higher degree by research candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University Library and, subject to the policy and procedures of The University of Queensland, the thesis be made available for research and study in accordance with the Copyright Act 1968 unless a period of embargo has been approved by the Dean of the Graduate School.

I acknowledge that copyright of all material contained in my thesis resides with the copyright holder(s) of that material. Where appropriate I have obtained copyright permission from the copyright holder to reproduce material in this thesis and have sought permission from co-authors for any jointly authored works included in the thesis.

Publications included in this thesis

No publications included.

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Submitted manuscripts included in this thesis No manuscripts submitted for publication.

Other publications during candidature No other publications.

Contributions by others to the thesis Chapter 2: Dr Jessica Waanders helped design and perform the high performance liquid chromatography experiments. Chapter 3: Dr Oressia Zalucki constructed UAS-HA syntaxin227. Deniz Ertekin helped perform the immunohistochemistry experiments and helped write the associated methods. Chapter 4: Dr Lucy Heap performed the watershed segmentation of the two-photon microscopy data mentioned in this chapter.

Statement of parts of the thesis submitted to qualify for the award of another degree No works submitted towards another degree have been included in this thesis

Research Involving Human or Animal Subjects No animal or human subjects were involved in this research

Acknowledgements

I am first and foremost most grateful to Dr Bruno van Swinderen who has been an extraordinarily patient and understanding supervisor. I am also thankful to my co- supervisor, Dr Victor Anggono, with whom I completed my first lab rotation.

Thank you to my committee members: Dr Nela Durisic, Dr Kai-Hsiang Chuang, and Dr Helen Cooper who always kept me on my toes with insightful questions and gave me plenty of advice that helped shape this project.

Thank you to all the members of the van Swinderen Lab: Dr Leonie Kirszenblat, Dr Martyna Grabowska, Dr Lucy Heap, Dr Kai Feng, Dr Michael Troup, Adam Hines, Rhiannon Jeans, and Deniz Ertekin. Your enthusiasm and quirky sense of humour made work enjoyable, and your expertise made this thesis possible. In particular, thank you to Dr Michael Troup for your guidance on this project. Thank you also to Dr Kunle Bademosi who taught me how to perfectly fillet a maggot, and Dr Lucy Heap whose MATLAB wizardry made chapter 4 possible. I would also like to give special thanks to Deniz Ertekin who gamely answered all of my questions no matter how stupid or pedantic.

This thesis would also not have been half as enjoyable without my MPhil cohort: Sam Armstrong, Julius Alpay, Clare Harris, Chai Chee Ng, and Maleeha Waqar. You have been an inspiring group to go on this journey with, and I’m excited to see your future successes. In particular Chai Chee, Maleeha, and Dr Consuelo Santamaria Ferrada have been an immeasurable support and have kept me relatively sane throughout this project. I could have not asked for a better home-away-from-home.

Finally, I would of course like to thank my parents and sister. California is a long way away but, even from there, you were still able to offer sage advice. Your constant encouragement and support is the reason I was able to get where I am now.

Financial support This research was supported by the University of Queensland Research Training Tuition Fee Offset Scholarship.

Keywords General anaesthesia, Drosophila, larva, behaviour, chemotaxis, anaesthetic, sedative, propofol, isoflurane, gaboxadol

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Australian and New Zealand Standard Research Classifications (ANZSRC) ANZSRC code: 110903, Central Nervous System, 40% ANZSRC code: 110301, Anaesthesiology, 30% ANZSRC code: 060801, Animal Behaviour, 30%

Fields of Research (FoR) Classification FoR code: 1109, Neurosciences, 100%

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Table of Contents

Abstract ...... i Declaration by author ...... ii Publications included in this thesis ...... iii Submitted manuscripts included in this thesis ...... iv Other publications during candidature ...... iv Contributions by others to the thesis ...... iv Statement of parts of the thesis submitted to qualify for the award of another degree ...... v Research Involving Human or Animal Subjects ...... v Acknowledgements ...... vi Financial support ...... vii Keywords ...... vii Australian and New Zealand Standard Research Classifications (ANZSRC) ...... viii Fields of Research (FoR) Classification ...... viii Table of Contents ...... ix List of Figures ...... xii List of Abbreviations (LoA) ...... xiii Chapter 1: Introduction ...... 1 1.1 A brief history of general anaesthesia ...... 1 1.2 General anaesthetics activate GABAergic sleep circuits ...... 4 1.3 General anaesthetics target presynaptic release mechanisms...... 6 1.4 Two-step hypothesis of general anaesthesia...... 8 1.5 Drosophila, sleep, and general anaesthetics ...... 9 1.6 Propofol ...... 13 1.7 Thesis outline ...... 15 Chapter 2: Designing an assay to measure larval Drosophila behavioural response to the general anaesthetic propofol ...... 16 2.1 Introduction ...... 16 2.2 Methods ...... 18 2.2.1 Fly stock maintenance ...... 18 2.2.2 Larval preparation ...... 18 2.2.3 Larval propofol and vehicle aversion assay ...... 18 2.2.4 Preparation of propofol and HL3 Ringer’s solution ...... 19 2.2.5 High performance liquid chromatography ...... 19 2.2.6 Larval mouth hook contraction assay ...... 20 2.2.7 Larval nociception assay ...... 20 2.2.8 Larval crawling speed assay ...... 20 ix

2.2.9 Larval chemotaxis assay ...... 20 2.2.10 Larval angle deviation ...... 21 2.2.11 Statistics ...... 21 2.3 Results...... 22 2.3.1 Larvae are averse to propofol...... 22 2.3.2 Propofol does not affect mouth hook contractions ...... 23 2.3.3 Propofol has minimal effect on larval nociception ...... 25 2.3.4 Propofol treatment affects crawling speed ...... 27 2.3.5 Larvae are attracted to methanol ...... 27 2.3.6 Larval chemotaxis is adversely affected by DMSO ...... 28 2.3.7 Propofol abolishes larval chemotaxis to methanol ...... 30 2.3.8 Larval chemotaxis recovers from propofol exposure ...... 30 2.3.9 Larvae become disoriented after propofol treatment ...... 32 2.3.10 Immersion in 10mM propofol leads to a relevant dose in larval tissue ...... 33 2.4 Discussion ...... 34 Chapter 3: Investigating pre- and postsynaptic targets for general anaesthetics using a chemotaxis assay ...... 37 3.1 Introduction ...... 37 3.2 Methods ...... 39 3.2.1 Fly stock maintenance and crosses ...... 39 3.2.2 Preparation of gaboxadol (THIP), propofol, and HL3 Ringer’s solution ...... 39 3.2.3 Larval crawling speed after isoflurane treatment ...... 39 3.2.4 Brain dissection and immunohistochemistry...... 40 3.2.5 Statistics ...... 40 3.3 Results...... 41 3.3.1 Knockdown of the RDL subunit does not confer resistance to propofol ...... 41 3.3.2 The sedative gaboxadol does not affect larval chemotaxis...... 42 3.3.3 Sytaxin227 and propofol inhibition of larval chemotaxis ...... 44 3.3.4 Comparing the effect of syntaxin227 and wild-type syntaxin1A overexpression on larval propofol sensitivity ...... 46 3.3.5 SyxH3-N protects against propofol inhibition of larval chemotaxis ...... 48 3.3.6 Isoflurane reduces larval crawling speed ...... 49 3.3.7 SyntaxinH3-N confers resistance to isoflurane ...... 50 3.4 Discussion ...... 52 Chapter 4: Developing a technique for imaging larval central nervous system activity ...... 55 4.1 Introduction ...... 55 4.2 Methods ...... 56 x

4.2.1 Larval CNS preparation ...... 56 4.2.2 Imaging ...... 56 4.2.3 Fluorescence analysis ...... 57 4.3 Results...... 57 4.3.1 A filleted larval preparation for two-photon imaging ...... 57 4.3.2 Imaging the larval brain under gaboxadol ...... 59 4.4 Discussion ...... 61 Chapter 5: Conclusions and future directions ...... 63 List of References ...... 68

List of Figures

Figure 1.1: The Meyer-Overton correlation ...... 2 Figure 1.2: General anaesthetics are a structurally diverse group of drugs ...... 3 Figure 1.3: EEG patterns during an awake state, general anaesthesia, and sleep ...... 5 Figure 1.4: Components of the SNARE complex ...... 7 Figure 1.5: Sleep/wake circuits in the human and fly brain ...... 10 Figure 1.6: Life cycle of Drosophila ...... 11

Figure 1.7: Structure of the GABAA receptor showing binding sites of key agonists ...... 14 Figure 2.1: Larvae are averse to propofol ...... 22 Figure 2.2: Propofol does not inhibit mouth hook contractions ...... 24 Figure 2.3: Larval response to mechanical stimuli after propofol treatment ...... 26 Figure 2.4: Propofol reduces larval crawling speed ...... 27 Figure 2.5: Larvae positively chemotax to a methanol target, but this behaviour is affected by DMSO ...... 29 Figure 2.6: Propofol abolishes larval chemotaxis to methanol ...... 31 Figure 2.7: Propofol disorients larval chemotaxis ...... 32 Figure 2.8: Using HPLC to determine propofol concentration in larval tissue ...... 33 Figure 3.1: The structure of syntaxin1A and its role in the SNARE complex ...... 38 Figure 3.2: RDL knockdown does not confer propofol resistance ...... 42 Figure 3.3: The sedative gaboxadol does not affect chemotaxis ...... 43 Figure 3.4: Effect of syntaxin227 on larval propofol sensitivity...... 44 Figure 3.5: Comparing the effect of syntaxin227 and syntaxin1A overexpression on larval propofol sensitivity ...... 46 Figure 3.6: SyntaxinH3-N effects larval propofol sensitivity ...... 49 Figure 3.7: Larval sensitivity to isoflurane ...... 51 Figure 4.1: Setup for calcium imaging in the larval central nervous system ...... 58 Figure 4.2: Larval central nervous system response to gaboxadol ...... 60

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List of Abbreviations (LoA)

BP band-pass CB central brain CaM calmodulin CNS central nervous system CS Canton-S dFB dorsal fan-shaped bodies DMSO dimethyl sulfoxide EEG electroencephalogram fMRI functional magnetic resonance imaging GABA γ-aminobutyric acid

GABAA γ-aminobutyric acid receptor type A GFP green fluorescent protein GRD GABA and glycine-like receptor of Drosophila HA histaminergic LC locus coreleus LCCH3 ligand-gated chloride channel homolog 3 LoA list of abbreviations LP long-pass MB mushroom bodies NA noradrenergic PBS Phosphate-buffered saline PMT photomultiplier tube PPL1 potocerebral posterior lateral 1 qPCR quantitative polymerase chain reaction RDL resistance to dieldrin REM rapid eye movement RNAi ribonucleic acid RNAi RNA interference ROI region of interest SEM standard error of the mean SNAP25 synaptosomal-associated protein of 25kDa SNARE soluble N-ethylmaleimide factor attachment protein receptor THIP 4,5,6,7-tetrahydroisoxazolo[5,4-c]pyridin-3-ol

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TM transmembrane TMN tuberomammillary nucleus UAS upstream activator VLPO ventrolateral preoptic nucleus VNC ventral nerve chord

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Chapter 1: Introduction

1.1 A brief history of general anaesthesia

In 1811, the English novelist Fanny Burney wrote a letter to her sister relating an unflinching account of her recent mastectomy:

“When the dreadful steel was plunged into the breast—cutting through veins—arteries— flesh—nerves—I needed no injunctions not to restrain my cries. I began a scream that lasted unintermittingly during the whole time of the incision—and I almost marvel that it rings not in my ears still!” (Burney, 1811).

Ms. Burney was one of the lucky ones; she survived her operation. But as her letter attests, surgery was an excruciating affair before the advent of modern anaesthesia. The anatomist John Hunter went so far as to describe surgery as “a humiliation of the futility of science” and denounced the surgeon as no better than “a savage armed with a knife” (Jacobsohn, 1994).

Humans have searched for a suitable way to dull the pain of surgery since the beginning of medicine itself. Early attempts often involved prescribing draughts containing anything from marijuana to opium, or simply inebriating the patient (Caton and Antognini, 2003). A particularly inventive method from the 18th century involved knocking the patient out with a sudden blow to the jaw (Bishop, 1960). Like many other fields of medicine, superstition hindered the development of effective anaesthetics. In the West, many doctors believed pain to be an important stimulus that kept the patient alive during surgery. In China, the 3rd century physician Hua Tuo concocted an herbal infusion known as máfèisàn, which may have been a general anaesthetic or powerful analgesic. However, much of his work was destroyed, and Confucian prohibitions against surgery halted further development of anaesthetics in East Asia (Veith, 1966). Nevertheless, possibly the first reliably documented surgery to be performed using general anaesthesia took place in 1804, when the Japanese physician Hanaoka Seishū removed a tumour using Dutch-imported surgery techniques and a recreation of máfèisàn (Izuo, 2004). However, the advent of modern anaesthetics did not come until the mid-19th century. On 16 October, 1846, the American dentist William Morton demonstrated the use of ether as a general anaesthetic during a tooth extraction performed in front of a crowd of doctors and medical students (Caton and Antognini, 2003). Since Morton’s ether demonstration, general anaesthetics have become

common place. But despite nearly two centuries of routine clinical use, the mechanisms through which general anaesthetics render patients temporarily unconscious and unresponsive are still not well understood.

Within months of Morton’s demonstration, scientists began to speculate on the mechanism underlying general anaesthesia. Physicians noticed that patients being administered a general anaesthetic would present an orderly sequence of clinical signs beginning with a disturbance in consciousness, then a loss of reflexes, and finally paralysis of the cardiac and respiratory systems (Caton and Antognini, 2003). This led French physiologist Pierre Flourens to theorise that some parts of the nervous system were more susceptible to general anaesthetics than others (Caton and Atognini, 2003; Flourens, 1847). However, establishing the site of action of anaesthetics proved more difficult. By the mid-19th century, it was widely accepted that anaesthetics acted primarily on the brain rather than the peripheral nerves. French physician Claude Bernard, however, wanted to find a cellular mechanism that could explain general anaesthetics’ physiological effects. He conducted a series of experiments and found that anaesthetics could affect many types of cells ranging from muscles to plants. He thus hypothesised that general anaesthesia resulted from the reversible coagulation of proteins in nerves and muscles (Leake, 1978). But the most enduring theory of general anaesthesia came at the end of the 19th century

Figure 1.1: The Meyer-Overton correlation The lipid solubility (quantified here as the oil-gas partition coefficient) of an anaesthetic is correlated with its potency. Only inhalational anaesthetics are shown, and potency is represented by the anaesthetic partial pressure (measured in atmospheres) required to prevent movement in response to a surgical incision.

Reproduced with permission from Campagna, J.A., Miller, K.W., and Forman, S.A. (2003). Mechanisms of actions of inhaled anesthetics. The New England Journal of Medicine 348, 2110-2124, Copyright Massachusetts Medical Society. 2

when Hans Meyer and Ernest Overton independently observed that the potency of a general anaesthetic is directly proportional to its lipophilicity (figure 1.1) (Sandberg and Miller, 2003). This correlation led to the development of many lipid-based theories which speculated that general anaesthetics disrupt the lipid bilayer of cells (Ueda et al., 1986; Mori et al., 1984; Cantor, 1997), an idea that persisted until the end of the 20th century. While these theories have been proven incorrect, many of our current hypotheses of general anaesthetic mechanisms have a basis in these early ideas. With a few caveats, the Meyer-Overton correlation holds mostly true to this day, and Bernard’s ideas about coagulating proteins bears a similarity to emerging ideas about general anaesthetics’ interaction with membrane-bound proteins.

Since Morton’s ether demonstration, a diverse array of molecules have been identified as general anaesthetics. These can either be gases such as isoflurane and xenon or liquids such as propofol and etomidate (figure 1.2). Despite the diversity of molecular structures,

Figure 1.2: General anaesthetics are a structurally diverse group of drugs all anaesthetics are lipophilic and (with a few exceptions) their anaesthetic potency is well aligned with their oil solubility as the Meyer-Overton correlation predicted (Sandberg and Miller, 2003). Any molecular theory of anaesthetics must take this correlation into account.

Most current theories accept that general anaesthetics interact with lipophilic regions in proteins to disrupt their function. This was first demonstrated in the 1980s when Franks and Lieb discovered that anaesthetics can reversibly inhibit firefly luciferase (Franks and Lieb, 1984). Since then, anaesthetising drugs have been found to affect the function of a

variety of proteins such as kinesin (Bensel et al., 2017), cytochrome C (Slater et al., 1993), various ion channels (Franks, 2008), and presynaptic release elements (Xie et al., 2013).

1.2 General anaesthetics activate GABAergic sleep circuits

Almost all general anaesthetics interact with γ-aminobutyric acid type A (GABAA) receptors. Exceptions are small, aprotic general anaesthetics such as xenon (de Sousa et al., 2000), cyclopropane, and butane (Raines et al., 2001). The GABAA receptor belongs to a superfamily of cys-loop ligand-gated ion channels and its structure consists of five subunits surrounding a central pore. There are nineteen possible GABAA subunits: α1 – 6,

β1 – 3, γ1 – 3, δ, ε, θ, π, and ρ1 – 3. As its name suggests, GABAA receptors are activated by γ-aminobutyric acid (GABA), the major inhibitory neurotransmitter in the central nervous system (CNS) (figure 1.7). Upon activation, GABAA receptors allow for the influx of chloride ions into the neuron. This results in the hyperpolarisation of the neuronal membrane and effectively inhibits neurotransmission (Garcia et al., 2010). In mammals,

GABAA receptors are widely expressed throughout the CNS (Laurie et al., 1992) and (in the adult brain) are involved in inhibition of arousal pathways and potentiation of sleep circuits (Sherin et al., 1998).

The maintenance of sleep is an active process that requires continuous inhibition of arousal pathways. One important neuronal population for promoting sleep is the GABAergic neurons of the ventrolateral preoptic area, a part of the hypothalamus. These neurons release GABA and galanin and thereby inhibit many arousal nuclei such as the tuberomammillary nucleus (Sherin et al., 1998). Sleep and general anaesthesia have similar characteristics (namely an increased arousal threshold, immobility, and loss of consciousness) and it is therefore unsurprising that anaesthetics have been found to act on the same mechanisms that promote sleep. Research in rodents has demonstrated that general anaesthetics inhibit wake-promoting neurons in the tuberomammillary nucleus and that the sedative component of general anaesthesia may be mediated by GABAA receptors in this endogenous sleep pathway (Nelson et al., 2002). There are also brain- wide similarities between general anaesthesia and sleep. A variety of imaging studies on humans have demonstrated similarities between the brain during non-rapid eye movement sleep and the brain under general anaesthesia (figure 1.3) (Franks, 2008). Observations that sleep deprivation can promote the efficacy of general anaesthetics and that patients addicted to arousal-inducing drugs (such as amphetamines) are more resistant to general

anaesthesia (Hernandez et al., 2005) give credence to the idea that sleep and general anaesthesia may have shared mechanisms. The sleep-potentiating model is further validated by studies in animals that possibly do not sleep. For example, in the nematode Caenorhabditis elegans, which probably does not sleep during its adult stage, certain behaviours (such as mobility) are more resistant to volatile anaesthetics compared to mammals (Crowder et al., 1996). This is consistent with the idea that GABAergic sleep- promoting circuits are important targets for general anaesthetics in animals that do sleep.

While general anaesthesia is similar to sleep in many ways, it is clearly a different form of sedation. Unlike sleep, consciousness, pain response, and motor functions are mostly lost during general anaesthesia (Song et al., 2018). Furthermore, electroencephalogram (EEG) data can reveal differences in brain activity between sleep and general anaesthesia. At low doses of general anaesthetic, the thalamus emits tonic bursts which are similar to those

Figure 1.3: EEG patterns during an awake state, general anaesthesia, and sleep (A) EEG pattern of an awake patient with their eyes open (left) and with their eyes closed (right). (B) EEG patterns during general anaesthesia have several phases depending on the depth of anaesthesia. Paradoxical excitation is followed by phases 1 and 2, then burst suppression, and finally an isoelectric, coma like state. (C) EEG patterns during stages of sleep: rapid eye movement (REM) sleep stage 1 non-REM sleep, stage 2 non-REM sleep, and stage 3 non-REM sleep.

Reproduced with permission from Brown, E.N., Lydic, R., and Schiff, N.D. 2010. General Anaesthesia, sleep, and coma. N Engl J Med. 363(27): 2638–2650., Copyright Massachusetts Medical Society. 5

found in slow wave sleep. But as the concentration of general anaesthetic is increased, these bursts become suppressed. These more widely separated burst patterns found in deeply anaesthetised patients bears more resemblance to the high-amplitude, low- frequency activity observed in comatose patients rather than sleep. Finally, at the most profound state of general anaesthesia, the EEG is isoelectric (figure 1.3) (Brown et al., 2010). Emergence from general anaesthesia is also characterised by delayed recovery of alertness, unlike the quick recovery after waking up from sleep (Song et al., 2018). These differences indicate there are other targets for general anaesthetics besides GABAergic sleep circuits.

1.3 General anaesthetics target presynaptic release mechanisms

While postsynaptic receptors embedded in sleep/wake circuits are undoubtedly a target for many general anaesthetics, this may not completely explain how general anaesthetics abolish behavioural responsiveness as observed under surgical concentrations. An alternative target might be presynaptic. There are a growing number of studies which have shown that clinically relevant concentrations of general anaesthetics also affect presynaptic proteins involved in synaptic release (Xie et al., 2013; Baumgart et al., 2015; Bademosi et al., 2018).

All neurons communicate via release of neurotransmitters which are held in vesicles in the axon terminal. Vesicle docking to the plasma membrane is facilitated by the Soluble NSF Attachment Protein Receptor (SNARE) complex, a group of proteins that is highly conserved across most eukaryotes (Kloepper et al., 2007). In response to calcium influx, the plasma membrane-bound protein syntaxin1A interacts with the vesicle-bound protein synaptobrevin as well as SNAP25 to form a four stranded helical bundle known as the ternary complex (figure 1.4). This complex acts as a scaffold and pulls the vesicular membrane and plasma membrane together until they fuse, thereby releasing the vesicle’s contents into the synaptic cleft (Wu et al., 1999; Poirier et al., 1998).

One of the first clues that there may be a presynaptic mechanism to general anaesthesia came from the purportedly-sleepless C. elegans. GABA receptors in C. elegans are resistant to drugs that are typically GABAergic in humans, including the general anaesthetic propofol (Bamber et al., 2003), and yet C. elegans is still immobilised by high concentrations of general anaesthetics (Crowder et al., 1996). Without this postsynaptic effect, other potential targets could become apparent through behavioural screens of 6

Figure 1.4: Components of the SNARE complex The SNARE complex facilitates vesicle fusion to the plasma membrane in a calcium- dependent manner. The core of the SNARE complex consists of four α-helices (one each from syntaxin and synaptobrevin, and two from SNAP25). Synaptobrevin has a transmembrane domain embedded in the vesicular membrane, while syntaxin has a transmembrane domain embedded in the plasma membrane. Synaptotagmin has two calcium-binding domains which interact with SNAP25. In response to an influx of calcium, the SNARE complex pulls the vesicular membrane and plasma membrane together until they fuse, thus releasing neurotransmitters into the synaptic cleft. mutants. Indeed, one such C. elegans mutant, md130, was found to be resistant to volatile anaesthetics. This mutant has a point mutation in unc-64, the gene that codes for neuronal syntaxin1A, a core protein in the SNARE complex (van Swinderen et al., 1999). Other mutations of SNARE complex protein such as reduction-of-function mutants of SNAP25 and synaptobrevin, as well as other syntaxin1A mutants were found to be hypersensitive to volatile anaesthetics (van Swinderen et al., 1999). This clearly demonstrated that genetic manipulations of presynaptic release elements could change an organism’s sensitivity to a general anaesthetics. However, it did not prove that general anaesthetics were interacting directly with SNARE proteins. It could be that these mutations mitigate the effects of general anaesthetics through other means, such as interfering with other anaesthetic targets. Subsequent studies however, seemed to demonstrate a more direct targeting of SNARE proteins by general anaesthetics. Inhibition of synaptic release machinery has been found for the inhalational anaesthetic isoflurane (Herring et al., 2009) as well as intravenous anaesthetics such as propofol and etomidate (Herring et al., 2011). 7

Notably, molecules with similar structure and lipophilicity but which are not general anaesthetics do not inhibit neurotransmitter release (Herring et al., 2009; Herring et al., 2011). While many general anaesthetics cause inhibition of synaptic release, their exact targets within the SNARE complex may differ. For instance, studies in rat pheochromocytoma cell (PC12) culture have demonstrated that knockdown of synaptotagmin1 prevents etomidate from inhibiting neurotransmitter release, but only has a small effect on the potency of isoflurane and propofol (Xie et al., 2013). So while it is becoming clearer that anaesthetics may interact with a variety SNARE complex proteins, it is still not clear how this interaction takes place. It is, for example, possible that synaptotagmin1 and SNAP25 form a hydrophobic pocket for binding of anaesthetics (Xie et al., 2013). Alternatively, since general anaesthetics are lipophilic, they may be interfering with the plasma membrane itself, thereby changing the lipid environment in which key SNARE proteins are embedded, resulting in the inhibition of SNARE complex formation (van Swinderen and Kottler, 2014). This view is supported by a recent study which found that propofol and etomidate restrict syntaxin1A mobility in the plasma membrane, trapping syntaxin1A in nanoclusters (Bademosi et al., 2018). The potential importance of hydrophobic pockets in proteins and movement of proteins in the lipid membrane could provide an elegant explanation for Meyer and Overton’s observation that the strength of a general anaesthetic is correlated with its lipid solubility. This presynaptic mechanism provides a possible explanation for the effect that general anaesthetics have on all animals, including those that do not sleep.

1.4 Two-step hypothesis of general anaesthesia

There now seems to be two hypotheses for the mechanism of general anaesthesia: one presynaptic and one postsynaptic. However, a view was recently put forward as a way to reconcile the presynaptic targets and sleep pathway targets of anaesthetics. Van Swinderen and Kottler hypothesised that general anaesthesia may be best characterised as a two-step process. At low doses, general anaesthetics potentiate sleep-promoting circuits, thereby producing unconsciousness, while at the higher doses necessary for surgery, general anaesthetics inhibit presynaptic release machinery brain-wide, thereby causing a total loss of behavioural responsiveness. The inhibition of synaptic processes that this hypothesis proposes could explain the slow recovery from general anaesthesia

compared to sleep. This slow recovery might be due to the general anaesthetic causing a loss of functional connectivity brain wide (van Swinderen and Kottler, 2014).

1.5 Drosophila, sleep, and general anaesthetics

The common fruit fly (Drosophila melanogaster) is a particularly useful model organism due to its fast life cycle, easy maintenance, and well-established array of genetic tools. Drosophila has already been used to probe for anaesthetic targets (Leibovitch et al., 1995; Zalucki et al., 2015b; Troup, 2018) and its flexibility as a model has allowed for anaesthetics to be studied at many different levels from the synapse (Bademosi et al., 2018) to the brain (Cohen et al., 2016).

The discovery that adult flies sleep (Hendricks et al., 2000; Shaw et al., 2000) was particularly important to the study of general anaesthesia in flies since endogenous sleep circuits have been demonstrated to be an important general anaesthetic target in the mammalian brain. It has long been known that Drosophila have a circadian rest-activity cycle. However, it was only within the past two decades that the restful state has been characterised as sleep (Shaw et al., 2000). In mammals, a period of inactivity must meet certain criteria to be considered sleep including consolidated circadian periods of immobility, stereotypical sleep posture or resting place, elevation of the arousal threshold, and state reversibility with stimulation (Campbell and Tobler, 1984). The restful state observed in adult Drosophila meets these criteria. Adult flies have been shown to rest more frequently during certain periods within the day. Furthermore, during rest, adult flies assume a supported posture and are largely immobile except for infrequent sporadic movements of the proboscis and caudal abdominal twitches (Hendricks et al., 2000).

Supporting the behavioural evidence for sleep, specific neurons and brain structures have been found that modulate sleep in the adult Drosophila brain. Activation of the dorsal fan- shaped body has been found to promote sleep in adult Drosophila (figure 1.5A) (Donlea et al., 2011) and dopaminergic neurons which innervate the dorsal fan-shaped body have been found to mediate arousal (figure 1.5C) (Ueno et al., 2012; Liu et al., 2012). Furthermore, analogues of mammalian GABAergic sleep circuits have been found in adult Drosophila (Parisky et al., 2008; Agosto et al., 2008; Chung et al., 2009). The GABA receptors in Drosophila, however, differ slightly from mammalian GABA receptors. As noted above, mammals have five major types of subunits (α, β, γ, δ, and ρ). In contrast, Drosophila only has three subunit types: RDL (Resistant to dieldrin) (Ffrench-Constant et al., 1993), GRD (Harvey et al., 1994), and LCCH3 (Henderson et al., 1993).

Figure 1.5: Sleep/wake circuits in the human and fly brain (A) In the human brain, noradrenergic (NA) and histaminergic (HA) neurons from the locus coreleus (LC) as well as the tuberomammillary nucleus (TMN) are wake- promoting (red lines). (B) GABAergic neurons originating from the ventrolateral preoptic nucleus (VLPO) (blue lines) inhibit the arousal centres of the brain to promote sleep. (C) In the fly brain, dopaminergic, wake-promoting neurons originating from the protocerebral posterior lateral 1 (PPL1) cluster (red) innervate the mushroom bodies (MB) and the dorsal fan-shaped bodies (dFB). (D) In the fly brain, activation of the dFB promotes sleep. It is thought that general anaesthetics like isoflurane promote sleep circuits in both human and fly brains.

Reprinted from BioEssays 36(4), van Swinderen, B. and Kottler, B. 2014. Explaining general anesthesia: A two-step hypothesis linking sleep circuits and the synaptic release machinery. BioEssays 36(4): 372-381., with permission from John Wiley and Sons.

Homomultimeric RDL receptors are the most common GABAA receptor arrangement in Drosophila and are expressed brain-wide in both adult and larval flies (Enell et al., 2007). Despite the difference in subunit composition, homomultimeric RDL receptors in

Drosophila play a role in sleep onset and duration similar to the mammalian GABAA receptor. RDL is expressed in the large ventral lateral neurons which are part of the circadian clock, and manipulation of RDL expression in these neurons changes sleep onset and duration (Parisky et al., 2008). Overexpression causes flies to fall asleep faster and sleep longer, while partial knockdown with RNAi leads to a small, but significant decrease in sleep time (Parisky et al., 2008).

Figure 1.6: Life cycle of Drosophila The common fruit fly (Drosophila melanogaster) undergoes holometabolan development consisting of four life stages: egg (embryo), larva, pupa, and imago (adult). The larva goes through three stages between moults known as instars. After the third instar, the larva pupates and its organs degenerate and restructure into their adult forms. Development time can depend on temperature or health of the fly.

Image: Weigmann et al., 2003, FlyMove

In parallel with mammalian studies, general anaesthetics have been found to act on sleep/wake circuits in adult Drosophila. There is a strong correlation between Drosophila strains with short sleep phenotypes and resistance to sevoflurane and isoflurane (Weber et al., 2009). Furthermore, activation of dopaminergic and octopaminergic wake-promoting neurons confers resistance to the volatile anaesthetic isoflurane. Conversely, activation of sleep-promoting neurons in the dorsal fan-shaped body results in hypersensitivity to isoflurane (Kottler et al., 2013). Susceptibility can also be modulated with arousal-inducing drugs. For example, methamphetamine fed flies are resistant to isoflurane (Kottler et al., 2013). The similarities in the influence of general anaesthetics on Drosophila and mammalian sleep circuits emphasise that these circuits are an important target for general anaesthetics in all animals that sleep.

While sleep has been fairly well characterised in adult Drosophila, there is comparatively scant evidence that larval Drosophila sleep. Larvae do express RDL receptors brain-wide (Enell et al., 2007) and display GABAergic plasticity at least in short-term olfactory

habituation (Larkin et al., 2010), but there is no clear evidence for RDL involvement in a larval sleep circuit. However, larval flies do have a circadian rhythm (Koštál et al., 2009), and a recent study argued that Drosophila larvae do have a sleep-like state required for development (Szuperak et al., 2018). While this state has some behavioural characteristics which resemble sleep, the underlying neural circuits that produce this behaviour seem to be different than the sleep circuitry of adult Drosophila. For example, the neurotransmitters dopamine and octopamine modulate arousal in adult flies (Andretic et al., 2005; Crocker et al., 2010). In contrast, while both dopaminergic and octopaminergic neurons are present in larvae, only octopamine modulates arousal in larvae (Szuperak et al., 2018). Furthermore, the circuits of the dorsal fan-shaped body (which are sleep promoting in the adult brain) do not fully develop until during the pupal stage (figure 1.5) (Riebli et al., 2013; van Swinderen and Kottler 2014). Despite this possible lack of sleep circuitry, larvae can still be immobilised with volatile anaesthetics such as isoflurane and halothane (Zalucki et al., 2015b).

In this regard, larval Drosophila could be compared to C. elegans, another animal potentially without GABAergic sleep circuits. Like Drosophila larvae, a sleep-like state has been proposed for C. elegans (Raizen et al., 2008). This sleep-like state happens just after moulting during a period of quiescence called lethargus (Raizen et al., 2008), and is associated with GABAergic synaptic plasticity (Dabbish and Raizen, 2011). It may be that lethargus is a developmental stage that accomplishes some sleep functions, but does not involve sleep-promoting mechanisms analogous to what happens in other animal brains. However, these periods of quiescence have not been observed in the adult stage and a GABAergic sleep pathway has not been identified. If general anaesthetics only acted through sleep-promoting pathways, it would stand to reason that C. elegans should be insensitive to general anaesthetics. But this is not the case. As discussed previously, C. elegans can be immobilised at high concentrations of volatile anaesthetics (Crowder et al., 1996). This suggests that general anaesthetics have other targets in C. elegans that are only activated at higher concentrations.

Rather than promoting sleep pathways, the effect of general anaesthetics on C. elegans and Drosophila larvae may be due to the inhibition of presynaptic release mechanisms. Larval Drosophila may therefore be particularly well situated for investigating the presynaptic effects of general anaesthetics. The absence of endogenous sleep circuits may expose presynaptic targets for anaesthetics. Furthermore, genetic manipulations of these presynaptic targets may yield more obvious behavioural phenotypes that would

otherwise be obscured due to the effects of a more sensitive GABAergic target. Drosophila larvae have been previously used to study volatile anaesthetics (Zalucki et al., 2015b), but have been left unexplored as a model for intravenous anaesthetics such as propofol.

1.6 Propofol

Propofol (2,6-diisoprophylphenol) is a dialkylphenol and is a sedative-hypnotic agent used for the induction of general anaesthesia (figure 1.2) (Trapani et al., 2000). It is used in a variety of clinical settings because it causes rapid loss of consciousness and has a short recovery period that is relatively smooth compared to other general anaesthetics. It is an oil at room temperature and, like other general anaesthetics, is insoluble in water. Doses of propofol are delivered as an oil in water emulsion which is administered either as a continuous infusion (6-12 mg/kg/hour) or as intermittent bolus injections (20-50mg). The dose of propofol required to induce anaesthesia in adult humans is between 2 and 2.5 mg/kg (Kotani et al., 2008). The blood propofol EC50 in adult humans is between 1.07 and 0.95 μg/mL and has a half-life of 116 +/- 34 min (Shafer et al., 1988).

Propofol is both a cardiovascular and respiratory depressant, and anaesthetic induction with propofol often results in hypotension (Kotani et al., 2008). Propofol-induced hypotension is mediated by an impairment of baroreflex regulatory mechanisms and the sympathetic nervous system (Ebert et al., 1992). In rare cases, propofol can cause propofol infusion syndrome in critically ill patients. The main features of this syndrome include cardiac failure, rhabdomyolysis, and renal failure (Vasile et al., 2003). Less serious side effects include pain upon injection likely due to direct interactions with the mustard-oil receptor TRPA1 channels (Matta et al., 2007).

Propofol is typically thought of as a GABA-acting general anaesthetic. Like most other general anaesthetics, propofol potentiates GABAA receptor activity and can directly activate chloride currents at high concentrations. While propofol can act on α, β and γ subunits of the GABAA receptor, the β subunit is thought to contribute the most to this interaction (figure 1.7) (Garcia et al., 2010). The efficacy of the interaction is dependent on the arrangement of propofol’s functional groups, with the hydroxyl group being particularly integral. Studies using propofol analogues have identified three key ligand-receptor interaction sites. The most important site involves the formation of a hydrogen bond with propofol’s hydroxyl group while the other two sites are hydrophobic pockets (figure 1.7)

(Krasowski et al., 2001). Given propofol’s affinity to GABAA receptors, it is unsurprising 13

that it is often considered to be the quintessential sleep-promoting anaesthetic. Propofol- induced loss of consciousness is characterised by the appearance of EEG slow waves which bear a resemblance to the slow waves during non-REM sleep (figure 1.3) (Murphy et al., 2011). Studies in rodents have found that propofol inhibits wake-promoting neurons in the tuberomammillary nucleus (Nelson et al., 2002). Further studies in rats found that propofol anaesthesia can help in the recovery from sleep deprivation (Tung et al., 2004) but this has not been successfully replicated with human patients in a clinical setting (Lewis et al., 2018).

More recently, it has been found that propofol has a variety of interactions with SNARE complex proteins. Knockdown experiments in PC12 cells have confirmed that propofol interact with SNAP-25, SNAP-23, and synaptotagmin I (Xie et al., 2013). However, it is not clear how this interaction takes place. A more recent study found that propofol restricts syntaxin1A mobility on the plasma membrane and it could be that propofol blocks the initial steps of SNARE complex formation (Bademosi et al., 2018).

Figure 1.7: Structure of the GABAA receptor showing binding sites of key agonists

A schematic of the β-subunit (left) and an assembled GABAA receptor (right). The β- subunit consists of 4 transmembrane domains (TM1-4) and the extracellular domain contains a cysteine-linked loop, the defining characteristic of the cys-looped ligand- gated ion channel superfamily. The most common GABAA receptor configuration is ααββγ, shown to the right (Chang et al., 1996). GABA primarily binds at the interface between the α and β subunits, and directly activates chloride currents. Gaboxadol, a sedative, also binds to this site (Bergmann et al., 2013). An important propofol binding site is located at the interface between transmembrane domains 1 and 2 of the β- subunit (Yip et al., 2013)

1.7 Thesis outline

This thesis developed an assay to measure the effect of the anaesthetic propofol on third instar larvae of the common fruit fly, Drosophila melanogaster. These assays were then used to investigate pre- and postsynaptic targets of general anaesthesia.

Chapter 2 investigates several possible behavioural endpoints to assess the effects of propofol on Drosophila larvae. I first devised a delivery method for propofol which involved immersing larvae in a propofol-laced Ringer’s solution using dimethyl sulfoxide (DMSO) as a solvent. High performance liquid chromatography (HPLC) was used to measure the concentration of propofol in the larval tissue. I next identified and assayed a series of increasingly complex behavioural end points to assess the effects of propofol on larvae: mouth hook contractions, nociceptive response, crawling speed, and chemotaxis.

Chapter 3 employs the chemotaxis assay and crawling speed assay to explore a pre- and postsynaptic target of propofol. An RDL knockdown line was tested to investigate the postsynaptic target and two syntaxin1A mutants were generated to explore propofol’s putative presynaptic mechanism.

Chapter 4 develops methods for calcium imaging of the larval CNS during gaboxadol perfusion. This technique may be used in future experiments to assess the effects of propofol and other drugs on larval CNS function.

Chapter 5 provides a conclusion discussing the caveats of the results from the previous chapters as well as potential future directions.

Chapter 2: Designing an assay to measure larval Drosophila behavioural response to the general anaesthetic propofol

2.1 Introduction

In a clinical setting, general anaesthesia is thought of as a reversible drug-induced loss of consciousness. However, general anaesthetics can have many other effects on biological functions and behaviours which vary depending on the depth of anaesthesia. Therefore, general anaesthesia can be described as a state of decreased responsiveness which can be assessed using a variety of behavioural endpoints. For example, in humans, implicit memory is particularly susceptible to general anaesthetics (Chortkoff et al., 1993), while movement responses to noxious stimuli and consciousness are more resistant (Zalucki and van Swinderen, 2016).

Since general anaesthetics started being used in medicine, doctors have used animal models to assess the effect of general anaesthetics on behaviour. One of the earliest of these researchers was the English physician John Snow. In a series of experiments, he placed small animals (such as guinea pigs, mice, and cats) in enclosed chambers and observed their reactions to different concentrations of inspired anaesthetic (in this case, chloroform). From these experiments, he was among the first to find a quantifiable relationship between the concentration of inspired anaesthetic and behavioural response (Snow, 1858). Subsequent studies on a variety of animals revealed a curious feature of general anaesthetics: most animals appear to be sedated by general anaesthetics at similar drug concentrations (Zalucki and van Swinderen, 2016). While it is difficult to make behavioural comparisons across species, the general trend seems to be that more complex behaviours (which involve multiple neural pathways or behavioural sequences) are more susceptible to general anaesthetics than simpler ones. Accordingly, in the nematode worm (Caenorhabditis elegans), pharyngeal pumping (the contraction and relaxation of pharyngeal muscles during feeding) is more resistant to general anaesthetics than chemotaxis (Crowder et al., 1996) and in humans, pain response is more resistant than task learning (Chortkoff et al., 1993). These commonalities strongly suggest the presence of a conserved target for general anaesthetics across the animal kingdom. Furthermore, this opens up the possibility for using animal models to probe for potential general anaesthetic targets that have relevance to humans.

The common fruit fly (Drosophila melanogaster) is a particularly useful model organism due to its fast life cycle, easy maintenance, and well-established array of genetic tools. Behavioural end points in adult flies have already been used as a model to probe for general anaesthetic targets (Karunanithi et al., 2018; Campbell and Nash, 2001). However, the larval stage remains relatively unexplored. While there have been several studies on the effects of general anaesthetics on the larval neuromuscular junction (Sandstrom, 2004; Zalucki et al., 2015b; Bademosi et al., 2018), there is a paucity of studies examining the behavioural effects of general anaesthetics (particularly propofol).

Larval Drosophila could be a potentially useful system because of the differences between the larval and adult nervous system. The first clue to this possible difference is the discrepancy between adult and larval sleep behaviour. While Drosophila adults have been found to fulfil the criteria for sleep (Shaw et al., 2000; Hendricks et al., 2000), evidence for larval sleep remains contentious (Szuperak et al., 2018; Zalucki and van Swinderen, 2016). In fact, many of the key structures thought to be involved in the sleep/wake circuits (such as the dorsal fan-shaped body) do not fully develop until during the pupal stage (Riebli et al., 2013). This is particularly relevant to investigating general anaesthetic targets because general anaesthetics are thought to activate endogenous sleep pathways (Garcia et al., 2010). An interesting contrast to other model organisms is the case of C. elegans which, probably lacking sleep circuitry, is remarkably resistant to the effects of volatile general anaesthetics on mobility- they are never entirely sedated by these drugs (Zalucki and van Swinderen, 2016). However, using Drosophila as a model organism could take this one step further by offering the possibility of studying these different brains in the same animal, or at least the same genetic strain. If larvae do indeed lack sleep circuits, it could make other general anaesthetic processes more accessible for study because these would not be confused with purely sedative mechanisms of general anaesthesia.

In this chapter, I outline several possible behavioural endpoints to assess the effects of the general anaesthetic propofol (2,6-diisoprophylphenol) on Drosophila larvae (figure 2.1A). Propofol is the quintessential sleep-promoting general anaesthetic and sedation under propofol has been shown to share similar characteristics to sleep (Franks, 2008; Murphy et al., 2011) and can even help in the recovery of sleep deprivation (Tung et al., 2004). It is an oil at room temperature and is insoluble in water (Trapani et al., 2000). Due to this insolubility, I first devised a delivery method for propofol which involved immersing larvae in a propofol-laced Ringer’s solution using dimethyl sulfoxide (DMSO) as a solvent. I verified the efficacy of this method using high performance liquid chromatography (HPLC)

to determine the concentration of propofol in the larval tissue. I then explored a variety of behavioural assays to investigate different anaesthetic endpoints for third instar Drosophila larvae. A chemotaxis assay was determined to be most suitable for use in further studies. Interestingly, even though the concentration of propofol in the larval tissue was comparable to the level required for sedation in mammals, propofol did not sedate the larvae at any of the concentrations tested.

2.2 Methods

2.2.1 Fly stock maintenance

Wild type strain Canton-S (CS) fruit flies (Drosophila melanogaster) were housed in plastic bottles on standard yeast and agar media. To induce egg laying, flies were kept in an incubator at 25OC. After two days, adults were cleared out of the bottles and the larvae were left to develop in the incubator for an additional 5 days at 25OC.

2.2.2 Larval preparation

Third instar Drosophila larvae were removed from the food media by first scooping the media from the bottle with a spatula and then mixing with tap water. The media and water mixture was agitated with a small paintbrush, and the larvae were then transferred with the paintbrush to a petri dish containing water. The larvae were again agitated with a paintbrush to remove excess food. Wandering third instar larvae were excluded due to potential behavioural differences from feeding third instar larvae.

2.2.3 Larval propofol and vehicle aversion assay

3w/v% agar solution was microwaved until the agar was melted completely. To make 10mL of propofol-laced agar at each concentration, propofol (≥97% Sigma-Aldrich) was diluted in polyethylene glycol (average Mn 400, Sigma Aldrich) to make a 100mM propofol stock solution. The appropriate volume of this stock solution was then added to 9mL of 3% agar and topped up to 10mL with glycol. 100µL of blue food colour (food colour 133, Queen Fine Foods) was added per 10mL of agar to increase larval visibility. 3mL of propofol-laced agar was then pipetted into a 15mL falcon tube and left to cool. Using a paintbrush, third instar larvae were selected from the distilled water and placed on the agar 18

surface. Each larva’s distance from the agar surface was recorded every 20minutes for one hour (figure 2.1B).

2.2.4 Preparation of propofol and HL3 Ringer’s solution

A 560mM propofol stock solution was prepared in DMSO. The appropriate volume of propofol stock solution was diluted in haemolymph-like (HL3) Ringer’s solution to make media for assays. Controls contained an equivalent volume of DMSO. Minimum HL3 Ringer’s solution was made according to Stewart et al. 1994 (70mM NaCl, 5mM KCl,

1.5mM CaCl2 · 2H2O, 20mM MgCl26H2O, 10mM NaHCO3, 5mM Trehalose, 115mM sucrose, and 5mM HEPES; pH 7.2; Π=343 ± 0.7).

2.2.5 High performance liquid chromatography

The HPLC protocol used was modified from Gardner et al. 2016. Third instar Drosophila larvae were exposed to 10mM of propofol in HL3 Ringer’s solution for 2 hours. Groups of 15 larvae were analysed independently to determine the tissue concentration of propofol. Following propofol treatment, 15 larvae were rinsed in distilled water, dried, and weighed. The larvae were then homogenised in 400µL solution containing a 3:1 volume ratio of acetonitrile and 0.02M Na2H2PO4 buffer. The homogenised tissue was centrifuged at 10,000 x g for 20 minutes at 4OC. The supernatant was filtered using a syringe filter and 25µL of supernatant was injected into an Agilent 1100 series HPLC instrument (Agilent technologies Pty. Ltd, Santa Clara, USA).

Propofol standards were prepared in the same solution as the tissue. 0.001mM, 0.01mM, and 0.05mM propofol were used to create a standard curve. The chromatography system consisted of a 250mm x 4.6mm ODS C18 column, and a fluorescence detector (part # G1321A, Standard FLD cell, Germany). The mobile phase was an isocratic mixture of A:

2-propanol B: acetonitrile (0.1% TFA), C: 0.02 Na2H2PO4 buffer. The flow rate was 1.5 mL/minute and the fluorescence detector was set at an excitation of 276nm and an emission of 310nm. Retention time for propofol was 5.5 minutes (figure 2.8A).

2.2.6 Larval mouth hook contraction assay

After collection, larvae were immersed in different concentrations of propofol (0mM, 1mM, 5mM, and 10mM). Every 30 minutes, each larva was observed, and their behaviour was recorded as with mouth hook contractions or no mouth hook contractions (figure 2.2A).

2.2.7 Larval nociception assay

Larvae were immersed in 1mL HL3 Ringer’s solution with different concentrations of propofol for 2 hours. Following Hoyer et al. 2018, a stimulation tool was built using an 18mm length of fishing line (6 lb test, diameter 0.23 mm) or tungsten wire (50.8 mm diameter). The fishing line was taped to a toothpick so that 8 mm of the fibre protruded from the end of the toothpick. The filament was calibrated to exert a force of 45-50mN (4.59-5.1g) using a balance. The assay was performed on a 100 x 10mm plastic petri dish containing 20mL of 3% agar with a thin film of 2ml of dH2O on top to ease larval movement. The tool described above was used to deliver a 1 second mechanical stimulus at a 45o to 60o angle to the larva’s dorsolateral A3-A6 regions (figure 2.3A). After a 3 second pause, the stimulus was reapplied a second time. For each stimulus, the response of each larva was scored: no response, direction change, U-shape, or roll (figure 2.3B).

2.2.8 Larval crawling speed assay

Larvae were immersed in 1mL HL3 Ringer’s solution with different concentrations of propofol or DMSO control for 2 hours. The assay was performed on a 100 x 10mm plastic petri dish containing 20mL of 3% agar. Three larva were placed in the middle of the plate and filmed with a Logitech webcam for 5 minutes. A piece of tracing paper was taped over the monitor and as the video was played back, the position of each larva was marked in 5 second intervals. Distance was calibrated with a 1cm scale bar in frame. Larval crawling speed was estimated by measuring the distance each larva travelled over the 5 minute assay (figure 2.4A).

2.2.9 Larval chemotaxis assay

Larvae were immersed in 1mL HL3 Ringer’s solution with different concentrations of propofol for 2 hours. Larvae that pupated or crawled up the side of the petri dish were removed. The assay was performed on a 100 x 10mm plastic petri dish containing 20mL of 20

3% agar. A Teflon container containing 20µL of methanol was placed on one side of the dish and another Teflon container containing 20µL of distilled water was placed at the other. These Teflon containers were custom built and consisted of a 5mm diameter pot capped with a lid perforated with nine holes. A piece of paper marking the odour zones was placed under the dish. There were three zones: a 1cm neutral zone bisecting the plate, an odour zone, and a control zone (figure 2.5A). During each assay, 30 larvae were placed in the centre of the dish with a paintbrush. The cover was then placed on the dish and the larvae were allowed to migrate for 5 minutes. At the end of the assay, the number of larvae in each zone was counted and a preference index was calculated. Preference Index = (Odour Zone - Control Zone) / Total. Larvae that had crawled onto the lid were not counted.

2.2.10 Larval angle deviation

A larval chemotaxis assay was set up as described above except with only 1 larva per plate and filmed with a logitech webcam. A variety of tracking software were tested, but a manual method was found to be the most straightforward and most accurate. A piece of tracing paper was taped over the monitor and as the video was played back, the position of each larva was marked in 5 second intervals. From these positions, angle deviations for each 5 second time point was determined. Angle deviation refers to the angle between the larva’s velocity vector and a vector pointing from the velocity vector’s origin to the methanol odour container. An angle of 0º indicates a larva is moving directly toward the methanol container.

2.2.11 Statistics

Where relevant, data is presented as means and the error bars indicate ±SEM. For some points, the error bars are shorter than the height of the symbol and have therefore been left out. Statistical analysis was performed in Graphpad Sofware Prism Version 7.02. Circular statistical analysis for angle deviation was performed in Matlab R2016b using the circstats package. Significance was set at p<0.05.

2.3 Results

2.3.1 Larvae are averse to propofol

Adult flies will readily eat propofol-laced food and are sedated after feeding, so this is an effective means of anaesthetic delivery (Gardner et al., 2016). Gardner et al. 2016 demonstrated that feeding adult flies 1mM propofol results in a clinically relevant concentration of about 1µg/mg in adult fly heads (Gardner et al., 2016). In order to draw equivalences with adult studies, I attempted to feed larvae propofol-laced agar in a similar manner. However, when placed on agar laced with propofol, larvae rapidly migrated out of the agar. To quantify this negative gustatory response, wild-type larvae were placed on agar food containing varying concentrations of propofol at the end of a 15mL falcon tube. The tube was placed on its side and the distance of each larvae from the agar surface was

Figure 2.1: Larvae are averse to propofol (A) The structure of propofol (2,6-diisopropylphenol) consists of a central benzene ring with one hydroxyl group flanked by isopropanyl groups at the 2- and 6- positions. (B) Propofol aversion assay design. Larvae were placed on top of propofol-laced or glycol-laced agar and allowed to migrate for 1 hour. (C) No significant difference was observed in migration for glycol-laced agar. A repeated measures two-way ANOVA with a Bonferroni multiple comparisons between 0% vs 5% and 10% glycol concentrations all yielded p>0.9, n.s. (N=10 for each condition, error bars show SEM). (D) Wild-type larvae rapidly migrate out of propofol-laced agar. Repeated measures two-way ANOVA with Bonferroni multiple comparisons for wild type (each condition compared to 0mM control): 1mM: p=0.0022, 2mM: p<0.0001, and 10mM: p=0.0040 (N=10 for each condition, error bars show SEM.)

measured at 1 minute intervals (figure 2.1B). Larvae were not found to be averse to glycol, which was used as a vehicle for propofol (figure 2.1C). However, when immersed in propofol-laced agar, the larvae rapidly migrated out of the agar at all concentrations tested (figure 2.1D). In an attempt to mask the taste of propofol, larvae were also tested on agar made up with apple juice, yeast paste, and varying amounts of sucrose instead of distilled water (data not shown). None of these seemed to mitigate larval aversion to propofol. Feeding was therefore abandoned as a way to deliver propofol and an alternative method to deliver the drug to larvae was necessary. For subsequent assays, larvae were instead immersed in a propofol-laced HL3 Ringer’s solution for 2 hours. Because propofol is a highly lipophilic molecule, DMSO was used as an emulsifier to keep the propofol in solution.

2.3.2 Propofol does not affect mouth hook contractions

Larvae feed almost constantly until they reach the wandering stage just before pupation. During feeding, the mouth hooks extend and retract to bring food into the mouth. These mouth hook contractions persist even when the larvae are removed from their normal agar food and placed in liquid. As one of the simplest larval behaviours, mouth hook contraction was predicted to be fairly resistant to general anaesthesia. To test this, larvae were immersed in different concentrations of propofol (0mM, 1mM, 5mM, and 10mM), and their behaviour observed. There was no difference in mouth contractions between the propofol- treated and control groups even after 4 hours immersed in solution (figure 2.2B; Chi- square test (3, N=200) = 7.56, p=0.560). From observation, the rate of mouth hook contractions was perhaps slower in the 10mM propofol treated group, but this was not quantified. Nevertheless, larvae survived remarkably well during prolonged propofol treatment. Mouth hook contractions persisted even after 24hours of propofol immersion (data not shown). Even more surprisingly, after 24 hours, between 10% and 24% of larvae across all test concentrations began to pupate while still immersed in the propofol solution. Of the 200 larvae tested, only four died (two in the 1mM treated group and two in the 10mM treated group). The persistence of mouth hook contractions even at very high concentrations of propofol make it not useful as an anaesthetic behavioural endpoint (at

least at the concentrations tested). However, the persistence of mouth hook contractions and commencement of pupation demonstrate that propofol treatment does not significantly affect larval survival or development. This could possibly indicate that propofol has no major target in fly larvae.

Figure 2.2: Propofol does not inhibit mouth hook contractions (A) Larval feeding behaviour consists of rhythmic contractions of the cephalopharyngeal skeleton which drives the contractions of the mouth hooks. Reprinted from Journal of Insect Physiology, 55 /33, Schoofs A, Niederegger S, Spieß P, From behavior to fictive feeding: Anatomy, innervation and activation pattern of pharyngeal muscles of Calliphora vicina 3rd instar larvae, 218-230, 2009, with permission from Elsevier. (B) After 4 hours of continuous immersion in solution, there was no difference in mouth contractions between the propofol treated and control groups even after 4 hours (N=50 for each condition, Chi-square test (3, N=200) = 7.56, n.s. p=0.560).

2.3.3 Propofol has minimal effect on larval nociception

In the wild, parasitoid wasps of the genus Leptopilina are a predator of Drosophila larvae. However, the adult wasps do not feed on the larvae. Rather, the female wasp uses its ovipositor to pierce the larval cuticle and inject its eggs into the larva’s body. After hatching, the wasp larvae proceed to eat the Drosophila larva from the inside. This reproductive strategy is very effective for the wasp; up to 70% of wild Drosophila larvae may be infected with wasp eggs (Fleury et al., 2004). In response to this predation pressure, Drosophila larvae have evolved several stereotyped behaviours to evade the ovipositor of the wasp. When attacked, a larva will roll around its anterior-posterior axis in a corkscrew-like motion (figure 2.3B). In an incomplete nociceptive response, the larva may bend its body in a U-shape or change crawling direction (figure 2.3B). These behavioural responses can be induced with noxious thermal or mechanical stimulation and is triggered by multi-dendritic class IV neurons (Robertson et al., 2013). After being immersed in a Ringer’s solution containing propofol or DMSO control for 2 hours, larvae were mechanically stimulated with a tool made of fishing line or a tungsten metal filament that mimicked the ovipositor of a parasitoid wasp (Hoyer et al., 2018) (figure 2.3A). Each larva was stimulated twice (3 seconds apart) and their behavioural response was scored. For the larvae stimulated with a wire filament, the 10mM propofol treated group was less likely to respond to the first mechanical stimulation compared to controls (Chi-squared test (2, N=100) = 9.035, p=0.0109) (figure 2.3B). However, none of the other stimulations yielded significant differences between propofol treated groups and controls (fishing line, 1st stimulus: Chi-squared test (3, N=100) =2.822, p=0.4199; fishing line, 2nd stimulus: Chi-squared test (3, 1.463) =1.463, p=0.6909; wire, 2nd stimulation Chi-squared test (2, N=100) =5.38, p=0.0679). Nociceptive response seemed to be too simple of a behaviour to be affected by propofol (at least at the concentrations tested). I next sought to assay more complex behaviours which are presumed to be more susceptible to general anaesthetics. Following previous work in C. elegans, coordinated movement and chemotaxis should be more vulnerable to general anaesthetics (Crowder et al., 1996).

Figure 2.3: Larval response to mechanical stimuli after propofol treatment (A) Larvae were stimulated in the dorsolateral region of abdominal segments A3-A6 using a filament either made of nylon fishing line or a tungsten wire. (B) Larvae have several different nociceptive responses after mechanical stimulation.

Image reproduced with permission from Bio-protocol, Hoyer, N., Petersen, M., Tenedini, F. M., and Soba, P. 2018. Assaying mechanonociceptive behavior in Drosophila larvae. Bio-protocol 8(4): e2736.

(C) Nociception assay with either fishing line or wire stimulation. Each larva was stimulated twice (3 seconds apart) and their behavioural response was scored (n=50 for each condition) (fishing line, 1st stimulus: Chi-squared test (3, N=100)=2.822, p=0.4199; fishing line, 2nd stimulus: Chi-squared test (3, 1.463)=1.463, p=0.6909; wire, 1st stimulation: Chi-squared test (2, N=100)=9.035, p=0.0109; wire, 2nd stimulation Chi- squared test (2, N=100)=5.38, p=0.0679). 26

2.3.4 Propofol treatment affects crawling speed

I hypothesised that propofol would reduce the crawling speed of the larvae. This is because previous studies have shown that propofol affects quantal release from the larval motorneurons (Bademosi et al., 2018) and that volatile anaesthetics reduce larval mobility (Zalucki et al., 2015b). Larval crawling speed was estimated by measuring the distance each larva travelled in 5 second intervals (figure 2.4A). After 2hour exposure to 10mM propofol, crawling speed was significantly reduced (Welch’s t test, p=0.0018) but the larvae were still mobile (figure 2.4B).

Figure 2.4: Propofol reduces larval crawling speed (A) Larvae were first exposed to propofol for 2hours and then crawling speed was estimated by measuring the distance each larva travelled in 5 second intervals. (B) Effect of 2hour exposure to 10mM propofol exposure on crawling speed is significant (Welch’s t test, * p=0.0018).

2.3.5 Larvae are attracted to methanol

It has been well demonstrated that general anaesthetics impair chemotaxis in C. elegans (Crowder et al., 1996). In C. elegans, chemotaxis is more vulnerable to general anaesthetics than behaviours such as pharyngeal pumping, egg laying, or coordinated movement (Crowder et al., 1996). It was predicted that Drosophila larval chemotaxis would also be more sensitive to general anaesthetics, as it is a complex behavioural endpoint that involves a level of decision making. A chemotaxis assay adapted from Apostolopoulou et al. 2013 was developed to measure larval olfactory response. A 10cm petri dish was coated in 3% agar and then two Teflon containers, one with 20µL of methanol and the other with 20µL of distilled water, were placed on either side of the dish (figure 2.5A). 30 larvae were placed at the centre of the dish and allowed to migrate for 5 minutes. At the

end of the assay, the number of larvae in each zone was counted and a preference index (PI) was calculated. Increased concentration of methanol led an increase in preference index (figure 2.5A; one-way ANOVA F (7, 34) = 14.38, p<0.0001). Larval attraction to many odorants peaks at a diluted concertation, making a bell shaped preference index curve (Khurana et al., 2013). However, larvae were found to be most attracted to pure methanol, with no peak at a lower concentration (figure 2.5B).

2.3.6 Larval chemotaxis is adversely affected by DMSO

One of the challenges of propofol delivery is that it is insoluble in water. DMSO was used as a water-soluble vehicle to achieve good propofol miscibility in Ringer’s solution. However, given that larvae were previously found to have a negative chemotactic response to propofol, it was important to investigate the possible effects of DMSO on larval chemotaxis. CS larvae were exposed to a DMSO Ringer’s solution for 2 hours prior to the chemotaxis assay (figure 2.5C). It was found that DMSO adversely affects larval chemotaxis to methanol (figure 2.5D; one-way ANOVA F (4, 37) = 13.04, p<0.0001). To minimise this effect, a solution of 1.8% v/v DMSO in HL3 Ringer’s solution was chosen as a vehicle for propofol exposure. This was the minimal amount of DMSO that could effectively dissolve 10mM propofol in solution (Sigma).

Figure 2.5: Larvae positively chemotax to a methanol target, but this behaviour is affected by DMSO (A) Chemotaxis assay layout. Larvae are placed in the centre of a petri dish coated in 3% agar and allowed to migrate for 5minutes. On one side of the petri dish is a Teflon container with 20µL of methanol diluted in distilled water, on the other side is another container with 20µL of distilled water. At the end of an assay, a preference index is calculated. (B) Increased methanol concentration increased larval chemotaxis (one way ANOVA F(7, 34) = 14.38, p<0.0001). (C) DMSO assay work flow. Third instar larvae were collected and exposed to DMSO for 2 hours. Then, their chemotactic response to 100% methanol was assayed using the chemotaxis assay described above. (D) Increased exposure to DMSO (chemical structure shown) decreases larval chemotaxis to a pure methanol odour target (one-way ANOVA F (4, 37) = 13.04, p<0.0001).

2.3.7 Propofol abolishes larval chemotaxis to methanol

CS fly larvae were treated with propofol by immersion in a solution of propofol in HL3 Ringer’s solution for 2 hours. Control animals were immersed in HL3 Ringer’s solution containing the equivalent volume of DMSO vehicle. After propofol exposure, larval chemotactic response to methanol was measured (figure 2.6A). Compared to the 0mM control, treatment with propofol significantly reduced larval chemotaxis to methanol starting at 2mM (figure 2.6B; Welch’s t-test compared to 0mM control 1mM: p=0.1050, 2mM: p=0.0202, 5mM: p=0.0035, 7mM: p=0.0001, 10mM: p<0.0001). At 5mM concentration, chemotaxis was abolished entirely and the preference index was not significantly different from zero (figure 2.6B; one sample t-test 5mM: p=0.1450, 7mM: p=0.9896, 10mM: p=0.6625). The same data was also analysed in a grouped analysis. Rather than calculating a preference index, the response of each larva is shown. This allows for a more complete picture of the distribution of larvae on the plate. Recapitulating the result obtained using the preference index, the number of larvae on the methanol odour side of the plate decreases significantly with increased propofol compared to 0mM propofol control (figure 2.6C; Fisher’s exact test 1mM: p=0.1227, 2mM: p=0.0115, 5mM: p=0.0005, 7mM: p<0.0001, 10mM: p<0.0001). Using the grouped analysis approach, it can also be shown that the number of larvae remaining in the centre of the plate does not change significantly compared to 0mM propofol controls (figure 2.6C; Fisher’s exact test 1mM: p=0.4744, 2mM: p=0.3702, 5mM: p=0.7670, 7mM: p=0.3299, 10mM: p=0.0565). This demonstrates that the larvae’s decreased attraction to methanol is not due to a significant number of larvae being immobilised. Rather, the larvae seem to be unable to orient themselves toward the methanol odour target after propofol treatment.

2.3.8 Larval chemotaxis recovers from propofol exposure

One of the hallmarks of general anaesthetics is their reversible effects. In order to ensure that propofol treatment was not damaging the larvae and that the reduction in chemotactic response was reversible, a recovery assay was performed. After exposure to 10mM propofol, larvae were tested with the chemotaxis assay and then left to recover for 4 hours on standard yeast and sugar media. After the recovery period, larvae were retested with the chemotaxis assay. Larvae were demonstrated to recover normal chemotaxis behaviour after 4 hours of recovery (figure, 2.6D; t-test p=0.4441). This shows that the effect of propofol on chemotaxis is reversible.

Figure 2.6: Propofol abolishes larval chemotaxis to methanol (A) Chemotaxis assay work flow. Third instar larvae were collected and exposed to propofol for 2 hours. Then, their chemotactic response to 100% methanol was assayed using the chemotaxis assay described in figure 2.5A. (B) Effect of 2 hours of exposure to different concentrations of propofol on CS fly larvae. Propofol concentration effects chemotaxis (Welch’s t-test compared to 0mM control n.s. p=0.1050, * p=0.0202, ** p=0.0035, *** p=0.0001, **** p<0.0001). Hash marks (#) indicate a dataset is not significantly different from zero (one sample t-test 10mM: p=0.6625, 7mM: p=0.9896, 5mM: p=0.1450). N represents the number of experiments per data set. (C) Grouped analysis of the same data as figure 2.6B. Here, the response of individual larvae is shown. Compared to 0mM control, the number of larvae on the odour side decreases significantly with increased propofol exposure (Fisher’s exact test: n.s. p=0.1227, * p=0.0115, *** p=0.0005, **** p<0.0001). However, the number of larvae remaining in the centre of the plate does not change significantly (Fisher’s exact test p>0.05). N represents the number of individual larvae per data set. (D) Larval chemotaxis recovers from 2hours of 10mM propofol treatment. After 4 hours of recovery on standard yeast and sugar media, there was no significant difference between propofol treated and non-propofol treated groups (t test, n.s. p=0.4441).

2.3.9 Larvae become disoriented after propofol treatment

As demonstrated previously, propofol treatment does not seem to immobilise a significant number of larvae, so their impaired chemotaxis to methanol is probably due to disorientation. A larva’s ability to orient toward an odour target can be quantified by measuring its angle deviation. Angle deviation refers to the angle between a larva’s velocity vector and a vector pointing from the velocity vector’s origin to the methanol odour container (figure 2.7A). This is a measure of how effectively each larva orients itself towards the methanol target during the assay. An angle of 0º indicates a larva is moving directly toward the methanol container. Angle deviation was measured in 5 second intervals over a 5 minute assay.

A Rayleigh test against a generated von Mises distribution with the same mean vector direction demonstrated that the angle deviations of larvae treated with 10mM propofol were uniformly distributed and that the mean vector was therefore not significant (Rayleigh test p=0.3592). In contrast, the angle deviations of larvae that were not treated with propofol had a significant mean vector (magnitude: 0.261532, direction: -18.554º, std: 61.8410º, Rayleigh test: p=1.0594 x 10-4) (figure 2.7B). This demonstrates that larvae lose the ability to orient themselves toward the methanol target after a 2 hour treatment with

Figure 2.7: Propofol disorients larval chemotaxis (A) Angle deviation (β) is the angle between the larva’s velocity vector (blue) and a vector pointing from the velocity vector’s origin to the methanol container (red). An angle of 0º indicates the larvae is oriented directly toward the methanol container. (B) Polar plot of binned angle deviations with and without propofol treatment. Untreated larvae (left) had a significant mean vector (magnitude: 0.261532, direction: -18.554º, std: 61.8410º, Rayleigh test against a generated von Mises distribution: p=1.0594 x 10-4). The red arrow represents the direction of the mean vector, but its magnitude is exaggerated for clarity. Angle deviations of larvae treated with 10mM propofol for 2 hours (right) were not significantly different from a uniform distribution (Rayleigh test against a generated von Mises distribution: p=0.3592).

10mM propofol. It is probable that chemotaxis is vulnerable to general anaesthetics because it is a higher order endpoint that involves the coordination of different parts of the nervous system.

2.3.10 Immersion in 10mM propofol leads to a relevant dose in larval tissue

Exposure to high amounts of propofol (>5mM) had been found to result in an impairment of more complex larval behaviours (chemotaxis, crawling speed). But the persistence of

Figure 2.8: Using HPLC to determine propofol concentration in larval tissue (A) HPLC workflow. (B) Example propofol fluorescence peak from the tissue lysate of larvae exposed to 10mM propofol for 2 hours (top) and a fluorescence peak from a 0.01mM standard for reference (bottom). The retention time for propofol was about 5.5 minutes.

many other behaviours (mouth hook contractions, nociceptive response, and mobility) at such a high concentration seemed remarkable. However, it was unclear how much propofol was actually reaching the larval tissue. I therefore used high performance liquid chromatography (HPLC) to evaluate the concentration of propofol in the tissue of larvae after 10mM propofol treatment. Groups of 15 larvae were treated with 10mM propofol by immersion in a 1mL solution of propofol in HL3 Ringer’s solution for 2 hours. The larvae were collected and homogenised. Propofol concentration was measured using a fluorescence detector and normalised to the weight of the sample tissue (figure 2.8A). Immersion in 10mM propofol for 2 hours resulted in an average concentration of 8.470 ± 0.8963 ng of propofol per mg of tissue (figure 2.8B). Given the high external concentration that the larvae were exposed to, the tissue concentration was much lower than expected. However, this lower tissue concentration makes sense considering many behavioural endpoints persisted after propofol exposure. Furthermore, this clinically relevant concentration makes it likely that impairment of behaviours such as chemotaxis was due to general anaesthetic effects rather than off target effects.

2.4 Discussion

Different behavioural endpoints in animals can model the loss of responsiveness we call general anaesthesia in humans. If the target mechanisms are similar, then it is reasonable to use animal models like Drosophila larvae. However, it is important to know which endpoints make for viable assays. In this chapter I have successfully delivered propofol into larval tissue and have identified some key behaviours in Drosophila larvae which are impacted at different concentrations of propofol. However, taking several factors into account, the chemotaxis assays seems to be the most suitable for future studies.

Exposing larvae to propofol by immersion in a Ringer’s solution with 10mM propofol resulted in an average concentration of 8.470 ng/mg in the larval tissue. This is within a similar order of magnitude to the 10 μg/g found in the brains of mice during propofol infusion for surgery (Shortal et al., 2018). In adult humans, about 2 to 2.5mg/kg of propofol is used for induction of general anaesthesia in clinical settings (Bryson et al., 1995). It is also important to note that the concentration in larvae was estimated using whole body lysates, and probably does not reflect the concentration within the central nervous system. Attempts to directly measure the concentration of propofol in the larval central nervous system were unsuccessful since the amount of tissues that could be reasonable collected

did not yield a propofol concentration high enough for the florescence detector to detect. Being highly hydrophobic, it is likely that a large portion of propofol is taken up in the fat tissue. This could explain why the larvae are not immobilised despite being exposed to a high concentration of propofol. A more interesting conclusion could be that, like C. elegans, larval mobility is extremely resistant to propofol, but this is not conclusive. Nevertheless, the immersion method of exposure was found to be more effective than feeding (larvae are adverse to eating propofol) or injection (cumbersome and often results in gut extrusion through the cuticle).

The nociception assay was eliminated due to the discrepancy in the results between this and previous studies. The study upon which this assay was based found that the 1st mechanical stimulation (with fishing line) resulted in 20% of the larvae rolling while the 2nd stimulation resulted in approximately 50% of the larvae rolling (Hoyer et al., 2018). This is a much greater response than what I observed. Changing the stimulant from nylon fishing wire to metal wire did not increase nociceptive response. Since I was unable to replicate the previous study, I decided to not expand upon this assay.

Phototaxis was considered as a possible behavioural endpoint. However, there is conflicting literature on whether or not third instar larvae are attracted or repelled by light. Work done in the 1980s reported that larvae are photophilic in early growth stages, but become photophobic later development (Manning and Markow, 1981). More recent studies have concluded the opposite: larvae are photophobic during early developmental stages. Later stages, however, remain controversial. Some have reported Drosophila remains photophobic throughout its larval stage (Yamanaka et al., 2013), while others have found that Drosophila larvae may become photophilic (Sawin-McCormack et al., 1995) or at least photoneutral with age (Godoy-Herrera et al., 1992). Furthermore, recent work has suggested that hydrotaxis and thigmotaxis dominate over phototaxis and therefore impact the results of a larval phototaxis assays (Humberg and Sprecher, 2018). Due to these factors, I chose not to fully develop a phototaxis assay.

Some assays were ruled out due to being cumbersome. Larval speed and angle deviation were tracked by hand. This method proved to be more reliable than a software tracking system (SOS track), but too slow to be scaled up for further investigation.

The chemotaxis assay was selected for further use in this thesis due to its efficiency and flexibility. The chemotaxis assay devised allows for an efficient, population level assessment of the effects of propofol. Furthermore, besides a preference index, the assay

can be used to assess other effects of general anaesthetics such as angle deviation and recovery. Pure methanol was chosen to be the odour target as this elicited the strongest chemotactic response. There was no peak in preference index at a lower concentration as there is with many other odours (Khurana et al., 2013).

It was also notable that exposure to DMSO (the vehicle used to ensure propofol dissolved in Ringer’s solution) negatively effects larval chemotaxis to methanol. This was anticipated since, while an excellent solvent and preservative, DMSO is not used in many clinical applications due to its toxicity. It is easily absorbed through the skin and there is some evidence that it is a developmental neurotoxin (Hanslick et al., 2009). When used for surgery, propofol is typically formulated with an emulsion of soybean oil. The resulting solution is an opaque white colour, giving propofol the moniker “milk of amnesia” (Baker and Naguib, 2005). Despite this, many studies using propofol in a research setting use DMSO as a solvent. Furthermore, DMSO can be used in microscopy studies since it is clear and can be imaged through, whereas soybean oil is opaque. Since further studies using two-photon microscopy are planned (chapter 4) and because there is precedence in the literature, I chose to use DMSO as a solvent despite the potential drawbacks.

Despite the effect of the DMSO vehicle, the chemotaxis assay demonstrated that an increase in propofol exposure led to a clear reduction in the preference index to methanol. While the assay demonstrated that treatment with sufficient propofol abolishes larval preference for methanol, it is important to note that this was not due to immobility. As shown in figure 2.6C, larvae move from the centre of the plate regardless of the amount of propofol exposure. This is further supported by the crawling speed assay. While treatment with 10mM propofol significantly reduced the crawling speed of the larvae, it did not immobilise them. This demonstrates that the abolishment of methanol preference is due to disorientation (as demonstrated by the angle deviation) rather than immobility.

Chapter 3: Investigating pre- and postsynaptic targets for general anaesthetics using a chemotaxis assay

3.1 Introduction

Many different proteins have been identified as possible general anaesthetic targets. However, only a few of these have compelling evidence to support their direct involvement in general anaesthetic-induced loss of consciousness. In the previous chapter, a series of behavioural endpoints for propofol were identified in Drosophila larvae. This chapter will use these behavioural endpoints to investigate pre- and postsynaptic propofol targets and make comparisons with isoflurane, a gaseous general anaesthetic, and gaboxadol, a GABAergic sedative drug.

As discussed in chapter 1, most general anaesthetics interact with GABAA receptors and consequently effect endogenous sleep circuits (figure 1.7) (Franks, 2008). In mammals,

GABAA receptors are widely expressed throughout the central nervous system (CNS) (Laurie et al., 1992) and, at least in the adult mammalian brain, are involved in the inhibition of arousal pathways and potentiation of sleep circuits (Sherin et al., 1998). In

Drosophila, GABA performs a similar function, but the structure of the GABAA receptor differs slightly. The main GABAA subunit in Drosophila is RDL (Enell et al., 2007). It is expressed in the sleep/wake circuits of adult flies (Chung et al., 2009), and can be activated by a variety of sedatives such as gaboxadol (McGonigle and Lummis, 2010). Studies in cell culture have demonstrated that the general anaesthetics isoflurane and propofol interact with the RDL receptor (Edwards and Lees, 1997; Pistis et al., 1999). However, it is not known if RDL is the primary target for general anaesthetics in larvae. As demonstrated in chapter 2, propofol affects larval crawling speed and chemotaxis. Given that RDL is widely expressed in the larval brain and is a known propofol target, it is expected that RDL plays a role in propofol’s mechanism of action in larvae.

General anaesthetics also affect the function of multiple SNARE proteins at clinically relevant doses (Xie et al., 2013). In particular, syntaxin1A, an important part of the core SNARE complex, has been shown to be a possible intermediary for general anaesthetics’ ability to inhibit synaptic release. In the nematode worm, C. elegans, a point mutation known as md130 results in two variants of syntaxin1A (known as md130A and md130B) which are truncated at residue 227 and therefore missing part of the H3 domain and all of the transmembrane domain. The two isoforms differ in the number of novel amino acids at the carboxy-terminus: 12 in the case of md130A and 9 in the case of md130B. Expression 37

of these mutants in C. elegans diminishes sensitivity to isoflurane (van Swinderen et al., 1999) and overexpression of md130A in mammalian neurosecretory (PC12) cells blocks the ability of isoflurane and propofol to inhibit synaptic release (Herring et al., 2009; Herring et al., 2011). A possible mechanism for this inhibition was proposed in a recent study which found that propofol restricts the movement of syntaxin1A on the plasma membrane (Bademosi et al., 2018). The co-expression of syntaxin227 (a truncated variant similar to md130, but without the novel residues) (figure 3.2B) abolishes the effect of propofol on both synaptic release and syntaxin1A mobility (Bademosi et al., 2018). Additionally, expression of syntaxin227 confers resistance to both propofol and isoflurane in adult flies (Troup, 2018). A deletion mutant, syntaxinH3-N, which is missing the N-terminus of the H3 domain (Wu et al., 1999), has also been shown to rescue propofol-induced changes in syntaxin1A mobility (Bademosi et al., 2018) and has been demonstrated to confer resistance to isoflurane and propofol in adult flies (Troup, 2018) (figure 3.1B).

In chapter 2, I established chemotaxis and crawling speed assays to investigate the effect of propofol on third instar larvae. In this chapter, I use these assays to investigate both pre- and postsynaptic targets of propofol and to make comparisons with isoflurane, a gaseous general anaesthetic, and gaboxadol, a GABAergic sedative.

Figure 3.1: The structure of syntaxin1A and its role in the SNARE complex (A) The SNARE complex facilitates the fusion of vesicles with the plasma membrane. Syntaxin1A consists of several domains: the HABC domains (dark green), the H3 domain (red), and the transmembrane domain (yellow). The H3 domain, also known as the SNARE motif, interacts with synaptobrevin (blue) and SNAP25 (light green) to form the α-helical bundle at the core of the SNARE complex. (B) Structure of wild-type syntaxin1A (top) compared to two different mutants. The grey brackets represent deleted residues. Syntaxin227 is a truncated form that lacks the C- terminus of the H3 domain and the entirety of the transmembrane domain. SyntaxinH3-N is a deletion construct with the N-terminus of the H3 domain removed.

3.2 Methods

3.2.1 Fly stock maintenance and crosses

Fly lines were maintained as described previously (see section 2.2.1).

RDLRNAi and syntaxin1A variants were expressed using the GAL4-UAS system under the control of a pan-neuoronal GAL4 driver (Brand and Perrimon, 1993). GAL4 is a yeast transcription factor which binds to UAS (upstream activation sequence) to drive gene transcription. GAL4 was linked to the pan-neuronal driver ElavC155 (embryonic lethal abnormal vision). To make each cross, virgin female ElavC155-Gal4 flies and male flies expressing the desired UAS construct were collected and then allowed to breed in a vial containing standard yeast and agar media for 3 days. UAS-RDLRNAi is Rdl8-10J (Liu et al., 2007). Virgin female ElavC155-Gal4 flies were crossed with male UAS-Rdl8-10J / tm6,tb and the larvae were selected against the tubby (tb) phenotype. UAS-HA syntaxin227 was constructed by Dr Oressia Zalucki (University of Queensland, Australia) from full length Drosophila syntaxin1A with an HA tag. SyntaxinH3-N (W2202; syxH3-N) was expressed constitutively along with endogenous wild-type syntaxin1A. SyntaxinH3-N was a gift from Hugo Bellen (Baylor College of Medicine, USA) and was first constructed by Wu et al., 1999.

See section 2.2.2 for larval preparation methods.

3.2.2 Preparation of gaboxadol (THIP), propofol, and HL3 Ringer’s solution

20mg/mL stock solution of gaboxadol (4,5,6,7-tetrahydroisoxazolo(5,4-c)pyridin-3-ol or THIP) (Sigma) was made up in distilled water. A 560mM propofol stock solution was prepared in DMSO (Sigma). The appropriate volume of each stock solution was diluted in HL3 Ringer’s solution to make media for assays. Minimum haemolymph-like solution (HL3) was made according to Stewart et al., 1994 (70mM NaCl, 5mM KCl, 1.5mM CaCl2 · 2H2O,

20mM MgCl26H2O, 10mM NaHCO3, 5mM Trehalose, 115mM sucrose, and 5mM HEPES; pH 7.2; Π=343 ± 0.7).

3.2.3 Larval crawling speed after isoflurane treatment

The assay was performed on a 100 x 10mm plastic petri dish containing 20mL of 3% agar. The dish was placed in a custom airtight chamber measuring 27.5 x 28.5 x 7cm

(5.48625L). Isoflurane was delivered with an MQV anaesthetic machine (Mediquip) via a small opening fitted with a hose end (figure 3.7A). A vacuum removed the isoflurane through a similar opening on the opposite side of the chamber. At the beginning of the assay, three larvae were placed in the middle of the plate. The chamber was sealed, and isoflurane was then pumped in at a rate of 3.5L/min. The larvae were allowed to equilibrate for 10minutes, and were then filmed with a Logitech webcam for 5 minutes. A piece of tracing paper was taped over the monitor and, as the video was played back, the position of each larva was marked in 5 second intervals. Distance was calibrated with a 1cm scale bar in frame. Larval crawling speed was estimated by measuring the distance each larva travelled over the 5 minute assay.

3.2.4 Brain dissection and immunohistochemistry

The CNS of third instar larvae (ElavC155-Gal4 > UAS-HA syntaxin1A and ElavC155-Gal4 > UAS-HA syntaxin227) were removed by gently pulling the mouth hooks with dissection forceps. After dissection, brains were transferred to a mini PCR-tube with 200µl of 1x phosphate-buffered saline (PBS) solution. All of the following steps were performed on a rotator with 37RPM at room temperature. Brains were fixed with 4% paraformaldehyde diluted in PBS-T (1x PBS, 0.2 Triton-X 100) for 20-30min, followed by 3 washes in PBS-T. After the washes, they were blocked with 10% goat serum (Sigma Aldrich, St. Louis, MO, USA) for 1hour, followed by overnight primary antibody incubation. On the second day, primary antibody was removed and brains were washed 3 times with PBS-T. Then the secondary antibody was added and the tube was covered with aluminium foil for overnight incubation. On day three, the secondary antibody was removed, and the brains were washed with PBS-T. Brains were then transferred to microscope slides and mounted on a drop of vectashield (Vector Laboratories, Burlingame, CA) for imaging.

3.2.5 Statistics

Where relevant, data is presented as means and the error bars indicate ±SEM. Statistical analysis was performed in Graphpad Sofware Prism Version 7.02. Where relevant, a Pearson normality test or, where there were <8 data points, a Shapiro-Wilk normality test was used to determine the normality of a data set. In one instance (figure 3.5D), a data set was found to be not normally distributed. In this case, a Kolmogorov-Smirnov test was used when comparing this set to others. In all other cases when comparing two data sets, 40

a Welch’s t-test (which does not assume equal standard deviations between data sets) was used. For comparisons between three or more genotypes within a single propofol concentration, a one-way ANOVA with a Tukey's multiple comparisons test was used.

3.3 Results

3.3.1 Knockdown of the RDL subunit does not confer resistance to propofol

Homomeric RDL receptors are the most common GABAA receptor in the Drosophila brain (Enell et al., 2007) and has been demonstrated to be potentiated by propofol in vitro (Pistis et al., 1999). Propofol is a well characterised GABAergic drug, so it was hypothesised that RDL would be a significant target for propofol in the larval brain. If this is the case, then eliminating the RDL subunit could provide some resistance to propofol since it would be less likely to bind to GABAA receptors and exert its inhibitory effect. RNAi was used to knock down expression levels of RDL. Resistance was assessed using the chemotaxis assay developed in chapter 2. No difference was observed between the response of RDL knockdown larvae and controls. RDL knockdown lines were compared with genetic controls (figure 3.2). One-way ANOVA for each propofol concentration yielded insignificant results (0mM: F(2,12)=2.941, p=0.0913; 5mM: F(2,12)=0.4566, p=0.6440; 10mM: F(2,13)=0.001852, p=0.9981). A Tukey’s multiple comparisons test between RDL knockdown and control genotypes in each propofol concentration also yielded insignificant results (p>0.4). This implies that there may be other propofol targets which are responsible for the abolishment of larval chemotaxis.

Figure 3.2: RDL knockdown does not confer propofol resistance

Effect of propofol exposure on RDL knockdown and control lines. The effect of RDL knockdown was not significant (one-way ANOVA: 0mM: F(2,12)=2.941, p=0.0913; 5mM: F(2,12)=0.4566, p=0.6440; 10mM: F(2,13)=0.001852, p=0.9981). Error bars depict SEM, N indicates number of experiments in each dataset (there were 30 larvae per experiment).

3.3.2 The sedative gaboxadol does not affect larval chemotaxis.

In the adult Drosophila brain, RDL plays a crucial role in the suppression of arousal pathways and is thus important for the control of sleep onset (Chung et al., 1999; Parisky et al., 2008; Agosto et al., 2008). General anaesthetics are thought to enhance this sleep- promoting pathway to sedate adult flies (Weber et al., 2009; Kottler et al., 2013). However, it is not clear if similar sleep/wake pathways exist in larvae. While RDL knockdown did not mitigate propofol’s effect on larval chemotaxis, this does not mean a GABAergic sleep pathway is absent in larvae since propofol has other targets besides GABAA receptors. A non-general anaesthetic sedative with high specificity for GABAA receptors could therefore be used to probe for these putative sleep/wake pathways.

Gaboxadol or 4,5,6,7-tetrahydroisoxazolo(5,4-c)pyridin-3-ol (THIP) was derived from muscimol, a psychoactive alkaloid produced by flytrap mushrooms (Amanita muscaria). It was originally developed as a sedative for human use, but pharmaceutical research has been discontinued (Krogsgaard-Larsen, 1984). However, gaboxadol has since found a use in a research setting. It is a GABAA agonist that directly activates chloride currents in

mammalian GABAA receptors and fly RDL receptors (McGonigle and Lummis, 2010). However, unlike propofol, gaboxadol is not a general anaesthetic. It promotes slow-wave sleep and reduces spindle activity during non-REM sleep in humans (Faulhaber et al., 1997) and can similarly induce sleep in adult Drosophila either through feeding (Dissel et al., 2015) or direct perfusion across the brain (Yap et al., 2018). Gaboxadol treatment could thus be a good way to more directly probe for a sleep-promoting RDL circuit in larvae.

A previous study found that feeding adult flies 0.1mg/mL of gaboxadol results in increased quiescence and sleep (Dissel et al., 2015). Similarly, perfusion of 0.2mg/mL gaboxadol directly across an adult fly’s brain results in immobility within 5 min (Yap et al., 2017). Larvae were immersed in a HL3 Ringer’s solution containing gaboxadol for 2 hours, the same amount of time as the propofol exposure in previous experiments. Larval response to gaboxadol was then measured using a chemotaxis assay. Following gaboxadol exposure, larvae were not immobilised. Furthermore, no difference in larval preference to methanol was found after 2hour exposure to 0.1mg/mL or 1mg/mL gaboxadol (figure 3.3B; one-way ANOVA, F(2,12)=1.104, p=0.3637). Since RDL knockdown did not protect against propofol and because a GABAergic sedative had no effect on chemotaxis, it is

Figure 3.3: The sedative gaboxadol does not affect chemotaxis (A) Structure of gaboxadol (4,5,6,7-tetrahydroisoxazolo(5,4-c)pyridine-3-ol or THIP). (B) Effect of gaboxadol (THIP) exposure on CS wild-type flies. Treatment with 0.1mg/mL or 1.0mg/mL gaboxadol did not lead to a significant change in the methanol preference index (one way ANOVA, p=0.3637). Error bars depict SEM, N indicates number of experiments in each dataset (there were 30 larvae per experiment).

likely that propofol may have other target mechanisms that contribute to the abolishment of chemotaxis.

3.3.3 Sytaxin227 and propofol inhibition of larval chemotaxis

Propofol also targets key proteins in the presynaptic release machinery such as syntaxin1A (Bademosi et al, 2018). A truncated isoform of syntaxin1A, md130, has previously been shown to rescue the effect of propofol on synaptic release (Herring et al., 2011). A similar truncated isoform, syntaxin227, rescues the effect of propofol on syntaxin1A mobility in mammalian cell culture (Bademosi et al., 2018) (figure 3.1B). In adult flies, syntaxin227 expression can confer resistance to propofol and isoflurane (Troup, 2018). Furthermore, syntaxin227 has been shown to protect synaptic release in the 1b boutons of the larval NMJ (Karunanithi et al., manuscript in preparation). I therefore sought to investigate whether this resistance could be observed at the behavioural level. Syntaxin227 was expressed under the control of a pan-neuronal Gal4 driver (ElavC155-Gal4 > UAS-HA syntaxin227.1). Fly lines with the Elav and syntaxin227 genotypes were crossed with W1118 wild-type flies as controls. Treatment with propofol led to a significant reduction in chemotaxis to methanol in all fly lines tested (two way ANOVA F(1,44)= 60.85, p <0.0001). However, no difference was observed between syntaxin227 larvae and genetic controls (figure 3.4A; one-way ANOVA: 0mM: F(2,14)=3.045, p=0.0798; 5mM: F(2,11)=0.08617, p=0.9181; 10mM: F(2,17)=0.344, p=0.7138). The response of synatxin227 larvae was also compared to the two genetic controls at each propofol concentration using a Tukey’s multiple comparison test. This also yielded insignificant results (p>0.25). Expression of HA-tagged syntaxin227 was verified with immunostaining of the larval CNS. The CNS was also immunostained with NC82, a monoclonal antibody that marks the Bruchpilot (Brp) protein which is localised in active zones and commonly used as a neuropil marker (Wagh et al., 2006). There is clear expression of HA-tagged syntaxin227 in cell bodies, however expression is less clear in the neuropil (figure 3.4B).

Figure 3.4: Effect of syntaxin227 on larval propofol sensitivity (A) Syntaxin227 mutant expressing larvae are not resistant to 10mM propofol compared to genetic controls. In all fly lines, treatment with 10mM propofol led to a significant reduction in the methanol preference index compared to 0mM controls (two way ANOVA F(1,44)=60.85, p <0.0001). However, the effects of the different fly lines on propofol response was not significant (one-way ANOVA: 0mM: p=0.0798, 5mM: p=0.9181, 10mM: p=0.7138). Error bars depict SEM, N indicates number of experiments in each dataset (there were 30 larvae per experiment). (B) Expression pattern of HA tagged syntaxin227 and NC82 in the larval CNS and expanded views of the central brain and the ventral nerve chord.

3.3.4 Comparing the effect of syntaxin227 and wild-type syntaxin1A overexpression on larval propofol sensitivity

Compared to its genetic controls, syntaxin227 expression did not result in a propofol- resistant phenotype. However, a more adequate genetic control may be the overexpression of wild-type syntaxin1A using the same Gal4-UAS system. Wild-type syntaxin1A was overexpressed under the control of a pan-neuronal Gal4 driver (ElavC155- Gal4 > UAS-HA syntaxin1A). Fly lines with the ELAV and UAS-HA syntaxin1A genotype were crossed with W1118 wild-type flies as controls. Treatment with 10mM propofol led to a significant reduction in chemotaxis to methanol in all fly lines tested (two way ANOVA F(2,49)=30.72, p <0.0001). However, no difference was observed between the response of larvae overexpressing syntaxin1A and genetic controls at any of the propofol concentrations tested (figure 3.5A; one-way ANOVA: 0mM: F(2,12)=0.395, p=0.6821; 5mM: F(2,15)=1.445, p=0.2668; 10mM: F(2,22)=1.58, p=0.2284). SynatxinWT overexpression was also compared to the genetic controls at each propofol concentration using a Tukey’s multiple comparison test. However this also yielded insignificant results (p>0.25). Overexpression of HA-tagged syntaxin1A was verified with immunostaining of the larval CNS. The CNS was also immunostained with NC82 to label the neuropil (figure 3.5C).

Both wild-type syntaxin1A and syntaxin227 were expressed using the same Gal4-UAS system. Expression of these proteins may have led to behavioural deficits that could mask resistance to propofol. Comparing syntaxin227 larvae to syntaxin1A overexpressing larvae could therefore be useful to control for these effects. Furthermore, these constructs were expressed along with endogenous syntaxin1A, so comparing these two lines is a good

Figure 3.5: Comparing the effect of syntaxin227 and syntaxin1A overexpression on larval propofol sensitivity (A) Larvae overexpressing syntaxin1A are not resistant to propofol. In all fly lines, treatment with 10mM propofol led to a significant reduction in the methanol preference index compared to 0mM controls (two way ANOVA F(2,49)=30.72, p <0.0001). However, the effects of the different fly lines was not significant on propofol response (one-way ANOVA: 0mM: p=0.6821, 5mM: p=0.2668, 10mM: p=0.2284). Error bars depict SEM, N indicates number of experiments in each dataset. (B) Compared to larvae overexpressing syntaxin1A, syntaxin227 larvae are resistant to treatment with 5mM propofol (Welch’s t-test, * p= 0.0102). Error bars depict SEM, N indicates number of experiments in each dataset (there were 30 larva per experiment). (C) Expression pattern of HA tagged syntaxin and NC82 in the larval CNS. NC82 is a monoclonal antibody that marks the Bruchpilot (Brp) protein which is localised in active zones.

way to more directly compare the effect of propofol on the truncated isoform versus wild- type syntaxin1A. Interestingly, compared to wild-type syntaxin1A overexpression larvae, syntaxin227 larval chemotaxis was resistant to 5mM propofol treatment (figure 3.5B; Welch’s t-test, p=0.0102). This result implies that the transmembrane domain or the H3 domain may be important for propofol to exert its effect.

3.3.5 SyxH3-N protects against propofol inhibition of larval chemotaxis

SyxH3-N is a syntaxin mutant with a deletion that removes the majority of the N-terminal region of the H3 domain (amino acids 204–250) (Wu et al., 1999). Like syntaxin227, this mutant been shown to rescue propofol-induced changes in syntaxin1A mobility on the plasma membrane (Bademosi et al., 2018) (figure 3.1B). However, in contrast with syntaxin227, syntaxinH3-N does not interact with SNAP25 (Wu et la., 1999; Bademosi et al., 2018). SyntaxinH3-N expression also confers resistance to isoflurane and propofol in adult flies (Troup, 2018). It therefore seems likely that this resistance could be observed in larval behaviour.

SynxtaxinH3-N was expressed constitutively (along with endogenous wild-type syntaxin1A) and W2202 wild-type flies were used as a genetic background control. Treatment with 10mM propofol led to a reduction in chemotaxis to methanol in all fly lines tested (two-way ANOVA F(1,44)=60.85, p <0.0001). However, at 5mM, syntaxinH3-N larvae were more resistant compared to genetic background controls (figure 3.6; Kolmogorov-Smirnov test, p=0.0377). A Kolmogorov-Smirnov test was used in lieu of a t-test since the W2202 dataset contained an outlier which made the data not normally distributed (D’Agostino & Pearson normality test K2=17.02, p=0.0002). This result implies that the N-terminus of the H3 domain may be important for propofol to exert its effect.

Figure 3.6: SyntaxinH3-N effects larval propofol sensitivity At 5mM propofol, larvae expressing syntaxinH3-N were more resistant than wild type (Kolmogorov-Smirnov test, p=0.0377). Error bars depict SEM, N indicates number of experiments in each dataset (there were 30 larvae per experiment).

3.3.6 Isoflurane reduces larval crawling speed

So far, I have focused on propofol, an intravenous general anaesthetic. However, if inhibition of synaptic release is a mechanism common to many general anaesthetics, it is probable that similar manipulations of syntaxin1A could confer resistance to other types of general anaesthetics such as isoflurane.

In order to draw comparisons with the propofol results, I attempted to create a chemotaxis assay for isoflurane. Isoflurane is a gas, which necessitated a different delivery method than propofol. Larvae where placed on a 100 x 10mm agar plate inside of an airtight chamber. Isoflurane was then pumped into the chamber with an anaesthetic machine (figure 3.7A). Unfortunately, because the assay had to be conducted in an airtight box, manipulating the larvae was challenging. In particular, preventing larvae from crawling toward the methanol target before they had been sufficiently exposed to isoflurane was problematic.

Measuring crawling speed proved to be the most straightforward assay to measure larval response to isoflurane. The assay was performed on a 100 x 10mm plastic petri dish

containing 20mL of 3% agar. At the beginning of the assay, CS wild-type larvae were placed in the middle of the agar plate and the correct concentration of isoflurane was pumped into the chamber. The larvae were allowed to equilibrate for 10minutes and were then filmed with a webcam for 5 minutes. Larval crawling speed was estimated by measuring the distance each larva travelled over the 5minute assay.

Isoflurane significantly reduced the crawling speed of CS wild-type larvae (one-way ANOVA F=11.05, p=0.0004). Even 0.25 volume% (v%) was enough to reduce crawling speed compared to the control (Dunnett’s multiple comparisons *p=0.0306). 0.5v% was sufficient isoflurane so that larval crawling speed was not significantly different from 0mm/s (figure 3.7B; one sample t test p=0.1393).

3.3.7 SyntaxinH3-N confers resistance to isoflurane

Previous work in our lab has demonstrated that adult flies expressing syntaxinH3-N are resistant to the general anaesthetic isoflurane (Troup, 2018). I hypothesised that larvae would be similarly resistant. Larvae constitutively expressing syntaxinH3-N (W2202; syntaxinH3-N) were exposed to 1 v% isoflurane and their crawling speed was measured over a 5 minute assay as described previously. Compared to genetic background controls, syntaxinH3-N expressing larvae had a higher crawling speed after exposure to 1v% isoflurane (Welch’s t-test, p=0.0275) (figure 3.7C). As shown previously, syntaxinH3-N larvae were also resistant to propofol, suggesting that propofol and isoflurane are acting on syntaxin1A in similar ways.

Figure 3.7: Larval sensitivity to isoflurane (A) Isoflurane delivery set up. Isoflurane was delivered with an MQV anaesthetic machine (Mediquip) to an air-tight chamber (27.5 x 28.5 x 7cm). The assay was performed inside the chamber on a 100 x 10mm plastic petri dish containing 20mL of 3% agar. Larvae were recorded for 5 minutes with a webcam and their average crawling speed was measured (see methods). (B) Effect of isoflurane on crawling speed of CS wild-type larvae. Isoflurane significantly affected larval crawling speed (one-way ANOVA F=11.05, p=0.0004; Dunnett’s multiple comparisons to 0v%: *p=0.0306, **p=0.0013, ***p=0.0002). Hash marks (#) indicate data is not significant from 0.0mm/s crawling speed (one sample t test p>0.05). Error bars depict SEM, N indicates number of animals in each dataset. (C) Compared to W2202 wild-type controls, the crawling speed of larvae expressing the syntaxinH3-N mutant is less affected by isoflurane. (Welch’s t-test n.s. p=0.5645, * p=0.0275). Error bars depict SEM, N indicates number of animals in each dataset.

3.4 Discussion

It is now generally accepted that general anaesthetics exert their effect by targeting proteins key to neuronal function. These drugs interact with a wide variety of proteins, however only a few are thought to be important for producing general anaesthesia. With a few exceptions, GABAA receptors have been established as an important general anaesthetic target and the activation of GABAergic sleep circuits has been proposed as a plausible explanation for the mechanism underlying general anaesthesia. This is supported in part by similarities between the induction of anaesthesia and sleep (Franks, 2008). More recent evidence has suggested a presynaptic mechanism involving synaptic release facilitated by the SNARE complex (Xie et al., 2013). It has also been shown that genetic manipulations of key proteins in the SNARE complex can alter an organism’s susceptibility to general anaesthetics (van Swinderen et al., 1999; Zalucki et al, 2015). In this chapter, Drosophila third instar larvae were used to investigate aspects of the pre- and postsynaptic mechanism of propofol.

General anaesthetics are known to affect sleep/wake pathways in mammals (Franks, 2008) and adult flies (Weber et al., 2009; Kottler et al., 2013) and GABAergic sleep circuits are thought to be the primary target for propofol. Interestingly, knockdown of RDL did not produce a propofol resistant behavioural phenotype. While not consistent with a GABAergic hypothesis for the mechanism of propofol, this result is in agreement with previous work in our lab which showed that RDL knockdown in adult flies does not produce an isoflurane resistant phenotype (Troup, 2018). It is possible that increased expression of other GABAA subunits such as GRD (Harvey et al., 1994) or LCCH3 (Henderson et al., 1993) may be compensating for the knockdown of RDL. Alternatively, the RDL receptor’s inherent resistance to propofol could explain this lack of a phenotype. Propofol’s potentiation of the RDL receptor is threefold less potent compared to its potentiation of the human GABAA (α6β3γ2L) receptor (Pistis et al., 1999). Consequently, a much higher concentration of propofol is required to potentiate the GABAA receptor in flies. It is therefore possible that the concentration of propofol delivered to larval tissue was insufficient to activate the RDL mechanism, and the reduction in chemotaxis was due to other mechanisms which require lower concentrations of propofol.

To further probe for potential sleep-promoting pathways in larvae, larvae were exposed to the GABAergic sedative gaboxadol. Feeding adult flies 0.1mg/mL gaboxadol results in increased quiescence and sleep (Dissel et al., 2017). In contrast, larvae immersed in ten

times that concentration for 2 hours remained mobile and larval chemotaxis was unaffected. This could indicate that larvae lack the sleep circuitry often implicated to be the main target of general anaesthetics. However, the difference in delivery method could also account for this apparent resistance in larvae compared to adult flies. An important caveat of this result is that it is not clear how much of the gaboxadol was ingested by the larvae, and it may be that insufficient gaboxadol was taken up in the central nervous system to have an effect. Furthermore, gaboxadol is a full agonist of the mammalian GABAA receptor, but is only a partial agonist of RDL receptors (McGonigle and Lummis 2010) which could lead to further resistance.

Given that manipulations of the GABAergic targets did not change larval sensitivity to propofol, it was likely that propofol was interacting with other targets to produce an anaesthetising effect. I tested the overexpression of wild-type syntaxin1A as well as two different syntaxin1A mutants. Overexpression of wild-type syntaxin1A did not affect sensitivity to propofol, using the chemotaxis endpoint. However, a truncated isoform of syntaxin1A, syntaxin227, was resistant compared to overexpression of the wild-type protein, and constitutive co-expression of a deletion mutation, syntaxinH3-N, resulted in resistance compared to genetic background controls. These results are consistent with previous studies which found that these syntaxin1A variants can confer resistance to propofol in adult flies (Troup, 2018) and rescue impairment of syntaxin1A protein mobility caused by propofol (Bademosi et al., 2018).

The effect of syntaxinH3-N on propofol resistance can also be generalized to isoflurane. Isoflurane and propofol are very different general anaesthetics both in structure (halocarbon vs. phenol) and state (gas vs. liquid), but the resistance of syntaxinH3-N larvae to both implies a similar mechanism of action. SyntaxinH3-N resistance in larvae mirrors previous work in our lab which demonstrated that adult Drosophila expressing syntaxinH3-N are resistant to isoflurane (Troup 2018), demonstrating that this rescue effect transcends life stage.

The resistance of these mutants may also give clues to how general anaesthetics affect syntaxin1A. It has been hypothesised that general anaesthetics prevent the formation or “zippering” of the SNARE complex by interfering with hydrophobic pockets in the constituent proteins (Hemmings et al., 2005; Nagele et al., 2005; Xie et al., 2013). Both mutants have a deletion in part of the H3 domain (figure 3.1B). The H3 domain is particularly important for SNARE complex formation as it directly interacts with other proteins such as synaptobrevin and SNAP-25 (Südhof and Rizo, 2011) (figure 3.1A). It 53

could be that general anaesthetics may be inhibiting the H3 domain’s interactions with other SNARE proteins. Recent studies have also shown that propofol restricts the movement of syntaxin1A proteins on the plasma membrane, further limiting its ability to form SNARE complexes (Bademosi et al., 2018). However, expression of either syntaxinH3-N or syntaxin227 rescues this mobility defect in PC12 cells (Bademosi et al., 2018). Taken together, these results may suggest that general anaesthetics could be impairing the formation of SNARE complexes via some mechanism involving the H3 domain of syntaxin1A, and that this effect confers resistance to general anaesthetics in any system involving neurotransmission (mammalian neurosecretory cells, fly larvae, fly adults, etc.). It remains unknown, however, whether the effects of propofol on larval behaviour stem from target processes in their central nervous system, or whether all synaptic neurotransmission across the animal is equally important.

Chapter 4: Developing a technique for imaging larval central nervous system activity 4.1 Introduction

The previous chapter used behavioural assays to investigate the effect of general anaesthetics and the sedative gaboxadol on larvae. However, understanding the effects of general anaesthetics and sedatives remain incomplete until the brain is considered. In humans, imaging techniques such as functional magnetic resonance imaging (fMRI) can be used to monitor changes in brain dynamics during drug treatment (Hudetz, 2012). However, many of these techniques are not suitable for use in Drosophila larvae. For an animal with a small central nervous system (CNS) like Drosophila, two-photon microscopy can capture the whole-brain activity in an individual animal in vivo. Here, we have paired this technique with the ultrasensitive protein calcium sensor GCaMP6. This is a genetically encoded calcium indicator that pairs the calcium binding protein calmodulin (CaM) with green florescent protein (GFP) in a way where they fluoresce in response to the calcium influx during depolarisation events. This fluorescence can be used as a proxy for neuronal activity (Chen et al., 2013). Furthermore, two-photon microscopy is ideal for in vivo samples because it allows for high-resolution, high-contrast fluorescence imaging much deeper in the tissue and with less phototoxicity compared to conventional confocal microscopy (Svoboda and Yasuda, 2006). Compared to the adult fly, not much has been published regarding imaging the larval CNS in vivo. Previous functional imaging of the larval CNS involved dissecting the brain from the animal and embedding it in low-melting point agar (Lemon et al., 2015). The larval CNS is surprisingly robust to this manipulation and, despite being removed from the body, still displays fictive behaviours such as forward and backward crawling (Lemon et al., 2015).

In the previous chapter, it was found that propofol treatment affected larval chemotaxis while gaboxadol treatment did not. The behavioural effects of these drugs should be reflected in changes in CNS activity. Imaging the brain could also potentially disambiguate the putative sleep effects from presynaptic effects. In this chapter, I develop a larval preparation for calcium imaging in order to measure the neural activity in the larval CNS during treatment with the sedative gaboxadol. While the results in this chapter are preliminary, the methods developed here are a starting point for understanding the neural substrates underlying the behavioural effects of sedatives in larvae. I focus on gaboxadol for two reasons: I sought brain imaging data to match the behavioural findings in chapter

3, and an acute perfusion protocol for gaboxadol (which is soluble in solution) was developed in anticipation of future experiments with propofol.

4.2 Methods

4.2.1 Larval CNS preparation

Fly rearing was carried out as described previously (see chapter 2 methods). The calcium indicator GCaMP6s was expressed under the control of a pan-neuronal driver (57c10- GAL4>UAS-GCaMP6s).

Third instar larvae were prepared based on a protocol described in Bademosi et al., 2018. Each larval dissection was carried out in Schneider’s Insect medium on a sylgard-coated plate. The mouthparts and posterior abdomen of the larva were pinned in place with minutien pins. A minutien pin was used to make a small hole through the cuticle along the dorsal midline. Iridectomy scissors were then used to cut along the dorsal midline and then laterally so that the body wall could be pinned to form a flat sheet. The intestines and other viscera were then gently scraped away to access the nervous system (figure 4.1B).

4.2.2 Imaging

Imaging was performed using a ThorLabs microscope with a Ti: Sapphire laser source (Mai Tai DeepSee, Spectra Physics) at 930nm. Laser power was controlled by a pockels cell with voltage modulated by an electro-optic modulator (Conoptics, 302A). A piezo (Precision Instruments) moved a 40x objective (water immersion 0.8NA, Nikon) through the Z plane to allow imaging in 3 dimensions. Imaging resolution was 512 x 256 pixels. A volume of 26 (10µm) slices plus 4 flyback slices were taken across 300 frames. GCaMP florescence was detected by a photomultiplier tube (PMT) following long-pass by a 594 dichroic beam splitter and band-pass by a 525/25 filter (figure 4.1A).

The larval CNS was maintained in 200µL of Schneider’s insect media throughout imaging. Baseline activity was recorded for 300 time points (5 minutes). Media was aspirated off while new media containing 0.2mg/mL gaboxadol was added. In order to not dry the brain, new media was added as old media was aspirated off. During each media change, the brain was washed three times. Activity during drug treatment was recorded for 300 time points. The drug-containing media was then aspirated off and new media was added. Recovery was then recorded for 300 time points (figure 4.2A).

4.2.3 Fluorescence analysis

Images were analysed by taking a standard deviation z-projection and selecting a region of interest (ROI) around the ventral nerve chord (VNC) and an ROI around the central brain (CB). ImageJ was used to measure the mean change in greyscale value of each ROI for each condition. An ROI with the same dimensions was used when comparing the same region across different time points (figure 4.2B). The mean fluorescence was normalised to the first 10 frames (10 seconds) of the baseline recording and averaged across larvae (n=4).

4.3 Results

4.3.1 A filleted larval preparation for two-photon imaging

While embedding the CNS in agar yields good images (Lemon et al., 2015), this method would not be ideal for delivering drugs to the brain. At first, an attempt was made to image the CNS in an intact larva. To prevent movement, both the head and tail were pinned to a sylgard-coated plate. Although some peripheral nerves could be identified, GCaMP expression was not strong enough to image the CNS through the cuticle. Instead, a dissection method based off of Bademosi et al. 2018 was developed (see above). This involved cutting the larva open entirely and pinning the cuticle flat onto a sylgard-coated plate. The viscera were then removed to give access to the CNS (figure 4.1B). This preparation allowed for in vivo imaging while also giving easy access so that drugs could be perfused across the CNS. Here, I focussed on gaboxadol because it is soluble in Schneider’s insect medium at the required concentration. Preliminary experiments with propofol revealed difficulty in performing these acute experiments at clinically-relevant concentrations without using an opaque emulsifier (intralipid) which precluded imaging. Future studies with propofol will require a drug washout protocol to address this issue.

Figure 4.1: Setup for calcium imaging in the larval central nervous system (A) Schematic of two-photon microscope. PMT (photomultiplier tube), BP (band-pass), LP (long-pass). Two-photon microscopy uses two low-energy photons from the same laser (red lines) to cause a fluorescent molecule to transition to a higher energy state (green lines).

Image adapted from Dr. Michael Troup, University of Queensland.

(B) Schematic of larval preparation. Larvae were pinned to a sylgard plate and cut down the dorsal midline. The cuticle was pinned down and the viscera removed to reveal the CNS. (C) Larval CNS expressing GCaMP6s clearly showing the central brain and the ventral nerve chord.

4.3.2 Imaging the larval brain under gaboxadol

As observed in chapter 3, gaboxadol did not affect larval chemotaxis. However, gaboxadol was delivered by immersing the larvae in a 1mg/mL gaboxadol solution which may have not delivered sufficient drug to the animal. In humans, gaboxadol promotes slow-wave sleep and reduces spindle activity during non-REM sleep (Faulhaber et al., 1997). Similarly, in adult flies, perfusion of 0.2mg/mL gaboxadol directly across the brain results in immobility within 5 min and an overall decrease in local field potential activity in the brain (Yap et al., 2017). Given that RDL (the main target for gaboxadol) is expressed widely in the larval CNS (Enell et al., 2007), gaboxadol should affect larvae similarly.

The calcium indicator GCaMP6s was expressed under the control of a pan-neuronal driver (57c10-GAL4>UAS-GCaMP6s) and two-photon microscopy was used to image CNS activity for 5 minutes during baseline, gaboxadol treated, and recovery conditions (figure 4.2A). Larvae were treated with 0.2mg/mL gaboxadol as this concentration was used in a previous study of the adult fly brain (Yap et al., 2017).

In each recording, ROIs were selected around the central brain and the VNC (figure 4.2B). To compare the activity of these regions during each condition, the change in average fluorescence was measured in each ROI. Fluorescence was normalised to the first 10 frames of the baseline condition. A one-way ANOVA with a Tukey’s multiple comparisons test was used to compare the average fluorescence in each condition. Surprisingly, treatment with 0.2mg/mL gaboxadol reduced the average fluorescence of both regions (central brain: p=0.0119, VNC: p=0.0336). However, the recovery condition was not significantly different from the gaboxadol treatment condition (central brain: p= 0.5674, VNC: p=0.3167) or the baseline condition (central brain: p=0.06, VNC: p=0.3167). A two- way ANOVA with a Sidak’s multiple comparison test was used to compare the central brain to the VNC in all conditions. There was no difference in change in mean fluorescence between the central brain and VNC for all conditions (F(1,6)=2.181, p=0.1902; baseline: p=0.9984, gaboxadol: p=0.6121, recovery: 0.3801).

Figure 4.2: Larval central nervous system response to gaboxadol (A) Experimental timeline indicating the time points of media changes. (B) A standard deviation z-projection of the larval brain. ROIs were selected around the ventral nerve chord (VNC) and the central brain (CB). (C) Example traces of the average greyscale intensity of the ROIs during baseline, 0.2mg/mL gaboxadol, and recovery conditions. Intensity is normalised to the first 10 frames of the baseline condition. (D) Treatment with 0.2mg/mL gaboxadol reduced the average fluorescence of both ROIs. However, the recovery condition was not significantly different from gaboxadol treatment (one-way ANOVA and Tukey's multiple comparisons test CB: * p=0.0119, n.s. p>0.06; VNC: * p=0.0336, n.s. p>0.3). There was no difference in fluorescence between the CB and VNC for all conditions (two-way ANOVA with a Sidak’s multiple comparisons test n.s. p>0.3). N=4 for each condition, error bars represent SEM. 60

4.4 Discussion

There is a potentially interesting comparison to be made between the effects of general anaesthetics and sedatives on the CNS. While the results presented in this chapter are preliminary, gaboxadol seems to affect the overall activity in the larval CNS. This is in contrast to the result of the behavioural assay in chapter 3, which found that gaboxadol did not affect larval chemotaxis. It therefore seems likely that larvae are susceptible to gaboxadol, but that the delivery method used in chapter 3 did not result in a concentration high enough in the CNS to result in a behavioural change. However, it could also be that gaboxadol is not acting as a sedative on the larval brain and that it causes a decrease in activity through some other mechanism.

Although the results presented here are preliminary, the methods developed could be used in the future to investigate the effects of general anaesthetics and soluble drugs on the larval CNS in vivo. A variety of dissection techniques were tested. Two often opposing factors were at play: the technique must keep the CNS alive, but also must restrain the animal enough to prevent too much movement during imaging. At first, an attempt was made to image the CNS in an intact larva using minutien pins to keep the animal in place. However, the fluorescence signal proved to be too weak to image the CNS through the cuticle. Next, a semi-intact dissection similar to a CNS-hoisting technique described previously (Iyengar, 2012) was attempted. This involved cutting and pinning the cuticle above the CNS to make it more accessible for imaging. However, this method often resulted in considerable movement in the z-direction due to continuous peristaltic contractions of the body wall muscles and cephalopharyngeal skeleton. Finally, a more complete dissection method was used where the whole cuticle was cut open and pinned flat (figure 4.1B). While more stable than the semi-intact dissection, this technique still had some drift in the z-direction. This was partially due to some peristaltic contractions of the body wall muscles, but also perhaps due to the movement of the objective in the media.

A more thorough analysis of these images was attempted using watershed segmentation to identify individual active neurons. However, the GCaMP fluorescence signal was not defined enough to segment individual neurons. This could be because the dissection was too invasive, however the larval brain has been shown to be fairly robust and can still show activity when removed from the body entirely (Lemon et al., 2015). Improving the dissection technique could produce better results with more clearly active neurons. This could also be due to the expression of GCaMP in the larvae. Using a different driver or a

different GCaMP localised to a different part of the cell may result in a more defined signal that can be segmented. To compare functional connectivity and neural activity dynamics between baseline, gaboxadol-treated, and recovery conditions, future experiments could find correlations between active neurons using graph theoretical analysis (Wang et al., 2010; Sporns, 2013).

An attempt was also made to measure CNS activity during propofol treatment. However, there was difficulty in delivering a clinical dose of propofol to the CNS. In the larvae tested, there did not seem to be a consistent decrease in activity during propofol treatment. This could possibly be due to the DMSO used to dissolve propofol. DMSO has been used as a solvent for propofol in previous studies in PC12 cells (Herring et al., 2011) and the larval NMJ (Bademosi et al., 2018), but it is not used to dissolve propofol in a clinical setting. Instead, a phospholipid stabilized soybean oil called intralipid is used. However, intralipid is opaque, so it is not ideal for imaging. If intralipid were used as a vehicle, imaging could be done immediately after propofol treatment. While the results presented in this chapter are preliminary, the methods developed here can be further optimised and used in the future to answer questions about the response of the larval CNS to propofol and other drugs.

Chapter 5: Conclusions and future directions

Despite their structural diversity, all general anaesthetics lead to reversible loss of consciousness. However, how they exert their influence are still not entirely known. A postsynaptic, GABAergic mechanism involving the potentiation of endogenous sleep circuits has been widely supported (Franks, 2008), but more recent evidence suggests that there may also be a presynaptic component as well. Adult Drosophila have been used as a convenient model organism to study general anaesthetic mechanisms (Karunanithi et al., 2018), but larval behavioural responses to general anaesthetics have remained relatively unexplored, especially for intravenous general anaesthetics. This thesis developed larval Drosophila as an animal model to probe intravenous general anaesthetic effects. The value of using Drosophila larvae for such studies is twofold: first, synaptic physiology is much better studied in larval preparations (Bademosi et al., 2017; Bademosi et al., 2018) and second, their central nervous system (CNS) is significantly simpler. It has been proposed that larvae may lack endogenous sleep circuitry (van Swinderen and Kottler, 2014), so it was thought that studying the larval stage could better dissect the presynaptic effects from the sleep effects of general anaesthesia. While this thesis could not rule out the existence of a larval sleep circuit, it successfully characterised larval behavioural endpoints to the intravenous general anaesthetic propofol. Furthermore, genetic manipulation of the SNARE protein syntaxin1A was found to result in resistance to propofol, whereas no such resistance was found when GABAergic function was downregulated.

As discussed in chapter 2, more complex behaviours were more susceptible to propofol, as was expected based on similar evidence in other animals (Zalucki and van Swinderen, 2016). Chemotaxis, which involves coordinated movement to navigate an odour gradient, was abolished with propofol treatment, whereas mouth hook contractions and mobility were not. However, it is difficult to relate these behaviours to the responses of other model organisms. The same can be said of comparing the response of adult and larval Drosophila, since the two life stages share very little in common in terms of morphology or behaviour. This is further complicated by the differences in drug delivery. Adult Drosophila can be fed propofol. In adult flies, oral ingestion of 1mM propofol leads to a concentration of approximately 243 ng/mg tissue (Gardner et al., 2016). This is far higher than the 8.470 ng/mg tissue achieved through the immersion method used for larvae (see chapter 2). It is therefore hard to make comparisons between the adult and larval stages’ susceptibility to propofol using the methods described here. However, an interesting point of comparison 63

can be made with C. elegans. While no formal study has been published on the behavioural response of C. elegans to propofol, it has been informally reported that propofol does not immobilise C. elegans (Forman, 2006). Despite exposing larvae to high concentrations of propofol for up to 24 hours, I was similarly unable to immobilise larvae with propofol. In both cases, this could be due to the difficulties of delivering propofol. Alternatively, since propofol is typically considered a GABAergic drug, the failure to immobilise C. elegans and larval Drosophila could indicate the lack of GABAergic sleep circuits in these animals.

In chapter 3, the putative GABAergic target was explored further. The main GABAA subunit in the Drosophila nervous system is RDL (Enell et al., 2007), a known propofol target (Pistis et al., 1999). Interestingly, knockdown of RDL receptors in larvae did not lead to a propofol-resistant phenotype. This is well aligned with previous studies in adult flies which found the same RDL knockdown did not confer resistance to general anaesthetics (Troup, 2018). In order to probe further for potential GABAergic sleep circuits, I exposed larvae to gaboxadol, a GABAA agonist with high specificity. This was inspired in part by work done in adult flies which become sedated after being fed 0.1mg/mL gaboxadol, (Dissel et al., 2017). Larvae, however, remained mobile and larval chemotaxis was unaffected even after immersion in ten times that concentration for two hours. This could provide further support for the view that that larvae may lack the sleep circuitry often implicated to be the main target of general anaesthetics. However, in chapter 4, preliminary results indicated that perfusion of 0.2mg/mL gaboxadol reduces activity across the larval CNS. This same concentration of gaboxadol has been found to decrease brain activity in adult flies (Yap et al., 2017). While this reduction in CNS activity does not necessarily mean that gaboxadol is acting as a sedative in larvae, it does suggests that treatment with sufficient gaboxadol should lead to observable behavioural changes. It is therefore possible that the immersion method used in chapter 3 resulted in insufficient gaboxadol in the CNS to have a behavioural effect. The concentration of gaboxadol in larval tissue could potentially be determined with HPLC. However, unlike propofol (which has a clear emission peak), fluorescence detection may not be viable for gaboxadol and tandem mass spectrometry may be more suitable (Madsen 1983; Kall et al., 2007).

While these results do not conclusively rule out the existence of a sleep circuit in Drosophila larvae, it could suggest that if sleep circuits do exist in larvae, they might not be GABAergic. It is also possible that GABA may have a different function in larvae compared to adults. In the developing mammalian brain, for example, GABA takes on an excitatory

role (Chen et al., 1996). However, even if GABAergic sleep circuits do exist in larvae, propofol could have caused the reduction in chemotaxis through mechanisms other than the potentiation of RDL receptors. RDL receptors are much more resistant to propofol than mammalian GABAA (α6β3γ2L) receptors. The EC50 (the concentration of propofol producing a half-maximal response) is 8.7µM propofol for the α6β3γ2L receptor and 25µM for the RDL receptor (Pistis et al., 1999). However, neither adult nor larval flies are extraordinarily resistant to general anaesthetics.

In chapter 3, it was found that larvae co-expressing a deletion mutant form of syntaxin1A (syntaxinH3-N) which is missing the N-terminus of its H3 domain, was resistant to both propofol and isoflurane. Conversely, expression of a truncated syntaxin1A variant (syntaxin227) was not more resistant compared to its genetic controls. These results could be explained by the difference in the way each mutant protein was expressed. SyntaxinH3-N was expressed constitutively whereas syntaxin227 and wild-type syntaxin1A were tagged with HA and expressed under the control of a pan-neuronal Gal4 driver. It could be that differences in expression method are responsible for differences in susceptibility to general anaesthetics. It is important to note, however, that syntaxinH3-N was expressed in a different background than syntaxin227 and wild-type syntaxin1A (W2202 versus W1118). Different background strains have different sensitivities to general anaesthetics (Campbell and Nash, 2001; Zalucki et al., 2015a), so it would be spurious to draw comparisons between mutants in different background strains. However, syntaxin227 larvae were found to be more resistant to propofol than larvae overexpressing wild-type syntaxin1A. Both syntaxinH3-N and syntaxin227 are missing part of the H3 domain which suggests that this domain may be an important site of interaction for propofol.

Previous studies demonstrated that syntaxin227 rescues propofol-induced inhibition of synaptic release in PC12 cells in vitro (Bademosi et al. 2018). This discrepancy between in vitro resistance and behavioural susceptibility could be due to the small size of the boutons in the larval CNS (which is responsible for many of the behaviours assayed here). Larger boutons allow for more neurotransmitter-containing vesicles. If general anaesthetics decrease the number of active release sites per synapse, then synapses with fewer release sites are more likely to be impacted. This would make the CNS particularly vulnerable to general anaesthetics. With this caveat in mind, it is possible that wild-type syntaxin1A overexpression is protective against general anaesthetics, but not sufficiently so to protect the central nervous system.

Another reason why resistance at a synaptic level may not have translated to behavioural resistance could be due to the assay design. The chemotaxis assay was a rather blunt tool which resulted in datasets with high variance. It is therefore likely that this assay was unable to detect small behavioural resistance. The chemotaxis assay also had several confounds that may have impacted its sensitivity to detect anaesthesia phenotypes. The foremost concern was that the larvae’s aversion to propofol out-competed their chemotactic response to methanol. As observed in chapter 2, larvae are highly averse to propofol, but it was not determined whether this response was gustatory or chemotactic. The propofol immersion technique used may have masked the smell of methanol, and it remains possible that the disorientation observed after propofol treatment could have been an olfactory effect. However, there are several lines of evidence which suggest that this artefact is unlikely, and that instead an anaesthetising effect was actually observed. First, syntaxinH3-N larvae are resistant to propofol, as determined by the chemotaxis assay. It is unlikely that this resistance is an olfactory effect since syntaxinH3-N larval chemotaxis was not significantly different from wild type in the absence of propofol. Furthermore, HPLC determined a clinically relevant concentration in larval tissue. Finally, syntaxinH3-N larvae were also found to be resistant to isoflurane. Resistance to another general anaesthetic with a different delivery mechanism gives further support that the chemotactic defect observed was an anaesthetic rather than olfactory response. As a future experiment, it would be useful to assess syntaxinH3-N larval resistance to propofol using a crawling speed assay (as was done for isoflurane), to fully exclude spurious effects on olfaction.

Chapter 4 developed techniques to image the larval CNS during treatment with gaboxadol. If the dissection technique is improved and propofol solubility issues resolved, two-photon imaging of the larval CNS could reveal differences in neural activity under propofol treatment that are responsible for the behavioural difference observed in previous chapters. Following studies in humans (Alkire et al., 1995) and adult flies (van Swinderen, 2006), it is probable that a clinical concentration of general anaesthetics would result in decreased activity across the whole larval CNS. Alternatively, particularly under low concentrations of general anaesthetic, there may be changes in connectivity between different brain regions. To compare functional connectivity and neural activity dynamics between baseline, propofol-treated, and recovery conditions, future experiments could find correlations between active neurons using graph theoretical analysis.

This thesis set out to develop larval Drosophila as a model organism to study behavioural responses to the intravenous general anaesthetic propofol. Since larvae may lack GABAergic sleep circuits, larvae seemed to be a good candidate to separate the presynaptic effects from the sleep effects of general anaesthesia. While it was found that a presynaptic mechanism involving syntaxin1A is a likely target for propofol, the question of the existence of a larval sleep circuit target remains unanswered. However, for future experiments, C. elegans may be a more suitable model to investigate how genetic manipulations affect behavioural responses to propofol. C. elegans behavioural assays are already well established (Crowder et al., 1996) and the genetic tools available in C. elegans allow for efficient expression of mutant proteins of interest. Nevertheless, this thesis did establish a series of behavioural assays which could be used in the future to further investigate the effects of intravenous general anaesthetics on larval behaviour. Furthermore, deletion of part of the H3 domain of syntaxin1A was found to result in resistance to propofol and isoflurane. These results suggest that propofol and isoflurane act through similar mechanisms and that this mechanism involves a presynaptic component that transcends life stage and brain size.

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