Genetic screen to Characterize Shank Interactors at the Drosophila Neuromuscular Junction

by

Ashley Hogg

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Cell and Systems Biology University of Toronto

© Copyright by Ashley Hogg 2018

ii

Genetic Screen to Characterize Shank Interactors at the Drosophila

Neuromuscular Junction

Ashley Hogg

Master of Science

Department of Cell and Systems Biology University of Toronto

2018

Abstract Mutations in Shank family genes are highly implicated in idiopathic autism spectrum disorders

(ASD) and therefore have been repeatedly studied using rodent models, however, study results

have been varied possibly due to redundancy from multiple Shank family genes. Drosophila

models offer a clearer approach as they possess only one Shank gene, allowing in-depth study of

Shank and its interactions with other factors. Using the Drosophila neuromuscular junction as a

model of glutamatergic brain synapses, a genetic screen was performed to identify novel

interactors of Drosophila Shank and characterize new molecular pathways to understand how

Shank functions at synapses. Results from this screen indicated that Ca-α1d may share a

relationship with Shank. Further characterization in a Shank overexpression background

confirmed an interaction that is not apparent in a Shank loss-of-function background. Studying

genes linked to ASDs in a simple model may reveal unappreciated signalling pathways

providing new directions in autism research.

iii

Acknowledgments

First, I would like to thank my supervisor Dr. Bryan Stewart for managing to be both

extremely supportive and easygoing at the same time. You provide an environment where your

students’ myself included can grow in our knowledge and be independent. Without your

guidance I may never have even considered graduate school as an option for my future. Your

enthusiasm about neurobiology is boundless and always encouraging. Thank you for this

opportunity!

To my committee, Dr. Joel Levine and Dr. Adriano Senatore thank you for taking time

out of your busy schedules to be a part of this project. Your direction was much appreciated.

Thank you for all your help over the last two years Katie, this is basically your thesis

project. Just kidding don't take my degree away. Your constant support and guidance was

immensely helpful along the way. Thank you for bringing me into the world of Shank and

sharing your baby with me. You will make an amazing supervisor; anybody would be lucky to

have you as lab mom.

To all the friends I made in these past 2 years, Abi, Maliha, Urfa, Gordy and Christine.

Thank you for always being there for a chat, science related or not. You guys really made this

work fun and offered up well-needed distractions. We did some dumb stuff over these two years

from someone lighting their hands on fire, ruining the family home of some gingerbread men,

playing with dry ice, and I think we can all admit running gels gave us some trouble. For

insurance purposes I won’t be able to say who did what. I wish you guys the best in your

futures.

iv

To the ever helpful fly community. Thank you Dr. Daniel Eberl for providing me with

the AR66 Ca-α1d mutant fly line (University of Iowa, Iowa City, IA). Many thanks to

Bloomington Drosophila Stock Center (Indiana University, Bloomington, IN; NIH

P40OD018537) for providing fly stocks.

I would also like to thank my parents. Even though they had no idea what was going on

basically the entire time they were always ready to sit through a practice presentation on Shiv

(it's Shank mom but good hustle). Your constant love and support saw me through this project

and it was nice to always have someone in my corner.

To Sal you came into this adventure part way through but probably still managed to take

the brunt of my complaining and science talk. You were genuinely interested in the work I do

unlike most people who’s eyes glaze over when I start talking neurobiology. You had endless

encouragement for me and always believed in me. I can’t tell you how much that meant. If you

were serious about wanting to read my thesis strap in because here we go!

v

Table of Contents

Acknowledgements................................................................................................................ iii-iv Table of Contents........................................................................................................................ v List of Tables ............................................................................................................................. vi List of Figures .......................................................................................................................... vii List of Abbreviations ......................................................................................................... viii-ix Chapter 1 – General Introduction ................................................................. 1-27

1.1 Synaptic transmission .................................................................................................... 1-5 1.2 Drosophila as a model organism ................................................................................. 6-12 1.3 Shank ......................................................................................................................... 13-19 1.4 Ca-α1d ....................................................................................................................... 20-22 1.5 Autism spectrum disorders (ASD) ............................................................................. 23-24 1.6 Shank and ASD .......................................................................................................... 25-26 1.7 Thesis aims and hypothesis ............................................................................................. 27

Chapter 2 – Materials and Methods ............................................................ 28-39 2.1 Fly stocks and strains ................................................................................................. 28-31 2.2 Dissection ........................................................................................................................ 32 2.3 Immunofluorescence .................................................................................................. 33-34 2.4 Image acquisition and analysis ........................................................................................ 35 2.5 Statistical analysis and sample size ................................................................................. 36 2.6 RNAi validation – RT-PCR ....................................................................................... 37-39 2.6.1 Primer verification .............................................................................................. 37-38 2.6.2 RT-PCR ................................................................................................................... 38 2.6.3 Gel image acquisition and analysis .......................................................................... 39 Chapter 3 – Results ....................................................................................... 40-62 3.1 RNAi screen (Phase one) ........................................................................................... 40-45 3.2 RNAi screen (Phase two) ........................................................................................... 46-51 3.3 Ca-α1d, a postsynaptic Ca2+ channel, interacts with Shank ...................................... 52-62 3.3.1 Ca-α1d heterozygotes in Shank overexpression ................................................ 54-55 3.3.2 Double versus single Shank and Ca-α1d heterozygotes .................................... 56-58 3.3.3 Visualization of possible colocalization of Ca-α1d and Shank with i immunohistochemistry ..................................................................................................59-62 Chapter 4 – Discussion .................................................................................. 63-73 4.1 Interpretation of genetic screen phase one ................................................................. 63-65 4.2 Interpretation of genetic screen phase two ................................................................. 66-68 4.3 Interaction between Ca-α1d and Shank is only apparent in a Shank overexpression b background ....................................................................................................................... 69-70 4.4 Immunohistochemistry interpretation .............................................................................. 71 4.5 Relation to the field .................................................................................................... 72-73 References ............................................................................................................................ 74-81

vi

List of Tables

Table 1. Shank interactors and Drosophila homologs

Table 2. Bloomington RNA interference lines

Table 3. VDR RNA interference lines

Table 4. Bloomington Ca-α1d mutant lines

Table 5. Ca-α1d mutant line

Table 6. Haemolymph-like saline solution (HL3) recipe

Table 7. Phosphate buffer (PBS) recipe

Table 8. Squishing buffer recipe

Table 9. TAE buffer recipe

Table 10. Phase one statistical and morphological results

Table 11. Phase one statistical significance results

Table 12. Phase two statistical and morphological results

Table 13. Phase two statistical significance results

Table 14. Ca-α1d heterozygotes in a Shank overexpression background statistical and

morphological results

Table 15. Ca-α1d heterozygotes in a Shank overexpression background statistical significance

results

Table 16. Ca-α1d and Shank loss-of-function heterozygote statistical and morphological results

Table 17. Ca-α1d and Shank loss-of-function heterozygote statistical significance results

vii

List of Figures

Figure 1. Phase one bouton number graph

Figure 2. Phase one sample neuromuscular junction confocal images

Figure 3. Phase two bouton number graph

Figure 4. Phase two sample neuromuscular junction confocal images

Figure 5. Genetic screen bouton counts for Ca-α1d only

Figure 6. Verification of Ca-α1d knock down with RT-PCR

Figure 7. Ca-α1d heterozygotes in Shank overexpression background graph

Figure 8. Ca-α1d heterozygotes in Shank overexpression background sample neuromuscular

junction confocal images

Figure 9. Ca-α1d and Shank loss-of-function heterozygote bouton number graph

Figure 10. Ca-α1d and Shank loss-of-function heterozygote sample neuromuscular junction

confocal images

Figure 11. Sample confocal images of CaV1.3 validation with differing Ca-α1d expression

Figure 12. Sample confocal images of interaction hypothesis investigation

viii

List of Abbreviations

ACh - Acetylcholine

ADHD - Attention deficit hyperactivity

disorder

ANK - Ankyrin repeats

ASD - Autism spectrum disorder

AZ - Active zone

Ca-α1d - Ca2+-channel protein α1 subunit

D

Cac - Cacophony

CNS - Central nervous system

dCSP - Drosophila anti-cysteine string

protein

DHP - Dihydropyridine

Dlg - Discs large

DMG - Damaged due to fragility (could not

analyze)

EPP - End plate potential

FNI - Fz2 nuclear import

Fz2 - Frizzled-2

GB - ghost boutons

GFP - Green fluorescent protein

GluR - Glutamate receptors

HL3 - Haemolymph-like saline solution

HRP - Anti-horeseradish peroxidase

LTHL - lethal

mGluR - Metabotropic glutamate eceptors

NGS - Normal goat serum

NMJ - Neuromuscular junction

NSF - N-ethylmaleimide-sensitive factor

PBS - Phosphate buffer

PBT - PBS plus Triton (0.1%)

ix

PCR - Polymerase chain reaction

PDZ - PSD-95/discs large/zonula

occludens-1

PSD - Postsynaptic density

RIM - Rab3-interacting molecules

RISC - RNA-induced silencing complex

RNAi - RNA interference

rtGEF - Rho-type guanine nucleotide

exchange factor

SAM - Sterile alpha motif

SH3 - Src homology 3

SNAP - Soluble NSF attachment protein

SNARE - Soluble NSF attachment protein

receptor

Tb - tubby

t-SNARE - Target membrane SNARE

v-SNARE - Vesicle membrane SNARE

shRNA - Small hairpin RNAs

shmiRNA - small hairpin micro RNAs

SSR - Subsynaptic reticulum

VAMP - Vesicle-associated membrane

protein

VDR - Vienna Drosophila resource center

w- - white

UAS - Upstream activation sequence

1

Chapter 1 – General Introduction

1.1 Synaptic transmission

At the most basic level, the nervous system is responsible for coordinating the activities of an

organism by transmitting signals. These signals serve to maintain homeostatic order and allow

the organism to respond to different internal and external cues. The nervous system is also

responsible for more complicated processes such as movement, sensory awareness, learning and

memory. To achieve the above functions, signals need to travel throughout the organism in a

timely and organized manner.

Synaptic transmission is the process by which a nerve cell passes information on to target cells

via synapses. A synapse is the space where the cell membranes of the cell sending the signal, the

presynaptic cell, and the cell receiving the signal, the postsynaptic cell, are in close apposition.

Both the pre- and postsynaptic cells contain an extensive array of molecular machinery which

carry out the signaling process including the release of neurotransmitter and reception and

integration of the signal.

Synapses can be classified into two functionally distinct groups: chemical or electrical,

depending on the type of signals they employ. An electrical synapse utilizes channels called gap

junctions, which connect the pre- and postsynaptic cells. These channels allow the direct

passage of an electrical charge from one cell to the next, whereby a voltage change in the

presynaptic cell induces a voltage change in the postsynaptic cell. At a chemical synapse,

neurotransmitter released from the presynaptic cell is received by receptors on the postsynaptic

cell to induce a change. More specifically, voltage changes in the presynaptic cell open voltage-

gated Ca2+ channels found in the nerve terminal. This triggers a series of biological events

which ultimately leads to exocytosis of synaptic vesicles containing neurotransmitter from the

nerve terminal. These chemicals diffuse across the synapse to bind receptors located on the

membrane of the postsynaptic cell. Once bound the neurotransmitter can elicit either an

electrical response from the postsynaptic cell or trigger a secondary messenger pathway, both of

which can influence the postsynaptic cell in an excitatory or inhibitory manner. Chemical

2

synapses are often classified according to the neurotransmitter they release. For example,

glutamatergic chemical synapses release glutamate, which elicits an excitatory response

(Silverthorn, 2007).

Knowledge regarding chemical transmission has come a long way since the 17th century when it

was believed that nerves produce movement in muscle through the use of “animal spirits”

distilled by heat from the heart (Descartes, 1637). By the 19th century nerves were shown to end

outside the muscle fibre and displayed the capacity to be electrically excitable (Krause, 1863). It

was du Bois-Reymond (1874) who first hinted that nerve endings use some sort of chemical

transmitter to stimulate the muscle. However, his theory was slightly off base in that he

mistakenly believed the nerve pierced the muscle and if the nerve in fact remained outside it

could only be using electricity as stimulation.

In the very early 1900s Elliot, with some help from Langley, laid the groundwork for the idea of

chemical transmission instead of electrical transmission. They also identified adrenaline as the

neurotransmitter which is released from sympathetic nerves to stimulate smooth muscle (Elliott,

1904; reviewed in Bennett, 2000). They termed the junction between the nerve and smooth

muscle the “myo-neural junction”. Around the same time (1905) Langley and Ehrlich suggested

the presence of transmitter receptors in the postsynaptic cell (reviewed in Bennett, 2000). Due to

the work of Loewi (1921), chemical transmission was established as an accepted hypothesis and

became the focus of countless researchers when he discovered “Vagustoff” (later identified as

acetylcholine (ACh)) was responsible for transmission between the vagus nerve and cardiac

muscle (reviewed in Bennett, 2000).

Some of the most basic physiological properties of neuron-to-target cell communication came

from Hodgkin and Huxley (1945). By inserting microelectrodes into giant squid axons they

determined that the inside of the cell was maintained at a more negative voltage when compared

to the outside of the cell. This later came to be known as a membrane potential, which is what

ultimately allows the propagation of action potentials.

Another key player in the field of synaptic physiology was Bernard Katz. In the mid to late 20th

century Katz, along with other researchers, used the frog neuromuscular junction (NMJ) to look

into the result of neuromuscular transmission called end plate potentials (EPP). EPPs are local

3

depolarizations of the muscle where neurotransmitter (ACh) has bound a receptor (Fatt and

Katz, 1951). Further research into this showed that EPPs result from the summation of “small

all-or-none units” called quanta (Del Castillo and Katz, 1954). They ultimately came to the

conclusion that neurotransmitter was released in discrete quantal units which summate to

depolarize the muscle. Neurotransmitter is released from the nerve terminal either when it leaks

spontaneously producing miniature EPP, or when triggered to do so by an action potential (Del

Castillo and Katz, 1954). A quantum is now known to represent the response to the vesicle

fusion and subsequent release of neurotransmitter held within a single vesicle (Heuser et al.,

1979).

After chemical transmission was recognized as the means of communication between a neuron

and its target cell, focus turned to understanding the mechanics of synaptic transmission. For

synaptic transmission to occur neurotransmitter from the presynaptic cell must be released into

the synapse. To accomplish this, secretory vesicles containing the neurotransmitter must fuse

with the plasma membrane of the presynaptic cell and this process requires specialized proteins

which form a soluble N-ethylmaleimide-sensitive factor (NSF) attachment protein receptor

(SNARE) complex (Söllner et al., 1993). The complex is made up of two proteins bound to the

target (plasma) membrane (t-SNAREs) called syntaxin and SNAP-25, and synaptobrevin (also

known as vesicle-associated membrane protein (VAMP)) which is found on the vesicle

membrane (v-SNARE) (Rothman, 1994). The SNARE complex is made up of a bundle of four

parallel α-helices connected by hydrophobic interactions; one helix comes from syntaxin and

synaptobrevin each and two helices come from SNAP-25. The zippering of these four helices

brings the vesicle and plasma membranes into close proximity. For vesicular fusion to occur the

repulsive forces generated by the negative charge of the phospholipid bilayers must be

overcome by the energy released during SNARE complex formation (Antonin et al., 2002). The

SNARE complex is responsible for both the docking and the fusion of the two membranes

(Weber et al., 1998)

Synaptotagmin, a Ca2+ sensing protein found bound to the vesicle, also plays a role in the fusion

process. It binds to syntaxin and acts as a sensor to initiate Ca2+ dependent neuronal vesicle

fusion (Hanson et al., 1997). Proof that this complex is responsible for the fusion of secretory

vesicles was provided by a study which used botulinum and tetanus toxins to proteolyse the

4

components of the SNARE complex, which prevented synaptic transmission (Niemann et al.,

1994; Montecucco and Schiavo, 1995). Once vesicular fusion is complete the SNARE complex

is reversibly disassembled by the ATPase activity of NSF and recycled for future use (Hanson et

al., 1997).

Also present bound to the secretory vesicles of the presynaptic cell are three members of the

Rab protein family: Rab5, Rab11 and most abundantly Rab3. Rab proteins are able to bind GTP

and regulate intracellular transport. Specifically, Rab3 regulates the release of neurotransmitter

by binding to RIMs (Rab3-interacting molecules) and inducing vesicular docking at the active

zone (Südhof, 2004). Rab3 participates in a cycle of association and dissociation with the

secretory vesicles that parallels vesicle exocytosis and endocytosis, where its dissociation from

the vesicle is dependent on Ca2+-triggered vesicle exocytosis. This means that when the vesicle

is at rest Rab is bound to it and once exocytosis begins Rab dissociates (Südhof, 2004; Südhof,

2014). Around the active zone, RIM can be found within a protein complex composed of several

different non-membrane proteins including Munc13. RIMs contain multiple domains that

interact with other RIMs and directly or indirectly with several other synaptic components. They

possess a zinc-finger domain on their N-terminal which interacts with both Rab3 and Munc13

(Südhof, 2004). Through these interactions RIM is able to perform many functions to regulate

neurotransmission including positioning vesicles near voltage-gated Ca2+ channels, recruiting

Ca2+ channels to the active zone (AZ), mediating plasticity and activating Munc13 (Südhof,

2014). Munc13 seems to serve an important role in secretory vesicle maturation. The loss of this

protein allows for the formation of normal excitatory synapses but the synaptic-vesicle cycle is

arrested leading to a shortage of releasable secretory vesicles (Augustin et al., 1999). Munc13

and RIMs are also considered the most important priming factors as they help get the vesicles

ready for Ca2+ triggered fusion (Südhof, 2014).

Membrane fusion between the vesicle and the presynaptic cell membrane allows the

neurotransmitter inside the vesicle to diffuse across the synapse and bind to the receptors on the

membrane of the target cell. Response to the released neurotransmitter can be either excitatory

or inhibitory depending on which ion channels are gated or regulated by the neurotransmitter

receptor. Over one hundred of these chemical messengers have been identified, including

GABA and glycine which generally illicit an inhibitory response, and glutamate and

5

acetylcholine which generally illicit an excitatory response. Whether or not an action potential,

in the case of a neuron, or a contraction, in the case of a muscle, is generated depends upon the

summation of the signals. Summation involves combining excitatory and inhibitory signals from

either multiple sources of simultaneous input (spatial summation) or repeated signals in rapid

succession from one source (temporal summation). For an action potential or muscle contraction

to be triggered in the postsynaptic cell summation of the signals must surpass a certain voltage

threshold. Depolarization of the postsynaptic cell results from excitatory neurotransmission and

makes the postsynaptic cell more likely to reach the voltage threshold required to illicit an

action potential or contraction. Hyperpolarization occurs after inhibitory neurotransmission and

makes it harder for the postsynaptic cell to reach the voltage threshold (Coolen et al., 2005).

6

1.2 Drosophila as a model organism

The very first publication that used Drosophila melanogaster as the experimental organism was

published in 1905 by Fredrick W Carpenter. However, research into Drosophila genetics really

took off when Thomas Morgan stumbled upon a male fly with white eyes in a population of red-

eyed flies during his experiments on evolution. Unbeknownst to Morgan he had just discovered

the first Drosophila mutation, white eye (w-), which would set the stage for future work into

Drosophila genetics and heredity (Morgan, 1910). Further work with Drosophila by Morgan

and colleagues would reveal novel features about Drosophila such as, that chromosomes and

genes were the conveyors of inheritance, male Drosophila were heterozygous for sex, and the

phenomena of deletions, sex linkage, cross-over events, and non-disjunction of X chromosomes

(Morgan, 1910, 1911; Morgan et al., 1915). Research using Drosophila took a back seat to

microbial genetics for several years until the 1970s when it re-emerged as a popular model

organism to study the nervous system.

The Drosophila melanogaster genome is 180-megbases long, consisting of approximately two

thirds euchromatin and one third heterochromatin. Up to 89% of the protein-coding genes in the

genome can be found in the euchromatin. It has a relatively small genome of about 13,000 genes

compared to the 30,000 found in humans. The chromosomal make-up of the organism consists

of the sex chromosomes (XY or XX), two larger autosomal chromosomes called 2 and 3 and

finally a smaller autosomal chromosome 4 (Celniker and Rubin, 2003). Only the autosomal

chromosomes may undergo recombination.

Fruit flies are a prominent organism in research for several reasons mostly revolving around

how easy they are to experiment with. They require minimal care, their life cycle is short, they

can be cultured in large numbers, and there is a low cost to maintain them. Their genome is easy

to manipulate and quite small which allows for genetic mapping (Keshishian et al., 1996;

Celniker and Rubin, 2003). Drosophila cells and tissues are amendable to many forms of

analysis, including imaging at single-cell resolution, electrophysiology, and molecular genetics

(Keshishian et al., 1996). Balancer chromosomes which prevent recombination are widely

available and allow recessive mutations that would otherwise be lethal, to be passed on to the

7

next generation without selection (Celniker and Rubin, 2003). Mutant flies can be generated or

obtained where a gene’s function has been knocked out or knocked down to determine the

impact of that gene on the organism. Overexpression of genes can also be generated by using

the UAS/GAL4 system created by Brand and Perrimon (1993). This system works through the

use of two separate transgenic elements. One element is the transcriptional activator isolated

from yeast called GAL4. The other element is the Upstream Activation Sequence (UAS)

promoter made up of five GAL4 binding sites placed upstream of the target gene. Gene

transcription can only occur if these two elements are combined and the progeny of that

combination will overexpress the target gene in the expression pattern of the GAL4 promoter

(Brand and Perrimon, 1993). For example, a mef2-GAL4 activator drives expression of the

target gene in the body wall muscles of Drosophila. This system has been critical to

understanding the role of specific genes in development and functioning of Drosophila.

Another tool useful in the Drosophila model is RNA interference (RNAi). RNAi is a reverse

genetic approach to perform gene knockdown in culture or in vivo. There are four different types

of RNAi used in vivo including synthetic siRNAs, small hairpin RNAs (shRNAs), small hairpin

microRNAs (shmiRNAs) and long dsRNAs. siRNAs and long dsRNAs were used in this

project. siRNAs are ~20 base pairs long. When expressed in the animal, one strand of the

siRNA becomes incorporated into the RNA-induced silencing complex (RISC). RISC is a multi-

subunit ribonucleoprotein complex responsible for cleaving mRNA and consequently RNA

interference. Via complementary base-pairing the incorporated siRNA strand directs RISC to

the mRNA to degrade it. Long dsRNA range from 200-500 base pairs long. Long dsRNA, once

expressed in the animal is cleaved by the enzyme Dicer into siRNAs and follows a similar

process as above (Perrimon et al., 2010).

Another factor in the popularity of Drosophila as a model is the amount of evolutionary

conservation in the genes that coordinate basic developmental processes such as establishing the

body axes, cell types and organ systems. The high level of conservation between the Drosophila

and human genomes allows for the investigation of the genes responsible for hereditary diseases

in humans. Studies have found that 75% of human disease-causing genes have orthologs in flies

meaning that approximately 700 human disease genes are conserved enough that they can be

8

studied in flies. Mutant flies that model characteristics of human diseases can be generated and

used for study (Bier, 2005). Work with disease genes in a simple model such as the fruit fly can

be used to determine the basic biology underlying human disease pathology.

As with most scientific tools there are some limitations to using Drosophila as a model

organism for vertebrates. First, you cannot use fruit flies to study any of the genes related to

biological processes that only take place within vertebrate species. This includes studying genes

related to development of a four-chambered heart, bone or mammary glands. Although the short

generation time, small genome and large offspring number of Drosophila are extremely

convenient for experimentation there are other organism available such as yeast which display

the above characteristics but to a greater degree. And of course there is always the question of

relevance of genetics studies performed in simple organisms with regard to human disease (Bier,

2005).

Despite these limitations the Drosophila NMJ has become a popular model for the study of

synapse assembly, function and plasticity. The developing synapses of Drosophila and

vertebrates are similar at the cellular and molecular level and structurally and functionally

similar proteins are expressed at both developing synapses. Many of the cellular components of

synaptic transmission are conserved between Drosophila and vertebrates as well such as the

SNARE complex (Keshishian et al., 1996). In particular, the 3rd instar larval NMJ is a useful

model synapse due to its accessibility and stereotypical structure. The NMJ model has been

extremely valuable in determining the roles of synaptic genes and discovering new ways in

which they interact.

NMJ assembly begins at the mid- to late-stages of embryo development when the growth cones

(axons) of motor neurons leave the central nervous system (CNS) through a common lateral

nerve exit point (Keshishian et al., 1996). After this they follow three peripheral nerve tracts

which innervate the body wall muscles, the intersegmental and segmental nerves which

innervate the dorsal and ventral regions, and the transverse nerve which innervates the mid-body

region (Johansen et al., 1989a reviewed in Keshishian et al., 1996). Axonal growth cones leave

their stereotypical paths and select a specific muscle to contact from a pool of potential muscles

9

by following molecular cues (Broadie and Bate, 1993) and passing through guidance choice

points which act independently from each other (Seeger et al., 1993 reviewed in Keshishian et

al., 1996). Each motor neuron specifically projects to one or more muscle fibres creating a

precise wiring pattern. A single body wall hemisegment receives innervation from its own CNS

segment and from its anterior CNS segment. This set up of motor neurons suggests there is

motor control of individual muscle fibres and groups of fibres. The pattern of the Drosophila

musculature consists of an arrangement of segmental repeats with 30 muscles in each

hemisegment (A2-A7) innervated by 35 motor neurons from the CNS (Keshishian et al., 1996).

Initial synapse formation occurs over several hours. First, the filopodia of the growth cone

extend towards the myopodia of the target muscle. Once contact has been made a morphological

transition occurs from the large flat growth cone to prevaricosities which contain immature

presynaptic specializations. Lastly, mature boutons are formed with synapses at the end of the

nerve terminals (Jin, 2002). New boutons and synaptic branches are formed in the second phase

of synapse development. As the animal progresses through its three larval stages the muscle size

increases drastically and so too must the motor neuron increase its number of boutons and

synapses to properly innervate the muscle (Harris and Littleton, 2015). Successful synapse

formation requires highly organized events to occur simultaneously on either side of the synapse

ensuring the precise apposition of the pre- and postsynaptic signalling apparatus.

The Drosophila third instar larval stage is the last and largest of the three larval stages. This

stage offers an NMJ easily accessible by dissection and musculature identifiable under a light

microscope. For this project, the NMJ innervating hemisegment A3 of muscle 6 and 7 was

examined via immunofluorescence and confocal microscopy. This NMJ is a popular choice for

use in studies due to its convenient and easily accessible location on the body wall. The

innervation comes from two different type I motor neurons, MN6/7-Ib and MNSN b/d-Is

(Hoang and Chiba, 2001). The branches of these two motor neurons come together to form a

single arbor with well characterized shape and size. Three classes of motor neurons can be

found in the fly, type I-III, which are classified based on the size of the boutons, anatomy of

their arbors and their function in innervation. Type I motor neurons innervate the body wall and

regulate contractions (Johansen et al., 1989). A single axon has the capability to innervate more

than one muscle, such as the motor neurons mentioned above which innervate both muscles 6

10

and 7. Ib and Is boutons differ in their size and degree of neurotransmission with Ib boutons

being slightly larger but generating a weaker synaptic response compared to Is (Kurdyak et al.,

1994).

NMJs are a unique type of chemical synapse formed by the apposition of a motor neuron and a

muscle fibre (Levitan and Kaczmarek, 2015). The Drosophila NMJ is glutamatergic, meaning

glutamate is the neurotransmitter held within and released from the synaptic vesicles. Upon

release glutamate binds to glutamate receptors on the membrane of the muscle and causes an

influx of Ca2+ ions which leads to depolarization, activating the muscle. The synaptic proteins

involved in vertebrae synaptic transmission that have been studied at the Drosophila NMJ are

70-80% conserved. Both mammalian central excitatory synapses and Drosophila NMJs utilize

ionotropic glutamate receptors and therefore the NMJ may act as a model to learn about

synaptic function and diseases which may affect the mammalian excitatory synapses

(Keshishian et al., 1996).

At the molecular level, the Drosophila NMJ is made up of two components used to produce

synaptic transmission, the presynaptic AZs and the postsynaptic density (PSD). The NMJ of the

Drosophila larva houses a substantial number of boutons that innervate the muscle (Harris et al.,

2016). Numerous AZs are found on each bouton which are specialized to enable

neurotransmitter (glutamate) release and recycling. Secretory vesicles can be found clustered

with voltage-gated Ca2+ channels at the AZs (Melom et al., 2013). The clustering allows for

locally high concentrations of Ca2+ and swift fusion of vesicles with the membrane. A

cytomatrix exists within the AZ to organize the proteins involved in synaptic assembly and

regulation. The cytomatrix is made up of Ca2+ channels and different scaffolding proteins such

as Brp, DRBP, liprin-α and Syd-1. Trans-synaptic protein partners exist that facilitate synaptic

organization , like Neurexin and Neuroligin (Harris and Littleton, 2015).

In general, the postsynaptic cell is responsible for assembling the proteins that will bind the

released neurotransmitter and facilitate signal transduction. On the postsynaptic cell of many

synapses there exists a submembranous electron dense domain that can be visualized by electron

microscopy. This domain is called the postsynaptic density (PSD) (reviewed in Harris and

Littleton, 2015). The PSD works to receive and integrate signals from the presynaptic cell and

11

transduce them to the postsynaptic cell to regulate synaptic function (Boeckers et al., 2002). At

Drosophila type-I NMJ boutons the PSD is not identifiable by an electron dense domain.

Instead, the membrane has an arrangement of elaborate folds and depressions called the

subsynaptic reticulum (SSR). Embedded in the SSR are the proteins typically found at a PSD

including glutamate receptors (GluR), signalling complexes, ion channels (like L-type Ca2+

channels), adhesion molecules and scaffolding molecules (such as Shank). Another scaffolding

protein at Drosophila PSDs is Discs large (Dlg), the homolog of mammalian PSD-95. In

mammalian neurons it organizes the PSD and regulates GluR trafficking. At the Drosophila

NMJ, Dlg is found throughout the SSR, where it plays a crucial role in the formation of the SSR

and recruits other PSD proteins such as ion channels and adhesion proteins (reviewed in Harris

and Littleton, 2015).

The GluRs found in the membrane of the SSR in Drosophila are ionotropic non-NMDA-type

receptors (reviewed in Harris and Littleton, 2015). They create an excitatory response once

bound to glutamate. These receptors are composed of four subunits which assemble into two

different types depending on which subunit they contain. A-type contains subunit IIA and B-

type contains subunit IIB and both types contain IIC, IID and IIE as their remaining three

subunits. Subunits IIC, IID and IIE are required for receptor formation and function (Schmid et

al., 2006) whereas subunits IIA and IIB are interchangeable but each imparts distinct synaptic

properties (DiAntonio, 2006). Null allele work has demonstrated that GluRs play an important

role in the excitability of larval NMJs and in synapse maturation (Schmid et al., 2006). Mutants

with either IIA or IIB removed are viable but do display deficits and mutants with both removed

are embryonic lethal (Peterson et al., 1997). Likewise, removal of any one of the IIC-IIE

subunits causes embryonic lethality (Featherstone et al., 2005; Qin et al., 2005).

Also present at the PSD in vertebrates and Drosophila is a spectrin skeleton made up of a lattice

network of α- and β-spectrin subunit heterotetramers (Pielage et al., 2006). These form a

network with actin filaments that underlies the plasma membrane, and in Drosophila repeats of

the lattice correspond to the size of a single AZ, suggesting that the spectrin lattice may play a

direct role in AZ organization. In the vertebrate system spectrin helps organize ion channels and

adhesion molecules into domains and is thought to play a role in clustering of neurotransmitter

12

receptors. In Drosophila α- and β- spectrin are required for specification of AZ size, spacing

and function during development. Null mutations of either spectrin subunit are lethal at

embryonic or early larval stages. Experimentally circumventing this lethality demonstrated that

removal of postsynaptic spectrin subunits contributed to a thinner SSR that was no longer

tightly wrapped around the bouton (Pielage et al., 2006).

The plasticity of the Drosophila NMJ is another feature that makes it a prominent model

synapse. There are a few ways in which the NMJ can be plastic, including altering its synaptic

connections, altering its size or complexity of boutons (Keshishian et al., 1996). This plasticity

is regulated using several mechanisms including 1) regulation of excitability, 2) anterograde,

retrograde and autocrine signalling between the pre- and postsynaptic cell and 3) signal

regulation by the extracellular matrix, cytoskeleton and vesicle trafficking pathways. Changes to

neuronal activity alter growth of the NMJ resulting in an increase in total bouton number (Harris

and Littleton, 2015). Additional boutons can form either when new boutons bud from mature

boutons or when novel boutons form from an existing branch of boutons or between two

boutons (Zito et al., 1999). Sometimes in response to elevated neuronal activity rapid bouton

budding can occur resulting in structures called ghost boutons. Ghost boutons are not fully

formed and therefore are incapable of neurotransmission because they lack the presynaptic

machinery and the postsynaptic apparatus (Atman et al., 2008; Piccioli and Littleton, 2014).

13

1.3 Shank

As mentioned above, the PSD plays a crucial role in the reception of incoming synaptic signals.

It is composed of several components such as GluRs, signalling complexes, ion channels,

adhesion molecules and scaffolding molecules such as Shank. Each component has a unique but

necessary function in this area to allow transduction of signals to the postsynaptic cell.

The Shank family of proteins are highly conserved scaffolding proteins that are important for

synaptic function. The vast majority of Shank research thus far has been conducted in mouse

models. In mice and humans, there are three members of the Shank family: Shank1, Shank2,

and Shank3 (Sheng and Kim, 2000). Both the mammalian and Drosophila Shank proteins are

large proteins with a molecular weight of approximately 200kDa (Sheng and Kim, 2000; Harris

et al., 2016). Within the postsynaptic cell Shank is uniformly enriched across the PSD but

concentrated in the deeper parts overlapping and below PSD-95 (Dlg). This was demonstrated in

both mammalian and Drosophila models (Sheng and Kim, 2000; Harris et al., 2016). When the

mammalian synapse is developing Shank is found in the growth cones or axons and dendrites

before concentrating at the PSD of the synapse between postnatal day 6-10 (Du et al., 1998;

Naisbitt et al., 1999).

All three mammalian Shanks are highly conserved with one another, each displaying the same

protein binding domain structure. The Shank3 binding motifs (from N- to C-terminal) include 5-

6 N terminal ankyrin (ANK) repeats, a Src homology 3 (SH3) domain, a PSD-95/discs

large/zonula occludens-1 (PDZ) domain, a region rich in proline and serine and a sterile alpha

motif (SAM) domain (Sheng and Kim, 2000; Boeckers et al., 2002; Jiang and Ehlers, 2013). In

mice and rats, the ANK, SH3 and PDZ domains have been shown to each bind several

functionally important proteins (Jiang and Ehlers, 2013). The SAM domain of Shank

multimerizes Shank in a tail-to-tail fashion, allowing for aggregation of multiple sets of proteins

(Sheng and Kim, 2000). The Drosophila Shank gene is highly conserved with all three

mammalian Shank genes and the proteins they encode have numerous conserved protein binding

motifs (Harris et al., 2016). Studies of Drosophila Shank have shown that it plays a comparable

role at the NMJ as seen in rodent models of Shank both in vivo and in vitro. This similarity is

14

likely due to its protein-protein interaction motifs which are conserved from invertebrates to

humans.

More than 30 different postsynaptic proteins have been shown to interact with Shank in

mammals including GluR, cell adhesion molecules, ion channels, cytoskeletal proteins,

scaffolding proteins and proteins involved in signalling cascades (Boeckers et al., 2002;

Kreienkamp, 2008; Grabruker et al., 2011). At the PSD Shank3 forms large sheets that act as a

backbone to organize the PSD complex and the proteins that bind its domains (Moessner et al.,

2007). Each Shank binds to multiple different scaffolding proteins such as GKAP (Vulcan),

Homer and Grip giving it the title of “master scaffolder”. By interacting with the above proteins

mammalian Shank indirectly interacts with NMDA receptors, metabotropic GluR (mGluR) and

AMPA receptors, three major classes of postsynaptic glutamate receptors. These interactions

may facilitate crosstalk between ionotropic (NMDA) and metabotropic (mGluR) signalling

pathways (Sheng and Kim, 2000). Overall, Shank plays a major regulatory role at the synapse

(Jiang and Ehlers, 2013). It regulates the actin cytoskeleton, abundance, and signalling of

ionotropic glutamate receptors and the formation, organization, transmission, and plasticity of

the synapse (Grabruker et al., 2011; Jiang and Ehlers, 2013).

15

Table 1. Mouse Shank interactors and their respective Drosophila homologs Binding domain

Mouse Shank interactor

Drosophila homolog

Homolog function References

Ankyrin repeats

α-Fodrin α-spectrin • Interacts with β-spectrin • Transports fat to larval fat body for storage • Connects Shank to membrane-associated cytoskeleton*

(Boeckers et al., 2002; Gramates et al., 2017)

SH3 Densin-180

Scribbled • Scaffolding protein that regulates apicobasal polarity • Interacts with Dlg • Organizes synaptic architecture

(Gramates et al., 2017)

SH3 GRIP1 Grip (Glutamate receptor binding protein)

• Scaffolding and signalling protein • Directs developing muscle • Scaffold protein for AMPA receptors and Eph receptors/ligands*

(Sheng and Kim, 2000; Gramates et al., 2017)

SH3 Cav1.3 Ca-α1d (Ca2+-channel protein α1 subunit D)

• α subunit of L-type voltage-gated Ca2+ channel in neurons • Mediates Ca2+ influx

(Gramates et al., 2017)

PDZ domain

ProSAPip1 CG15365 • Not well characterized

PDZ domain

β-Pix rtGEF (Rho-type guanine nucleotide exchange factor)

• Regulates postsynaptic structure and muscle development, protein localization and epithelium growth

(Gramates et al., 2017)

PDZ domain

PSD-95 Dlg1 (Discs large 1)

• Scaffolding protein • Recruits PSD proteins • Role in SSR formation • Interact with NMDA receptor*

(Boeckers et al., 2002; Harris and Littleton, 2015)

PDZ domain

GKAP1/3 Vulcan • Recruits Shank to postsynaptic sites* • Couples with NMDA receptors via MAGUK proteins*

(Boeckers et al., 2002)

Proline-rich region

Homer1 Homer • Links mGluR to other intracellular signalling proteins • Role in assembling excitation-Ca2+ coupling signaling complexes*

(Sheng and Kim, 2000; Gramates et al., 2017)

Proline-rich region

Cortactin Cortactin • Regulates actin cytoskeleton organization and controls rearrangement in response to stimulus

(Gramates et al., 2017)

* = Mouse protein function

16

All three members of the mammalian Shank family can be found in the rat brain and somewhat

in other areas except Shank1 which is brain specific (Lim et al., 1999). For example, Shank2 can

be found in the kidney and liver and Shank3 can be found in the heart and spleen (Du et al.,

1998; Lim et al., 1999). Within the brain, the Shank proteins have distinct but partially

overlapping expression patterns. The Shank1 protein can be found mostly in the cortex,

hippocampus, and amygdala and less so in the thalamus and substantia nigra. Shank 2 is found

in many brain regions including the cortex, hippocampus, cerebellum, olfactory bulb and central

gray. Shank3 localization overlaps with Shank 2 in the cortex and hippocampus but they are

expressed in different cells in the cerebellum. Shank2 is expressed in the Purkinje cells and

Shank3 is expressed in the granule cell layer (Boeckers et al., 2002). From immunoreactivity

work, Shank was found in a punctate pattern at excitatory synapses but not inhibitory synapses

(Naisbitt et al., 1999). Shank was only found at the synapses and not in the cell bodies or

dendrites of neurons (Lim et al., 1999; Sheng and Kim, 2000).

Mutations in the Shank genes produce major defects in synapse morphology. Generally,

mutations in Shank3 contribute to loss of dendritic spines, reduced spine volume, and a thinner

PSD. These features combined generate a phenotype of impaired synapse maturation and

function. Shank mutants also exhibit abnormal social behaviour in mice. Mouse behavioural

studies have found defects in Shank3 contribute to reduced social interaction and affiliation

behaviours, and reduced performance in learning and memory tasks (Jiang and Ehlers, 2013).

Harris et al. (2016) used transgenics and null mutants to characterize the single homolog of

Shank in Drosophila. The function of Shank in the postsynaptic compartment at the NMJ was

shown to be dose-dependent, with an optimal level of Shank required for normal synaptic

development to occur. When Shank levels were increased or decreased beyond this optimal

level, the same adverse phenotypes were observed: 1) a decrease in synaptic bouton number, 2)

an increase in the number of immature boutons, called “ghost boutons” (GB), and 3) disruption

of the SSR (Harris et al., 2016). These phenotypes indicate defects in the development and

maturity of the NMJ.

17

Another study by Wu et al. (2017) generated a different Shank null mutant which had an 8210

bp deletion, including exons which encode amino acids 57-1871. They found that Shank

localized in the axons, including the neutropil of the CNS, and not the postsynaptic cell. In these

null mutants, no defects were seen at the NMJ, but defects were found in calyx boutons and in

olfactory responses mediated by the calyx. The defects in olfactory acuity could be partially

rescued by presynaptic Shank expression implying a presynaptic role for Shank (Wu et al.,

2017).

Overexpression of Shank can be created by utilizing the UAS/GAL4 system mentioned

previously. When the strong muscle driver mef2-GAL4 was used to overexpress UAS-Shank in

the postsynaptic cell at the NMJ, Shank levels were elevated 7 fold (+ 1) above control levels.

Consequently, bouton number was reduced by 29% and ghost bouton number increased 6-fold.

When a more moderate muscle driver (24B-GAL4) was used in this system the bouton number

was reduced by 21% but no significant increase in ghost boutons was seen (Harris et al., 2016).

Shank levels can be reduced through the use of a null allele generated by Harris et al. (2016).

This null mutant carries a deletion from the middle of the first intron to the 3’ untranslated

region (UTR) removing 97% of the Shank gene’s coding region. The ShankD101 null mutants

exhibited a 24% reduction in bouton number and a 4-fold increase in ghost boutons. Animals

heterozygous for ShankD101 exhibited more mild defects including 15% reduction in bouton

number with no appearance of ghost boutons, demonstrating that loss of a single copy of Shank

is enough to produce defects. Therefore, the degree of morphological defects at the NMJ

depends on the level of Shank expression, with ShankD101 and mef2 Shank animals producing the

most severe phenotypes. When Shank expression was driven presynaptically using a neuronal

driver (C155-GAL4), no increase in Shank levels were seen at the NMJ and no adverse

phenotype was generated suggesting that Shank acts in the postsynaptic cell. In regards to SSR

disruption, the overall area was unchanged but the SSR had fewer infoldings and made fewer

connections with the presynaptic terminal when Shank levels were altered (Harris et al., 2016).

The SSR is where synaptic components like scaffolding proteins, adhesion molecules, and

glutamate receptors localize near the synaptic cleft (Johansen et al., 1989). Consequently, flaws

in the SSR can affect the assembly and regulation of the synaptic signalling apparatus (Johansen

18

et al., 1989). Hence, altered levels of Shank produce both presynaptic (altered bouton number

and ghost boutons) and postsynaptic (disrupted SSR) abnormalities.

That same paper by Harris et al. (2016) also showed a role for Shank in regulating the

internalization of Frizzled-2 (Fz2) as part of the Fz2 nuclear import (FNI) signaling pathway. In

this pathway, Wnt1 is secreted by the presynaptic cell and binds to the Fz2 receptor embedded

in the postsynaptic membrane. The receptor is internalized then cleaved and the fragment enters

the nucleus and interacts with RNA binding proteins. Both overexpression and loss of Shank

lead to a deficiency of the Fz2 fragments in the nucleus. Impairment of the Wnt FNI signaling

pathway was associated with the excess GB phenotype seen in Shank mutants but was not found

to contribute to other Shank phenotypes. Thus, it is not known which pathways are affected to

create the altered bouton number and SSR phenotypes.

19

1.4 Ca-α1d

Ca2+ channels can be found conserved in a wide range of species and play a role in many crucial

processes such as membrane excitability, synaptic transmission, and differentiation (Tsien et al.,

1988). In response to a depolarization event, voltage-gated Ca2+ channels facilitate the influx of

extracellular Ca2+ into the muscle cell cytosol where it can influence different processes (Chorna

and Hasan, 2012). Voltage-gated Ca2+ channels display the same heteroligomeric configuration

consisting of several proteins designated as α1, α2, β, δ and γ subunits (Catterall, 1991). The α1

subunit is the pore-forming ion selective subunit and the others are responsible for regulating the

channel’s function (Dolphin, 2012). Both the Drosophila and vertebrate α1 subunits possess

four repeat domains (I-IV), each consisting of 6 hydrophobic domains (1-6) that span the

membrane. These are arranged so all the positively charged side chains are on the same side of

an α-helix to act as the voltage sensor (Stühmer et al., 1989).

The mammalian α1 subunit genes are grouped into 3 families (Cav1, Cav2, and Cav3). The same

can be seen in Drosophila where the genome encodes three α1 subunits (Dmca1D, Dmca1A,

and Ca-α1T) which are classified as Cav1-, Cav2-, and Cav3-type channels, respectively (Eberl et

al., 1998; Ren et al., 1998).

The Dmca1D channel hereby referred to as Ca-α1d, and the Dmca1A channel, hereby referred

to as Cacophony (Cac), are present on opposing sides of the synapse. Ca-α1d regulates inward

currents in the larval body wall muscles (Ren et al., 1998) and Cac is found in the presynaptic

neuron where it participates in neurotransmitter release (Worrell and Levine, 2008). Ca-α1d is

encoded by the 1(2)35Fa gene found on the left arm of chromosome 2 and Cac is encoded by

the cac gene on the X chromosome (Eberl et al., 1998; Chorna and Hasan, 2012). The Ca-α1d

protein is strongly expressed in both the developing and adult nervous system (Eberl et al.,

1998) as well as the adult muscles (Chorna and Hasan, 2012). However, it also plays numerous

roles all over the organism such as in the heart and gut (Eberl et al., 1998; MacPherson et al.,

2001). Both channels are also found in the Malpighian tubules where they play a role in

epithelial fluid transport (MacPherson et al., 2001). Null alleles of either Dmca1D, cac or the

20

accessory subunits (α2-γ) cause embryonic lethality (Chorna and Hasan, 2012). Therefore

Dmca1D and cac are not functionally redundant (Eberl et al., 1998).

Due to the importance of Ca2+ signaling to development and physiological functions, mutations

in genes related to Ca2+ signalling, such as Ca2+ channels, are often homozygous lethal early in

development (Chorna and Hasan, 2012). Two null alleles of Drosophila Ca-α1d, Ca-α1d[X10],

Ca-α1d[X7], cause embryonic lethality (Eberl et al., 1998). The X10 allele contains a pre-

mature stop codon which produces a shortened protein, missing the last 2 transmembrane

domains and its carboxytail, ultimately forming a non-functional channel. The specific mutation

of the X7 allele thus far is undefined but it produces the same severity of mutation as X10

resulting in late embryonic death. The phenotype that is observed before death includes trachea

that do not fill with gas, no heartbeat and slow to nonexistent movement. The embryos are

unable to hatch because of the absence of movement, which likely requires L-type Ca2+

channels, and therefore death is the outcome (Eberl et al., 1998).

The Ca-α1d[AR66] allele is a hypomorphic missense (point) mutation in the Ca-α1d gene

which causes most animals to die as pupae with a few that make it to adulthood. Animals

possessing the AR66 allele are able to hatch as larvae demonstrating the protein retains partial

function after mutation. This point mutation substitutes the wild-type cysteine codon (TGT) to a

tyrosine (TAT). The mutation is found within a transmembrane domain involved in determining

the rate of channel activation and peak current. This mutation does not cause gross disruption of

nervous system development but does delay overall development by 1-2 days. AR66 mutants

are indistinguishable from wild-type siblings until the late pupal stage. About 50% manage to

eclose but their wings do not expand and they die in the food. The other 50% manage to open

the puparium but cannot exit the pupal case and die. Therefore, the AR66 mutant Ca2+ channel is

not sufficient to fulfill the adult’s Ca2+ signaling requirements provided by Ca-α1d. All three

mutants displayed no abnormalities in their embryonic nervous system. Based on the absence of

muscle contractions in the pharate larva the defect caused by the mutant alleles is likely related

to muscle physiological and not to the nervous system (Eberl et al., 1998).

21

When the Drosophila Ca-α1d subunit was compared to the α1 subunits of different vertebrate

Ca2+ channels the similarity between their amino acid sequences ranged from 63.4-78.3% with

Ca-α1d being most similar to the α1 subunit of Cav1.3, an L-type Ca2+ channel (Zheng et al.,

1995). Like L-type vertebrate Ca2+ channels, Ca-α1d channels are dihydropyridine (DHP)

sensitive (Eberl et al., 1998; Chorna and Hasan, 2012).

Cav1.3 has been shown to interact with Shank3. The C-terminus of Cav1.3 is rich in proline and

contains five SH3 domain-binding sites. Cav1.3 C-terminus was shown to bind to both the SH3

and PDZ domain of Shank. This binding facilitates Ca2+ channel clustering at the synapse in rat

hippocampal neurons in vitro (Zhang et al., 2005). However, it has not been determined if this

interaction also occurs between the Drosophila homolog of Cav1.3, Ca-α1d, and Drosophila

Shank.

22

1.5 Autism Spectrum Disorders (ASD)

Autism is a developmental neuropsychiatric syndrome with a range of symptoms often

identifiable before the age of three (Moessner et al., 2007; Geschwind, 2011). Autism case

studies were first observed and described by L. Kanner in 1943. Since then, the term autism has

expanded to autism spectrum disorders (ASDs) and encompasses other disorders including

autism, Asperger syndrome, pervasive developmental disorder (not otherwise specified) and

childhood disintegrative disorder. ASD patients are a population made up of individuals that

vary clinically in regards to disruption in their cognition and behaviour rather than individuals

with a distinct clinical disorder (Geschwind, 2011). There are three characteristics that form the

foundation of an autism diagnosis, including impaired reciprocal social interactions, impaired

communication abilities and restricted behaviour and interests (Moessner et al., 2007), though

the clinical presentation of these deficiencies greatly varies in human patients (Jiang and Ehlers,

2013). These phenotypes may appear in two distinct manners: onset may be gradual, where

delayed development becomes apparent over time, or development may progress normally until

regression begins (Zwaigenbaum, 2001; Werner et al., 2005; Martinez-Pedraza and Carter,

2009).

The National Autism Spectrum Disorder Surveillance System estimates the prevalence of

autism is approximately 1 in 66 Canadian children (Canada, 2018). The prevalence is also

heavily male biased, with approximately four males diagnosed for every female (Werling and

Geschwind, 2013). This ratio is consistently seen across time and population. The presentation

of symptoms also differs between males and females which may play a role in their differing

levels of diagnosis (Werling and Geschwind, 2013). Males typically show more outward

behavioral issues, like aggressive behaviour, hyperactivity, repetitive behaviours and interests,

and reduced prosocial behaviours, whereas females display more internal behavioural issues,

such as anxiety, depression and other emotion-based issues (Mandy et al., 2012; Solomon et al.,

2012). The more disruptive nature of the male presentation of autism may be contributing to the

increased level of diagnosis of males. Interestingly, the ratio of 4:1 changes when you examine

it within different severities of the disorder. For instance, in patients with low intellectual

disability, the ratio is six boys for every one girl whereas with moderate to severe disability the

23

ratio is closer to 1.7 to 1 (Fombonne, 1999). Higher levels of intellectual disability in females

may be the motivating factor to seek diagnosis. Differences in genetics or hormone levels could

also be contributing to this ratio. There is mounting evidence for heritable loci with differing

penetrance based on sex as well as hypotheses about excess fetal testosterone during

development contributing to autism (reviewed in Werling and Geschwind, 2013). There is no

dependable internal or external marker that can predict or confirm ASDs. Diagnosis of an ASD

is evidence-based as the disorder is extremely heterogeneous (Wang et al., 2011). Evaluation is

necessary to confirm diagnoses, disregard similar conditions, identify comorbidity if any and

determine the severity of impairment (Sanchack and Thomas, 2016).

ASDs are caused by an interplay of genetic and environmental influences (Geschwind, 2011).

Until rather recently, not much was known about the neurobiological basis underlying ASDs.

Based on twin and family studies the heritability of ASD is estimated to be around 90%

(Moessner et al., 2007). However, no one gene or mutation accounts for the majority of ASD

cases, the most common genetic causes only accounting for 1-2% of cases. Furthermore, these

genes are linked to a range of cellular mechanisms, including cell adhesion, synaptic vesicle

release, neurotransmission, synaptic structure, RNA processing to protein translation

(Geschwind, 2011). On the other hand, several environmental factors contribute to an increased

risk of developing an ASD. These factors include gestational diabetes, maternal bleeding during

pregnancy, medications taken during pregnancy, maternal infection, preterm birth and low birth

weight. Exposure to some medications and synthetic chemicals in utero has been implicated in

an increased risk of developing ASDs, such as exposure to valproate, certain antidepressants, or

organophosphates during the first trimester. Also, an immune response triggered by a maternal

infection during gestation can disrupt fetal brain development. Above is just a brief list of some

of the environmental factors that can play an additive or multiplicative role in increasing the risk

of a child born with an ASD.

24

1.6 Shank and ASD

More than 1000 genes contribute to the risk of ASDs, and even the genes most commonly

linked to ASDs only account for 1-2% of cases (Betancur, 2011). Shank is one of the most

common monogenic causes of ASDs (Durand et al., 2007; Moessner et al., 2007), with

haploinsufficiency of SHANK3 considered one of the most predominant mechanisms underlying

pathogenesis (Betancur and Buxbaum, 2013; Jiang and Ehlers, 2013). Mutations in both

SHANK1 and SHANK2 have also been linked to ASD (Wang et al., 2011; Sato et al., 2012).

Evidence from human case studies has demonstrated that altered gene dosage of SHANK3

contributes to ASDs as well as severe cognitive deficits, particularly related to language and

speech (Durand et al., 2007). Several groups have looked into the genetics behind specific ASD

cases and many have found alterations, such as frameshifts or deletions, in SHANK3 are present

(Durand et al., 2007; Moessner et al., 2007). Chromosomal rearrangements (deletions and

duplications) occurs in 3-6% of cases of ASDs. The SHANK3 gene is found at 22q13.3 and

deletions in this area are associated with Phelan-McDermid syndrome. This syndrome is

characterized by newborns with low muscle mass, overall developmental delay, possible

accelerated growth, delayed speech, autistic behavior and dysmorphic features (Durand et al.,

2007). Overexpression of SHANK3 may also result in ASD as demonstrated in both mice and

human studies (Bozdagi et al., 2010). Large duplications of the SHANK3 genomic region in

humans is implicated in a variety of neuropsychiatric disorders, such as attention deficit

hyperactivity disorder (ADHD), schizophrenia, and ASDs (Durand et al., 2007; Failla et al.,

2007; Moessner et al., 2007). Similarly, mice with Shank3 duplications display abnormal

behaviors associated with ASDs such as manic behavior, convulsions, and defects in neuronal

excitatory/inhibitory balance (Han et al., 2013). Therefore, it appears conservation between

species exists where an optimal dose of Shank is required for normal synapse function.

Two separate mouse studies, one by Bozdagi et al. (2010) and the other by Wang et al. (2011),

found similar results when they deleted different portions of the Shank3 gene. Bozdagi et al.

disrupted Shank at the exon coding for the ankyrin repeats and Wang et al. disrupted Shank at

exons 4-9. They both found that deletions in the Shank gene led to decreased or abnormal social

interaction and communication, and a decrease in synaptic plasticity, specifically related to

25

dendritic spine remodelling. Bozdagi et al. also found a reduction in both glutamatergic synaptic

transmission as well as AMPA receptor-mediated transmission (2010). They also observed an

increase in presynaptic release which may be an attempt to compensate for the lower

transmission. Wang et al. also found mice with Shank3 deletions displayed abnormal motor

behaviours, decreased learning and memory abilities, more repetitive behaviours, and altered

protein composition at the PSD, namely, reduced GKAP and Homer1 levels (2011).

The relationship between ASDs and Shank demonstrates a tangible link between the

pathophysiology of ASDs and synaptic dysfunction (Jiang and Ehlers, 2013). However,

different mutations in Shank could be acting through different cellular mechanisms to alter

protein-protein interaction to cause synaptic dysfunction and lead to the varied clinical

presentation of ASDs (Jiang and Ehlers, 2013). Redundancy between the 3 Shank genes in

mammals has made it challenging to generate full knock-outs to analyze Shank function, and

different published mutations in Shank have variable outcomes on synaptic function and

behaviour (Jiang and Ehlers, 2013). Drosophila is an advantageous model as the fly genome

encodes only a single member of the Shank family. By knocking out this single Shank gene, we

may be able to better understand the underlying molecular deficits that arise from loss of Shank.

Fly models are key to understanding the deficits responsible for these heterogeneous disorders.

26

1.7 Thesis aims and hypothesis

Both loss or overexpression of Shank at the Drosophila NMJ leads to a decrease in the number

of synaptic boutons, indicating a defect in synaptic growth. The purpose of this project is to

explore the mechanism of how overexpression of Shank affects synaptic growth through a

genetic screen for Shank interactors. Using RNAi, I knocked down the expression of the

Drosophila homologs of proteins known to interact with Shank in mammals, in order to

determine if we can modify the NMJ defects in flies overexpressing Shank by changing the

expression of putative interactors. This screen will allow us to identify novel proteins which

may interact with Drosophila Shank directly or indirectly. Further, we can characterize new

molecular pathways to understand how Shank functions at synapses and potentially find a

pathway related to Shank which explains the lowered bouton phenotype seen in Shank-altered

flies. Aim 1 involved conducting a genetic screen to identify novel Shank interactors. To

accomplish this, RNAi lines were used to determine which of the candidate genes produce a

synergistic effect when knocked down in combination with Shank overexpression. Aim 2

involved in-depth analysis of one candidate gene, Ca-α1d, which emerged from preliminary

results of the screen as a likely Shank interactor. To accomplish this, null mutants of Shank and

loss of function mutants for the candidate were used to create double heterozygous and

homozygous mutants to further test for genetic interactions at the NMJ. We also investigated

whether Shank and Ca-α1d colocalized at synapses and/or affected each others’ localization to

the postsynaptic membrane via immunohistochemistry. The ultimate purpose is to contribute

some knowledge about Drosophila Shank, a crucial yet unappreciated PSD protein known to be

associated with ASDs.

27

Chapter 2 – Material and Methods

2.1 Fly stocks and strains

All Drosophila melanogaster stocks were raised on standard media at room temperature (21-

22oC).

The white (w-) line was used as the wild-type control genotype for screen experiments.

A recombinant line (UAS-Shank-GFP,mef2-GAL4/TM6,Tb,Sb) was created which utilizes the

GAL4/UAS system to drive overexpression of the Shank gene. To do this a pre-existing line

which causes Shank overexpression (UAS-Shank-GFP) (Harris et al., 2016) was crossed to the

mef2-GAL4 drive. Progeny of this cross (UAS-Shank-GFP/mef2-GAL4) were subsequently

crossed to a balancer line (TM6,Tb,Sb). Recombination was recognized if larva expressed GFP.

Three individual male flies exhibiting GFP and “tubby” (Tb) phenotype (balancer) were selected

and crossed to TM6,Tb,Sb to generate stable stocks. Third instar larvae were selected from each

stock, dissected and examined more closely for GFP expression using confocal microscopy. The

line displaying the best expression was selected and expanded.

All crosses producing larva to be dissected were raised on standard media at 25oC. For every

cross, the number of males and females per vial was kept consistent. Each cross consisted of 10

virgin (unmated) female Drosophila and 5 male Drosophila.

Control crosses for the screen were as follows:

1) UAS-Shank-GFP,mef2-GAL4 x w-

2) w-

3) UAS-Shank-GFP x w-

4) mef2-GAL4 x w-

5) UAS-nsyb-RNAi x UAS-Shank-GFP,mef2-GAL4

6) UAS-nsyb-RNAi x mef2-GAL4

28

nsyb-RNAi was selected as a control as nsyb is involved in mediating neurotransmission at

presynaptic terminals (Deitcher et al, 1998) and is expected to have no effect when knocked

down postsynaptically.

Experimental crosses involved crossing obtained RNAi lines first to mef2-GAL4 to examine the

impact of the RNAi on its own. Next, the same RNAi lines were also crossed to the above

recombinant line (UAS-Shank-GFP,mef2-GAL4/TM6,Tb,Sb) to examine the impact of the RNAi

on the neuromuscular junction of flies with Shank overexpression.

29

Table 2. Obtained Bloomington siRNA lines and their stock numbers Drosophila gene (RNAi) Bloomington stock number Label

scribbled 35748 (attP2) scrib(a)

58085 (attP40) scrib(b) α-spectrin 31209 (attP2) α-spec(a)

56932 (atP40) α-spec(b) Grip 41978 (attP2) Grip(a)

40930 (attP40) Grip(b) Ca-α1d 25830 (attP2) Ca-α1d(a)

33413 (attP2) Ca-α1d(b) CG15365 36856 (attP2) CG15365(a) RtGEF 32947 (attP2) RtGEF(a) dlg1 31181 (attP2) dlg(a)

39035 (attP40) dlg(b) vulcan 40925 (attP40) vlc(a) homer 41908 (attP2) homer(a)

56921 (attP40) homer(b) Cortactin 32871 (attP2) Cortactin(a)

Table 3. – Obtained VDR long dsRNA lines and their stock numbers Drosophila gene (RNAi) Vienna Drosophila Resource

center stock (VDR) number Label

CG15365 103369 (II) CG15365(b) RtGEF 100583 (II) RtGEF(b) vulcan 46230 (II) vlc(b)

Table 4. – Obtained Bloomington Ca-α1d mutant lines and their stock number Drosophila Ca-α1d alleles Bloomington stock number Label Ca-α1d[X7] 4275 X7 Ca-α1d[X10] 25141 X10

Table 5. Drosophila Ca-α1d alleles Source Label w;l(2)35FaAR66/cyo, en11 Eberl et al., 1998 AR66

30

The Ca-α1d X10 and AR66 alleles were = separately crossed with the above recombinant line to

remove one copy of Ca-α1d in animals overexpressing Shank (Ca-α1d/+;UAS-Shank-

GFP,mef2-GAL4/+). All three of the above Ca-α1d mutant lines were also used to create double

heterozygotes, where each Ca-α1d allele is crossed with ShankD101 (Ca-α1d/ShankD101) and

compared to the single heterozygotes (Ca-α1d/w- and ShankD101/w-).

31

2.2 Dissection

Larvae were collected at the third instar

(wandering) stage and placed in zero Ca2+

HL3 (Table .6) (adapted from Stewart et al.,

1994) on a magnetic dissection tray.

Magnetic pins were placed posteriorly and

anteriorly in the posterior spiracles and the

head region respectively, to hold the larva in

place dorsal side up. A small incision was

made near the center of the larva between the

two tracheae. One blade of the scissors was

inserted into the incision to create a

longitudinal incision anteriorly and then

posteriorly creating an incision spanning the

entire length of the organism. A horizontal incision was made slightly anterior to the posterior

pin. All organs and fat were then removed using forceps and scissors. Another horizontal

incision was made slightly posterior to the anterior pin. The four corners created by the

horizontal incisions were pinned down using magnetic pins to expose body wall muscles 6/7 in

abdominal segment 3.

Dissected larvae in zero Ca2+ HL3 solution were

fixed by removing the HL3 from dissection tray and

adding 4% formaldehyde solution diluted in zero

Ca2+ HL3 for 15 minutes. The dissections were then

washed twice with phosphate buffer (PBS) 10X

plus 0.1% Triton X-100 (PBT). Dissections were

transferred into a 1.5mL microcentrifuge tube

containing PBT to be stored in the fridge until

staining.

Table 6. Composition of haemolymph-like

saline solution (HL3)

Reagent Mass (mg)

NaCl 818

KCl 75

NaHCO3 168

MgCl2 813

Sucrose 7870

Trehalose 375

HEPES acid 238

CaCl+ 0

Reagents dissolved in 200 mL of distilled water

Table 7. Composition of phosphate

buffer (PBS) 10X

Reagent Mass (g)

NaCl 80

KCl 2

Na2HPO4 14.4

KH2PO4 2.4

Reagents dissolved in 1 L of milli-Q

double distilled water

32

2.3 Immunofluorescence

Dissections were washed 3 times for 20 minutes in a PBT-filled microcentrifuge tube on a

rotator. PBT was removed and dissections were blocked in PBT plus 2% normal goat serum

(NGS) for one hour on a rotator. Next, all but 100µL of the block was removed and fresh

blocking solution plus primary antibody were added to the tube containing the dissections and

allowed to incubate overnight in the fridge on a rotator.

The dissections were removed from the fridge and washed 3 more times with PBT for 20

minutes each on a rotator. The secondary antibody was added in a similar fashion to the primary

except it was diluted with PBT. The dissections incubated for 2 hours at room temperature, in

the dark, on a rotator. The dissections were washed for a final 3 times for 10-20 minutes using

PBT, in the dark, on a rotator. The stained larvae were removed from the tube and placed prone

on a glass slide with Vectashield® mounting medium for fluorescence (Vector Laboratories

Inc). A glass coverslip was placed over the slide and held in place using nail polish.

To observe bouton number, the following antibodies were used: anti-Cysteine String Protein

(dCSP) (1:100, mouse) (Developmental Studies Hybridoma Bank); goat anti-horseradish

peroxidase (HRP) conjugated to FITC (1:1000) (ICN Biomedical); Alexa Fluor® 546 goat anti-

mouse IgG (1:500) (Invitrogen). Anti-HRP was added to visualize the Drosophila neuronal

membrane.

To observe the L-type Ca2+ channel Ca-α1d, a rabbit CACH3/CaV1.3 polyclonal, with a biotin

conjugate, was used as the primary antibody (1:1000) (Bioss). This antibody is raised against

the human CaV1.3 α1-subunit. However, it recognizes an epitope that is highly conserved within

the Drosophila homolog of this subunit. The secondary antibody used was Alexa Fluor® 488

goat anti-rabbit IgG (H+L) (1.25:500) (Invitrogen). Accompanying these were the primary

monoclonal antibody anti-discs large (Dlg) 4F3 (1:10,000, mouse) (Parnas et al., 2001) and

secondary antibody Alexa Fluor® 546 goat anti-mouse IgG (1:500) (Invitrogen) used to

visualize the postsynaptic scaffold.

33

To observe Shank localization, a rabbit anti-Shank polyclonal raised against amino acids 51-148

of the Shank peptide was used as the primary antibody (Harris et al., 2016). The secondary

antibody used was Alexa Fluor® 546 goat anti-rabbit IgG (1.25:500) (Invitrogen). Lastly, Anti-

HRP was added to visualize the Drosophila neuronal membrane.

34

2.4 Image acquisition and analysis

Images of the body wall muscles 6/7 third segment were collected using a Nikon D Eclipse C1

confocal microscope (Nikon Instruments) through a 40x/0.95 air objective lens. Images were

taken after adjusting the zoom, gain and offset using the EZ-C1 3.91 imaging software. Z-stack

images were taken and then combined into a single plane using the ImageJ software to permit

analysis.

Bouton counting was automated as described in Schindelin et al. (2012) using a FIJI/ImageJ

plugin (version 2.0.0-rc-32/1.49v). This program required the use of the dCSP antibody in the

red channel to mark boutons (Nijhof et al., 2016). All of the images were reviewed by hand to

ensure the program did not miscount. If the program did in fact miscount, the program count

was discarded and the boutons were counted by hand.

35

2.5 Statistical analysis and sample size

Statistical analysis of the bouton count data was conducted using GraphPad Prism 5 software.

One-way ANOVA tests were performed to determine if there were statistically significant

differences between the bouton number of different experimental groups. A p-value of less than

0.05 was used as the significance level for all experiments. Sample sizes ranged from 8-21

images per experimental group.

36

2.6 RNAi validation – RT-PCR and analysis

2.6.1 Primer verification

RT-PCR was conducted to test the effectiveness

of the RNAi lines (Ca-α1d and Homer) at

knocking down their target gene products. First,

the designed primers (Sigma-Aldrich) were

verified on genomic DNA to ensure they annealed

to the correct sequence and amplified the correct

product. To do this a single OreR fly was frozen

in a PCR tube then mashed with 50µL of

squishing buffer. The mashed fly was incubated at

37oC for 30 minutes then heated to 95oC for 3 minutes. The tube was centrifuged to pellet the

squished fly and the supernatant (DNA template) was moved to a fresh tube.

PCR (Techne Touchgene Gradient) of the DNA template was conducted using the Platinum™

Hot Start PCR Master Mix (2X) (Invitrogen) reagents and protocol. Using the following primers

pairs:

1) Ca-α1d forward primer:

a. Sequence: GCATCGATTCTATGGGCATTGC

b. Melting point: 69.3oC

2) Ca-α1d reverse primer:

a. Sequence: TTGGTACACCCGATACAAGTCG

b. Melting point: 65.7oC

3) Homer forward primer:

a. Sequence: CGAACAACCGATTTTCACCTGCC

b. Melting point: 72.1oC

4) Homer reverse primer:

a. Sequence: GGTCATTGGACTTTTCACGTAGGC

b. Melting point: 68.4oC

Table 8. Composition of squishing buffer

Reagent Volume (µL)

1M Tris-HCl (pH 8) 818

0.5M EDTA 75

5M NaCl 168

Proteinase K (20µg/µL) 813

Reagents dissolved in 486.5 µL of

distilled water

37

Once the PCR was complete the products were analyzed using gel electrophoresis. The gel was

made using 150mL of TAE 1X buffer and 1.5g of agarose powder (BioShop®). Approximately

50mL of the melted gel was poured into the casting tray and 1 µL RedSafe (INtRON

biotechnology) was used to label nucleic acids. Once the gel was cast it was placed in a BIO-

RAD PowerPac Basic™ electrophoresis apparatus and submerged in TAE 1X buffer. To

monitor band length 1 kb DNA Ladder (New England BioLabs Inc.) was used with Gel Loading

Dye Purple (6x) (New England BioLabs Inc.) and

added to the first well. Into each subsequent well,

10 µL of PCR product and 2 µL of loading dye

were added. The voltage was set to 80V and run for

40 minutes. Images of the finished product were

taken on a BIO-RAD Molecular Imager Gel Doc™

XR+ and optimized using the Image Lab™

software.

2.6.2 RT-PCR

After the primers had been validated on genomic DNA, RNA was isolated from Drosophila

tissue using an RNeasy® Mini Kit (50) (Qiagen). In an RNA free environment (tools, supplies,

gloves, counter) RNA was extracted from 3 adult male Drosophila for each line being

investigated, which were OreR, Ca-α1d(a) RNAi, and Homer(b) RNAi. The RNA was

extracted according to the protocol of the RNeasy® Mini Kit. RT-PCR was performed on the

extracted RNA using a SuperScript™ III One-Step RT-PCR System with Platinum™ TaqDNA

Polymerase (Invitrogen) reagents and protocol with the above primer pairs. Along with the RT

reaction, a DNA contamination control was run by omitting the 2x reaction mix and superscript

III RT/Platinum™ TaqMix and adding 12.5 µL PCR master mix. A no template control was

also run by omitting the RNA template from the reaction. A gel was run with RT-PCR reaction

products in the same manner as above, using all the same equipment (2.6.1).

Table 9. Composition of TAE 50X buffer

Reagent Mass or volume

Tris 242 g

Glacial Acetic Acid 57.1 ml

0.5M EDTA (pH 8) 100 ml

Reagents combined in 1 L distilled water

38

2.6.3 Gel image acquisition and analysis

Images of the gel were obtained in the same manner as above (2.6.1). The images of the gel

were transferred from ImageLab to Image J. Image J was used to generate lane profile plots and

to obtain the magnitude of each peak. The magnitude of the plotted peaks of OreR Ca-α1d were

compared to peaks of Ca-α1d(a) RNAi experimental groups to generate an approximate

estimate of gene product knockdown. Peak magnitude indicates the amount of Ca-α1d cDNA

present after RT-PCR.

39

Chapter 3 – Results

3.1 RNAi screen (Phase one)

In order to identify gene products that interact with Shank at the NMJ, I first knocked down

each candidate gene and determined the number of boutons per NMJ. To do this, each UAS-

RNAi line was crossed to the strong driver mef2-GAL4 to drive RNAi expression in the

muscles. This allowed us to evaluate the effect of each knockdown on the NMJ, as well as,

providing a baseline to compare to in the next phase of the screen, in order to determine if RNA

knockdown would modify the Shank overexpression phenotype. After immunohistochemistry

and confocal imaging, the bouton count results can be seen below (Figure 1 and Table 10).

Figure 1. Graph of bouton number for each RNAi construct combined with mef2-GAL4. The square data points indicate a control group and the circle data points indicate an RNAi, mef2-GAL4 cross. Indicated by the asterisks (*) are the significant results. A p-value less than 0.05 was used as the significance level. All other values were not significantly different from the mean value of the mef2/w- genotype. LTHL = lethal, DMG = too damaged to analyze. * p<0.05, *** p<0.001, ANOVA

40

The asterisks (*) indicate a statistically significant change in bouton number when compared to

the bouton number of the mef2-GAL4/w- control group, which was the mef2-GAL4 driver

crossed to w- (mef2-GAL4/+). Two experimental groups exhibited a significantly lowered

bouton count: Ca-α1d(a)/mef2-GAL4 and homer(b)/mef2-GAL4. The mean bouton number +

S.E.M for Ca-α1d(a)/mef2-GAL4 was 60.11 + 4.535 (p=.0003) and for homer(b)/mef2-GAL4 it

was 55.83 + 5.326) (p<.0001) boutons. These lines were compared to the mef2-GAL4/+ control

which had a mean bouton number + S.E.M of 97.33 + 4.396. All mean values measured in the

experiment are listed in Table 10. Interestingly, the second construct of the Ca-α1d RNAi(b),

produced lethality (LTHL) at early larval stages and was exempt from analysis. This may

indicate that between the two Ca-α1d-RNAi constructs, (b) produces a stronger knockdown of

the gene product or had off-target effects. In contrast, only one construct (b) of the homer-RNAi

produced a significant effect on bouton number, while the other construct (a) did not exhibit a

phenotype. These results indicate that Ca-α1d and homer may play an important role at the

NMJ, particularly Ca-α1d as both of its constructs produced results. Two other knockdowns,

Cortactin(a)/mef2-GAL4 and Grip(b)/mef2-GAL4, had musculature that was too easily damaged

(DMG) to be examined properly despite multiple attempts. This may also indicate that they play

an important role at the NMJ or are important for muscle integrity. Several of the experimental

groups as seen in the table below had normal bouton counts but displayed varying degrees of

disturbed morphology such as: α-spec(a & b), CG15365(a), dlg(a), homer(a & b), RtGEF(a),

scrib(a & b), vlc(a & b) (Figure 2). The nsyb-RNAi was used as a control for the whole RNAi

system. The protein is known to be exclusively presynaptic so you would expect postsynaptic

KD using the mef2-GAL4 driver to produce no effect as shown (Figure 1 and table 10).

41

Genotype n Mean SEM Overall morphology

mef2-GAL4/+ 12 97.33 4.396 Normal

Shank-GFP/+ 12 103.60 5.2 Normal

nsyb 10 90.70 4.578 Normal

α-spec(a) 11 108.80 6.471 More branching, highly convoluted shape,

several long thin branches

α-spec(b) 7 95.57 15.63 Less branching, highly convoluted, coiled

shape

Ca-α1d(a) 9 60.11 4.535 Less branching, fewer boutons, bigger bulbous

boutons, smaller overall

Ca-α1d(b) N/A N/A N/A LETHAL

CG15365(a) 10 117.40 7.228 More branching, convoluted, more boutons

CG15365(b) 14 85.07 5.863 Normal

Cortactin(a) N/A N/A N/A DAMAGED

dlg(a) 11 117.2 4.958 More boutons, more branching, disturbed shape

dlg(b) 10 87.30 4.971 Normal

Grip(a) 11 74.91 4.791 Normal

Grip(b) N/A N/A N/A DAMAGED

homer(a) 11 81.91 4.874 Slightly disturbed branching, smaller in length

homer(b) 12 55.83 5.326 Less branching, smaller overall

RtGEF(a) 11 75.45 3.679 Slightly less branching

RtGEF(b) 12 92.50 5.477 Normal

scrib(a) 11 84.09 5.178 Disturbed branch distribution

scrib(b) 9 78.00 7.785 Abnormal branch distribution

vlc(a) 12 86.92 3.817 More and bigger boutons, less branching

vlc(b) 9 82.44 7.762 Slightly smaller length

Table 10. Phase one statistical and morphological results. Shading indicates genotypes that were crossed to w-. Genotypes without shading were all crossed to the mef2-GAL4 driver.

42

Statistical comparisons to control (mef2-GAL4/+)

ANOVA (Dunnet’s Multiple

Comparison Test)

Summary p-value

Shank-GFP/w- ns .9952

nsyb ns .9950

α-spec(a) ns .7985

α-spec(b) ns .9997

Ca-α1d(a) *** .0003

Ca-α1d(b) ns N/A

CG15365(a) ns .1578

CG15365(b) ns .6543

Cortactin(a) ns N/A

dlg(a) ns .1457

dlg(b) ns .9217

Grip(a) ns .0669

Grip(b) ns N/A

homer(a) ns .4263

homer(b) *** <.0001

RtGEF(a) ns .0769

RtGEF(b) ns .9991

scrib(a) ns .6316

scrib(b) ns .2205

vlc(a) ns .8659

vlc(b) ns .5481

Table 11. Phase one level of statistical significance results

43

44

Figure 2. Sample confocal images for each genotype demonstrating the range of morphological changes produced when each RNAi was crossed to the mef2-GAL4 driver. Each image is representative of the average morphological and bouton phenotype produced.

45

3.2 RNAi screen (Phase two)

To discover novel Drosophila Shank genetic interactors, the same RNAi’s were used as above

in combination with Shank overexpression. Animals overexpressing Shank were created by

recombining UAS-Shank and mef2-GAL4. RNAi’s were then crossed one by one to UAS-Shank-

GFP,mef2-GAL4 flies. We counted boutons per NMJ and compared this result to UAS-Shank-

GFP,mef2-GAL4 crossed to w-, which was reduced compared to controls as expected (see

Figure 3) and to RNAi knockdown alone (Figure 3 and table 12). If the addition of Shank

overexpression improved or worsened the impact of the RNAi’s from phase one of the screen

this would suggest that the candidate gene could be a Shank interactor and would be interesting

to study further. The results of this phase of the screen can be seen below (Figure 3 and Table

12).

None of the RNAi’s, when used in a Shank overexpression background, significantly altered

bouton count. However, three of the RNAi’s, Ca-α1d(a), Ca-α1d(b), and α-spectrin(b),

produced lethality at early larval stages indicating a potential genetic interaction between the

candidate gene and Shank. Since α-spectrin knockdown alone produced no bouton phenotype

(data from phase one, figure 1 and table 10), and lethality when combined with Shank

overexpression this could suggest a potential interaction with Shank. Similarly, as the Ca-

α1d(a) knockdown resulted in a decrease in bouton number alone, the observed lethality may

denote further synaptic defect in a Shank overexpression background indicating that Ca-α1d and

Shank might interact. Knockdown of Cortactin(a), Grip(a), Grip(b), and rtGEF(a) gene

products in combination with Shank overexpression caused the body wall musculature to

become fragile and therefore easily damaged during dissection consequently no data could be

collected for these groups. This fragility may suggest an interaction between these gene products

and Shank with respect to the health or integrity of the muscle tissue, though it is unknown at

this time whether Shank plays such a role in muscle biology. The mean bouton number for the

experimental groups can be seen below (Table 12)

46

Figure 3. Graph of bouton number for each RNAi construct used in a Shank overexpression background. The square data points indicate a control group and the circle data points indicate RNAi KD in Shank overexpression background. A p-value less than 0.05 was used as the significance level. All values without an asterisks (*) were not significantly different from the mean value of the UAS-Shank-GFP,mef2-GAL4/w- genotype. LTHL = lethal, DMG = too damaged to analyze. *p<0.05, ANOVA.

47

Genotype n Mean SEM Overall morphology

UAS-Shank-GFP/+ 12 103.6 5.2 Normal

mef2-GAL4/+ 12 97.33 4.396 Normal

UAS-Shank-GFP,mef2-

GAL4/+

15 76.40 4.308 Less branching, fewer boutons,

skinnier branches?

UAS-Shank-GFP,mef2-

GAL4/nsyb

11 59.18 3.266 Less branching, fewer boutons

α-spec(a) 9 85.22 5.587 Less branching

α-spec(b) N/A N/A N/A LETHAL

Ca-α1d(a) N/A N/A N/A LETHAL

Ca-α1d(b) N/A N/A N/A LETHAL

CG15365(a) 20 70.95 4.743 Less branching

CG15365(b) 13 62.38 5.549 Less branching

Cortactin(a) N/A N/A N/A DAMAGED

dlg(a) 8 67.75 4.455 Slightly less branching

dlg(b) 8 75.5 5.782 Slightly less branching

Grip(a) N/A N/A N/A DAMAGED

Grip(b) N/A N/A N/A DAMAGED

homer(a) 12 67.42 8.141 Less branching, thinner arbor

homer(b) 8 62.13 6.526 Less branching

RtGEF(a) N/A N/A N/A DAMAGED

RtGEF(b) 8 61.25 12.26 Less branching

scrib(a) 9 73.78 6.994 Less branching, thin arbor

scrib(b) 17 72.79 4.030 Less branching

vlc(a) 17 79.71 5.784 Less branching, more convoluted

vlc(b) 11 59.18 5.522 Less branching

Table 12. Statistical and morphological results. Shading indicates genotypes that were crossed to w-. Genotypes without shading were all crossed to UAS-Shank-GFP,mef2-GAL4 (to generate Shank overexpression).

48

Statistical comparisons to Shank overexpression (UAS-Shank-GFP,mef2-GAL4/+)

ANOVA (Dunnet’s Multiple Comparison Test) Summary p-value

UAS-Shank-GFP/+ ** .0094

mef2-GAL4/+ ns .0871

nsyb/UAS-Shank-GFP,mef2-GAL4 ns .2576

α-spec(a) ns .9618

α-spec(b) N/A N/A

Ca-α1d(a) N/A N/A

Ca-α1d(b) N/A N/A

CG15365(a) ns .9950

CG15365(b) ns .4617

Cortactin(a) N/A N/A

dlg(a) ns .8942

dlg(b) ns .9999

Grip(a) ns N/A

Grip(b) ns N/A

homer(a) ns .9238

homer(b) ns .6220

RtGEF(a) N/A N/A

RtGEF(b) ns .5471

scrib(a) ns .9996

scrib(b) ns .9991

vlc(a) ns .9994

vlc(b) ns .2703

Table 13. Phase two level of statistical significance results

49

50

Figure 4. Sample confocal images for each genotype demonstrating the range of morphological changes produced when each RNAi is applied in a Shank overexpression background. Each image is representative of the average morphological and bouton phenotype produced.

51

3.3 Ca-α1d, a postsynaptic Ca2+

channel, interacts with Shank

Ca-α1d is the pore-forming ion

selective subunit of a Ca2+ channel that

regulates inward Ca2+ currents in the

larval body wall muscle. Knockdown

of Ca-α1d in muscle resulted in a

significantly lowered bouton count and

disturbed NMJ morphology (Figure 2

and 8), and when combined with Shank

overexpression, resulted in lethality

(Figure 5). Thus, Ca-α1d appears to be

both important at the synapse and to

have a potential relationship with

Shank.

In order to confirm that our Ca-α1d

RNAi tools were knocking down Ca-

α1d transcript as expected, we performed RT-PCR. We designed primers flanking the 7-9th

intron of Ca-α1d in order to detect transcript (579 bp) and distinguish transcript from genomic

DNA (1204 bp). We detected the 579 bp transcript in control (OreR) animals and observed a

decrease in band intensity in Ca-α1d-RNAi animals indicating successful knockdown of the

transcript. Analysis indicated the band intensity decreased by approximately 50%. While RT-

PCR is not a truly quantitative experiment this analysis indicates there is a consistent and

substantial knockdown of product (Figure 6). The OreR template contained some genomic DNA

contamination but this can be disregarded as the band size differs from that of the Ca-α1d

transcript.

Figure 5. Condensed graph of Ca-α1d data from the RNAi line crossed to mef2-GAL4 and recombinant line compared to w- and UAS-Shank-GFP,mef2-GAL4/w- respectively. A p-value less than 0.05 was used as the significance level. LTHL = lethal * p<0.05, *** p<0.001, ANOVA

52

To further elucidate the relationship

between Shank and Ca-α1d three

Drosophila mutant lines were obtained

Ca-α1d[X10], Ca-α1d[X7] and Ca-

α1d[AR66] (Eberl et al., 1998). The

X10 and X7 null mutations are

considered severe and result in late

embryonic death. The AR66 mutation is

much less severe where about half of

the Drosophila die as pupae while the

rest make it to adulthood (Eberl et al.,

1998).

In order to validate and test for the

potential interaction between Shank

and Ca-α1d in a different manner the

three above lines were used to conduct

the following experiments:

1. The Ca-α1d[X10] and AR66

alleles were separately crossed

with the UAS-Shank-

GFP,mef2-GALF4 to remove

one copy of Ca-α1d in animals overexpressing Shank (Ca-α1d/+;UAS-Shank-

GFP,mef2-GAL4/+) (Figure 7).

2. Double heterozygotes were created, where each Ca-α1d mutant was crossed with

ShankD101 (Ca-α1d mutant/ShankD101) and compared to the single heterozygotes (Ca-

α1d mutant/+ and ShankD101/+) (Figure 9).

Figure 6. Image of RT-PCR gel results to demonstrate a decrease of gene product through the use of Ca-α1d(a) RNAi. DNA contamination reactions used Taq without superscript to test for presence of genomic DNA contamination. Used Ca-α1d forwards and reverse primers. Decrease can be seen when comparing lanes B & C to lanes E & F

53

3.3.1 Ca-α1d heterozygotes in Shank

overexpression

To examine the potential relationship

between Ca-α1d and Shank, one copy

of the Ca-α1d gene was removed and

replaced with the mutant allele (X10

or AR66) in a Shank overexpression

background (Ca-α1d mutant/+;UAS-

Shank-GFP,mef2-GAL4). We

observed a statistically significant

decrease in bouton number in Ca-α1d

mutant/+;UAS-Shank-GFP,mef2-

GAL4 compared to UAS-Shank-

GFP,mef2-GAL4 alone, for both the

Ca-α1d[X10] and Ca-α1d[AR66]

allele. Since Ca-α1d[X10]/+ and Ca-

α1d[AR66]/+ heterozygotes do not

have bouton number defects on their own, this result suggests a genetic interaction, consistent

with the potential interaction detected in the RNAi screen.

Genotype n Mean SEM Morphology

UAS-Shank-GFP,mef2-GAL4/+ 15 76.40 4.308 Less branching, fewer

boutons, skinnier branches?

Ca-α1d[X10]/+ 11 90.64 5.013 Normal

AR66/+ 12 97.00 7.165 More branching

Ca-α1d[X10]/UAS-Shank-

GFP,mef2-GAL4

8 46.88 5.439 Less branching

AR66/UAS-Shank-GFP,mef2-GAL4 9 49.78 4.468 Less branching

Figure 7. Graph comparing bouton number of single versus double Shank and Ca-α1d heterozygotes. The square data points indicate a control group and the circle data points indicate a double heterozygote and triangular data points indicate a single heterozygote. All values without an asterisks (*) were not significantly different from the mean value of the UAS-Shank-GFP,mef2-GAL4/+ genotype. * p<0.05, ANOVA.

Table 14. Statistical and morphological results of Ca-α1d heterozygotes in a Shank overexpression background experimentation

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Statistical comparisons to Shank overexpression (UAS-Shank-GFP,mef2-GAL4/+)

Dunnett’s Multiple Comparison Set Summary p-value

Ca-α1d[X10]/UAS-Shank-GFP,mef2-GAL4 ** .0049

AR66/UAS-Shank-GFP,mef2-GAL4 * .0102

Figure 8. Sample confocal images for each genotype demonstrating the NMJ morphology of Ca2+ channel heterozygotes in a wild-type or Shank overexpression background. Each image is representative of the average morphological and bouton phenotype produced.

Table 15. Statistical significance results of Ca-α1d heterozygotes in a Shank overexpression background experimentation

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3.3.2 Double versus single Shank and Ca-α1d heterozygotes

So far, all of our analyses have focused

on the defects that arise when Shank is

overexpressed. However, loss of Shank

produces very similar NMJ defects,

including a decrease in bouton number.

By partially reducing the expression of

Shank and Ca-α1d, a sensitized

background was produced that may

reveal genetic interactions between the

genes. Using the Ca-α1d[X7], [X10] and

[AR66] mutant alleles, and the Shank

null allele D101, we created single and

double heterozygous combinations. All

of the Ca-α1d double heterozygotes (Ca-

α1d mutant/ShankD101) were compared to

ShankD101 single heterozygotes

(ShankD101/+) and the Ca-α1d single

heterozygotes (Ca-α1d mutant/+). If a

worsening of the phenotype (decreased bouton count) was observed in double heterozygotes

compared to either single heterozygote, this would provide more evidence of the genetic

interaction identified in the original screen. However, no significant changes to bouton count

between any of the experimental groups was apparent (Figure 9).

Figure 9. Graph comparing bouton number of Ca-α1d mutant lines in Shank overexpression background. The square data points indicate a control group and the triangular data points indicate a Ca-α1d mutant with wild-type background and circle data points indicate a Ca-α1d mutant with Shank underexpression. All values without an asterisks (*) were not significantly different from the mean value of the ShankD101/+ genotype. * p<0.05, ANOVA.

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Genotype n Mean SEM Morphology

ShankD101/+ 15 79.67 4.747 Normal

Ca-α1d[X7]/+ 13 99.46 4.769 Normal

Ca-α1d[X7]/ShankD101 18 81.72 4.357 Normal

Ca-α1d[X10]/+ 11 90.64 5.013 Normal

Ca-α1d[X10]/ShankD101 21 77.14 3.812 Normal

AR66/+ 12 97.00 7.165 More branching

AR66/ ShankD101 15 90.73 5.257 Normal

Tukey’s Multiple Comparison Test Summary p-value

ShankD101/+ vs Ca-α1d[X7]/ShankD101 ns >.9999

ShankD101/+ vs Ca-α1d[X10]/ShankD101 ns >.9999

ShankD101/+ vs AR66/ ShankD101 ns .7432

Ca-α1d[X7]/+ vs Ca-α1d[X7]/ShankD101 ns .1714

Ca-α1d[X10]/+ vs Ca-α1d[X10]/ShankD101 ns .5364

AR66/+ vs AR66/ ShankD101 ns .9889

Table 16. Ca-α1d and Shank loss-of-function heterozygote statistical and morphological results

Table 17. Ca-α1d and Shank loss-of-function heterozygote statistical significance results

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Figure 10. Sample confocal images for each genotype demonstrating the NMJ morphology of Ca2+ channel and Shank single and double heterozygotes. Each image is representative of the average morphological and bouton phenotype produced.

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3.3.3 Visualization of possible colocalization of Ca-α1d and Shank with immunohistochemistry

Our data indicates that a relationship may exist between Ca-α1d and Shank as decreasing Ca-

α1d, either with RNAi knockdown or with mutant alleles, worsens the Shank overexpression

phenotype. One possible hypothesis regarding this relationship is that Shank overexpression

may partially impair an aspect of Ca-α1d potentially how it is packaged and incorporated into

the plasma membrane or function of Ca-α1d, leading to the apparent synaptic defects. In this

scenario, normal Shank expression would allow for normal functioning or expression of Ca-α1d

but when Shank levels are overexpressed Ca-α1d functions below optimal levels. When Shank

is overexpressed in combination with RNAi knockdown or loss of function mutations of Ca-α1d

the combination would further exacerbate the problem and explain the enhanced phenotype. An

alternative hypothesis is that Ca-α1d normally inhibits Shank levels at the synapse. In this case,

reducing Ca-α1d levels would result in loss of Shank level regulation leading to Shank

overexpression, and would explain the enhanced defects in a Shank overexpression background.

One way to test these hypotheses would be to perform immunostaining of both Shank and Ca-

α1d and see how their distribution patterns change in Shank and Ca-α1d mutant backgrounds.

For Ca-α1d there are currently no tools available to visualize the protein in flies. A

commercially available antibody against CaV1.3, the human homolog of Ca-α1d was used

instead. The amino acid sequence of the epitope against which this antibody was raised is highly

conserved between the two species so we hypothesized that it might bind to the Drosophila

protein. To validate the CaV1.3 antibody, we tested it on animals expressing Ca-α1d RNAi in

different parts of the organism using drivers such as C155 and mef2. The C155 driver is pan-

neuronal, driving expression in all neurons including the motor neurons of the NMJ. Imaging of

these dissections revealed punctate staining throughout the muscle with an apparent

concentration of signal at NMJs. However, no change in the signal was observed with either

neuronal or muscle knockdown of Ca-α1d, indicating the staining was not specific enough for

use in confirming the above hypotheses. Example images can be seen below (Figure 11).

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An antibody against Drosophila Shank (Harris et al, 2016) was used to examine Shank levels in

Ca-α1d knockdown animals compared to controls (mef2-GAL4/+). KD was generated using Ca-

α1d(a) and mef2-GAL4 muscle driver. Based on the above hypothesis it was expected that when

Ca-α1d was knocked down Shank intensity at the boutons would increase compared to controls

or the distribution may change. However, after imaging Shank intensity and distribution

appeared to be the same in both experimental groups. Example images can be seen below

(Figure 12). Therefore, we have no additional evidence at this time to further characterize the

potential relationship between Shank and Ca-α1d.

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A B C

D E F

G H I

Figure 11. Confocal images of Drosophila NMJ with differing Ca-α1d expression to validate use of CaV1.3 antibody. A-C NMJ of control (w-) animal with wild-type channel expression. D-F NMJ of animal with reduced channel expression in the motor neurons. G-I NMJ of animal with reduced channel expression in the muscle. The green channel (A,D,G) indicate staining of the Ca-α1d channel, the red channel (B,E,H) indicates staining of the post-synaptic membrane and C,F,I indicates a composite of the two channels.

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Figure 12. Confocal images of Drosophila NMJ with differing Ca-α1d expression to test interaction hypothesis. A(‘) NMJ of control (mef2-GAL4/+) animal with wild-type channel expression. B(‘) NMJ of animal with reduced channel expression in the muscle. The green channel indicates staining of the neuronal membrane and the red channel indicates Shank distribution. Shank can be seen throughout the muscle with enrichment in the boutons. Shank staining in the nuclei is non-specific (arrows) (Harris et al., 2016)

A A’

B B’

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Chapter 4 – Discussion

The aim of this thesis was twofold. The first goal was to use a genetic screen to identify

potential novel interactors of Shank and observe how overexpressing Shank altered synaptic

growth. This will help us understand how Shank functions at the synapses to characterize a new

molecular pathway which relates to the bouton phenotype displayed by flies with altered levels

of Shank. The second goal was to conduct in-depth analysis of the candidate gene that emerged

from the screen as a likely potential interactor of Shank. Drosophila larval NMJs were used as a

model system to accomplish the above goals. Ca-α1d appeared to be the candidate gene most

likely to have an interaction with Shank. In this thesis, an interaction between Ca-α1d was

confirmed and aspects of said interaction were characterized.

4.1 Interpretation of genetic screen phase one

Phase one of the genetic screen involved separately driving the expression of several RNAi’s in

the muscle to KD the transcript of candidate genes. This was done in order to observe the effect

KD of these specific proteins had on NMJ appearance and bouton count and provided a

reference for future comparison in phase two of the screen.

During phase one of the genetic screen, two candidate genes appeared to be important for

overall NMJ health. KD of Ca-α1d or homer lowered the bouton count per NMJ significantly

below control levels. This ultimately means the motor neuron is making fewer connections with

the postsynaptic muscle and therefore providing less innervation. These genes could be

necessary for NMJ development or interfere in some way with NMJ plasticity. Muscle size

increases drastically between each larval stage consequently motor neuron innervation must

increase as well to ensure proper innervation of the body wall. During development, motor

neurons exhibit plasticity by growing and forming more boutons in response to a demand

determined by the amount of larval movement and activity (Sigrist et al., 2003). If KD of these

genes disrupts the plasticity of the NMJ, the larva would crawl the normal amount signalling

motor neuron growth, but the motor neuron would be unable to respond to the activity signals,

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resulting in smaller than normal NMJs. Alternatively, KD of these genes could affect larval

crawling directly. For example, KD of Ca-α1d reduces the Ca2+ current in the body wall

muscles, limiting the ability to crawl. Plasticity could be normal, but the demand for motor

neuron growth could be diminished due to the absence of crawling, generating smaller NMJs.

These possibilities could explain the lowered bouton count seen in phase one of the screen and

could be explored with live imaging. Another potential explanation for the decrease in bouton

number is that these genes are crucial for NMJ growth during development and when knocked

down the NMJ is not able to form properly. homer, for example, is known to link GluRs to other

signalling proteins (Gramates et al., 2017). Furthermore, GluRs play an essential role in synapse

maturation (Schmid et al., 2006). Without homer, the GluRs may not be linked to the proper

signalling proteins during development and this could explain why the NMJs did not mature

with the proper amount of boutons. During development Ca2+ acts as a second messenger in

several intracellular signalling cascades and axon guidance mechanisms. Specifically, at the

NMJ Ca2+ influx activates two different signaling pathways dependent on Ca2+ that contribute to

the removal of off-target neuromuscular contacts (Vonhoff and Keshishian, 2017). Ca-α1d

could play a role in retrograde pathways during development, where changes in the Ca2+ current

in the muscle enact changes in the presynaptic motor neuron affecting synaptic development. A

few papers have also examined the role Ca-α1d may play in motor neurons, such as aCC and

RP-2, where the expression of this channel was altered using both RNAi and the AR66

mutation. They found that Ca-α1d was responsible for the majority of the Ca2+ currents recorded

from the cell body of these neurons and decreasing its expression reduced the detected current

(Worrell and Levine, 2008).

Phase one also generated one lethal hit when Ca-α1d was knocked down with the (b) construct.

Lethality was apparent at different larval stages. Some larva died as second instars on the sides

of the tube others died as first instar larvae. This could indicate that the (b) Ca-α1d construct is

stronger than (a) and Ca2+ currents could be too low to allow proper movement. This would

cause the first instar to be unable to hatch from the cuticle and second instar larva may not be

able to move to the food and starve on the sides of the tube or couldn’t climb out of the food to

pupate, resulting in death. It is also possible this RNAi may have bound complimentary

sequences and had off-target effects.

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Two experimental groups from phase one were too damaged to be dissected and analyzed:

cortactin and Grip(b). This fragility makes sense when the function of these two proteins is

considered. Cortactin is responsible for regulation of the actin cytoskeleton (Gramates et al.,

2017). When the Cortactin transcript was knocked down in the screen it may have led to the

disorganization of the actin cytoskeleton ultimately lowering the integrity of the muscle and

contributing to the apparent fragility. Similarly, Grip is a scaffolding protein which directs

developing muscle. Its loss may impede proper muscle development again making the animals

too fragile for analysis.

Lastly, phase one also produced several genotypes that had consistent morphological changes

with no statistically significant change in bouton count. These genotypes include: a-spec(a&b),

CG15365(a), dlg(a), homer(a&b), RtGEF(a), scrib(a&b), vlc(a&b).

Assuming the number of AZs are the same as well we cannot be sure if this change in

morphology altered functionality of the protein. Potential follow-up to answer this question

would involve electrophysiology work to see if synaptic function is altered.

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4.2 Interpretation of genetic screen phase two

In phase two of the genetic screen, the same transcripts were knocked down in the muscle but

with the addition of a Shank overexpression background. If the overexpression of Shank greatly

improved or worsened the bouton phenotypes seen from the candidate genes in phase one of the

screen this may suggest a potential interaction between said gene and Shank.

Phase two generated three experimental groups that were lethal. KD of α-spectrin with the (b)

construct and KD of Ca-α1d with either the (a) or (b) construct were lethal at early larval

stages.

For Ca-α1d, phase two showed an enhancement of the phenotype previously seen from phase

one. This may indicate a potential interaction between these two genes. The lethal phenotype

was beyond what was seen in flies with only Shank overexpression or only Ca-α1d KD

indicating this is likely not an additive result. If two genes interact in some way when one is

knocked down or not functioning adequately the other is there to offset this change. However, if

both gene levels are altered this compensation mechanism is removed and a worsened

phenotype is displayed. Mutant Drosophila larvae harboring very small NMJs are known to be

viable to adult stages in some cases (Banovic et al., 2010) so the lethality displayed here is

likely not a result of bouton count being too low but a more complex mechanism, such as a

crucial interaction between Shank and Ca-α1d, the mechanism of which we still do not

understand. Another factor to consider is that the flies with above normal levels of Shank are

viable and reach adulthood. Other than the decreased amount of mature boutons and SSR

disruption, the AZ and GluRs of the synapse remain unaffected in these Shank mutants. This

seems to indicate that the lethality seen in Shank overexpressing flies with Ca-α1d channels

knocked down is likely not an artifact of two seemingly healthy lines coming together and being

tipped over a threshold to generate lethality. Lastly, in a way the other candidate genes from the

genetic screen act as a control against an additive effect being indicated. In the screen, several

genes important to NMJ development or health were knocked down in combination with Shank

overexpression without causing death or in most cases a bouton phenotype. The above evidence

indicates a potential relationship between Shank and Ca-α1d but more follow-up is required.

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The lethal phenotype from α-spec KD is also compelling evidence of a potential interaction

with Shank. α-spectrin in combination with β-spectrin form the spectrin skeleton which is

required for proper synaptic development. The skeleton binds to actin filaments underlying the

plasma membrane and helps to organize the proteins localized there (Pielage et al., 2006). In

phase one of the genetic screen, there was no bouton phenotype seen although morphology was

highly disturbed consistent with disruption of the spectrin skeleton. However, in phase two

when α-spectrin was knocked down in a Shank overexpression background it was lethal. Again

this enhancement of the negative phenotype also appeared to not be an additive effect. This

offers another potential avenue of study into a novel interactor of Drosophila Shank. Two γ-ray

induced α-spectrin mutants already exist, l(3)dre3rg41 and l(3)dre3rg35, which contain premature

stop codons (Lee et al., 1993). The same experimental procedure used in the Ca-α1d

experiments with the X10 and X7 mutants can be applied with the above α-spectrin mutants.

Similar to phase one, the same muscle fragility was seen with cortactin and Grip. Since in both

phases, the body wall muscles were too damaged to analyze it is not possible to interpret if there

is an interaction. Additionally, in phase two KD of RtGEF generated larval body wall muscles

that were too fragile to analyze. Again this makes sense considering the role RtGEF as a

regulator of postsynaptic structure and muscle development (Gramates et al., 2017). Its KD

could be generating disorganized synapses with improperly formed muscle making it quite

delicate as seen above. This fragility generated by KD of RtGEF in a Shank overexpression

background but not in phase one of the screen may suggest an interaction between the gene and

Shank with respect to the health or integrity of the muscle tissue, though it is unknown at this

time whether Shank plays a role in muscle biology. Although it is difficult to interpret whether

this is an additive effect.

Alteration of Shank levels consistently contributes to the same phenotype: decreased bouton

count per NMJ, increased ghost bouton count per NMJ and disruption of the SSR (Harris et al.,

2016). Ghost boutons are incompletely formed bouton buds that never had the postsynaptic

apparatus assemble around them. The Ab used to highlight the boutons and count them in the

genetic screen, dCSP, highlights the synaptic vesicles. Therefore, it is possible that ghost

boutons could have been counted as normal boutons and a ghost bouton phenotype was

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overlooked. For this project only the bouton count phenotype was of concern however in the

future follow-up could be done to more closely look for a ghost bouton phenotype. To do this

double labelling with Dlg and HRP would be performed. HRP labels the presynaptic part of the

bouton and an absence of Dlg staining indicates the bouton is, in fact, a ghost bouton.

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4.3 Interaction between Ca-α1d and Shank is only apparent in a Shank

overexpression background

Based on preliminary results from the screen Ca-α1d was selected as the candidate to follow-up

on as it appeared likely it shared a relationship with Shank. It was seen that KD of Ca-α1d in the

muscle significantly lowered bouton count below control levels. When the KD occurred in a

Shank overexpression background the result was lethality. This enhancement of the phenotype

led me to believe the effect being seen was not additive as discussed above. Several experiments

were conducted to examine this potential interaction with different tools. Two Ca-α1d

homozygous lethal null mutants and one mutant line containing a point mutation that was ~50%

viable were obtained to conduct these experiments.

First, one copy of the wild-type Ca-α1d gene was removed using the X10 and AR66 mutants,

creating heterozygotes, in a Shank overexpression background. The heterozygotes with wild-

type Shank levels showed no bouton phenotype, but there was a statistically significant decrease

in bouton number in Ca-α1d heterozygotes in a Shank overexpression background, suggesting a

genetic interaction with Shank. This confirms what was previously seen in the screen results.

These genes could be interacting in a few ways. At the gene level, one gene can positively or

negatively impact the transcription of another gene. For example, overexpression of the Shank

gene could cause less transcription of the Ca-α1d gene to occur. Alternatively, Ca-α1d could

inhibit transcription of the Shank gene, and when Ca-α1d expression is knocked down with

mutants or RNAi, this could result in Shank overexpression. This same relationship could be

seen at the translation level with Shank inhibiting Ca-α1id translation or Ca-α1d inhibiting

Shank translation. The two proteins could also be binding directly or indirectly and affecting

each other’s activity by altering delivery to the PSD, changing the degradation rate, or affecting

the functioning of the protein. When you consider the role of these proteins and their

localization it is more likely they are interacting at the protein level.

Thus far, all experimentation has been conducted in a Shank overexpression background but as

previously explained, decreasing the levels of Shank results in the same adverse NMJ

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phenotype. To examine the relationship between Shank and Ca-α1d when Shank levels are

reduced, double and single Ca-α1d and Shank heterozygotes were used. After bouton number

analysis no significant alteration in bouton count was present. Due to the discovered relationship

from previous experiments, it was expected that when one wild-type copy of the Ca2+ channel

and one copy of Shank were removed an enhancement of the Shank phenotype would be seen

however this was not the case.

To examine this relationship in another manner a double mutant could be utilized. By

recombining the AR66 allele with the ShankD101allele (AR66, ShankD101/CyO-GFP) further

testing for an interaction could be conducted in a loss-of-function background. Based on the

results from previous Shank loss-of-function experiments, however, it is not expected that any

significant relationship between Shank and Ca-α1d will be apparent.

As you will recall there is a dose-dependent relationship between Shank and NMJ phenotype.

Both overexpression and loss of Shank lower bouton count, although it is still not understood

how different doses of Shank lead to the same phenotypic outcome. It appears as though the

genetic interaction between Shank and Ca-α1d is only apparent in the case of Shank

overexpression. For this project Shank was overexpressed in the RNAi genetic screen and the

previous experiment with the Ca-α1d heterozygotes and those were the experiments where a

significant relationship between Shank and Ca-α1d was seen. The precise nature of the

relationship between Ca-α1d and Shank may offer some insight into the mechanism by which

increased Shank levels alter the synapse. Therefore, the Shank overexpression phenotype can be

modified by reducing Ca-α1d through two different methods (RNAi and loss-of-function

mutations). However, Ca-α1d does not modify Shank loss of function. Repeating the genetic

screen in a Shank loss-of-function background may help reveal other potential novel Shank

interactors and mechanisms underlying Shank loss-of-function.

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4.4 Immunohistochemistry interpretation

Immunohistochemistry using an antibody raised against the mammalian epitope of CaV1.3 was

used to observe the localization of its Drosophila homolog Ca-α1d. This was done to determine

if Ca-α1d levels regulate the levels of Shank at the synapse as hypothesized. The antibody

against the mammalian CaV1.3 did show concentrated staining at NMJs when applied in wild-

type Drosophila. Pre- and postsynaptic KD of Ca-α1d was expected to show a decrease in the

signal at the NMJ. However, the signal level remained consistent among all treatments

indicating the staining was not specific enough to accurately highlight Ca-α1d localization. To

properly visualize the localization of Ca-α1d an antibody could be raised against Ca-α1d or an

HA tag could be applied to the protein.

A relationship appears to exist between Ca-α1d and Shank as reducing the levels of Ca-α1d

using different tools enhances the Shank overexpression phenotype. One theory to explain this

would be if Ca-α1d is inhibiting Shank levels at the synapse. If this is true reducing the levels of

Ca-α1d would worsen the Shank overexpression phenotype, as seen in the above experiments.

To test this hypothesis immunohistochemistry was conducted to visualize Shank localization,

first in control animals with wild-type Ca-α1d expression then in animals with lowered

expression in the muscles. It was expected that when Ca-α1d was knocked down Shank

intensity at the boutons would increase compared to controls or the distribution may change.

However, after analysis Shank intensity and distribution appeared to be the same in both

experimental groups. Therefore, there is no evidence to support this theory. This visualization

only confirmed that Shank is present at the boutons in its expected amount and distribution

however it does not establish if Shank is functioning normally. We would have to know more

about Shank’s function to test this accurately.

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4.5 Relation to the field

The majority of Shank research thus far has been completed in mammalian models, mostly in

mice. This is troubling however because in mammals there are three members of the Shank

family that are all highly conserved with one another. The presence of multiple forms of the

same protein creates the problem of redundancy and leaves findings from rodent models

inconsistent. Drosophila, however, possess only one copy of the Shank gene which is highly

conserved with all three of the mammalian genes. Therefore, the Drosophila model can be used

to investigate how Shank functions and influences synapse biology without the limitation of

redundancy. This model is also highly applicable to several genetic tools and manipulations that

can be taken advantage of.

Thus far only two groups have published work examining Drosophila Shank, both of which

mainly focused on the results of Shank loss-of-function. Although, the overexpression and loss

of Shank result in the same phenotype my work contributes to our knowledge of what occurs at

the synapse when Shank levels are raised. One of the papers mentioned above focused on

identifying a pre-synaptic role of Shank using null mutants. The other demonstrated a post-

synaptic role for Shank and a role in the Wnt signalling pathway. Currently, no group has

demonstrated any proteins that are known to interact with Shank unlike in rodent models where

several binding partners are known for each domain of Shank. My project offers a very

promising candidate for further study to begin the work of deciphering the interactors of Shank.

Further, I believe I have shown there are key differences in the mechanism behind the

phenotype generated from Shank loss or elevation that need to be considered. This project also

only focused on several genes whose homologs are known to interact with Shank in mice,

making them more likely to share an interaction. The success of this project may act as a

catalyst for a larger scale and more fruitful Drosophila genetic screen in the future.

Of the hundreds of mutations and copy number variations identified in individuals with autism

spectrum disorders (ASDs), Shank mutations represent the most prevalent cause and one of the

few monogenic causes. Understanding the molecular pathways that are disturbed in ASD is

crucial to elucidating its pathogenesis. By using a simple model to study autism-linked genes

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aspects of the signaling pathways related to this disorder may be uncovered. But before we can

use Drosophila Shank to provide new directions in ASD research we must first understand the

role it plays at the synapse and the proteins it interacts with.

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References

Antonin W, Fasshauer D, Becker S, Jahn R, Schneider TR (2002) Crystal structure of the

endosomal SNARE complex reveals common structural principles of all SNAREs. 9:107–

111.

Augustin I, Rosenmund C, Sudhof T, Brose N (1999) Munc13-1 is essential for fusion

competence of glutamatergic synaptic vesicles. Nature 400:457–461.

Banovic D, Khorramshahi O, Owald D, Wichmann C, Riedt T, Fouquet W, Tian R, Sigrist S,

Aberle H (2010) Drosophila Neuroligin 1 Promotes Growth and Postsynaptci

Differentiation at Glutamatergic Neuromuscular Junctions. Neuron 66:724–738.

Bennett MR (2000) The concept of transmitter receptors: 100 years on. Neuropharmacology

39:523–546.

Betancur C (2011) Etiological heterogeneity in autism spectrum disorders: more than 100

genetic and genomic disorders and still counting. Brain Res 1380:42–77.

Betancur C, Buxbaum JD (2013) SHANK3 haploinsufficiency : a “common” but

underdiagnosed highly penetrant monogenic cause of autism spectrum disorders. Mol

Autism 4:4–6.

Bier E (2005) Drosophila, the golden bug, emerges as a toll for human genetics. Nat Rev 6:9–

23.

Boeckers TM, Bockmann J, Kreutz MR, Gundelfinger ED (2002) ProSAP/Shank proteins - A

family of higher order organizing molecules of the postsynaptic density with an emerging

role in human neurological disease. J Neurochem 81:903–910.

Bozdagi O, Sakurai T, Papapetrou D, Wang X, Dickstein DL, Takahashi N, Kajiwara Y, Yang

M, Katz AM, Scattoni ML, Harris MJ, Saxena R, Silverman JL, Crawley JN, Zhou Q, Hof

PR, Buxbaum JD (2010) Haploinsufficiency of the autism-associated Shank3 gene leads to

deficits in synaptic function , social interaction , and social communication. Mol Autism

1:15 Available at: http://www.molecularautism.com/content/1/1/15.

Brand AH, Perrimon N (1993) Targeted gene expression as a means of altering cell fates and

generating dominant phenotypes. Development 118:401–415 Available at:

http://www.ncbi.nlm.nih.gov/pubmed/8223268.

Broadie K, Bate M (1993) Development of the embryonic neuromuscular synapse of Drosophila

74

melanogaster. J Neurosci 13:144–166.

Canada G of (2018) Surveillance of autism spectrum disorder (ASD). :1–1 Available at:

https://www.canada.ca/en/public-health/services/diseases/autism-spectrum-disorder-

asd/surveillance-autism-spectrum-disorder-asd.html [Accessed June 26, 2018].

Carpenter F (1905) The reactions of the pomace fly (Drosophila ampelophila Loew) to light,

gravity, and mechanical stimulation. Am Nat 39:157–171.

Catterall W (1991) Functional subunit structure of voltage-gated calcium channels. Science (80-

) 253:1499–1500.

Celniker SE, Rubin GM (2003) The Drosophila menalnogaster genome. Annu Rev Genomics

Hum Genet 4:89–117.

Chorna T, Hasan G (2012) The genetics of calcium signaling in Drosophila melanogaster.

Biochim Biophys Acta 1820:1269–1282.

Coolen A, Kuhn R, Sollich P (2005) Theory of Neural Informaion Processing systems. London,

UK: Oxford University Press.

Del Castillo J, Katz B (1954) Quantal components of the end-plate potential. J Physiol 124:560–

573.

Descartes R (1637) Discours de la Methode. Manchester: The University Press.

DiAntonio A (2006) Glutamate receptors at the Drosophila neuromuscular junction. Int Rev

Neurobiol 75:165–179.

Dolphin A (2012) Calcium channel auxiliary alpha2delta and beta subunits: Trafficking and one

step beyond. Nat Rev Neurosci 13:542–555.

Du Y, Weed S, Xiong W, Marshall T, Parsons J (1998) Identification of a novel cortactin SH3

domain-binding protein and its localization to growth cones of cultured neurons. Mol Cell

Bio 18:5838–5851.

Du Bois-Reymond E (1874) Experimental kritik der Entladungs hypothese uber die Wirkung

von Nerv auf Muskel. Sitzungsber Phys Math Kl:519–560.

Durand CM et al. (2007) Mutations in the gene encoding the synaptic scaffolding protein

SHANK3 are associated with autism spectrum disorders. Nat Genet 39:25–27.

Eberl DF, Ren D, Feng G, Lorenz LJ, Vactor D Van, Hall LM (1998) Genetic and

Developmental Characterization of Dmca1D, a Calcium Channel a1 Subunit Gene in

Drosophila melanogaster. Genetics 148:1159–1169.

75

Elliott TR (1904) On the action of adrenalin. J Physiol 31:401–467.

Failla P, Romano C, Alberti A, Vasta A, Buono S, Castiglia L, Luciano D, Di Bendetto D,

Fichera M, Galesi O (2007) Schizophrenia in a patient with subtelomeric duplication of

chromosome 22q. Clin Genet 71:599–601.

Fatt P, Katz B (1951) An analysis of the end-plate potential recorded with an intra-cellular

electrode. J Physiol:320–370.

Featherstone D, Rushton E, Rohrbough J, Liebl F, Karr J, Sheng Q, Rodesch C, Broadie K

(2005) An essential Drosophila glutamate receptor subunit that functions in both central

neuropil and neuromuscular junction. J Neurosci 25:3199–3208.

Fombonne E (1999) The epidemiology of autism: a review. Psychol Med 29:769–786.

Geschwind DH (2011) Genetics of autism spectrum disorders. Trends Cogn Sci 15:409–416.

Grabruker A, Schmeisser M, Schoen M, Boeckers TM (2011) Postsynaptic ProSAP/Shank

scaffolds in the cross-hair of synaptopathies. Trends Cell Biol 21:366–373.

Gramates L, Marygold S, dos Santos G, Urbano J, Antonazzo G, Matthews B, Rey A, Tabone

C, Crosby M, Emmert D, Falls K, Goodman J, Hu Y, Ponting L, Schroeder A, Strelets V,

Thurmond J, Zhou P, Consortium the F (2017) FlyBase at 25: looking to the future.

Nucleic Acids Res 45:D663–D671.

Han K, Holder JJ, Schaaf C, Lu H, Chen H, Kang H, Tang J, Wu Z, Hao S, Cheung S, Yu P,

Sun H, Breman A, Patel A, Lu H, Zoghbi H (2013) SHANK3 overexpression causes

manic-like behaviour with unique pharmacogenetic properties. Nature 503:72–77.

Hanson PI, Heuser JE, Jahns R (1997) Neurotransmitter release - four years of SNARE

complexes. Curr Opin Neurobiol 7:310–315.

Harris KP, Akbergenova Y, Cho RW, Baas-Thomas MS, Littleton JT (2016) Shank Modulates

Postsynaptic Wnt Signaling to Regulate Synaptic Development. J Neurosci 36:5820–5832

Available at: http://www.jneurosci.org/content/36/21/5820.abstract?etoc.

Harris KP, Littleton JT (2015) Transmission, development, and plasticity of synapses. Genetics

201:345–375.

Heuser J, Reese T, Dennis M, Jan Y, Jan L, Evans L (1979) Synaptic vesicle exocytosis

captured by quick freezing and correlated with quantal transmitter release. J Cell Biol

81:275–300.

Hoang B, Chiba A (2001) Single-cell analysis of Drosophila larval neuromuscular synapse. Dev

76

Biol 229:55–70.

Hodgkin BYAL, Huxley AF (1945) Resting and action potentials in single nerve fibres. J

Physiol 104:176–195.

Hong C, Ganetzky B (1994) Spatial and temporal expression patterns of two sodium channel

genes in Drosophila. J Neurosci 14:5160–5169.

Jiang Y hui, Ehlers MD (2013) Modeling Autism by SHANK Gene Mutations in Mice. Neuron

78:8–27 Available at: http://dx.doi.org/10.1016/j.neuron.2013.03.016.

Jin Y (2002) Synaptogenesis : insights from worm and fly. Development 12:71–79.

Johansen J, Halpern ME, Johansen KM, Keshishian H (1989) Stereotypic morphology of

glutamatergic synapses on identified muscle cells of Drosophila larvae. J Neurosci 9:710–

725.

Kanner L (1943) Autistic disturbances of affective contact. Nerv Child 2:217–250.

Keshishian H, Broadie K, Chiba A, Bate M (1996) The Drosophila Neuromuscular Junction : A

Model System for Studying Synaptic Development and Function. Annu Rev Neurosci

19:545–575.

Krause W (1863) Ueber die Endigung der Muskelnerven. Z Natl Med 18:136–160.

Kreienkamp H (2008) Scaffolding proteins at the postsynaptic density: shank as the architectural

framework. Handb Exp Pharmacol 186:365–380.

Kurdyak P, Atwood H, Stewart B, Wu C (1994) Differential physiology and morphology of

motor axons to ventral longitudinal muscles in larval Drosophila. J Comp Neurol 350:463–

472.

Lee J, Coyne R, Dubreuil R, Goldstein L, Branton D (1993) Cell Shape and Interaction Defects

in alpha-Spectrin Mutants of Drosophila Melanogangster. J Cell Biol 123:1797–1809.

Levitan I, Kaczmarek L (2015) The Neuron: Cell and Molecular Biology, 4th ed. New York,

NY: Oxford University Press.

Lim S, Naisbitt S, Yoon J, Hwang J, Suh P, Sheng M, Kim E (1999) Characterization of the

Shank family synaptic proteins. Multiple genes, alternative splicing, and differential

expression in brain and development. J Biol Chem 274:29510–29518.

Loewi O (1921) Uber humorale Ubertragbarkeit der Herznervenwirkung. Arch Ge Physiol

189:239–242.

MacPherson MR, Pollock VP, Broderick KE, Kean L, O’Connell FC, Dow JA, Davies SA

77

(2001) Model Organisms: New Insights Into Ion Channel and Transporter Function. L-type

calcium channels regulate epithelial fluid transport in Drosophila melanogaster. Am J

Physiol Cell Physiol 280:394–407.

Mandy W, Chilvers R, Chowdhury U, Salter G, Seigal A, Skuse D (2012) Sex differences in

autism spectrum disorder: evidence from a large sample of children and adolescents. J

Autism Dev Disord 42:1304–1313.

Martinez-Pedraza F, Carter A (2009) Autism spectrum disorders in young children. Child

Adolesc Psychiatr Clin N Am 18:645–663.

Melom JE, Akbergenova Y, Gavornik JP, Littleton JT (2013) Spontaneous and evoked release

are independently regulated at individual active zones. J Neurosci 33:17253–17263.

Moessner R, Marshall CR, Sutcliffe JS, Skaug J, Pinto D, Vincent J, Zwaigenbaum L,

Fernandez B, Roberts W, Szatmari P, Scherer SW (2007) Contribution of SHANK3

mutations to autism spectrum disorder. Am J Hum Genet 81:1289–1297 Available at:

http://www.sciencedirect.com/science/article/pii/S000292970763777X.

Montecucco C, Schiavo G (1995) Structure end function of tetanus and botulinurn neurotoxin.

Q Rev Biophys 28:423–472.

Morgan T (1910) Sex limited inheritance in Drosophila. Science (80- ) 32:120–122.

Morgan T (1911) An attempt to analyze the constitution of the chromosomes on the basis of

sex-limited inheritance in Drosophila. J Exp Zool 11:365–413.

Morgan T, Sturtevant A, Muller H, Bridges C (1915) The mechanism of mendelian heredity.

New York: Johnson Reprint Corporaiton.

Naisbitt S, Kim E, Tu J, Xiao B, Sala C, Valtschanoff J, Weinberg R, Worley P, Sheng M

(1999) Shank, a novel family of postsynaptic density proteins that binds to the NMDA

receptor/PSD-95/GKAP complex and cortactin. Neuron 23:569–582.

Niemann H, Blasi J, Jahn R (1994) Clostridial neurotoxins: new tools fo dissecting exocytosis.

Trends Cell Bio 4:179–185.

Nijhof B, Castells-Nobau A, Wolf L, Scheffer-de Gooyert JM, Monedero I, Torroja L,

Coromina L, van der Laak JAWM, Schenck A (2016) A New Fiji-Based Algorithm That

Systematically Quantifies Nine Synaptic Parameters Provides Insights into Drosophila

NMJ Morphometry. PLoS Comput Biol 12:1–25.

Parnas D, Haghighi AP, Fetter RD, Kim SW, Goodman CS (2001) Regulation of postsynaptic

78

structure and protein localization by the Rho-type guanine nucleotide exchange factor dPix.

Neuron 32:415–424.

Pereda AE (2014) Electrical synapses and their functional interactions with chemical synapses.

Nat Rev Neurosci 15:250 Available at: http://dx.doi.org/10.1038/nrn3708.

Perrimon N, Ni J, Perkins L (2010) In vivo RNAi : Today and Tomorrow. Cold Spring Harb

Perspect Biol 2:1–12.

Peterson S, Fetter RD, Noordermeer J, Googman C, DiAntonio A (1997) Genetic analysis of

glutamate receptors in Drosophila reveals a retrograde signal regulating pre-synaptic

transmitter release. Neuron 19:1237–1248.

Piccioli Z, Littleton JT (2014) Retrograde BMP signalling modulates rapid activity-dependent

synaptic growth via presynaptic LIM kinase regulation of cofilin. J Neurosci 34:4371–

4381.

Pielage J, Fetter RD, Davis GW (2006) A postsynaptic Spectrin scaffold defines active zone

size, spacing, and efficacy at the Drosophila neuromuscular junction. J Cell Biol 175:491–

503.

Qin G, Schwarz T, Kittel R, Schmid A, Rasse T, Kappei D, Ponimaskin E, Heckmann M,

Sigrist S (2005) Four different subunits are essential for expressing the synaptic glutamate

receptor at neuromuscular junctions of Drosophila. J Neurosci 25:3209–3218.

Ren D, Xu H, Eberl DF, Chopra M, Hall LM (1998) A Mutation Affecting Dihydropyridine-

Sensitive Current Levels and Activation Kinetics in Drosophila Muscle and Mammalian

Heart Calcium Channels. J Neurosci 18:2335–2341.

Rothman J (1994) Mechanisms of intracellular protein tranport. Nature 372:55–63.

Sanchack K, Thomas C (2016) Autism Spectrum Disorder: Primary Care Principles. Am Fam

Physician 94:972–979.

Sato D et al. (2012) SHANK1 deletions in males with autism spectrum disorder. Am J Hum

Genet 90:879–887.

Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S,

Rueden C, Saalfeld S, Schmid B, Tinevez J-Y, White DJ, Hartenstein V, Eliceiri K,

Tomancak P, Cardona A (2012) Fiji: an open-source platform for biological-image

analysis. Nat Methods 9:676–682 Available at:

http://www.nature.com/doifinder/10.1038/nmeth.2019.

79

Schmid A, Qin C, Wichmann C, Kittel R, Mertel S, Fouquet W, Schmidt M, Heckmann M,

Sigrist S (2006) Non-NMDA-type glutamate receptors are essential for maturation but not

for initial assembly of synapses at Drosophila neuromuscular junctions. J Neurosci

26:11267–11277.

Seeger M, Tear G, Ferres-Marco D, Goodman C (1993) Mutations affecting growth cone

guidance in Drosophila: genes necessary for guidance toward or away from the midline.

Neuron 30:409–426.

Sheng M, Kim E (2000) The Shank family of scaffold proteins. J Cell Sci 113:1851–1856.

Sigrist S, Reiff D, Thiel P, Steinert J, Schuster C (2003) Experience-dependent strengthening of

Drosophila neuromuscular junctions. J Neurosci 23:6546–6556.

Silverthorn DU (2007) Human Physiology: An integrated Approach, 4th ed. San Francisco:

Pearson/Benjamin Cummings.

Söllner T, Whiteheart S, Brunner M, Erdjument-Bromage H, Geromanos S, Tempst P, Rothman

J (1993) SNAP receptors implicated in vesicle targeting and fusion. Nature 362:310–324.

Solomon M, Miller M, Taylor S, Hinshaw S, Carter C (2012) Autism symptoms and

internalizing psychopathology in girls and boys with autism spectrum disorders. J Autism

Dev Disord 42:48–59.

Stewart B, Atwood H, Renger J, Wang J, Wu C (1994) Improved stability of Drosophila larval

neuromuscular preparations in haemolymph-like physiological solutions. J Comp Physiol

175:179–191.

Stuhmer W, Conti F, Suzuki H, Wang X, Noda M, Yahagi N, Kubo H, Numa S (1989)

Structural parts involved in activation and inactivation of the sodium channel. Nature

339:597–603.

Sudhof T (2004) The synaptic vesicle cycle. Annu Rev Neurosci 27:509–547.

Südhof TC (2014) The Molecular Machinery of Neurotransmitter Release (Nobel Lecture).

Angew Chem Int Ed 53:12696–12717.

Sullivan K, Scott K, Zuker C, Rubin G (2000) The ryanodine receptor is essential for larval

development in Drosophila melanogaster. Proc Natl Acad Sci 97:5942–5947.

Tsien R, Lipscombe D, Madison D, Bley K, Fox A (1988) Multiple types of neuronal calcium

channels and their selective modulation. Trends Neurosci 11:431–438.

Vonhoff F, Keshishian H (2017) In vivo Calcium Signaling during Synaptic Refinement at the

80

Drosophila Neuromuscular Junction. J Neurosci 37:5511–5526.

Wang X, Mccoy PA, Rodriguiz RM, Pan Y, Je HS, Roberts AC, Kim CJ, Berrios J, Colvin JS,

Bousquet- D, Lorenzo I, Wu G, Weinberg RJ, Ehlers MD, Philpot BD, Beaudet AL,

Wetsel WC, Jiang Y (2011) Synaptic dysfunction and abnormal behaviors in mice lacking

major isoforms of Shank3. Hum Mol Genet 20.

Weber T, Zemelman B V, Mcnew JA, Westermann B, Gmachl M, Parlati F, Rothman JE (1998)

SNAREpins: Minimal Machinery for Membrane Fusion. Cell 92:759–772.

Werling D, Geschwind DH (2013) Sex differences in autism spectrum disorders. Curr Opin

Neurol 26:146–153.

Werner E, Dawson G, Munson J, Osterling J (2005) Variation in early developmental course in

autism and its relation with behavioral outcome at 3-4 years of age. J Autism Dev Disord

35:337–350.

Worrell JW, Levine RB (2008) Characterization of Voltage-Dependent Ca 2+ Currents in

Identified Drosophila Motoneurons In Situ. J Neurophysiol 100:868–878.

Wu XS, Gan G, Zhang XZ, Sun XJ, Wang Q, Gao XZ, Li XM, Jin S, Huang XJ, Thomas XU,

Jiang Y, Li XY, Tian R, Zhang XYQ (2017) A Presynaptic Function of Shank Protein in

Drosophila. J Neurosci 37:11592–11604.

Zhang H, Maximov A, Fu Y, Xu F, Tang T, Tkatch T, Surmeier DJ (2005) Association of

Cav1.3 L-Type Calcium Channels with Shank. J Neurosci 25:1037–1049.

Zheng W, Linda M, Ag P, Carolina N (1995) Cloning and Characterization of a Calcium

Channel alpha1 Subunit from Drosophila melanogaster with Similarity to the Rat Brain

Type D lsoform. J Neurosci 15:1132–1143.

Zito K, Parnas D, Fetter RD, Isacoff E, Goodman C (1999) Watching a synapse grow:

Noninvasive confocal imaging of synaptic growth in Drosophila. Neuron 22:719–729.

Zwaigenbaum L (2001) Autistic spectrum disorders in preschool children. Can Fam Physician

47:2037–2042.