an investigation of vesicular trafficking and secretion in type iii ...

AN INVESTIGATION OF VESICULAR TRAFFICKING AND SECRETION IN TYPE III

NEUROMUSCULAR JUNCTIONS OF D. MELANOGASTER

A DISSERTATION

PRESENTED TO THE FACULTY OF THE GRADUATE SCHOOL OF

CORNELL UNIVERSITY

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

BY

CASSANDRAVICTORIA INNOCENT

JANUARY 2015

© 2015 Cassandravictoria Innocent

FORWARD

The protective external cuticle of insects is ideal for many functions: it is practical as a

point for muscle attachments, its impermeability is a sensible way to prevent

desiccation; and its rigidity serves well as a shield against dangers it may encounter.

Nevertheless, cuticle has a major constraint: it does not grow along with the

developing animal. Therefore, to counteract this shortcoming, the insect life cycle is

punctuated by a series of molts. For Drosophila melanogaster, molting will occur a

total of two times over the length of its life as a larva. During the molt, a new and

larger cuticle is produced and will replace the older, outgrown cuticle that encases it.

Replacement of the previous cuticle ends in a process termed ecdysis – a

stereotyped sequence of shedding behaviors. Following each ecdysis, the new cuticle

must expand and harden. Studies from a variety of insect species indicate that this

cuticle hardening is regulated by the neuropeptide Bursicon (Fraenkel and Hsiao,

1966; Luo et al., 2005; Bestman and Booker, 2003; Weber, 1985). Therefore, the

canonical view of Bursicon is that its developmental involvement begins only at the

end of the ecdysis hormonal cascade, coinciding with the animal’s final ecdysis, when

the adult fly emerges (eclosion). However, previous work in this lab has offered

evidence that the canonical release of Bursicon is preceded by another, equally

predictable release event.

i

The research presented here makes use of this new observation – namely, that

Bursicon is released at various predictable points in the juvenile larval stages – and

attempts to exploit it.

Ecdysis is the mechanical program of stereotyped cuticle-shedding behaviors and is

initiated by the secretion of ecdysis triggering hormone (ETH) from INKA cells (Park

et al., 2002; O’Brien and Taghart, 1998) followed by a positive feedback loop

between ETH and eclosion hormone (EH) (Arakanea et al., 2008; Clarke et al., 2004;

Bestman and Booker, 2003) and ending with CCAP triggering the animal’s forward

escape (FE) (Loveall and Deitcher, 2010; Arakanea et al., 2008; Gammie and Truman.

1997). In the same way that many other nerve terminals co-package neuropeptides

and neurotransmitters within the same vesicles (Zhang et al., 2011;, Salio et al., 2006,

Hökfelt et al., 1999, Purves et al., 2001) so too does CCAP neurons (NCCAP),

copackaging both CCAP and another neurohormone, Bursicon. Because it is

important for melanization of the cuticle (Luo et al., 2005; Karsai et al., 2013; Loveall

and Deitcher, 2010; Woodruff et al., 2009), Bursicon is released stereotypically and

found exclusively at type III boutons (Honneger et al., 2008; Loveall and Deitcher,

2010). For this reason Bursicon can be specifically used as a tool to study trafficking

and secretion as an in vivo peptidergic marker.

ii

Therefore, the purpose of this study is to elucidate the process of secretory granule

trafficking and secretion occurring in type III NMJ boutons. It is an aim meant to

elaborate on the mechanism and timing of hormonal release from Bursicon-positive

motor neurons at the larval neuromuscular junction. The key findings here indicate

that these peptidergic motor neuron terminals express the same core exocytotic

machinery as is required for classical neurotransmitter release. Moreover, this effort

involves a series characterization and perturbation experiments meant to explore

factors underlying peptidergic vesicle secretion, and confirm which components are

employed by the mechanism. To do so first a preliminary registry of which molecules

are present, active and implicated in type III vesicular trafficking and secretion is

assembled from antibody-labelled immunocytochemistry stainings. Next, because

the process of neuropeptidergic delivery is essential for development (Žitňan et al.,

2003; Park et al. 2002; Dulcis et al., 2005), it has been considerably important to

individually perturb elements of the system in a cell-specific manner in order to

deduce function from the observed effects.

Subsequently, results of the first aim can be used in efforts to identify, define and

more fully understand peptidergic vesicle secretion in this model organism and

beyond. To this end, we can use the fact that peptidergic contacts secrete hormones

at the neuromuscular junction (NMJ) in two waves of release (coinciding with larval

iii

ecdysis) to our advantage. In consequence, understanding the factors governing

proper trafficking, aggregation and docking of secretory vesicles can lend

themselves to greater developments in such methods as drug delivery and targeting

as well as processes involving long-term hormonal regulation. For that reason, it is a

mechanism worthy of attention and deserves to be more fully uncovered.

iv

AN INVESTIGATION OF VESICULAR TRAFFICKING AND SECRETION IN TYPE III NEUROMUSCULAR JUNCTIONS OF D. MELANOGASTER (ABSTRACT)

Cassandravictoria Innocent, PhD

Cornell University, 2015

Bursicon is a neurohormone packaged in and secreted by type III synaptic contacts.

In comparison to the depth of information describing type I and type II

neuromuscular junctions (NMJs), type III NMJs are notably ill-defined. As such, this

lab has provided evidence identifying bursicon as a reliable marker in an aim to

characterize type III NMJs. Firstly, a comprehensive registry of proteins defining type

III NMJs can be compiled through antibody-labeled colocalizations using bursicon as

the type III marker. Candidates to be stained with bursicon include established

presynaptic markers such as the SNARE proteins. Positional overlap in these dual

antibody-staining profiles will be confirmed by sequential-excitation confocal

microscopy. This proteomic approach establishes an immunohistological staining

profile characterizing type III NMJs. Secondly, this effort in effect also produces a

directory identifying proteins involved in vesicle delivery in type III NMJs and can be

used to reveal the machinery by which neuropeptides are trafficked and secreted.

While what is known about the mechanisms of vesicle trafficking employed by

neurotransmitter-releasing type I synapses continues to be expanded, information

on these same properties in peptidergic type III NMJs idles. Therefore, using a GFP-

v

variant fusion protein able to mimic bursicon’s packaging and transport in vesicles,

this lab has perturbed in vivo delivery of a fluorescent neuropeptide to describe type

III synaptic release. In doing so we aim to elucidate the dynamic process specifically

employed by type III NMJs when trafficking secretory vesicles through boutons in

parallel and add to the breadth of knowledge gathered about the mechanism used

during peptide release.

vi

BIOGRAPHICAL SKETCH

Cassandravictoria Innocent wants to have a positive impact on the people around

her. Those changes may be tiny but she sincerely hopes that the changes are real.

Her motivation comes from a desire to experience and to learn and to pass it along

for the better. Therefore, she tries to spend her time wisely by absorbing science, in

planes flying, bending épeé blades and encouraging others. Lastly, she is defined by

the company she keeps and loves how defined she is.

vii

This dissertation is fondly dedicated to the memory of my third grade

teacher, Mr. Aaron Pohl. The milestones of my life always surprised

me, but never did surprise him. So after graduating HS, and again

after graduating college, I eagerly told him. Several years later, I can

only pray he hears about this now (and is still not surprised).

viii

ACKNOWLEDGMENTS

First I thank my advisor, Dr. David L Deitcher. For a dissertation it is common for the

writer to say “I could not have asked for a better advisor.” And while it is true in my

case, I will go on to say it again, with emphasis: I could not have asked for a better

advisor. He accepted me into his bright yellow lab in 2010 and for the next 4.5 years

continued to provide an incredibly safe space for me to grow – both intellectually

and personally. His compassion is as awesome as his laugh, and I will never forget

either.

Next I would like to warmly thank the remaining members of my special committee.

Dr. Douglas Knipple and Dr. Jeffrey Roberts introduced me to the complex and

fascinating worlds of insect biology and protein biochemistry, respectively. And after

taking their classes I knew I wanted to learn more under their advisement. Lastly, I

must especially thank Dr. Ronald Booker for hearing out my ideas on science and on

life, and telling me I am wrong on both. I treasure his insight and thank him dearly

for it.

Furthermore, while other professors at Cornell have impacted me to no end, I will try

to thank them here, though this list is inexhaustible: My deepest gratitude goes to

Dr. Eric Alani. This man has been invaluable to my evolution here at Cornell in

ix

countless of ways. He cares about students in a way that cannot be defined in words,

and for that I owe him more than I can express. Additionally, I have also had the

immense luck of knowing Dr. Jun Kelly Liu, Dr. Debra Nero, Dr. James Blankenship,

Dr. Tom Fox, Dr. Jian Hua and numerous other professors. Their collective kindness

and tutelage paved the avenue for me to get this far. Here I will also take a moment

to thank Dr. Frank Scalzo: in giving me a chance you gave me a world of opportunity;

Dr. Christine Labno: your encouragement on all fronts has been a delight; Dr.

Michelle Nearon: you are an inspiration; and Carol Bayles: you showed me the magic

of microscopy.

To my friends and family: the love and support you have given to me is the

fundamental reason I am here today. Dr. Nathaniel Frank saw more in me than I

could even pretend to see and so with his, “No… you absolutely CAN,” my interest in

applying to graduate school was spurred. My parents, Jean, Michelle and Emmanuel:

you showed me with your own lives how to succeed and backed every one of my

decisions in making my priorities your priorities. My siblings Jean, Gregory, Tanisha

and Emmanuela: you are my pride and backbone. My friends: you delivered words of

encouragement, boxes filled with white envelopes, couches stained with tears and

dried with laughter – I cannot thank any of you enough. Shoutouts go to Nadia

Rodriguez, Jeffrey Nelson, Krissy Sugatan, Bobby Smith II, Linda Ng, Christopher

Vredenburgh, Christina Black, Michaela Brangan, Lily Cui, James Macmillen, Aziza

x

Glass, Ferdinand Kohle, Kristy Lawton, Amath Gomis, Elizabeth Wayne, Santiago

Palacio, Natasha Udpa, Erin Johnson, Andres Pina, Mari Yamaguchi, Haya al Thani,

Ghanee Smith, and Abiola Davis. Also, I love you Milutin Stojanovic.

Last but not least, I received various sources of funding during my tenure here. In

particular I am grateful for the Wimsatt Fund as provided by the Genetics, Genomics

and Development (GGD) department, the Cellular and Molecular Biology training

grant as provided by the Neurobiology and Behavior (NBB) department, the Provost

Fellowship as provided by the Graduate School and the SUNY Minority Fellowship as

provided by State University of New York.

xi

TABLE OF CONTENTS

Forward i

Abstract v

Biographical sketch vii

Dedication viii

Acknowledgements ix

Table of Contents xii

List of Figures xiii

Chapter One: Introduction 1

Chapter Two: Secretory protein distribution and perturbed vesicle

release from type III boutons at the Drosophila melanogaster larval NMJ 34

Chapter Three: Implications & Conclusions 72

xii

LIST OF FIGURES

Fig. 1 Developmental transitions in Drosophila melanogaster 5

Fig. 2 Signature ecdysis behavior 9

Fig. 3 Type III Bouton Proteome Contain a List of SNARE Proteins 37

Fig. 4

The Active Zone marker Bruchpilot & the SNARE-associated

Complexin protein both Fail to Colocalize with Bursicon at type III

Boutons

39

Fig. 5 Type III Boutons are Not Glutamatergic 41

Fig. 6 A RNAi Knockdown of Syntaxin (Syx1A) Results in Over-filled Bouton

Phenotype. 44

Fig. 6 B ImageJ Analysis Following RNAi Knockdown of Syntaxin (Syx1A-

RNAi) 45

Fig. 6 C RNAi Knockdown of Syntaxin (Syx1A) Results in Larval Death at

Developmentally Significant Stages 46

Fig. 7 RNAi Knockdown of nSynaptobrevin (nSyb) is Insufficient to

Perturb Release at Type III NMJs 47

Fig. 8 A Proposed Bouton Capture 48

Fig. 8 B Dense Core Vesicle Capture and Refilling at type III Boutons 49

Fig. 9 A Exogenous/Mistimed release via photo-activation of UAS-

Chrimson and Signal Recovery 51

Fig. 9 B Neuropeptide Fluorescence Lowered in NCCAP-GAL4>UAS-

Chrimson Larvae Post Red LED Stimulation (RLS) 52

Fig. 9 C Exogenous/Mistimed release of CCAP/Bursicon Reveals Even

Refilling of Type III Boutons 54

Fig. 10 Tracheal Stainings From 2nd Instar Larvae 57

xiii

CHAPTER ONE:

Introduction

1.1 Brief Overview

The fast-acting kinetics of neurotransmitter biology has historically drawn much

attention to type I neuromuscular junctions (NMJ). Of the three classes, the type I

bouton is the most well described – its properties and defining characteristics have

been studied across a variety of model organisms and under various experimental

conditions (Arakanea et al., 2008; Huage et al., 2007; Donini et al., 2001; Bestman and

Booker, 2003; Gammie and Truman, 1997). In contrast however, the type III NMJ

remains understudied. As an example, little is known about type III bouton trafficking

dynamics – namely, how type III boutons traffic and secrete large dense core vesicles

(LDCVs) along a series of boutons. To this end fruit flies are especially amenable to

answering such a question.

Drosophila melanogaster is small, produces multiple generations in a relatively short

amount of time, and are easily raised under a range of laboratory conditions.

Additionally, the fruit fly has only 4 pairs of chromosomes and has an extensive

collection of mutants commercially available to the scientific public (Flybase

Consortium, 2014). Further still, because the Drosophila genome sequence was

1

completed and published (Spradling et al., 1999), it lends itself to being a wealthy

atlas of information in which the number of scientific investigations are boundless.

As a model organism the fruit fly has obtained notable popularity in scientific

inquiries in fields concerning ecdysis, neurotransmitter dynamics and synaptic

organization “due to its amenability to molecular manipulations” (Loveall,

Dissertation 2010). We therefore sought out to use a varied array of techniques to

study the mechanism underlying peptidergic vesicle secretion. Approaches used in

this study include the use of null mutants, knocking down specific suspected Soluble

NSF Attachment REceptors (SNAREs) through the use RNA interference (RNAi),

epifluorescent microscopy, confocal microscopy and exploiting light–drivable

channelrhodopsin to optogenetically control peptidergic neurons.

1.2 Fruit Fly Life Cycle

The life cycle of a fruit fly is divided into four major life stages: fertilized egg, larva,

pupa and adult. As a result, Drosophila undergoes complete metamorphism (termed

holometabolism), exchanging juvenile body parts for adult ones during the imaginal

molt of the pupal stage (Figure 1). Drosophila flies develop a stiff exoskeleton to

serve as protective armor in its adult stage. Prior to this, however, the larva has a

rigid outer cuticle to shield itself from varying hazards. Because of this inherent

2

rigidity the cuticle must be molted (Fraenkel and Hsiao, 1966). The process of

exchanging the old, outgrown cuticle for a larger one is known as ecdysis.

Before this, however, the juvenile stage lasts from egg hatching until pupariation

(formation of the pupa) and is itself punctuated by a series of developmental weight

milestones, with 2 molts interspersed. The individual juvenile larval stages are termed

instars, of which there are three: 1st -, 2nd - and 3rd instar (or L1, L2 and L3

respectively). Weight or mass checkpoints determine when the larva will pupate and

begin metamorphosis. When the critical size is met a hormonal cascade that ends in

metamorphosis begins (Testa et al., 2013). The timing of a metamorphic molt is

particularly important as initiating it ends the growth phase and establishes the final

body size of the adult insect (Nijhout, 1981). There is, however, a lag between when

the critical size is met and when the ecdysteroid - in this case, 20-hydroxyecdysone

or 20E (Winbush et al., 2011) - concentration has reached a critical threshold,

signaling the larva to stop feeding and end its body growth (Testa et al., 2013;

Winbush et al., 2011, Zdarek and Fraenkel, 1969).

3

Figure 1 - Developmental transitions in Drosophila melanogaster.

[1] Hatching of the 1st instar larva from the embryo is not typically

considered as an ecydis. Ecdysis (as portrayed in Figure 1)

occurs at the transition from a 1st instar larva into a 2nd instar larva

[2], and again at the transition from a 2nd instar larva into a 3rd

instar larva [3]. Pupariation, the transition of a 3rd instar larva into a

prepupa [4], is not an ecdysis but rather the formation of a

puparium from the 3rd instar cuticle. As the puparium tans [5], the

developing prepupa undergoes pupal ecdysis (pupation) [6] to form

a pharate adult. After the final ecdysis (eclosion) [7], the newly

hatched adult fly has a soft cuticle and unexpanded wings. Shortly

after hatching, the wings expand and the cuticle hardens [8] in a

process regulated by the hormone Bursicon. (Loveall and Deitcher, 2010)

4

5

1.3 Wild type Drosophila Behavior

Certain behaviors of adult Drosophila can involve very detailed movements and

includes such activities as courtship and mating (Lawton et al., 2014; Kimura et al.,

2014; Saleem et al., 2014). Actions like these will involve specialized control and

coordination of fine motor movements. Drosophila larval crawling however is a

rhythmic behavior consisting of a series of periodic strides (Ohyama, et al., 2013;

Heckscher et al., 2012; Nichols et al., 2012; Jakubowsk et al., 2012). If removed from a

food source, normal wild-type larva will crawl in search for food, an act termed

foraging (Ohyama, et al., 2013; Jakubowsk et al., 2012).

Larval locomotion (foraging or crawling; interchangeably) is divided into two phases.

First, the head and tail lift off from the crawling surface. This act adjusts the animal’s

center of mass. Then, when the head is anchored on the surface in a new direction, a

body wall wave propels the abdominal sections in the direction of the head

(Heckscher et al., 2012). In order to reverse, “abdominal body wall movements are

powered by phase-shifted contractions of dorsal and ventral muscles; and ventral

muscle contractions occur concurrently with contraction of lateral muscles in the

adjacent anterior segment” (ibid). As such, larval crawling involves less fine motor

control than some adult behaviors, yet it remains critical to the animal’s survival.

When not crawling these animals will exhibit ~40-50 body wall contractions each

6

minute (Nichols et al., 2012). Important for food scavenging, crawling can also serve

as a passive defense response against infection and parasitism. One such case is

exemplified when fruit fly larvae evade predatory wasps: if a wasp’s stinger

penetrates the Drosophila larval cuticle, the larva will instinctively respond in

“nocifensive escape behavior”. Nocifensive escape behavior is an evasive maneuver

described as “a corkscrew-like rolling around the anterior/posterior axis, in response

to noxious thermal or mechanical stimulation” (Robertson et al., 2013) and is an

effective but also very predictable wild-type reaction in response to danger (ibid).

Further still, the undulation and peristaltic motions found in this response are also

found in other behavioral responses, with one example described below.

1.4 Description of Ecdysis and Eclosion

A mechanical program of rhythmic movements where a larva detaches the old and

more rigid outer layer from new, pliable cuticle beneath in order to escape the

outgrown sheath (Bestman and Booker, 2003), ecdysis is an undulation reminiscent

of the aforementioned larval crawling movements. However, ecdysis lacks

transportation - the animal remains in the same location as when the movements

begin. The mechanics of this larval transitional behavior involve squeezing, pinching,

thrusting and twisting movements and are described as a series of dorsal-ventral

7

muscle contractions in all abdominal segments done to effectively detach the old

and new cuticles from each other (Park et al., 2002; Heckscher et al., 2012) (Figure 2).

Ecdysis begins with the secretion of ecdysis triggering hormone (ETH) from INKA

cells (Park et al., 2002; O’Brien et al., 1998). Next follows a positive feedback loop

consisting of ETH and eclosion hormone (EH) (Clarke et al., 2004). The appearance of

these modulating hormones set the stage for ecdysis with apolysis, the separation of

the epidermis from the old cuticle (Brena et al., 2012; Sugumar et al., 2013). Here, the

cycling between ETH and EH serves two purposes: 1) it promotes epidermal cell

proliferation and the secretion of a cuticulin layer meant to buffer the new, pliable

cuticle from enzymes digesting the endocuticle, a component of the old cuticle

whose contacts must be severed (Ito et al., 2008; Talbo et al. 1991; Mitchell and

Petersen, 1989) and 2) it stimulates the secretion of crustacean cardioactivepeptide

(CCAP) to be released in vesicles co-packaged with bursicon from CCAP neurons

(NCCAP) (Woodruff et al., 2008). Bursicon therefore canonically ends the ecdysis

hormone cascade when discharged at the neuromuscular junction (NMJ) (Loveall

and Deitcher, 2010), now able to activate its receptor rickets (rk).

8

Figure 2 - Signature ecdysis behavior. In order to separate old from new

cuticle, shed outgrown mouthparts and detach old tracheal trunks larvae

must perform a series of stereotyped movements involving thrusts,

pinching, stretching and twisting. This is physically exertion and may tax

the animal. Bouts of rest are interspersed for minutes at a time between

periods of active ecdysis. In total the sequence takes up to 30 minutes

from onset to forward escape (FE). (Park et al., 2002)

9

When 20E is released from the ring gland the larvae will cease feeding, begin

wandering and will initiate the imaginal molt soon thereafter (Winbush et al., 2011;

Gaziova et al., 2004; Roth et al., 2004). At this time, the stiff cuticle will dry out to

form the puparium so that adult structures may form within the pupal case. While

inside, additional hormones provoke stimulate such developmental milestones as

head eversion to occur. In particular, the secretion of juvenile hormone (JH) and 20E

together will instigate head eversion, a series of abdominal muscle contractions

meant to push tissues forming the thorax and head capsule toward the anterior end

(Tögel et al., 2013; Riddiford et al., 2010; Hammock and Newitt, 1986). Once the

metamorphosis is complete the adult cased within will break free of the cuticle in a

final ecdysis, termed eclosion.

1.5 Peptidergic Neurons

Neurons comprise the core framework of the eukaryotic nervous system.

Neurosecretion, the process by which vesicles are delivered to the synapse, is the

fundamental line of communication between a neuron and its target. In Drosophila

melanogaster the ventral ganglia project axons from the central nervous system

directly onto muscles of the animal’s body wall, creating a neuromuscular junction

(NMJ). The synaptic boutons resulting from these NMJ contacts contain

neuropeptides packaged into dense core vesicles (DCVs). These vesicles are then

10

secreted at the NMJ to regulate the activity of other neurons or into the hemolymph

to act on other tissues. Neuropeptides thus influence a multitude of processes and

behaviors within an animal’s development. To this end one regulating transcription

factor of note, DIMM, was identified by its pattern of peptidergic neuron expression

via an enhancer-trap screen (Hewes et al. 2003). DIMM is particularly important in

this conversation because it promotes the differentiation of neurosecretory

properties in many neurons (Chen et al., 2014; Hamanaka et al. 2010; Park et al. 2008;

Hewes et al. 2003).

1.6 Dense Core Vesicle Formation and Transport

In a eukaryoktic cell the process of protein synthesis and packaging occurs within the

cell body. After transcription, ribosomes lining the endoplasmic recticulum (ER)

translate mRNA into protein. The protein product is then packaged by the Golgi

complex into vesicles and trafficked throughout the cell. In the case of endocrine

neurons, when vesicles are filled with neuropeptides they must be shipped along the

axon to the synapse (Siegel et al., 1999; Shakiryanova et al., 2005). It is a process

requiring lengthy intracellular transport to reach target secretion sites. This delivery

occurs by active transport as the matured vesicles are tethered to molecular motors

(kinesins) able to ‘walk’ along microtubules to their destination at the boutons

(Siegel et al., 1999; Shakiryanova et al., 2005; Kang et al., 2014). Because these

11

messengers are signaling agents meant to trigger a downstream response, the dense

core vesicles bind to the synaptic membrane in order to release its contents. In

contrast to neurotransmitter synaptic vesicle secretion (which occurs on the

millisecond timescale), peptidergic secretion caters to long range, long term

consequences after numerous action potentials (Shakiryanova et al., 2005).

1.7 Vesicular Fusion Machinery

The process of fusing vesicles to synaptic membrane is facilitated by the coupling of

fusion proteins. The canonical view of vesicular trafficking and secretion holds that

Soluble NSF Attachment REceptors (SNAREs) are the molecular latches by which

neurotransmitter-filled vesicles dock at and fuse to the synaptic membrane (Söllner

et al., 1992; Vavassori et al., 2014; Stevens et al., 2011; Zhang et al., 2011; Jorquera et

al., 2012; Vilinski et al., 2002; Xu et al., 2002). In particular, vesicle-associated-SNAREs

(v-SNAREs Synaptobrevin or VAMP) dimerize with target plasma membrane-

associated-SNAREs (t-SNAREs SNAP25 and Syntaxin 1) to form a coiled-coil four-

helix SNAREpin (Cao et al., 2014; Xu et al., 2002). Once v- and t-SNAREs have

complexed, the vesicle is docked at the release site and the bi-lipid membranes fuse,

discharging the contents into the synapse. For neurotransmitters a recycling

mechanism is employed where reuptake occurs via pumps at the synaptic cleft

(Podufall et al., 2014). In contrast, however, neuropeptides are not recycled but

12

instead made anew at the ER and trafficked to the synapse. Still, many factors

regarding peptidergic and DCV trafficking and secretion remain elusive.

In addition to the aforementioned v- and t-SNAREs (alternately R- and Q-SNAREs,

respectively), other proteins and molecules play critical roles at the synapse as well.

Ca2+ creates an environment conducive to SNARE complex formation, allowing DCV

fusion (Kasai et al., 2010, Cao et al., 2014); Synaptotagmin acts as a Ca2+ sensor for

release following an action potential (Vavassori et al., 2014; Xu et al., 2002);

Glutamate, considered to be the major mediator of excitatory signals, has been

implicated in functions including cognition, memory and learning (Johansen et al.,

1989); Discs Large a major postsynaptic NMJ scaffolding component (Ataman et al.,

2006); bruchpilot (BRP) an active zone (AZ) protein known to both cluster Ca2+

channels and aggregate synaptic vesicles (SVs) at the AZ (Ehmann et al., 2014); Rop,

essential for its role in late stages of secretion (Bate and Broadie, 1995); and

Complexin (Cpx) is a cytosolic protein which has been implicated as a co-factor in

the SNARE-clamp model (Cho et al., 2014; Giraudo et al., 2006).

1.8 The Developing Larval Brain

In comparison to mouse or humans the insect brain has few nerve cells. With only

approximately 100,000 neurons (Chiang et al., 2011) the Drosophila central nervous

13

system (CNS) an ideal system in which to explore neurobiological questions

(Cardona et al., 2010; Clarac et al., 2007; Boyan et al., 2011). Not all neurons found in

the adult brain exist at the time of an egg hatch and so some later neural

connections will form as the animal develops. At time of laying, the average length

of a fertilized egg is ~0.5mm (Markow et al., 2009; Ashburner et al., 2005) eventually

maturing to an adult size of about 2.5mm (Kamakura 2011).

The juvenile stages for the Drosophila life cycle is characterized by rapid growth.

Under optimal conditions, a larvae must increase its mass nearly two hundred-fold

over the approximate 4–5 days between newly hatched L1 to wandering L3

(Nagarajan et al., 2014, Church et al., 1966). During this phase the animal’s neural

architecture is also undergoing a notable change. Newly hatched L1 larvae are

reported to have ~1,500 differentiated primary nerve cells per lobe (Cardona et al.,

2010; Larsen et al., 2009; Ito and Hotta, 1992). Between late embryogenesis and late

1st instar this number remains fixed until mitosis resumes (Ito and Hotta, 1992). At

this point the neuroblasts will undergo ~40-75 additional mitotic divisions to

produce secondary neurons (Larsen et al., 2009; Dumstrei et al., 2003).

Of the nerve cells comprising the larval CNS, approximately 42-46 neurons will be

14

CCAP-immunoreactive (Veverytsa et al., 2012; Karsai et al., 2013). Specifically, pairs

have been found to originate from the following places:

“2 pairs with somata in the brain, 1 pair with somata in the subesophageal

neuromeres (sog) sog1 and −3, 1 pair in thoracic neuromere t1-t2 and

abdominal neuromere a5–9 and 2 pairs in sog2, t3 and a1–4. In t3-a4 -the

ventral ganglion…[] two pairs…[] exit via the respective segmental nerve.”

(Karsai et al., 2013).

It is these two NCCAP cells from the ventral ganglion which will project onto muscles

12 and 13 to form type III neuromuscular junctions (NMJs) (Loveall and Deitcher,

2010; Zhao et al., 2008; Dulcis et al., 2005; Luo et al., 2005). After the L3 wandering

stage the larvae will pupate for metamorphosis. During this metamorphosis the

larval wiring will differentiate into that of an adult brain, allowing two additional cells

to become CCAP- and bursicon-immunoreactive. These cells are located in a5–a7

and a9 (Karsai et al., 2013; Veverytsa et al., 2012). Therefore, by the time of pupal

metamorphosis the animal will have nearly 54 CCAP neurons in total.

1.9 Synaptic Bouton Categories

Morphology of synaptic boutons vary greatly. As such the synaptic boutons resulting

from these NMJ contacts come in three categories and are characterized by their

contents. Type 1 big (Ib) is the largest of the boutons. The majority of its contents

15

are clear, small synaptic vesicles (SSVs) containing the neurotransmitter glutamate

(Wong et al., 2012; Bulgari et al., 2014; Budnik and Gramates, 1999). Type 1 small (Is)

follow suit with content while differing in size as Ib boutons range from 3-5µm while

Is boutons range in size from 1-1.5µm (Johansen et al., 1989). Type II NMJs are

defined by having small, clear vesicles (SVs) of glutamate as well as dense-core

vesicles (DCVs) containing nerumodulators such as octopamine (Budnik and

Gramates, 1999). Lastly, type III contacts – described as ‘insulin-immunoreactive’

terminals (Gorczyca et al., 1993) – are intermediate in bouton size, between types I

and II terminals. Moreover, peptidergic type III boutons consist mostly of DCVs but

do contain a small cache of SSVs. The fast-acting kinetics of neurotransmitter

biology has historically drawn much attention to type I NMJ terminals; of the three

classes the type I bouton properties and defining characteristics have been studied

under a multitude of experimental conditions in a variety of animals (Arakanea et al.,

2008; Huage et al., 2007; Donini et al., 2001; Bestman and Booker, 2003; Gammie and

Truman, 1997). In contrast however, little is known about type III synaptic terminal

bouton trafficking dynamics – namely, how type III boutons circulate its peptidergic

vesicles along a series of boutons for impending discharge. And so, given that

endocrine cells generate substantially more vesicles when compared to transmitter-

releasing cells (Bulgari et al., 2014), information such as how exactly peptidergic

granules are trafficked and docked at the synaptic membrane will remain to be

16

studied.

1.10 Bursicon as a Marker and Tool

Neuropeptides are molecules secreted into the hemolymph via the synapse by

neuroendocrine cells in response to their excitation. As signaling agents,

neurohormones and neuropeptides are involved in a wide range of physiological

events and, as a feature, are shared across diverse taxa from insects to mammals.

One such molecular messenger is bursicon (burs/pburs), a 30kDa-sized

heterodimeric cysteine knot protein related functionally to human bone morphogens

(Luo et al., 2005), distantly related to vertebrate glycoprotein hormones (Claeys et al.,

2005) and found in the insect Drosophila melanogaster. As previously stated, the life

cycle of a fruit fly is marked by multiple developmental milestones as the fly

develops from fertilized egg to reproductively viable adult. One of these

developmental landmarks is termed eclosion, the moment when an adult insect

emerges from its pupal case. Through head ligation studies bursicon was implicated

as the post-eclosion hormone that initiates the melanization and sclerotization of

adult cuticle (Fraenkel et al. 1966). Melanization and sclerotization – or darkening

and hardening – are governed by bursicon and crucial for fly survival post-eclosion

since the two events are germane to issues of immunity, thermoregulation and even

17

mobility (e.g. – flight, by way of stiffening the wing).

Bursicon is exclusively secreted from the type III boutons projected by NCCAP cells

onto muscle fibers 12 and 13 in the Drosophila larval body plan (Loveall and

Deitcher, 2010). This exclusivity gives bursicon the distinct advantage to serve as a

marker defining these boutons. Through a series of colocalizational studies between

bursicon and suitable candidates, a preliminary registry can be compiled in efforts to

establish an immunohistological staining profile. This directory will identify proteins

involved in neuropeptide trafficking and secretion. An additional advantage is

bursicon’s two-staged release. Initially only thought to be associated in adult insect

cuticle tanning and wing expansion, recent evidence from this lab uncovered a pre-

ecdysis release, prompting us to speculate the role of such an early discharge. In

experiments where either bursicon or its receptor rickets is knocked down the

developing larva has a greater mortality rate as when compared to wild type.

Moreover, those surviving to adulthood are left physically impaired (Loveall and

Deitcher, 2010) and unable to liberate themselves from the puparium. Further still, in

silencing proteins we suspect are involved in peptidergic vesicle docking, we observe

larval death at times critical to ecdysis. Lastly, bursicon’s exclusivity to type III

boutons lends itself for use to directly study the process of secretory granule

trafficking under a variety of conditions. While what is known about the mechanisms

18

of neurotransmitter-filled vesicle trafficking and employed by type I boutons

continues to be expanded (Wong et al., 2012), information on trafficking of large

dense core vesicles in neuropeptidergic type III boutons remains elusive.

1.11 Channelrhodopsins

Channelrhodopsins (ChR) are light-reactive proteins. First described in green algae

(Sineshchekov et al., 2005; G N et al., 2013), they are members of the G-Protein

Coupled Recepter (GPCR) family. By definition they will undergo a conformation

change in response to absorption of a particular wavelength. Specifically, light causes

Retinal-A to disassociate from the rest of the GPCR. Because channelrhodopsins are

channels, this conformational change ‘opens’ the gate without the need of a second

messenger. This lends itself to ultrafast kinetics: channelrhodopsins react within 50

milliseconds of illumination (G N 2013). In this permissive state cations will enter a

cell, depolarizing it in the process (Klapoetke et al., 2014; G N, 2013; Sudo et al.,

2013; Zhang et al., 2007).

Channelrhodopsins thus offer precise control over targeted cells. Given a genetically

defined neuron type, one can study their roles in vivo and in an isolated manner. For

this reason the optogenetics of channelrhodopsin offers greater spatiotemporal

resolution as compared to alternate forms of cellular control. As an example,

19

pharmacological methods are specific to the cell type but can have significant off-

target effects (G N et al., 2013). This issue is circumvented in using

channelrhodopsins as they are genetically targeted to an individual class of cells,

exclusively to those cells, and only in the presence of the desired wavelength of light.

Lastly, if a peptidergic nerve cell were under optogenetic regulation, action

potentials resulting from optical stimulation would presumably trigger secretion of

vesicles at the synapse.

1.12 ANF-EMD, the Bursicon mimic

ANF-EMD is a rat neuropeptide (Atrial Natriuretic Factor) tagged with a GFP-variant

(EMeralD). It is a fusion protein able to be targeted to secretory granules in the same

manner that endogenous neuropeptides are handled (Rao et al., 2001). This is an

important feature as protein modifications can alter how a cell processes an

exogenous protein. Additionally, because it is GFP-tagged, there is the unique added

benefit that the process of its packaging, trafficking and secretion can be monitored

in vivo. As an example, when packed in LDCVs the fusion protein is able to form

bright puncta. Therefore, because the fluorescent intensity is so reliably bright, ANF-

EMD has the distinct advantage to be used in single granule release studies meant to

analyze the path taken by an individual ANF-EMD-filled vesicle upon trafficking to

and docking at the synaptic boutons. Further still, when concentrated at nerve

20

terminals, a comparison between glutamate-containing clear vesicles in type I

boutons and LDCVs of type III boutons revealed that 60% of the tagged

neuropeptide was released from NMJs of either category upon stimulation (ibid).

This result confirms that the fluorescent peptide can dependably be used to evaluate

and contrast differences observed in the mechanisms controlling vesicle secretion

occurring at different bouton types. Altogether, these characteristics lend it to being

a valuable tool for the study of vesicular secretion.

21

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33

CHAPTER TWO:

Secretory protein distribution and perturbed vesicle release from type III boutons at

the Drosophila melanogaster larval NMJ

2.1 Abstract

Bursicon is a neurohormone packaged in and secreted by type III synaptic contacts.

In comparison to the depth of information describing type I and type II

neuromuscular junctions (NMJs), type III NMJs are notably ill-defined. As such, this

lab has provided evidence identifying bursicon as a reliable marker in an aim to

characterize type III NMJs. Firstly, a registry of proteins defining type III NMJs can be

compiled through antibody-labeled colocalizations using bursicon as the type III

marker. Candidates to be stained with bursicon include established presynaptic

markers such as the SNARE proteins. Positional overlap is a protein-based search

that can reveal the neuroanatomical distribution of whichever proteins we target via

fluorescent antibody-labelling and the dual antibody-staining silhouettes will be

confirmed by sequential-excitation confocal microscopy. This approach establishes

an immunohistological staining profile characterizing type III NMJs. Secondly,

creating an immuno-fingerprint for type III NMJs also produces a directory

identifying the proteins involved in secretory vesicle (SV) delivery at these boutons

and can be used as a prototypical example to understand factors involved in granule

34

secretion in general. Therefore, using a GFP-variant fusion protein able to mimic

Bursicon’s packaging and transport in vesicles, we can perturb in vivo delivery of a

fluorescent neuropeptide to describe type III synaptic release. Moreover, various

methods of perturbation are used to assess the function of ecdysis-relevant proteins

at these boutons. This experimental design elucidates the dynamic process of the

vesicle trafficking system specifically employed by type III NMJs and adds to the

breadth of knowledge gathered about the mechanism used during peptide release.

2.2 Research Design

In order to evaluate how secretory granules get trafficked and captured at type III

peptidergic boutons our lab first aimed to verify which proteins can be found at the

terminal. SNAREs previously implicated in neurotransmitter delivery at type I boutons

offered a valid starting point. Because bursicon is found exclusively at type III NMJs

(Honneger et al., 2008; Loveall and Deitcher, 2010) it therefore has the distinct

advantage to serve as a marker defining these boutons. To that end, our lab decided

to resolve which proteins comprising the type III NMJ synaptic proteome also co-

localized with bursicon, the type III bouton marker. The approach of positional

overlap is established via antibody immunoreactivity and fluorescent microscopy

techniques.

35

Once an immunohistological staining profile defining type III terminals is established

we can then assess the factors critical to SV capture via perturbational studies.

Dense core granule release of bursicon (endogenous) or of ANF-EMD (exogenous)

occurring at the type III NMJ are compared and contrasted in the same cell type.

Furthermore, release dynamics can be contrasted between natural developmental

timing and induced release by optogentic stimulation. Lastly, mistimed dense core

vesicle release studies offer valuable insight into the tractability (or rigidity) of the

hormone cascade governing a behavioral program such as ecdysis.

2.3 Type I and type III Boutons Have Similar SNARE Inventory but are Not

Glutamatergic

The canonical view of vesicular trafficking and secretion holds that Soluble NSF

Attachment REceptors (SNAREs) are the molecular latches by which

neurotransmitter-filled vesicles dock at and fuse to the synaptic membrane (Stevens

et al., 2011; Zhang et al., 2011; Jorquera et al., 2012; Vilinski et al., 2002). Fluorescent

antibody-labelled Bursicon is shown to positively colocalize with a variety of SNARE

proteins and other stereotypical synaptic candidates including the secretory protein

Rop, and SNAREs Syntaxin, neuronal Synaptobrevin, Snap 24 and Snap 25 (Figure 3).

Because Snap 24/25 antibody cross-reacts with Snap 24 protein in addition to Snap

25, Drosophila mutants had to be used. Snap 25 null flies (Snap 25mx124) (Vilinsky et

36

Figure 3 - Type III Bouton Proteome Contain a List of SNARE

Proteins. (A-D) Confocal images of a Drosophila larval

neuromuscular junction (muscles 12 & 13) showing the synaptic

terminals of NCCAP motor neurons that form type III boutons.

Release sites (green) were labeled with an antibody recognizing the

type III-exclusive protein Bursicon (Loveall et al., 2010). (E-H)

Colocalized proteins (red) were labeled by the antibody identified.

(L, panel inset) Magnification of representative type III boutons

(arrowhead) with Rop protein localized to the perimeters.

37

al., 2002; Rao et al., 2001) were crossed to a line containing a deficiency spanning the

Snap 25 region. One-quarter of the resulting F1 progeny expressed Snap 24 protein

only. These Snap 25 null larvae were dissected and antibody-labelled to reveal only

the Snap 24 protein staining pattern when the Snap 24/25 cross-reactive antibody

was used. As a result Snap 24 was revealed to be found at type III NMJs in low

concentrations, as compared to Snap 25 in wild type animals. Additionally, cysteine

string protein (CSP), a protein regulating vesicle exocytosis by affecting presynaptic

Ca2+ channels (Chamberlain et al., 1999), was found at the type III NMJ (data not

shown). In contrast to these colocalizations with Bursicon, the active zone marker

Bruchpilot and the SNARE-linked Complexin both failed to colocalize with Bursicon

at type III synapses (Figure 4).

Furthermore, though synaptic contacts were generally presumed to be glutamatergic

(Bate and Broadie, 1995; Johansen et al., 1989) our lab has evidence disputing this

expectation. Using the yeast-based GAL4/UAS system we drove expression of ANF-

EMD, a GFP-variant tagged rat neuropeptide atrial natriuretic factor targeted to

secretory granules like endogenous neuropeptides, under the control of vesicular

glutamate transporter (VGLUT-GAL4), limiting the GFP signal to glutamatergic neurons

(Figure 5). After staining with anti-Bursicon antibody (shown in red) no overlap of red

and green signals were observed in any dissected animals, implying that type III

boutons are distinctively not glutamatergic.

38

Figure 4 - The Active Zone marker Bruchpilot & the SNARE

-associated Complexin protein both Fail to Colocalize with Bursicon

at type III Boutons. Two representative images showing Bursicon-labelled

type III contacts (green) with non-overlapping proteins, Bruch Pilot and

Complexin, respectively (red).

39

2.4 Syntaxin Knockdown (Syx1A) Results in Over-filled Bouton Phenotype &

Larval Death

The t-SNARE Syntaxin is shown to be present in the cell body and axons of neurons.

However, to determine if it plays a role in peptidergic vesicle trafficking and/or

release we employed the use of RNAi. After Syntaxin RNAi (Syx1A-RNAi) was

specifically expressed in NCCAP, titrated anti-Bursicon antibody was used to evaluate

any perceivable effects on Bursicon abundance at the type III NMJs in 2nd instar

larvae. Comparing the average observed fluorescent intensity of control animals

(UAS-only) to knockdown animals at the same developmental stage revealed

boutons with greater anti-Bursicon fluorescent intensity, signifying an abundance of

Bursicon (Figure 6 A, B). Additionally, when NCCAP-GAL4>UAS-Syx1A-RNAi animals

were reared on shallow grape plates to better monitor their progress, it was

observed that larvae died at particular developmental stages. For these animals it

was observed that larvae would die disproportionately at either double mouth hooks

(dMH) or double ventral plates (dVP) – two developmental markers known to

foreshadow the onset of ecdysis between larval molts (Figure 6 C). The penetrance of

this larval death phenotype is incomplete, however, as some viable NCCAP-

GAL4>UAS-Syx1A RNAi adults can be recovered from the cross.

40

Figure 5 - Type III Boutons are Not Glutamatergic. Two examples

showing GFP-tagged vesicular glutamate with Bursicon-labelled type III

contacts (red). (E-F) Glutamatergic boutons (arrowheads) and type III

boutons (arrows) are situated closely together but are distinct. n=3

animals.

A B

C D

E F

41

2.5 Knockdown of Synaptobrevin is Insufficient to Perturb Release at type III

NMJs

Synaptobrevin is a v-SNARE and n-Synaptobrevin null mutants are embryonic lethal

(Deitcher et al.,1999). Therefore, to determine if Synaptobrevin plays a role in

peptidergic vesicular trafficking, we opted to silence neuronal Synaptobrevin (UAS-

nSyb-RNAi) in NCCAP only. Because the fruit fly life cycle is punctuated with two larval

molts, 2nd instar of the NCCAP-GAL4>UAS-nSyb-RNAi genotype survive to adulthood,

seemingly unaffected by the RNAi. To confirm that knockdown had indeed occurred

NCCAP-GAL4>UAS-nSyb-RNAi animals were dissected, stained for both bursicon and

nSynaptobrevin (nSyb), and the type III NMJs were inspected. Simultaneous and

sequential confocal laser excitation then verified that when nSynaptobrevin was

considerably diminished, Bursicon presence at type III terminals remained unaffected

(Figure 7).

2.6 Timed Dissections Confirm Peak Bursicon Levels

In addition to a registry listing which proteins are involved in trafficking and

secretion at type III NMJs, our lab sought to describe and propose a model of

refilling. Here, refilling can be observed both endogenously (via bursicon

immunocytochemistry) or in vivo exogenously (via use of the ANF-EMD protein) as

42

the paths taken by the fluorescent granules can directly be monitored over time. As

an example, the model of the refilling which may occur in type III boutons is based

on the capture of vesicles in type I boutons, as proposed by a Wong et al., 2012

paper. If bouton capture occurs in the same manner, then for type III boutons: 1)

vesicles will accumulate in the most distal bouton first and 2) the remainder will fill

equally with inefficient capture occurring at each bouton as the vesicles are cycled

back towards the soma before another pass (Figure 8 A).

To begin this aim we first decided to further refine the timing of how type III boutons

discharge their contents over the course of the animal’s development. Therefore, in

order to describe the progression of this refilling process, wild-type and NCCAP-

GAL4>UAS-ANF-EMD were staged and dissected, with the dissections being timed

to developmentally significant stages including dMH and dVP. In agreement with a

study performed earlier in this lab, comparing both sets of animals confirms that an

early, preliminary discharge occurs in type III boutons. This discharge is shown to

occur hours before ecdysis is predicted to begin and occurs when using either the

endogenous protein, Bursicon or with the Bursicon-mimic, ANF-EMD is used for in

vivo imaging of live tissue preparations. Thus, this purging event can be measured by

Bursicon immunoreactivity (data not shown) or ANF-EMD fluorescence (Figure 8 B).

43

Figure 6 A - RNAi Knockdown of Syntaxin (Syx1A) Results in

Over-filled Bouton Phenotype. (A-D) Drosophila larval ventral

ganglion showing NCCAP cell bodies and Syntaxin knockdown

staining. (E-H) NCCAP NMJ boutons stained with the type III bouton

marker Bursicon. n≥25 boutons.

Vent

. Gan

gl.

UAS-Syx1A-RNAi only NCCAP-GAL4 > UAS-Syx1A-RNAi

Burs SyxI

Type

III N

MJ

Burs Burs Burs Burs

Burs SyxI

44

Figure 6 B – ImageJ Analysis Following RNAi Knockdown of

Syntaxin (Syx1A-RNAi). NCCAP NMJ boutons stained with the type III

bouton marker Bursicon (left panel) and the inverted image on which

ImageJ threshold quantification was performed (right panel). Two

representative animals are shown: (A) a driverless control and (B) active

silencing of Syntaxin at the NMJ. n≥25 boutons.

45

Figure 6 C - RNAi Knockdown of Syntaxin (Syx1A) Results in

Larval Death at Developmentally Significant Stages. (A-C) Wild-

type Drosophila larval stages. (D) Dead NCCAP-GAL4>UAS-Syx1A-

RNAi animals showing dMH (arrowheads) and dVP (arrows), two

phenotypic markers of development.

46

Figure 7 - RNAi Knockdown of nSynaptobrevin (nSyb) is Insufficient

to Perturb Release at Type III NMJs. Confocal images of Drosophila 2nd

instar neuromuscular junctions (muscles 12 & 13). Bursicon (green) and

nSyb (red) overlap is shown in yellow. Top row (A-C) Control larvae

(driverless; UAS-nSyb-RNAi construct only). Bottom row (D-F) Type III

boutons in NCCAP-GAL4>UAS-nSyb-RNAi larvae. n≥60 boutons.

47

Figure 8 A – Proposed Bouton Capture. (Top) Schematic to determine

type III bouton capture against developmentally-timed dissections.

(Bottom) Proposed refilling rationale. Boutons in series would refill ‘back

end first’ with inefficient capture occurring at each bouton as the vesicles

are cycled through before another pass. Here white signifies empty/no

boutons captured, black indicates a bouton filled to capacity, and the

diminishing levels of blue represent a bouton capture gradient).

48

Figure 8 B – Dense Core Vesicle Capture and Refilling at type III

Boutons. Confocal images of NCCAP type III boutons in NCCAP-

GAL4>UAS-ANF-EMD larvae. n=16 animals (2 larvae per stage)

Pro

gressio

n O

ver Time

49

2.7 Exogenous/Mistimed release of CCAP/Bursicon Reveals Even Refilling of

type III Boutons

Chrimson is a red light-drivable channelrhodopsin able to provoke neural spiking

and release on demand. The UAS-Chrimson construct has a yellow fluorescent

protein (YFP) tag embedded into its sequence, making it visible in the 500-550 hv

(green) wavelength at all times. When under the specific promoter control of CCAP

neurons, NCCAP-GAL4>UAS-Chrimson larvae have yellow-green marked type III

boutons and will develop normally from fertilized egg to fertile adult. However, when

mid-3rd instar animals are exposed to a red LED light source, action potentials are

induced and NCCAP activity is triggered, releasing vesicles docked at the synapse

(Figure 9 A). Anti-Bursicon antibody (green) was titrated to levels able to produce a

faint signal. This enabled us to monitor bouton refilling in the background of the

UAS-Chrimson construct by assessing the increase in green fluorescence above the

established background UAS-Chrimson levels (Figure 9 B). Under these conditions it

was observed that after inappropriately-timed release of CCAP- and Bursicon-filled

vesicles was initiated, type III boutons refilled in an even manner across the length of

the boutons (Figure 9 C). Additionally, 2nd instar animals were stimulated via red

LED-pulse then left to feed and not dissected. These animals would reach adulthood

but have increased odds of dying at various larval stages (not limited to dMH

50

Figure 9 A – Exogenous/Mistimed release via photo-activation of

UAS-Chrimson and Signal Recovery (A-C) NCCAP-Gal4>UAS-Chrimson

type III contacts immunolabelled with anti-bursicon. Mid-3rd instar larvae

were dissected and fluorescent images taken via wide-field microscopy at

various time points: (A) prior to red LED stimulation (RLS) via Chrimson, (B)

immediately after RLS and (C) several minutes post RLS. n=3 animals

51

Figure 9 B – Neuropeptide Fluorescence Lowered in NCCAP-

GAL4>UAS-Chrimson Larvae Post Red LED Stimulation (RLS).

Comparison of mean GFP intensity pre- and post-photoactivation via red

LED. Images used for analysis of bouton refilling were acquired from mid-

3rd instar animals, only. Animals dissected between the appearance of

double mouth hooks (dMH) and double ventral plates (dVP) were 2nd

instar and were omitted from this analysis.

52

or dVP) when compared to UAS-Chrimson (no driver) larvae.

2.8 Mistimed Activation of ETH Release Does Not Perturb type III Bouton

Capture

C929//DIMMED (Hewes et al., 2003) is a reliable tracheal INKA cell promoter.

Furthermore, INKA cells are responsible for the production of ETH. Therefore,

because ETH and EH are presumed to initiate the ecdysis cascade (Arakanea et al.,

2008; Hewes et al., 2003; Park and Trigg et al., 2008; Park and Taghert et al., 2008)

our lab aimed to test whether exogenous activation of INKA cells would lead to any

perturbation of steady-state levels for neuropeptides downstream of ETH/EH, such

as Bursicon. Using UAS-Chrimson as the light-sensitive trigger, C929//DIMMED-

GAL4>UAS-Chrimson larvae were exposed to a single 1h-, 2h- or 4h-long red LED

pulse. Post stimulation, larvae were dissected immediately. Upon observation these

C929//DIMMED-GAL4>UAS-Chrimson 3rd instar animals have the particular

phenotype of elongating their bodies on the anterior-posterior axis while under the

red LED light source. It should be noted that this stretched-out state resembles the

initial stage of ecdysis – where undulation meant to detach old cuticle from new is

53

Figure 9 C - Exogenous/Mistimed release of CCAP/Bursicon Reveals

Even Refilling of Type III Boutons. (A-C) NCCAP type III boutons in NCCAP-

GAL4>UAS-Chrimson larvae. The motor neurons that form type Is, Ib or III

boutons were labeled by anti-HRP immunoreactivity (red) and type III

boutons were marked by anti-Burs (green). Faint Chrimson-YFP signal

remains visible after Bursicon is secreted (middle row). Bursicon (green)

levels slowly begin to replenish, as indicated by an increase in green

fluorescent intensity along the boutons (bottom row).

54

punctuated by periods of stationary rest. Additionally, this inertia will last for as long

as they remain under the red LED light, suspending all activities including feeding for

the duration of exposure. However, after several days of rest these animals would

eventually die. Driverless control animals continued feeding, in stark contrast to the

C929//DIMMED-GAL4>UAS-Chrimson larvae. Further still, when exposed to a red LED

pulse both wild-type and driverless control animals perform evasive maneuvers if

injured. Specifically, when being pinned down before a dissection, these larvae

began stereotyped “corkscrew-like rolling around the anterior/posterior axis, in

response to noxious thermal or mechanical stimulation” (Robertson et al., 2013). In

contrast, however, C929//DIMMED-GAL4>UAS-Chrimson 2nd instar showed no

reaction to being pinned down after being exposed to the appropriate light stimulus.

In light of these behavioral deficits we decided to investigate whether elements of

the ecdysis hormonal cascade were compromised. This reasoning came about

because some of the mechanical movements we were no longer observing

(aforementioned above) are shared by the programs of ecdysis and wound evasion

in nocifensive escape behavior, alike. To address this we first aimed to determine if

provoked release of ETH, the hormone known to initiate the ecdysis hormonal

cascade, could trigger an early ecdysis event. As such, when inspected,

immunocytochemistry revealed that ETH staining colocalized with the

55

C929//DIMMED-GAL4 pattern (as predicted) (Figure 10). Because Bursicon is the final

hormone of the ecdysis cascade we decided to monitor its discharge at the boutons

after stimulating C929//DIMMED-GAL4>UAS-Chrimson larvae. Yet, after a period of

up to 2hours post LED stimulation, no change in anti-Bursicon antibody intensity

could be detected in type III boutons. However, an experiment meant to confirm that

ETH is indeed released from the INKA cells post LED stimulation had not yet been

performed. Therefore, these results must be taken as preliminary.

56

Figure 10 – Tracheal Stainings From 2nd Instar Larvae. Two representative

images showing INKA cells (mass at tracheal branch point; bright green) in

C929//DIMM-Gal4>UAS-Chrimson.

57

2.9 Methods

Fly stocks

UAS-Chrimson (55135) stocks were reared on 0.05mM all-trans Retinal-A ≥ 98%

(Sigma-Aldrich) supplemented standard media vials in the dark. All other stocks were

reared on standard media (sans Retinal-A) at 25°C in 12 h:12 h LD cycle. Where

required, larvae were reared on grape juice and glucose plates (3% agar) augmented

with yeast to improve their development. Except where mentioned, all stocks are

available from the Bloomington Drosophila Stock Center (accompanied by stock

number). The stock w1118 (6326) was used as the wild-type strain. NCCAP-GAL4 flies

were used to drive expression in type III boutons of UAS-ANF-EMD as a stock of

NCCAP-GAL4>UAS-ANF-EMD (a generous gift from BJ Loveall). Additional GAL4

drivers used in this study include: CCAP-GAL4 (25686); Bursicon-GAL4 (26719); and

C929//DIMMED (25373). Additional UAS lines used in this study include: Syx1a-RNAi

(25811); Dicer 2 (24650); Dicer 3 (25651); and nSyb-RNAi (49202). To document

double mouth hook (dMH) or double ventral plate (dVP) phenotypes, whole-larva

bright field images were taken on a Leica MZFLIII microscope and saved with Leica

IM50 (version 1.20) software (Loveall and Deitcher, 2010).

58

Immunocytochemistry

All animals were wandering 3rd instar larvae or were 2nd instar larvae displaying

double mouth hooks (dMH) or double ventral plates (dVP) – as noted. Individual

larvae were dissected in cold Ca2+-free HL3 (Loveall and Deitcher, 2010) and filleted.

For vesicle membrane protein experiments, preparations were fixed for one hour at

room temperature in 4% paraformaldehyde (PFA) or Bouin’s Fixative. Fillets were

thoroughly washed in PBSTx (PBS + Tris + 0.3% Triton X-100), and incubated in

PBSTx + 10% normal donkey serum (NDS) for 1 hour. Tissues were then incubated in

primary antibody overnight at 4°C, quickly rinsed and then washed 3 × 15 minutes in

PBSTx. Subsequently, preparations were incubated for 4 hours in secondary antibody

at room temperature, thoroughly rinsed and washed in PBSTx, and finally mounted

on slides in Vectashield (Vector Laboratories) and stored in the dark at 4°C. The one

exception was that larval tracheae processed for ETH-IR were mounted on poly-

lysine-D - coated cover slips, dehydrated through an ethanol series into xylene, and

mounted in DPX mountant (EMS). Primary antibodies used include rabbit anti-

Bursicon (BURS; 1:5000, or 1:20,000 for bursicon release analysis) (a generous gift

from Benjamin White), rat anti-nSynaptobrevin (N-SYB; 1:200; a generous gift from

Hugo Bellen), mouse anti-DCSP2 (CSP; 1:100; Developmental Studies Hybridoma

Bank), goat anti-HRP (1:400; Jackson ImmunoResearch Laboratories), rabbit anti-

DVGLUT (1:5000); rabbit anti-CCAP (1:5000; a generous gift from John Ewer), anti-

59

Syntaxin (SYXI; 1:20), anti-Synaptotagmin (SYT; 1:2000; a generous gift from Noreen

Rent), anti-Complexin (CPX; a generous gift from J Troy Littleton), anti-Rop (ROP;

1:50; Developmental Studies Hybridoma Bank), anti-Discs Large (DLG1; 1:1000;

Developmental Studies Hybridoma Bank); anti-Bruchpilot (NC82; 1:25;

Developmental Studies Hybridoma Bank), anti-SNAP 25 (1:200; Rao et al., 2001), anti-

SNAP 24/25 (1:200; Vilinski et al., 2002) and rabbit anti-ETH (1:180,000; a generous

gift from John Ewer). Larvae sacrificed at different stages were stained with

Horseradish peroxidase (HRP), a synaptic marker known to label types I, II and III

boutons at the NMJ in addition to anti-burs or anti-CCAP antibodies to reveal type III

boutons, whenever appropriate. For secondary antibodies, the relevant species was

used with excitations of either 488 nm or 594 nm (1:1000; Invitrogen). For

fluorescence imaging, histological preparations were viewed on a Nikon Eclipse

E600FN microscope at 40×. Images were collected with a SPOT2 camera (Diagnostic

Instruments, Inc.) as 8-bit monochrome with the SPOT32 software (version 2.2).

Optogenetics

Promoter-driven UAS-Chrimson larvae were reared in the dark on appropriate food.

Prior to dissection in cold Ca2+-free HL3, larvae were gently washed to remove food

particles and other debris. Next, unpinned animals were pulsed with a red LED light

source for 45sec – 100 sec durations. This was accomplished by connecting the red

60

LED light source to a wave function generator and was a gift from Ron Hoy and Gil

Menda. Animals were then immediately dissected under low-light conditions or

returned to the dark to be dissected at a later time point.

Fluorescence intensity measurements

The GFP-tagged 'atrial natriuretic factor' reporter, UAS-ANF-EMD, was expressed

with CCAP-GAL4 to monitor in vivo neuropeptide release from bursicon-releasing

NCCAP in CCAP>ANF-EMD progeny. Images from live fluorescent tissue were

collected as described above for histological preparations. To measure ANF-EMD

release from type III boutons in larvae, we chose six developmental stages that

include the 2nd larval ecdysis. To obtain larvae approaching this ecdysis, it was

necessary to collect NCCAP-GAL4>UAS-ANF-EMD embryos on grape juice plates with

3% agar, supplemented with yeast. The recognition of these stages is described in

the Results. Staged animals were dissected in Ca2+-free HL3 saline with a dorsal -

longitudinal incision and splayed open with pins on magnetic plates. For each stage

a distinct set of four animals was selected and all visible type III boutons were

photographed with the same exposure setting. Since NCCAP-GAL4-expressing

boutons occur bilaterally in NMJ 12 of the T3-A4 segments, the maximum number of

type III boutons that can be analyzed per animal is 5 pairs, although we were not

always able to visualize this maximum number. With ImageJ software, we empirically

61

determined a threshold at which we could select type III bouton area in animals

preceding ecdysis (at the 'double mouth hook stage'), and with this threshold value

we measured mean pixel value in type III boutons at all stages. To calculate average

fluorescence intensity, we multiplied the area (selected by the threshold) and mean

pixel value, and took the average of these values for each stage. Comparisons

between stages to determine percent release were calculated as [fluorescence

intensity0]-[fluorescence intensity1]/[fluorescence intensity0]. Single-factor ANOVA

tests were also performed to determine if the fluorescence intensity from adjacent

stages differed significantly from each other. As a control against large-scale

vesicular release at the NMJ, the pan-neural elav-GAL4 driver was used to express

ANF-EMD at all NMJs. Confirmation of bursicon release by BURS-IR was performed

identically (Loveall and Deitcher, 2010).

Imaging

Images acquired via Leica SP2 Confocal, a Zeiss LSM 710 or Nikon Eclipse E600FN

microscope at 40×. Equal exposure settings were maintained while comparing

experimental and control samples, or comparing samples across different time

points. All images were cropped and monochrome images were given their color

identities with ImageJ or Adobe Photoshop CS (version 8.0). Figures were finalized

with ImageJ or Adobe Illustrator CS (version 11.0).

62

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CHAPTER THREE:

Implications & Conclusions

In this study we explored the utility afforded by having Bursicon and its mimic, ANF-

END, be reliable in vivo markers for type III NMJs and be tools to directly study

peptidergic vesicle secretion. Monitoring bouton fluorescence intensity in larvae

following release can provide a spatiotemporal representation of secretory granule

refilling. To this end experimental studies using a fluorescent fusion protein able to

mimic bursicon’s packaging, transport and release can shed light on the vesicle

trafficking system employed by type III boutons, and extended to other peptidergic

contacts. Next a series of experiments aimed at exploiting the advantages presented

by using bursicon were carried out to assess secretory granule trafficking under an

assortment of perturbed conditions and circumstances. Approaches included the use

of null mutants and post-transcriptional silencing via RNA interference (RNAi).

However, because the process of neuropeptidergic delivery is essential for

development (Žitňan et al., 2003; Park et al. 2002; Dulcis et al., 2005), it has been

considerably important to examine perturbation of the system in a cell-specific

manner in order to deduce function. Here, optogenetic control becomes invaluable.

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Optogenetics is the use of light-sensitive proteins as an inducible switch to control

the spiking of a neuron. It is a method which reliably provides genetic control with

spatiotemporal resolution. Examples of these include channelrhodopsin and other

members of the rhodopsin family (Ernst et al., 2014; Lim et al. 2014). Yet despite its

popularity, the use of optogenetic tools has expanded at a slower pace for

Drosophila than it has in a variety of other model organisms (Klapoetke et al., 2014;

Zhang et al. 2007), possibly due to light not being able to penetrate the larval fruit fly

cuticle in a dependable way (Klapoetke et al., 2014). To circumvent this issue we used

an atypical channelrhodopsin: Chrimson, a red-light drivable opsin with an excitation

peak at ~600 nm – about 45 nm red shifted from previous channelrhodopsins (ibid).

Intracellular recordings from muscles in 3rd instar larvae expressing Chrimson in

motor neurons demonstrated the channelrhodopsin’s ability to elicit action

potentials in larval Drosophila motor neurons (ibid). Thus, we optogenetically

activated neuroendocrine cells in order to stimulate secretion and produce certain

stereotypic behavior associated with the ecdysis signaling pathway. Further still,

because UAS-Chrimson is tagged with YFP it too can be used as a reliable marker to

outline type III terminals when it is under the promotion of a type III NMJ-specific

driver such as NCCAP-GAL4 or Burs-GAL4.

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At type III boutons we were able to individually visualize Snap 24, Snap 25, Rop,

Syntaxin, and nSynaptobrevin. Together these proteins comprise a short list of

factors which are directly involved with vesicle docking in that they are all

components of the SNARE vesicle delivery model. However, not all pre-synaptic

proteins co-immunostained with Bursicon.

Bruchpilot is considered to be a key active zone (AZ) component, critical for its role

in neurotransmitter release (Ehmann et al., 2014; Liu et al. 2011; Shahidullah et al.,

2013). Our lab provides evidence of no overlap when Bruchpilot is co-

immunostained with the type III NMJ marker, Bursicon. Another AZ-associated

marker, Complexin, is a cytosolic protein found at types 1s and 1b boutons. Known

to bind Snap 25 and Syntaxin – two components of the SNARE complex in the

‘clamp fusion’ model (Bykhovskaia et al., 2013; Iver et al., 2013) – Complexin also

shows no colocalization with Bursicon at peptidergic boutons. In contrast to

neurotransmitter secretion however, peptidergic secretion is not associated with the

AZ of the synapse. In particular, LDCVs are known to release from sites away from

the AZ (Shakiryanova et al., 2005). As such, finding no colocalization between

Bursicon and either Bruchpilot or Complexin stands to reason. Moreover Discs Large,

a post-synaptic marker, was stained as a negative control against pre-synaptic

Bursicon (data not shown).

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Lastly, our focus on proteins found at type I terminals led to questioning the

assumption that all bouton types are glutamatergic. To this end our lab provides

evidence that glutamate, an excitatory neurotransmitter (Johansen et al., 1989;

Grygoruk et al., 2010; Dickman et al., 2012; Daniels et al., 2011), is not associated with

peptidergic vesicles by way of showing that its transporter, vesicular glutamate, does

not co-immunostain with Bursicon. Together these absences may indicate

specialization between bouton types within a cell because the vesicle-associated

proteins of type I and type III contacts are similar but not identical. Since the list is

associated with what is currently known to be found at glutamatergic type I boutons,

and understanding that type III boutons are not glutamatergic, it stands to reason

that some, but not all proteins would be respresented in both neurotransmitter-

associated contacts (containing small, clear synaptic vesicles) as well as in

neuropeptidergic contacts (containing large, dense core vesicles).

SNAREs which do show fluorescent antibody overlap with Bursicon can also provide

much information on the properties of peptidergic secretion. For instance: Normal

development occurs in spite of dampening the nSynaptobrevin signal at these

boutons. Therefore, if nSyb does play a part in vesicular secretion employed by

neuroendocrine cells such as NCCAP, then this role has yet to be uncovered. Currently,

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the silencing performed in this study did not produce a discernable phenotype.

Therefore, additional studies incorporating Dicer may prove useful in bolstering the

knockdown efficiency against nSynaptobrevin.

In contrast, the over-filled boutons observed when silencing the t-Snare Syntaxin

suggest a function for Syntaxin in vesicle secretion. To that end our lab proposes the

following scenario: in the absence of vesicles at the synapse a neuron will create and

target newly-formed secretory granules to the synapse. However, if vesicle secretion

is perturbed by removing a molecule necessary for vesicle fusion (Syntaxin), it is

likely the neuron will continue to target vesicles to the terminals. If the cell persists in

producing new secretory granules while secretion is ineffective, the result is an over-

accumulation of secretory granules at the bouton.

Another consequence of reducing the expression of Syntaxin at type III junctions is

the increased likelihood that a larva will die at either dMH or dVP. The importance of

this observation is that both dMH and dVP are visual, developmental markers which

stereotypically precede the onset of ecdysis cuticle shedding behavior and suggests

that a critical quantity of either CCAP or Bursicon or a combination of the two must

be met to advance onto the next developmental stage. Moreover, Bursicon has been

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shown to release at a time point between dMH and dVP – namely before the post-

ecdysis time established in the literature (Loveall and Deitcher, 2010). As such, if this

pre-ecdysis Bursicon discharge is meant to prepare the animal for an impending

ecdysis, perhaps disabling the release mechanism diminishes the levels of secretory

granule actually released to dangerously low concentrations. If this is the case then

the animals that survived were presumably affected at a time when there is a long-

enough window to compensate for the lowered rate of secretion, before either the

first or second Bursicon purge. Finally, because cell death is also not observed in

these conditions, it is feasible to propose that the deleterious effects of an

incapacitated CCAP-Bursicon discharge can be reversed over time.

Alternatively, reducing Syntaxin expression may result in animals dying at dMH/dVP

stages because this was done specifically in CCAP neurons (NCCAP). As mentioned

previously, NCAAP co-package peptides within the same granule; in this case secretory

vesicles are filled with both CCAP and Bursicon, together. Therefore, disabling

Syntaxin will reduce the amount of CCAP or Bursicon discharged when the animal

deems it necessary. Remembering that these neurohormones are part of the ecdysis

hormone cascade, their functions are ecdysis-related where CCAP is involved for

muscle contractions and Bursicon for hardening of the cuticle. Therefore, NCAAP-

Gal4>Syx1A-RNAi animals may have a higher likelihood of dying at the dMH/dVP

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stage developmental stages because either the proper muscle contractions are not

occurring during ecdysis (from lack of CCAP being discharged), or from the cuticle

fragility (from lack of Bursicon being discharged), or a combination of the two.

Lastly, although findings of this study suggest even distribution and docking of

newly-targeted granules into empty boutons as they refill in series (like pearls on a

string), further studies are needed to determine if this is simply an artifact of

exogenous release. Here, previous studies are able to offer valuable guidance and

insight. In particular, a Wong et al., 2012 paper provides evidence that bouton

capture occurs in a distal to proximal manner in type I boutons, with accumulation

occurring initially in the most distal bouton. Furthermore this capture is termed

‘inefficient’ as only a fraction of the transient vesicles are captured during each

passage. In contrast, type III DCV capture has been shown to occur with

accumulation occurring in the most proximal bouton first (Bulgari et al., 2013).

However, it is crucial to note that both of these studies were done via

photobleaching by way of Simultaneous Photobleaching And IMaging (SPAIM) or

Fluorescence Recovery After Photobleaching (FRAP), respectively. This is to say that

the single puncta of fluorescence (representing an individual vesicle) is not in an

empty bouton. Instead, the fluorescence being monitored is in a field of other

vesicles, but these other vesicles were made invisible post-photobleaching. The

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importance of this is that the presence of other vesicles may interfere with the path

an individual vesicle may take. In contrast, the study proposed here is differs in that

the boutons are devoid of vesicles after an endogenous purge (such as what occurs

with Bursicon some hours before ecdysis). Therefore, observing the path of newly

targeted secretory granules in the absence of other vesicles may yield entirely

different results and can add further insight to the process of vesicle trafficking.

In the case that refilling is as described, however, another difficulty to be addressed

is the timing of the hormones involved in the ecdysis program. It can be reasoned

that susceptibility to undergo ecdysis has a window of opportunity. Specifically,

inappropriately timed spiking of INKA cells led to mistimed ETH release. As a

consequence, animals were prompted to begin the ecdysis sequence. However these

animals must then stop short of the full ecdysis program because elements

downstream of ETH could not be appropriately triggered. This outcome itself can be

due to either an ETH threshold not being met or because elements of the cascade

cannot be triggered outside of their preprogramed times.

Comparing secretion under endogenous versus inducible conditions at the same

motorneuron contacts can further reveal the importance of bouton-specific

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properties. Key challenges for follow-up experiments include expanding the list of

candidate molecules necessary and sufficient for peptidergic vesicle trafficking and

secretion, determining how quickly Bursicon-filled vesicles are targeted to restock

type III boutons and establishing whether the same pattern of bouton refilling which

occurs at type I contacts (Wong et al., 2012) are employed by all the bouton classes.

Moreover, improvements can be made on the recruitment of vesicles to the synapse

for release. In particular, it has been reported that in optogenetically-controlled

peptidergic neurons co-misexpression of DIMM encourages production of vesicles

(Hamanaka et al. 2010). Therefore this enhancement can be utilized specifically to

study refilling at the bouton. As an example: If type I contacts were modified to have

as many vesicles as type III, then it would be valuable to see if vesicle trafficking

would remain the same in these modified type I terminals or if they would instead

mimic what occurs at type III. Finally, results of this study and in the studies

proposed for the future can be used in efforts to identify, define and more fully

understand the mechanics occurring in peptidergic vesicle secretion. Such findings

may lead to greater developments in a variety of fields of study including offering

insight into processes involving hormonal regulation.

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