Treatment with the α-Synuclein Oligomer Prone Mutants

E35K and E57K Leads to Significant Intracellular

Aggregation and Inclusion Formation

by

Sri Dushyaanthan Sri Renganathan

A thesis submitted in conformity with the requirements

for the degree of Master of Science

Institute of Medical Science

University of Toronto

© Copyright by Sri Dushyaanthan Sri Renganathan 2015

ii

Treatment with the α-Synuclein Oligomer Prone Mutants E35K and

E57K Leads to Significant Intracellular Aggregation and Inclusion

Formation

Sri Dushyaanthan Sri Renganathan

Master of Science

Institute of Medical Science

University of Toronto 2015

Abstract

Alpha-synuclein (α–syn) is a key neuronal protein that undergoes pathogenic conformational

changes and accumulates in Parkinson’s disease. Currently α–syn oligomers have gained a lot of

attention as being a particularly toxic version of the protein. This work aims to characterize the

aggregation seeding properties of these oligomers. Cells were treated with either oligomeric

(E35K or E57K) or fibrillar versions of α–syn. Intracellular changes were assessed using

cellular fractionation, fluorescent imaging, and in vitro protein fragment complementation

assays. The results from this study highlight the ability of the E35K and E57K treatments to

induce changes in α–syn solubility to a greater extent than fibrillar forms of the protein.

Exposure to either E35K or E57K also caused the formation of α–syn inclusions that appeared

to be transmissible to naïve neighbouring cells. This study highlights interesting properties

potentially attributable to α–syn oligomers that merit further investigation.

iii

Acknowledgements

First and foremost, I would like to thank Dr. Anurag Tandon for giving me the opportunity to

work in his lab. I can say without a doubt that this experience has helped me grow both as a

person and as a scientist.

Thank you to my committee members Dr. William Trimble, Dr.Avi Chakrabartty, and Dr. Paul

Fraser for your guidance through all the aspects of this project.

I would also like to take this opportunity to thank all the past and present members of the

Tandon lab. Ms. Tammy Langman, Dr. Ana Giassi, Dr. Filsy Samuel, Dr. Sobia Iqbal, and Ms.

Maria Marano. It was a privilege to work with all of you and each one of you made this journey

so much more memorable.

I would also like to thank from the bottom of my heart, Dr. Anna Weichert and Dr. Jean Sevalle.

Both of you were not only amazing mentors, but also unbelievable friends, who provided

comfort, support, and guidance during the many challenges I faced. I will never forget the help

you provided.

Last, but definitely no least, thank you to my parents, Maleni and Renga, for always believing in

me and raising me to be the person I am today.

iv

Contributions

The author of this thesis wrote every chapter of this document, which was then edited by his

supervisor and program advisory committee members. The author conducted all the experiments

by his own hands except for the following:

Creation and purification of monomeric versions of phosphorylated and unphosphorylated wild-

type and mutant alpha-synuclein was conducted by Mr. Kyung Han, a member of the Fraser lab.

The primary sequence for the luciferase specific antibodies was selected by Dr. Paul Fraser, and

the peptides were created by Ms. Ling Wu, a member of the Fraser lab. Inoculations of animals

and collection of the antibodies was conducted by Ms. Tammy Langman.

The electron microscopy sample preparation and image acquisition was conducted by Ms. Yan

Chen a member of the Fraser Lab.

The PCMV6a construct used to express the alpha-synuclein luciferase fusion proteins was

originally designed by Dr. Jean Sevalle, and member of the Hyslop lab.

v

Table of Contents

Acknowledgements ..................................................................................................................... iii

Contributions .............................................................................................................................. iv

Table of Contents ......................................................................................................................... v

Abbreviations ............................................................................................................................ viii

List of Tables ............................................................................................................................... xi

List of Figures ............................................................................................................................ xii

Chapter 1 Introduction ............................................................................................................... 1

1.1. Neurodegenerative diseases ................................................................................................ 1

1.1.1. Proteopathies................................................................................................................ 1

1.2. History of Parkinson’s Disease ........................................................................................... 2

1.2.1. The Work of James Parkinson ..................................................................................... 2

1.2.2. The Inclusions Known as Lewy Bodies ...................................................................... 2

1.2.2.1. Lewy Body Propagation ....................................................................................... 3

1.3. Genetics .............................................................................................................................. 4

1.3.1. Synuclein, Alpha (Non A4 Component Of Amyloid Precursor) (SNCA) .................. 4

1.4. Alpha Synuclein (α-syn) ..................................................................................................... 7

1.4.1. Synuclein Proteins ....................................................................................................... 7

1.4.2. Synuclein Structure ...................................................................................................... 8

1.4.3. Posttranslational Modifications ................................................................................. 11

1.4.4. What is the Function of α-Syn? ................................................................................. 14

1.4.5. Alpha-Synuclein Aggregation ................................................................................... 16

1.4.5.1. Mutations ............................................................................................................ 16

1.4.5.2. Metals ................................................................................................................. 17

1.4.5.3. Pesticides and Herbicides ................................................................................... 18

1.4.5.4. Toxic α-Syn Species ........................................................................................... 19

1.4.6. α-Syn Oligomers ........................................................................................................ 21

1.4.6.1. E35K and E57K Lysine Mutants ........................................................................ 24

1.5. Alpha-Synuclein Migration .............................................................................................. 26

1.5.1. In Vitro Models .......................................................................................................... 27

1.5.2. In Vivo Models .......................................................................................................... 29

1.5.3. α-Syn Release ............................................................................................................ 31

vi

1.5.3.1. Exocytosis ........................................................................................................... 31

1.5.3.2. Exosomal Release ............................................................................................... 34

1.5.4. α-Syn Uptake ............................................................................................................. 37

1.5.4.1. Diffusion ............................................................................................................. 37

1.5.4.2. Endocytosis ......................................................................................................... 38

1.5.4.3. Exosomal Uptake ................................................................................................ 39

Chapter 2 Hypothesis and Aims ............................................................................................... 41

Chapter 3 Material and Methods ............................................................................................ 43

3.1. Vector Design ................................................................................................................... 43

3.2. Cell Culture ....................................................................................................................... 43

3.3. Stable Cell Line Generation ............................................................................................. 44

3.4. Gaussia Assay ................................................................................................................... 44

3.5. Coelenterazine Preparation ............................................................................................... 46

3.6. Luminescence Assay ........................................................................................................ 46

3.7. Electron Microscopy ......................................................................................................... 46

3.8. Monomer purification and phosphorylation ..................................................................... 48

3.9. Fibril and Oligomer Treatment ......................................................................................... 48

3.10. Cell Death Assay ............................................................................................................ 48

3.11. Fractionation ................................................................................................................... 49

3.12. Western Blotting ............................................................................................................. 49

3.13. Fluorescent Imaging ....................................................................................................... 50

3.14. Co-seeding ...................................................................................................................... 51

3.15. Cyto ID Staining ............................................................................................................. 51

3.16. Quantification of Punctate Structure Formation ............................................................. 53

3.17. Normalization and Statistical Analysis ........................................................................... 53

Chapter 4 Results ....................................................................................................................... 55

4.1. Construct Expression ........................................................................................................ 55

4.2. Characterization of α-Syn Species .................................................................................... 57

4.3. Dose Response with Wild-type α-Syn Fibrils .................................................................. 59

4.4. Cell Viability .................................................................................................................... 61

4.5. Aggregation Induction Following Treatment with Varying α-Syn Species ..................... 61

4.6. Luminescence Assay ........................................................................................................ 70

4.7. Fluorescent Imaging of E35K and E57K Treated Cells ................................................... 70

vii

4.8. Fluorescent Protein Fragment Complementation ............................................................. 74

4.9. Propagation of E35K and E57K Induced Morphological Changes .................................. 77

Chapter 5 Discussion ................................................................................................................. 84

5.1. Study Aims ....................................................................................................................... 84

5.2. Fibril Aggregation ............................................................................................................ 84

5.2.1. Soluble Fraction Analysis .......................................................................................... 84

5.2.2. Insoluble Fraction Analysis ....................................................................................... 85

5.3. E35K and E57K Induced Aggregation ............................................................................. 88

5.4. Luciferase PCA ................................................................................................................. 91

5.5. Intercellular Propagation .................................................................................................. 95

Chapter 6 Conclusion ................................................................................................................ 99

Chapter 7 Future Directions ................................................................................................... 101

References ................................................................................................................................. 106

viii

Abbreviations

PD Parkinson’s Disease

AD Alzheimer’s Disease

ALS Amyotrophic Lateral Sclerosis

MS Multiple Sclerosis

HD Huntington’s Disease

MSA Multiple System Atrophy

DLB Dementia with Lewy Bodies

PSP Progressive Supranuclear Palsy

SNpc Substantia nigra pars compacta

LC Locus coeruleus

LN Lewy neurites

LB Lewy bodies

α-syn Alpha-synuclein

LRRK Leucine-rich repeat kinase

UCHL1 Ubiquitin carboxyl-terminal esterase L1

NAC Non-beta-amyloid component

CD Circular dichroism

FTIR Fourier-transform infrared spectroscopy

RBCs Red blood cells

STEM Scanning transmission electron microscopy

SE-AUC Sedimentation equilibrium analytical

ultracentrifugation

E. coli Escherichia coli

PLKs Polo-like-kinases

SIAH Seven in absentia homologue

CSP α Cysteine string protein

SNARE Soluble N-ethylmaleimide-sensitive factor

attachment protein receptors

ix

SNAP-25 Synaptosomal-associated protein 25

HSP-70 Heat shock protein 70

DDT Dichlorodiphenyltrichloroethane

PCA Protein fragment complementation assay

hGluc Humanized gaussia luciferase

Luc 1 N-terminal luciferase fragment

Luc 2 C-terminal luciferase fragment

GFP Green fluorescent protein

CHIP Carboxyl terminus of Hsp70- interacting

protein

C.elegans Caenorhabditis elegans

CNS Central nervous system

PFF Preformed fibrils

Thio S Thioflavin S

3-MA 3-Methyladenine

ER Endoplasmic reticulum

HMGB1 High-mobility group protein B1

MVE Multivesicular endosome

ILV Intraluminal vesicles

GTP Guanosine-5'-triphosphate

chGal3 mCherry labelled glaectin-3

ROS Reactive oxygen species

LAMP-1 Lysosomal associated membrane protein-1

PrP Prion protein

Aβ Amyloid-β

RNA Ribonucleic acid

WT Wild-type

PCR Polymerase chain reaction

DMEM Dulbecco’s modified Eagle medium

PBS Phosphate buffered saline

EM Electron microscopy

PIC Protease inhibitor cocktail

x

EDTA Ethylenediaminetetraacetic acid

MOPS 3-(N-morpholino)propanesulfonic acid

TBST Tris buffered saline and tween 20

DAPI 4',6-diamidino-2-phenylindole

PFA Paraformaldehyde

HBSS Hanks buffered saline solution

FBS Fetal bovine serum

RLU Relative luminescence unit

SDS Sodium dodecyl sulfate

IAPP Islet amyloid polypeptide

SCN- Thiocyanate

SNCA Synuclein, Alpha (Non A4 Component Of

Amyloid Precursor)

CBD Corticobasal degeneration

CJD Creutzfeld-Jakob disease

FTLD Frontotemporal lobar degeneration

GSS Gerstmann-Straussler-Scheinker syndrome

xi

List of Tables

Table 1. Mutations Associated with Parkinson Disease ................................................................ 6

Table 2. Oligomers within proteopathies..................................................................................... 20

Table 3. Overview of in vitro and in vivo α-syn propagation studies .......................................... 32

xii

List of Figures

Figure 1. α-Syn structure and membrane binding ......................................................................... 9

Figure 2. α-Syn posttranslational modifications .......................................................................... 13

Figure 3. Gaussia luciferase protein fragment complementation assay ...................................... 22

Figure 4. Mechanisms of release and uptake of α-syn ................................................................ 33

Figure 5. Exosomal biogenesis .................................................................................................... 36

Figure 6. Double promoter vector ligation .................................................................................. 45

Figure 8. Co-seeding outline........................................................................................................ 52

Figure 9. Expression and functionality of endogenous α-syn luciferase constructs .................... 56

Figure 10. Characterization of α-syn fibrils and oligomers ......................................................... 58

Figure 11. α-Syn wild-type fibril dose response.......................................................................... 60

Figure 12. Cell viability ............................................................................................................... 62

Figure 15. Luciferase 1 and 2 antibodies ..................................................................................... 67

Figure 16. Analysis of Triton-X 100 insoluble fraction using luciferase 1 and 2 antibodies ...... 69

Figure 17. Quantification of aggregation using luminescence .................................................... 71

Figure 18. Changes in distribution of α-syn following exposure to E35K or E57K ................... 73

Figure 19. Fluorescent protein fragment complementation assay ............................................... 75

Figure 20. E35K and E57K induced redistribution of α-syn venus proteins ............................... 76

Figure 21. Propagation of E35K and E57K induced morphological changes ............................. 81

Figure 22. Percentage of recipient cells with punctate morphology ........................................... 82

Figure 23. Cell viability following co-seeding ............................................................................ 83

Figure 24. Conformational Specificity required for the Luciferase PCA .................................... 92

Figure 25. Reorganization of the fluorescent PCA fragments ..................................................... 96

Figure 26. Exosome isolation protocol ………………………………………………………..103

1

Chapter 1

Introduction

1.1. Neurodegenerative diseases

Neurodegenerative diseases target cells within the central nervous system causing loss of

structure and function. As the initial deterioration process commences, individuals begin to

notice mild symptoms such as memory impairments and problems with coordination. Gradually

these symptoms begin to increase in severity until the patient loses all ability to function

independently and ultimately these diseases result in fatal outcomes.

The most common neurodegenerative diseases are Alzheimer’s Disease (AD), Parkinson’s

Disease (PD), Amyotrophic Lateral Sclerosis (ALS), Multiple Sclerosis (MS), and Huntington’s

Disease (HD). In the United States roughly 5.4 million people suffer from AD, and

approximately 50,000 new cases of PD are diagnosed each year (de Lau & Breteler, 2006; Thies

et al., 2013). Since the risk of these disorders increases with age, as the baby boomers enter the

latter stages of their life, the incidence of neurodegenerative disorders is expected to soar (Thies

et al., 2013). In addition to degrading the quality of life, these disorders will also cause a

tremendous burden on our already stressed healthcare system. Therefore, finding treatments and

cures is of the utmost urgency.

1.1.1. Proteopathies

Since disorders such as PD, AD, ALS, and HD share similarities in their pathological cascade,

they have been classified under the umbrella term proteopathies (Walker & LeVine, 2000). This

refers to a class of diseases in which abnormalities in protein structure and function results in

negative cellular and physiological outcomes (Walker & LeVine, 2000). Frequently, these

proteins fail to adopt their appropriate structural conformation, and this misfolded form often

triggers aggregation of other proteins, organelle degradation or cell death pathways. This thesis

2

will focus on the proteopathy known as Parkinson’s disease, a neurological disorder that results

in cellular, motor, and cognitive impairments.

1.2. History of Parkinson’s Disease

1.2.1. The Work of James Parkinson

Dr. James Parkinson first described the symptoms associated with this disease in the monograph

titled An Essay on the Shaking Palsy in 1817. His observations were based on six patient cases

that presented abnormal movement patterns (Parkinson, 2002). His clinical notes indicated that

patients’ initially noticed weakness in their limbs followed by a mild trembling of the hands and

arms (Parkinson, 2002). During the latter stages patients exhibited disturbances in sleep, bodily

functions, speech, and motor impairments such as postural instability, shuffling gait, and

tremors (Parkinson, 2002).

Today, it is well established that in addition to motor deficits (rigidity, bradykinesia, postural

instability, shuffling gait, and tremors), PD patients also suffer from a range of non-motor

symptoms such as cognitive impairments, autonomic dysfunctions, sleep disorders, and sensory

deficits (Chaudhuri et al., 2006). Unfortunately, certain motor and non-motor symptoms are also

common to other disorders such as Multiple System Atrophy (MSA), Dementia with Lewy

Bodies (DLB), and Progressive Supranuclear Palsy (PSP) (Christine & Aminoff, 2004).

Therefore, the presence of both clinical signs and pathological findings are required in order to

unequivocally confirm a diagnosis of PD.

1.2.2. The Inclusions Known as Lewy Bodies

Despite identifying the clinical symptoms associated with PD, Parkinson’s work lacked

pathological data. It wasn’t until 1912, when Friedrich (Fritz) Heinrich Lewy provided the first

accounts of the neuroanatomical changes associated with PD. After examining 25 individuals,

Lewy described characteristic inclusions, which appeared within the dorsal motor nucleus of the

vagus, basal nucleus of meynert, the globus pallidus, lateral nucleus of the thalamus, and the

3

periventricular nucleus of the thalamus. Later Konstantin Tretiakoff also reported observing

similar inclusions within the substantia nigra pars compacta (SNpc) of PD patients, and noted a

correlation between the presence of these aggregates and nerve cells loss within the SNpc.

Today it is well established that loss of dopaminergic and noradrenergic neurons within the

ventrolateral SNpc and locus coeruleus (LC) respectively, in addition to the formation of

inclusions are cardinal signs associated with PD (Ehringer et al., 1960; Gaspar et al., 1991;

Hughes et al., 1992). These inclusions can develop as thread-like structures, called Lewy

neurites (LN), within the cellular processes or as globular aggregates, termed Lewy bodies (LB),

within the neuronal perikarya (Spillantini et al., 1998). Morphologically Lewy bodies may be

divided into two different categories. The brainstem type appears as single or multiple

intracytoplasmic inclusions that are spherical eosinophilic masses with a dense core and

peripheral halo (reviewed in Wakabayashi et al., 2007). They are typically found within the

brainstem, diencephalon, and peripheral autonomic regions. The cortical type on the other hand

has a less defined structure, which lacks a conspicuous halo or central core, and they are

restricted to the cerebral cortex and amygdala (reviewed in Wakabayashi et al., 2007). These

Lewy bodies are customarily 200-600nm in length and are composed primarily of an aggregated

protein called alpha-synuclein (α-syn) (Spillantini et al., 1998). In addition to α-syn, these

inclusions are also composed of neurofilaments, cytosolic proteins, α-syn binding proteins, cell

signalling molecules, phosphorylases, lipids, immunoglobulins, cytoskeletal proteins,

components of the ubiquitin-proteasomal system, and metallic cations (reviewed in

Wakabayashi et al., 2007). Some of these constituting elements have even been closely linked

with PD induction. For example, metals such as iron and aluminum have been considered

instigators of idiopathic PD (refer to section 1.4.4.2), while oligomeric or fibrillar forms of α-

syn are believed to induce cell death and propagate this pathology (refer to section 1.4.5).

However, despite the prominence this pathology has gained over the years due to its association

with PD and other disorders, the exact role these inclusions play in disease pathogenesis is still

ambiguous.

1.2.2.1. Lewy Body Propagation

It is believed that Lewy Body formation commences before the onset of clinical motor

symptoms. This pathology has been observed to originate in the glossopharyngeal, vagal, and

4

olfactory centers prior to invading the midbrain and neocortical areas (Braak et al., 2003). The

pathology within the anterior olfactory regions exhibited less invasive characteristics than the

aggregates originating in the brainstem. Therefore, Braak and colleagues proposed a six-stage

model for the ascending course of this pathology. Stages 1 and 2 characterize lesion formation

within the boundaries of the medulla and pontine tegmentum in areas such as the olfactory,

glossopharyngeal, vagal, caudal raphe, and gigantocellular reticular nuclei. Damage to these

regions is hypothesized to account for the gustatory, olfactory, and digestion abnormalities that

antedate the motor deficits in certain PD patients (Braak et al., 2003). Stages 3 and 4 involve

the rostral progression from the lower brainstem into the midbrain, basal prosencephalon, and

mesocortex. These stages typically involve pathology within substantia nigra and temporal

mesocortex resulting in motor and postural irregularities. Finally, stages 5 and 6 involve the

invasion of higher order sensory association areas such as the prefrontal and neocortex.

Currently, detection of disease onset during the early stages still poses a great challenge due to

lack of overt consistent clinical presentations. However, many in the field are still hopeful that

better understanding and identification of α-syn aggregation might provide an avenue for early

diagnosis and treatment.

1.3. Genetics

1.3.1. Synuclein, Alpha (Non A4 Component Of Amyloid Precursor) (SNCA)

In addition to being the main component of Lewy bodies, α-syn is also central to the

pathophysiology of familial PD. Point mutations in SNCA, the gene that encodes for α-syn,

causes early-onset PD with dementia. Over the years three distinct mutations have been noted

within the N-terminal region, the Alanine (A) to Threonine (T) (A53T), Alanine to Proline (P)

(A30P), and Glutamic acid (E) to Lysine (K) (E46K) substitutions (Kruger et al., 1998;

Polymeropoulos et al., 1997; Zarranz et al., 2004). These autosomal dominant mutations

promote beta sheet formation within the protein’s secondary structure, which increases its

propensity for aggregation (Kruger et al., 1998; Polymeropoulos et al., 1997; Zarranz et al.,

2004). Even though it is still unclear exactly how these mutations trigger the onset of PD, it is

5

Locus Gene Protein Protein Function Mutations Reference

PARK 1&4 SNCA α-syn Synaptic signalling?

Oxidative stress?

(AD);A53T; A30P; E46K;H50Q;G51D

Gene duplication or triplication

(Kruger et al., 1998;

Lesage et al., 2013;

Polymeropoulos et al.,

1997; Proukakis et al.,

2013; Zarranz et al.,

2004)

PARK 2 Parkin Parkin Ubiquitin E3 Ligase (AR) > 100 point mutations (Kitada et al., 1998)

PARK 3

SPR

Sepiapterin

reductase

Catalyzes the

reduction of

carbonyls

(AD) (?)

(Sharma et al., 2011)

PARK 5

UCHL1

UCHL-1

Ubiquitin

hydrolase or ligase

(AD) S18Y

(Ragland et al.,2009)

PARK 6

PINK

PINK

Mitochondrial kinase

(AR) > 40 point mutations and

deletions

(Valente et al., 2004)

PARK 7

DJ-1

DJ-1

Oxidative stress

protection

(AR) > 10 point mutations

(Bonifati et al., 2003)

PARK 8

LRRK2

LRRK2

Protein kinase

(AD) >40 missense variants

(Zimprich et al., 2004)

PARK 9

ATP13A2

ATPase

type13A2

Lysosomal ATPase

(AR) >5 point mutations

(Ramirez et al., 2006)

PARK 11

GIGYF2

GRB10

interacting

GYF

protein 2

(?)

(AD) 7 missense mutations

(Lautier et al., 2008)

6

PARK 14

PLA2G6

Phospholipase

A2

Phospholipid

remodelling

(AR) 2 missense mutations

(Paisan-Ruiz et al., 2009)

PARK 15

FBXO7

F-box

protein 7

Phosphorylation

dependent

ubiquitination

(AR) 3 point mutations

(Di Fonzo et al., 2009)

PARK 17

VPS35

VPS35

Transport of proteins

from endosome to

trans-golgi

AD) (?)

(Zimprich et al., 2011)

PARK 19

DNAJC6

Auxilin

Tyrosine proteins

phosphatase

(AR) (?)

(Edvardson et al., 2012;

Koroglu et al., 2013)

PARK 20

PARK21

SYNJ1

DNAJC13

Synaptojanin1

HSP40

Lipid phosphatase

Receptor mediated

endocytosis

(AR) missense mutation

(AD) missense mutation

(Krebs et al., 2013; Quadri et

al., 2013)

(Vilarino-Guell et al., 2014)

Table 1. Mutations Associated with Parkinson Disease

Table outline autosomal dominant (AD) and autosomal recessive (AR) factors associated with PD. (?) denotes areas that have yet to be fully

characterized. Table was adapted from information reviewed by (Dugas et al., 2010) and (Lesage & Brice, 2009).

7

believed that perturbations in protein structure play an influential role. Other modification such

as genomic duplication or triplication of chromosome 4q21, which encompasses the SNCA locus

also results in either late or early-onset PD (Chartier-Harlin et al., 2004; Singleton et al., 2003).

Individuals with alterations in gene dosage exhibited a 1.8-fold increase in soluble α-syn, and

their brain lysates contained high molecular mass species of this protein (Miller et al., 2004).

However, despite the aggressive phenotype exhibited by these patients, cases involving either

SNCA overexpression or missense mutations are exceedingly rare and only make up a small

portion of PD incidences. The rest of the cases are either sporadic or result from mutations in

other genes. To-date, at least 13 different genes in addition to SNCA have been associated with

autosomal dominant and recessive forms of PD. Table 1 summarizes other loci and genes

associated with this disorder.

1.4. Alpha Synuclein (α-syn)

Alpha-synuclein has long been thought to be a key player in PD. Not only is it found in Lewy

bodies, but mutations and changes in α-syn expression result in aggressive PD phenotypes.

Therefore, understanding the function and role of this protein within the brain is crucial to

elucidating its involvement in the pathogenesis of this disease. Thus, the next few sections will

examine in detail the structure and endogenous role of this protein.

1.4.1. Synuclein Proteins

In 1988 the first synuclein sequence was isolated from the electric organ of the Pacific electric

ray (Torpedo Californica) (Maroteaux et al., 1988) ( Figure 1). Currently three homologs of this

protein have been discovered in humans: alpha-, beta-, and gamma-synuclein (reviewed in

Levadan et al., 1998). These proteins are 127 to 140 amino acids in size and have similar

domain organizations. All three proteins consist of an N-terminal region that contains 7

imperfect repeats and an acidic C-terminal tail (reviewed in Levadan et al., 1998). α- and β-

synuclein contain a stretch of hydrophobic amino acid residues, called the non-beta-amyloid

component (NAC), in a central domain which separates the amphipathic N-terminal from the

acidic C-terminal tail (reviewed in Levadan et al., 1998). To date, only α- and β-synuclein have

8

been implicated in neurodegenerative diseases, but in terms of PD, there has been no evidence

thus far directly linking β-synuclein to this disease (Mori et al., 2002). Only α-syn has been

associated with the formation of Lewy bodies and the initiation of PD through mutations in the

SNCA gene. Therefore, the remainder of this thesis will focus on the structure and function of

this protein within the brain, and its potential role in PD pathogenesis.

1.4.2. Synuclein Structure

α-Syn possesses an unusually large stokes’ radius (34 Å), and sediments at a much slower rate

than similar sized globular proteins (Weinreb et al., 1996). This key discovery was the first

piece of evidence that suggested α-syn might possess an elongated structure. Following these

results, circular dichroism (CD) and fourier-transform infrared spectroscopy (FTIR) studies

provided additional information indicating that α-syn lacked a secondary structure (Uversky et

al., 2001a; Uversky et al., 2002). The overall conformation was only slightly affected when

boiled or subjected to other chemical denaturants (reviewed in Uversky et al., 2009; Weinreb et

al., 1996). These findings suggested that α-syn might exist in a natively unfolded state at a

neutral pH (Uversky et al., 2001a), a logical assumption given the protein’s low overall

hydrophobicity and high net charge, which would preclude partial folding (reviewed in

Munishkina et al., 2003; reviewed in Uversky et al., 2009; Uversky et al., 2001a). However,

perturbations such as high temperature or low pH cause α-syn to adopt a partially folded

structure possessing β-sheet conformations (reviewed in Munishkina et al., 2003). These

changes are believed to influence α-syn’s structure by either modulating the charge of the

protein (i.e. neutralizing negative repulsion) or increasing its hydrophobicity, which increases its

propensity to aggregate.

In addition to these extrinsic factors, interaction with lipids has also been shown to induce

conformational changes. The amino acid repeats within α-syn’s N-terminal region allows it to

reversibly bind to membranes (Davidson et al., 1998; Fusco et al., 2014; Zarbiv et al., 2014).

This repeat region exhibits similarities to the lipid binding domains within apolipoproteins,.

Anything that disrupts this sequence, such as the A30P mutation, drastically reduce α-syn’s

membrane affinity (Bussell & Eliezer, 2003; Zarbiv et al., 2014). Human α-syn exhibits a 57%

9

Figure 1. α-Syn structure and membrane binding

a) Schematic illustrates the different regions within α-syn. The PD associated mutations are

found within the N-terminal region. Phosphorylation of serine (S) 129 is one of the most

common modifications found on aggregated α-syn within Lewy bodies. b) The conformational

change undergone by α-syn upon membrane binding. The N-terminal region assumes a helical

structure predominantly facilitated by the KTKEGV repeats, while the C-terminal region retains

its unstructured characteristics.

10

increase in α-helicity upon binding to membranes or small vesicles (Davidson et al., 1998). The

lipid content and vesicle surface area strongly dictates the strength of this interaction. Increases

in the ratio of lipid to protein or surface area to volume further promotes α-syn binding, which is

concurrently accompanied by an increase in α-helicity (Zhu & Fink, 2003). Stabilization of the

alpha helix due to association with lipid rich vesicles has also been shown to negatively

correlate with α-syn fibrillization. Thioflavin T assays indicated that fibrillization lag-time was

dramatically increased when α-syn was bound to vesicles (Zhu & Fink, 2003). Furthermore, it

also appears that α-syn exhibits specificity for certain membrane lipids relative to others. GM1

gangliosides often found within lipid rafts and caveolae (specialized neuronal membrane

domains that contain the caveolin family of proteins and sphingolipids) are abundantly

expressed within the neurons and makeup 10% of the total lipids within synaptic regions

(Martinez et al., 2007). GM1 containing small unilaminar vesicles (SUV) induce greater α-

helical transformations relative to other brain sphingolipids and either inhibit or eliminate α-syn

fibrillization depending on the GM1 content within these SUVs (Martinez et al., 2007). The

functional role of α-syn’s membrane association is still up for debate, but some have suggested

it might have a neuroprotective effect because membrane bound α-syn is less likely to undergo

aggregation. Taken altogether α-syn’s membrane binding ability provides further support for its

conformational pliability and sensitivity to extraneous influences.

Interestingly, recent studies have called into question much of the previously published work

regarding the structural and membrane binding characteristics of α-syn. It has been shown that

cytoplasmic α-syn exists as a tetramer 55–60 kilodaltons (kD) in size (Bartels et al., 2011;

Dettmer et al., 2013; Gould et al., 2014; Luth et al., 2015). A variety of experimental approaches

such as non-denaturing conditions, cross-linking experiments, scanning transmission electron

microscopy (STEM), and sedimentation equilibrium analytical ultracentrifugation (SE-AUC)

consistently yielded tetrameric forms of α-syn. This conformational state was observed in

different cell lines as well as post-mortem tissue (Bartels et al., 2011; Dettmer et al., 2013;

Gould et al., 2014; Luth et al., 2015). Furthermore, tetrameric versions of α-syn exhibited

unique properties such as the ability to undergo helical transformations in the absence of

vesicles or lipid membranes. In fact, the presence of these lipid structures had no effect on the

conformational state of these tetramers. These findings were also recapitulated using an

Escherichia coli (E. coli) model (W. Wang et al., 2011). The authors reported isolating

oligomeric forms of α-syn under non-denaturing conditions, which existed in a helical state in

11

the absence of vesicles (W. Wang et al., 2011). These studies argue that α-syn naturally exists in

a tetrameric state and that this conformation even precludes it from aggregating (Bartels et al.,

2011; Dettmer et al., 2013; W. Wang et al., 2011). In a recent paper the Selkoe lab has even

demonstrated that these tetrameric isoforms can be purified from brain homogenate (Luth et al.,

2015). However, it appears that these tetramers dissociates when extensive purification steps

are incorporated into the isolation process, which could be due to the loss of lipids that might

have stabilized the complex.

In response to the aforementioned claims, studies were conducted in order recapitulate these

findings, but unfortunately were unable to isolate this elusive tetramer (Burre et al., 2013;

Fauvet et al., 2012). The study by Fauvet et al. 2012 even created monomeric and dimeric α-syn

standards in order to better characterize the size of the α-syn obtained from mammalian cell

lines, E.coli, RBCs, murine, and human tissue (Fauvet et al., 2012). It was discovered that α-

syn isolated from these different sources co-eluted and co-migrated with the monomeric

standard, and this pattern was consistent under denaturing and nondenaturing conditions (Burre

et al., 2013; Fauvet et al., 2012). Even when isolating α-syn from RBCs, similarly to the initial

study by Bartel et al. 2011, the full-length monomeric version of the protein was the dominant

species (Fauvet et al., 2012). Due to these conflicting results the native structure of α-syn still

remains uncertain and this topic continues to be an area of intense debate.

1.4.3. Posttranslational Modifications

α-Syn like many other proteins undergoes posttranslational modifications, which can affect its

structure and aggregation kinetics (see Figure 2.). Phosphorylation of α-syn, specifically on the

serine and tyrosine residues within the C-terminal region, is a common posttranslational

modification found on aggregated α-syn within Lewy bodies. Of the four sites within the acidic

C-terminal tail, the Serine 129 residue has gained the most attention (Barrett et al., 2015;

reviewed in Oueslati et al., 2010). First observed post-mortem in patients with PD, MSA, and

DLB, Ser 129 phosphorylation has now been consistently recapitulated in cell culture and

animals models making it one of the hallmark pathological features of synucleinopathies

(Barrett et al., 2015; reviewed in Oueslati et al., 2010). Unfortunately, the potential enzymes

responsible for this modification in vivo still remain elusive, but a number of potential

12

candidates such as Casein Kinase I & II, polo-like-kinases (PLKs), G-protein-coupled receptor

kinases (1,2,5 and 6), and LRRK are currently under investigation (reviewed in Oueslati et al.,

2010). In vitro, this modification appears to inhibit the fibrillization process while in vivo the

results are a bit more conflicted. In rodent models, phosphorylation doesn’t significantly affect

aggregation, but cells overexpressing proteins with this modification exhibited greater viability

in comparison to those overexpressing wild-type α-syn (Azeredo da Silveira et al., 2009;

Gorbatyuk et al., 2008; McFarland et al., 2009). However, experiments conducted in

Drosophila models depicted very contradictory results. In these organisms, phosphorylated α-

syn had a greater predisposition for forming oligomers, which compromised cell viability, while

inhibiting phosphorylation not only negated the toxic effects, but also promoted fibrillization

(Chen & Feany, 2005; Chen et al., 2009). This discrepancy could be due to differences in the

intracellular mechanisms, but nonetheless further research is needed in order to get a better

consensus regarding the role of phosphorylation in vivo.

In addition to being phosphorylated, much of the aggregated α-syn within Lewy bodies also

undergoes ubiquitination. This modification customarily occurs on the lysine residues within the

N-terminal region (K10, K21, K23, K32, K34, K43, and K96) where K21 and K23 are the sites

most frequently modified (reviewed in Oueslati et al., 2010). Currently, the E3 ubiqutin ligases

Parkin, ubiquitin carboyx-terminal hydrolase L1 (UCH-L1) and seven in absentia homologue

(SIAH) are considered to be the most likely enzymes involved in this modification (reviewed in

Oueslati et al., 2010). Some of these proteins (Parkin and UCH-L1) have not only been

implicated with familial forms of PD (see table 1.1), but have also been found within Lewy

bodies (Parkin and SIAH) (reviewed in Wakabayashi et al., 2007). In vivo, this modification

seems to play a neuroprotective role. Increased ubiquitination in Drosophila and rats attenuates

α-syn induced neurotoxicity, while mutations that inhibit this process completely negate any

protective effects (Haywood et al., 2004; Yang et al., 2003). Thus, even though ubiquitinated α-

syn is predominantly found within Lewy Bodies, this modification might actually serve a

neuroprotective role in PD.

Another common posttranslational modification that α-syn undergoes is truncation. However,

unlike phosphorylation and ubiquitination, which were mainly observed on aggregated α-syn

found within Lewy bodies, truncation can occur under normal conditions (reviewed in Oueslati

et al., 2010). Both diseased and healthy brains appeared to possess equal amounts of truncated.

13

Figure 2. α-Syn posttranslational modifications

a) Schematic depicts the potential α-syn serine (S), threonine (T), and tyrosine (Y)

phosphorylation sites observed in individuals with PD. b) The major lysine (K) ubiquitination

sties identified in Lewy bodies (upper arrows) and in vitro models (lower arrows). c) Known α-

syn truncation sties, which were identified from Lewy body extracts. The arrows in indicate the

last residue in the truncation, but when residue is not known a range is given. Figures were

adapted from data (reviewed in Oueslati et al., 2010)

14

α-syn (W. Li et al., 2005). In vitro studies examining these smaller isoforms revealed that they

were more prone to form fibrils (Hoyer et al., 2004; Li et al., 2005; Murray et al., 2003).

Cleavage of residues between amino acids 110-120 exhibited an enhanced rate of fibrillogenesis

and even seeded the aggregation of full-length α-syn species. Studies have argued that the C-

terminal, where a majority of the truncation occurs, participates in long-range interactions with

the N-terminal region, which inhibits aggregation. Cleavage of residues within this region

appears to diminish this shielding effect (reviewed in Oueslati et al., 2010). In vivo, α-syn

truncation has also been reported to cause aggregation and neuronal loss. In Drosophila α-syn

(1-120) resulted in increased oligomer and proteinase K resistant inclusions, which were

accompanied by a faster loss of dopaminergic neurons (Periquet et al., 2007). Similarly in

rodents, α-syn (1-120) caused inclusion formation within the SN and olfactory bulb in addition

to decreasing striatal dopamine (Tofaris et al., 2006). Truncated versions of α-syn have also

been found in electrodense-laminated structures within dystrophic neurites in the Thy-1

transgenic mouse model, and antibodies that target these smaller isoforms even helped

ameliorate PD-like pathology (Games et al., 2013; Games et al., 2014). Even though increasing

evidence suggests that the C-terminal region is an important regulator of aggregation in vitro, in

vivo this relationship is still very tenuous requiring further research.

1.4.4. What is the Function of α-Syn?

Genetic studies have demonstrated that α-syn is crucial to the pathogenesis of familial PD. Not

only is the onset of the disorder attributed to mutations in this protein, but α-syn is also the main

component of Lewy bodies, one of the main symptoms of this disease. Despite the importance

of this protein, its normal function is still poorly understood. Evidence suggests that α-syn might

regulate neurotransmission, a likely possibility given its predominant localization within

presynaptic terminals (reviewed in Bendor et al., 2013). However, the exact nature of this

relationship is still intensely debated. It has been argued that α-syn promotes neurotransmitter

release (Cabin et al., 2002; Murphy et al., 2000). Studies using α-syn knockout mice and

antisense oligonucleotides demonstrated that loss or reduction in α-syn expression caused a

striking deficiency in distal pool synaptic vesicles in addition to impairments in synaptic

response to a repetitive stimulation (Cabin et al., 2002; Murphy et al., 2000). While another

group demonstrated a potential inhibitory role by showing greater dopamine release in α-syn

15

knockout mice relative to wild type after paired pulse stimulation (Abeliovich et al., 2000).

Subsequent studies using chromaffin cells from mice overexpressing either wild type or mutant

A30P α-syn provided similar results. The authors noticed that the frequency and number of

exocytic events per stimulus was lower for both WT and A30P α-syn overexpressing cells while

transmitter accumulation and vesicle fusion kinetics was unaltered (Nemani et al., 2010). Based

on these results they proposed that α-syn could potentially inhibit the vesicle-priming step that

occurs prior to secretion (Nemani et al., 2010). Due to these conflicting findings there still isn’t

a consensus regarding how α-syn might regulate synaptic release.

α-Syn was also reported to possess a very strong association with cysteine string protein (CSP

α) (Chandra et al., 2005). CSP α knockout mice experience progressive synaptic degeneration

and eventually die within two months (Fernandez-Chacon et al., 2004). However,

overexpression of α-syn within these knockout mice appears to attenuate the degeneration

process while loss of α-syn exacerbates the knockout phenotype (Chandra et al., 2005; Sharma

et al., 2012). CSP α helps maintain the soluble N-ethylmaleimide sensitive factor attachment

protein receptor (SNARE) complex, which is involved in synaptic vesicle fusion. Deletion of this

protein induced selective reductions in synaptosomal-associated protein 25 (SNAP-25), a

protein within the SNARE complex, along with its chaperones HSC70 and heat shock protein

70 (HSP 70), which subsequently impaired SNARE complex assembly. Neurodegeneration in

CSP α knockout mice seems to be primarily due to defective SNAP-25 function (Sharma et al.,

2012). Overexpression of α-syn appeared to correct decreases in HSC70 and HSP70 in addition

to modestly improving SNARE complex assembly. These results indicate that α-syn might act

through an alternative mechanism in order to maintain the integrity of the presynaptic terminal.

α-Syn has been shown to directly promote SNARE assembly by binding to SNARE specific

protein synaptobrevin-2 via its C-terminus, and helped maintain SNARE mediated fusion (Burre

et al., 2014; Burre et al., 2010). Experiments in triple knockout mice lacking all three synuclein

proteins showed age dependent neurological impairments, deficits in SNARE complex

assembly, and premature death (Fernandez-Chacon et al., 2004). Thus it appears that

maintenance of neuronal synapses is another function facilitated by α-syn in addition to its

regulatory role in vesicle release.

16

1.4.5. Alpha-Synuclein Aggregation

In a majority of PD cases there is a redistribution of α-syn from its customary monomeric form

to large detergent insoluble aggregates found in Lewy bodies. Consequently, it is important to

understand the factors that affect α-syn’s predisposition to aggregate, if we hope to unlock the

mysteries surrounding this pathology. Furthermore one of the major topics of this thesis is also

α-syn aggregation, and therefore the following sections will review some of the factors that can

influence α-syn’s aggregation kinetics.

1.4.5.1. Mutations

α-Syn possesses an inherent ability to aggregate when incubated at 37°C at a pH of 7.4, but this

process is very time consuming (reviewed in Bendor et al., 2013). Therefore, factors such as

agitation or increases in α-syn concentration are required to shift the structural equilibrium from

a monomeric state to a partially folded intermediate in order to drastically increasing the rate of

fibrillization (reviewed in Fink, 2006). In vitro, α-syn aggregation exhibits a sigmoidal growth

pattern, which entails an initial lag phase, followed by exponential growth that eventually leads

to a plateau (reviewed in Fink, 2006). The transition from the lag phase, which represents α-syn

in a monomeric form, to the exponential growth phase is the most time consuming step. The

classic α-syn mutations associated with familial PD have been known to shift this equilibrium

away from the monomeric state in favour of more aggregated isoforms (reviewed in Fink, 2006;

reviewed in Uversky et al., 2003; Uversky et al., 2009). Interestingly, these mutations do not

alter the monomeric structure of α-syn nor the rate of transition from the natively unfolded

monomer into a partially folded intermediate, which is the first step in the aggregation cascade

(Li et al., 2001; Narhi et al., 1999). These mutations mainly hinder α-syn’s propensity to form

alpha helices while concurrently increasing its ability to form β-sheet structures, a conformation

that is enriched in many aggregates (Li et al., 2001). Based on these findings, it has been

hypothesized that the increased susceptibility to form β-sheet could result in faster aggregation.

By stabilizing the β-sheet structural conformation, these mutations were thought to promote the

transition from the partially folded intermediate to larger aggregates. In vitro experiments with

the A53T mutation revealed a much shorter lag phase before fibrillization relative to wild-type

α-syn (Li et al., 2001). The CD spectra collected at different times points to monitor the

17

conversion from the random coil conformation to β-sheet structures indicated a faster and more

extensive transition for these mutant isoforms (Li et al., 2001; Narhi et al., 1999). When these

mutations were expressed in animal models using the mouse prion promoter, motor

dysfunctions and neuropathology were observed much earlier in mice expressing human A53T

α-syn (Gomez-Isla et al., 2003; M. K. Lee et al., 2002; Luk, Kehm, et al., 2012b). These animals

exhibited abnormal inclusions containing α-syn and ubiquitin within the midbrain, brainstem,

and cerebellum. Brain homogenates from these animals also contained detergent insoluble high

molecular weight α-syn, normally indicative of aggregated species (Gomez-Isla et al., 2003; M.

K. Lee et al., 2002; Luk, Kehm, et al., 2012b). These transgenic animals also exhibited motor

phenotypes such as resting tremors, rigidity, and dystonic posturing much earlier than cohorts

expressing wild-type α-syn (Gomez-Isla et al., 2003; Luk, Kehm, et al., 2012b). However,

despite the influential role of the A30P and A53T mutations in PD pathogenesis, these amino

acid changes are extremely rare and absent from idiopathic cases. Therefore, it is important to

consider alternative factors that could potentially induce PD in these individuals. In recent years,

strong arguments have been made for the importance of environmental influences in the

pathogenesis of PD. For example, numerous studies have implicated factors such as heavy metal

ions, pesticides, and herbicides as potential instigators of PD (Gomez-Isla et al., 2003).

1.4.5.2. Metals

Metals play an essential role in facilitating a broad spectrum of physiological functions within

brains such as neurotransmitter synthesis, storage, and release. However, fluctuations in the

concentration of these cations have also been suspected to influence the pathophysiology of

different neurodegenerative diseases. Epidemiological studies conducted on individuals from

highly industrialized areas have long since indicated an increased risk of PD from chronic

exposure to metals (reviewed in Chin-Chan et al., 2015; reviewed in Uversky, 2003). For

example, a retrospective cohort study of individuals from metropolitan Detroit revealed a

significant association between individuals with idiopathic PD and long-term exposure to metals

such as manganese and copper (Rybicki et al., Gorell et al., 1997; 1993; Zayed et al., 1990).

These findings correlate well with post-mortem studies that have reported high levels of

aluminum and iron within the brains of PD patients, and these metals were predominantly

localized within Lewy bodies (Hirsch et al., 1991). Iron in particular has been known to induce

18

free radical formation by accelerating the autoxidation of molecules such as catecholamines

normally located within the SN (Hirsch et al., 1991). These radicals can go on to damage

biological molecules, cause mitochondrial dysfunction, excitotoxicity, and raise calcium levels

triggering cell death. Ions such as Cu2+ and Al3+ can also interact with α-syn and facilitate

aggregation and induce structural perturbations (Oestreicher et al., 1994). Several mono, di, and

tri valent cations (Li+, K+, Na+, Cs+, Ca2+, Co2+, Cd2+, Cu2+, Fe2+, Mg2+, Mn2+, Zn2+, Co3+, Al3+

and Fe3+) can accelerate α-syn fibrillization and among these metals Al3+,Cu2+, Fe2+, Co3+ , and

Mn2+ seem to have the greatest effect (reviewed in Uversky et al., 2003). It is hypothesized that

these cations can bind to the acidic C-terminal tail of α-syn and mask the negative charge and

thereby inhibit the columbic repulsion, which can increase α-syn’s propensity for aggregation

(reviewed in Uversky et al., 2003). Consequently, metal ions might play a very important role in

the pathogenesis of PD due to their toxic influences resulting in either oxidative stress or α-syn

fibrillization.

1.4.5.3. Pesticides and Herbicides

Pesticides and herbicides such as paraquat, rotenone, dichlorodiphenyltrichloroethane (DDT),

dieldrin, and diethyldithiocarbamate are another common environmental agent thought to induce

the onset of idiopathic PD (Uversky et al., 2001b). In vitro studies demonstrated that pesticides

such as rotenone were able to halve the lag-time in the fibrillization process (Tanner, 1989).

When administered in rats, rotenone blocked mitochondrial complex I uniformly throughout the

brain, which resulted in progressive nigrostriatal dopaminergic degeneration (Uversky et al.,

2002). These animals developed cytoplasmic inclusions within nigral neurons that contained α-

syn and ubiquitin (Uversky, Li, Bower, et al., 2002). The neuropathology was also accompanied

by motor and postural deficits, the severity of which depended on the extent of the dopaminergic

lesions. Herbicides such as paraquat also induced similar in vitro and in vivo effects. However,

unlike rotenone, paraquat also appeared to influence intracellular α-syn expression in addition to

inducing aggregation. Two days post administration of the herbicide, α-syn levels rose

dramatically (Uversky, Li, Bower, et al., 2002). This overexpression eventually led to α-syn

deposits that stained positively for β-sheet structure within the SNpc similarly to the Lewy

bodies found in PD patients (Manning-Bog et al., 2002). These toxin models provide insight

into additional factors that might also play a role in PD pathology onset.

19

1.4.5.4. Toxic α-Syn Species

Abnormal protein accumulation is a common phenomenon across many different

neurodegenerative disorders, and in some cases these inclusions are believed to be the toxic

culprit that cause cell death. For PD, Lewy bodies are considered to be one of the hallmark

symptoms of the disorder, but the exact role these inclusions play within the neurodegenerative

process is unclear. Some have suggested that Lewy bodies are non-toxic and are associated with

a neuroprotective function (reviewed in Kalia et al., 2013). This is a likely possibility given that

Lewy bodies have been observed in the brains of aged individuals without clinical features of

PD. In a study that conducted over 1200 autopsies, approximately 12% of the cases possessed

incidental Lewy bodies (Forno, 1969; Saito et al., 2004). These individuals were asymptomatic

for the classical PD motor deficits, and their brains did not exhibit any degeneration.

Furthermore, patients with PARK 2 gene mutations possess all the clinical symptoms of PD, but

not all of them possess Lewy body pathology. Therefore, these reports provide evidence that

Lewy pathology alone cannot explain the pathogenesis of the disease. Other factors such as

environmental toxins, mutations, and age related cellular dysfunctions are all elements that

makeup this complex neurodegenerative process. In fact, studies are now looking at smaller α-

syn species, called oligomers, as another potential toxic culprit. It has been noted oligomeric

isoforms play an important role in PD and other proteopathies (see Table 2.). For example,

higher levels of α-syn oligomers have been observed within the cerebral cortex, cerebrospinal

fluid, and blood of patients who suffer from PD or other synucleinopathies (Aasly et al., 2014;

Paleologou et al., 2009; Wang et al., 2015). In transgenic mice with the A53T mutation, the

level of α-syn oligomers is also much greater than cohorts overexpressing wild-type α-syn

(Bezard & Przedborski, 2011)The consistent appearance of these oligomers is an indication that

these smaller α-syn isoforms might play integral role within the disease process, and this has led

to increased research regarding their properties and toxic capabilities.

20

Proteopathies Toxic Protein Reference

Synucleinopathies

(PD, DLB, MSA)

Alpha-synuclein

(Karpinar et al., 2009; Liu et al., 2009;

Paleologou et al., 2009; Sharon et al.,

2003; Winner et al., 2011)

Amyloidopathy

(AD, Downs Syndrome)

Beta-amyloid (McLean et al., 1999; Pike et al., 1991;

Walsh et al., 2002)

Tauopathies

(AD, FTLD, CBD, PSP)

Tau (Berger et al., 2007; Lasagna-Reeves et

al., 2011; Lasagna-Reeves et al., 2012;

Maeda et al., 2007; Maeda et al., 2006)

Prionopathies

(CJD,GSS, FFI, Kuru)

Prion protein (Biasini, Turnbaugh, Unterberger, &

Harris, 2012; Minaki, Sasaki, Honda, &

Iwaki, 2009; Silveira et al., 2005;

Simoneau et al., 2007)

Trinucleotide repeat

disorders

(HD)

Poly Q protein (Lajoie & Snapp, 2010; Legleiter et al.,

2010; Sathasivam et al., 2010; Schaffar

et al., 2004; Shirendeb et al., 2011)

Table 2. Oligomers within proteopathies

Table highlights current evidence that examines the role of oligomers within different

proteinopathies. Table was adapted from information reviewed in Kalia et al. 2013. AD,

Alzheimer disease; ALS, amyotrophic lateral sclerosis; CBD, corticobasal degeneration; CJD,

Creutzfeld-Jakob disease; DLB, dementia with Lewy bodies; FTLD, frontotemporal lobar

degeneration; GSS, Gerstmann-Straussler-Scheinker syndrome; HD, Huntington disease; MSA,

multiple system atrophy; PD, Parkinson disease; PSP, progressive supranuclear palsy;

21

1.4.6. α-Syn Oligomers

Due to growing interest in the role of α-syn oligomers in PD pathogenesis, an increasing number

of in vitro and in vivo models have been used in order to better understand their properties.

Recently, studies have adopted and modified a classic protein fragment complementation assay

(PCA) in order to better understand and characterize the behaviour of α-syn oligomers (Outeiro

et al., 2008). This PCA was designed to study the dynamics of protein-protein interactions

within cells (Remy & Michnick, 2006). The concept involved fusing two proteins of interest to

complementary fragments of a reporter. If the proteins interact, the reporter fragments are

brought together refolding into the native structure, which reconstitutes its activity. Based on the

signal emitted by the reporter, protein-protein interactions can be monitored and quantified. The

reporter protein utilized for this assay was humanized Gaussia luciferase (hGluc). This

bioluminescent protein catalyzes the oxidation of the substrate coelenterazine in a reaction that

emits a blue light approximately 488nm in length. The signal generated by this reporter is

100-fold higher than other luciferases and can even be utilized in live cells. In a recent study, the

complementary amino (luc1) and carboxy (luc 2) domains generated by Remy and Michnick

were fused to wild-type α-syn (Outeiro et al., 2008) (see Figure 3.). These individual α-syn

fusion constructs (α-syn-luc1 and α-syn-luc 2) were inert by themselves, but upon α-syn

oligomerization the two halves were brought together, which led to the emission of a

bioluminescent signal. Utilizing the same concept, this group also created an additional PCA

that employed complementary fragments of green fluorescent protein (GFP). The utility of

fluorescent PCAs arises from the irreversible nature of the interaction between the

complementary reporter fragments. The reconstitution of the reporter permanently traps the

proteins of interest (α-syn) in their interacting conformation. This allows for better visualization

and study of transient protein interactions or states. Therefore, the authors claimed that using

this assay they were better able to monitor oligomerization by stabilizing this transient

conformation.

Experiments that utilized this GFP PCA revealed that α-syn oligomerization led to increased

cytotoxicity, but more importantly this assay also allowed researchers to test proteins and

substances that could potentially reverse or alleviate this toxicity. They discovered that

upregulation of proteins such as Hsp70 and CHIP (carboxyl terminus of Hsp70- interacting

protein) counteracted the effect of these oligomers (Outeiro et al., 2008). As members of the

22

Figure 3. Gaussia luciferase protein fragment complementation assay

Outeiro et al modified a pre-existing protein fragment complementation in order to study α-syn oligomerization. a) i) The enzyme gaussia

luciferase was split and the two complementary halves were attached to wild-type α-syn creating either α-syn-luc1 or α-syn-luc2.

Individually these two halves are inert. ii) Dimerization of α-syn brings the two halves together reconstituting the active luciferase enzyme.

iii) Upon addition of the substrate coelenterazine, a photo-oxidation reaction that is catalyzed by the active gaussia enzyme occurs releasing

a 475nm wavelength of light. Quantifying the amount of released light is used as a way to assess dimerization. b) Schematic illustrates the

chemiluminescent mechanism of coelenterazine. Coelenterazine reacts with oxygen and gaussia luciferase to yield an energy rich 1,2

dioxetane compound, which loses carbon dioxide creating a coelenteramide anion in the excited state. This excited molecule eventually loses

a photon and gets protonated creating coelenteramide

23

molecular chaperone system, these proteins manage aggregated intracellular species by either

refolding them or directing them toward proteasomal degradation (Danzer et al., 2011; Tetzlaff

et al., 2008). Thus, co-transfecting either CHIP or Hsp70 with the complementary α-syn-GFP

constructs reduced oligomerization, denoted by a decreased fluorescent signal, and mitigated

cytotoxicity (Tetzlaff et al., 2008). In addition to these fluorescent and bioluminescent reporters,

α-syn oligomeric conformations have also been stabilized using either genetic mutations or

chemical reagents (Danzer et al., 2007; Karpinar et al., 2009; Lazaro et al., 2014; Rockenstein et

al., 2014; Winner et al., 2011). Since similar artificial oligomers were utilized in this study (see

results section 4.5), the concluding portion of this section will focus on this aspect of α-syn

aggregation. The genetic approach involved inducing selective mutations within the α-syn

sequence that caused the protein to form oligomers instead of fibrils. The A56P, A76P, and the

triple A56P/A76P/A30P mutants demonstrated markedly different aggregation kinetics relative

to A30P and A53T mutants (Karpinar et al., 2009). The proline mutations inhibited β-sheet

formation, and thus make it more difficult for the A56P, A76P, and the triple A56P/A76P/A30P

mutants to form large aggregates. Fluorescently tagged proline mutants expressed within human

embryonic kidney (HEK) cells failed to undergo any significant aggregation unlike their A53T

and A30P counterparts. However, these oligomeric mutants still exhibited toxicity comparable

to cells transfected with the A53T mutant. Similar results were also observed in vivo in

Caenorhabditis elegans (C.elegans) and Drosophila models. When expressed within the

dopaminergic cells of C.elegans and Drosophila the proline mutants failed to aggregate, but

caused severe neurodegeneration (Karpinar et al., 2009). Worms and flies with the triple proline

mutation A56P/A76P/A30P exhibited the greatest dopaminergic loss. The neurodegeneration

induced by the oligomeric mutants also resulted in behavioural deficits. Worms with either the

A56P or A56P/A76P/A30P mutation exhibited reduced searching behaviour in the presence of

food, while flies exhibited a diminished climbing response, which is another behaviour

dependent on dopaminergic neurons (Karpinar et al., 2009). Another study also employed a

similar approach in order to study the toxicity of α-syn oligomers within their mammalian

model (Rockenstein et al., 2014; Winner et al., 2011). They mutated the glutamic acids residues

to lysine within α-syn’s N-terminus in order to inhibit salt bridge formation, an essential process

in fibrillization. These new oligomers exhibited greater toxicity in comparison to those observed

in the previous study. When injected into the substantia nigra of rats, these oligomers induced

greater dopaminergic cell loss than either the A30P or A53T mutants. In vitro the E35K and

24

E57K mutants caused increased caspase-3 activation relative to the classic fibril forming

mutants. Much of the toxic effects of these lysine mutants could be attributed to their ability to

induce calcium influxes, an occurrence also noted by an earlier study that chemically induced

the formation of oligomers. This study created oligomers by treating monomeric α-syn with

different buffers and purification techniques, which induced the formation of varying

populations of oligomers that differed in structure and toxicity (Danzer et al., 2007). Monomeric

α-syn exposed to sodium phosphate buffer during the oligomerization process resulted in a

heterogeneous population of globular, protofibrillar, and annular structures (Danzer et al., 2007).

These oligomers formed pores within synthetic vesicles and cell membranes causing an influx

of calcium from extracellular sources (Danzer et al., 2007; Winner et al., 2011). The

incorporation of either agitation or ultrafiltration created yet another distinct population of

oligomers that were more globular and lacked any pore forming abilities (Danzer et al., 2007).

However, these oligomers exhibited a remarkable propensity for recruiting cytoplasmic α-syn

into aggregates. When exogenously added to the culture media these oligomers could enter cells

and induce aggregation. This seeding ability observed in this study correlates well with an

emerging hypothesis in field, which suggests that PD pathology can be spread from a diseased

neuron to a healthy neighbour. Many believe the potential cell-to-cell transmitter to be α-syn,

but its exact structural conformation is still up for debate. It is possible that this transmitted

species could be oligomeric in nature, and due to better characterization methods we are getting

closer to determining its conformational identity.

1.4.6.1. E35K and E57K Lysine Mutants

Since the E35K and E57K lysine mutants will play a significant role within this project, this

section will review in greater detail their structural characteristics and findings from previous

studies.

Oligomers are defined as two or more repeating units of a protein of interest. They are

considered a precursor to fibrils and typically do not adopt rigid secondary motifs (Giehm et al.,

2011). These structures possess a wreath-like conformation with diameters ranging from 100-

180 nm. Unfortunately, they are very transient and often difficult to isolate. The role of the

lysine point mutations on residue 35 and 57 within the N-terminal region was to better stabilize

25

these structures. The E35K and E57K mutations inhibit the formation of salt bridges, which

form between β-sheets and stabilize this secondary conformation (Winner et al., 2011). By

inhibiting β-sheet structure formation these mutations are supposed to impede the transition into

fibrils and help retain α-syn within an oligomeric state. Interestingly, the study by Winner et al.

2012 indicated that in addition to inserting these point mutations, a specific monomer

purification protocol needed to be implemented in order to successfully facilitate the formation

of oligomeric structures. If this protocol was not incorporated, despite the presence of the lysine

mutations, the α-syn tended to form fibrillar structures. This unique oligomer purification

protocol excluded the streptomycin sulphate precipitation step. Streptomycin sulphate is

typically employed in order to precipitate DNA and ensures a cleaner monomer solution.

However, the authors argued that by excluding this step, the resulting solution was less

conducive for large scale α-syn self assembly because the DNA created a negatively charged

environment, which repelled the negative charges on α-syn (Winner et al., 2011). The study

demonstrated that α-syn purified using this protocol resulted in circular structures under EM,

which possessed a radius of 100 nm. These structures possessed significantly less thioflavin-T

binding capabilities relative fibrillar forms of the protein even after extended incubation periods.

The CD spectra revealed that after a 30 day incubation period these oligomer prone mutants

failed to exhibit any significant β-sheet structural motifs unlike their fibrillar counterparts.

Furthermore, the oligomer specific antibody A11, which exhibited little to no affinity for WT

fibrils, also recognized the structure formed from these lysine mutants.

The acute toxicity resulting from either cellular expression or viral injection, into the SN of

rodents, was significantly greater for the E35K and E57K oligomer prone mutants relative to the

fibril forming mutants A53T and E46K. These results were also nicely recapitulated using a

long-term model, which examined the chronic effects resulting from oligomer exposure. Mice

overexpressing the E57K mutant exhibited greater synaptic toxicity relative to WT and non-

transgenic littermates especially within the frontal cortex (Rockenstein et al., 2014). These mice

also exhibited a reduction in synaptic protein such as MAP2, synaptophysin, and synapsin 1.

These cellular changes were also accompanied by behavioural deficits that positively correlated

with the expression level of these mutants indicating a potential dose-dependent effect. Even

though the exact mechanisms by which these oligomers exert their effect is unclear, the authors

postulated that by impairing localization of synapsin 1 these oligomers indirectly impair vesicle

clustering within the active zone. By stabilizing these transient oligomeric conformations these

26

mutations allow us better study the role of these isoforms in pathology induction and

propagation using both in vitro and in vivo models.

1.5. Alpha-Synuclein Migration

One of the major topics addressed in this thesis is the intercellular migration of α-syn.

Therefore, the subsequent sections will review the history and development of this concept of α-

syn migration in addition to addressing the potential cellular mechanisms that could facilitate

the release and uptake of this protein.

Heiko Braak initially alluded to the migratory capabilities of α-syn in his staging model, which

indicated that Lewy bodies have a topographically predictable progression (Braak et al., 2003)

(see section 1.2.2.1). He proposed that within the central nervous system (CNS), these

inclusions originated in the brainstem and anterior olfactory nucleus spreading rostrally and

caudally. The damage caused by this pathology as it progressed through these extrastriatal

regions was hypothesized to induce dysfunctions such as hyposmia, cardiac denervation,

constipation, and sleep disorders, which were proposed to antedate the motor impairments.

Gastrointestinal dysfunctions in particular are of great interest because not only are they

considered to be early indicators of PD, but Heiko Braak and others believe that the

gastrointestinal tract could potentially be an entry point for pathogens that can instigate

idiopathic PD (reviewed in Hawkes et al., 2007). Since the enteric system is heavily innervated

by the vagus nerve, it is hypothesized that the α-syn aggregates induced by ingested pathogens

could propagate along these vagal efferents in order to access the CNS (Abbott et al., 2001;

Cersosimo et al., 2013; reviewed in Hawkes et al., 2007; Shannon et al., 2012). Following entry

into the brain these aggregates are predicted to follow the topographical progression outlined in

Braak’s staging model (see section 1.2.2.1). However, at the time this proposal was put forth

there was very little evidence clearly documenting α-syn’s migratory capabilities. It was only

after the transplant studies documented by two independent groups that work surrounding this

topic gained momentum (Kordower et al., 2008; Li et al., 2008). They published results that

strongly suggested the interneuronal propagation of α-syn pathology from diseased neurons to

newly grafted cells. They indicated that autopsies conducted on patients who received fetal

dopaminergic cell transplants revealed Lewy body pathology in these newly transplanted cells

27

similar to that of the older host neurons. Since these cells were relatively young, observing such

drastic Lewy pathology was unprecedented. Therefore, it was hypothesized that aggregates

from older neighbouring neurons were transmitted into these newly implanted cells and

subsequently acted as seeds that accelerated the pathology (reviewed in Brundin et al., 2008;

Kordower et al., 2008; Li et al., 2008). This rationale was termed the Prion Hypothesis of PD

because the authors believed that α-syn exhibited prion-like characteristics. Based on their

results, misfolded α-syn appeared to migrate between neighbouring cells inducing aberrant

conformational changes to healthy cytoplasmic α-syn. Following the proposal of this hypothesis

numerous different studies have attempted to ascertain its validity using a variety of in vitro and

in vivo models.

1.5.1. In Vitro Models

Initially in vitro cellular models provided the easiest way to characterize α-syn’s seeding and

interneuronal migratory capabilities. One of the first studies involved the addition of preformed

wild-type α-syn fibrils (PFF) to the culture media in order to determine if these aggregates could

induce the recruitment and aggregation of endogenous α-syn monomers (Luk et al., 2009a). In

the PBS control, α-syn was predominantly localized as puncta within presynaptic terminals,

while those treated with PFF, exhibited large Lewy body-like perinuclear inclusions. This

phenotype was most likely elicited by the PFF because treatment with either α-syn monomers or

non-specific exogenous proteins failed to produce analogous outcomes. The cytoplasmic

inclusions induced by the PFF were very similar to Lewy bodies found in patients. They were

detergent insoluble and contained posttranslational modifications such as phosphorylation of the

Ser129 residue and ubiquitination (Luk et al., 2009a; Sacino et al., 2013; Volpicelli-Daley et al.,

2011). The cores of these inclusions were primarily composed of exogenously added PFF, while

the periphery consisted of post-translationally modified endogenous α-syn (Luk et al., 2009a;

Sacino et al., 2013; Volpicelli-Daley et al., 2011). The authors discovered that recruitment and

modifications of endogenous proteins by these aggregates exhibited a temporal sequence

(Sacino et al., 2013; Volpicelli-Daley et al., 2011). A 24-96 hr incubation period with the PFF

was required before aggregates started to appear (Luk et al., 2009a; Volpicelli-Daley et al.,

2011). Immunoblots indicated that initial changes were very mild; however, as time progressed

a significant decrease in detergent soluble α-syn was accompanied by a concomitant increase in

28

high molecular weight α-syn species within the detergent insoluble fraction (Sacino et al., 2013;

Volpicelli-Daley et al., 2011). Furthermore, older neurons appeared to be more susceptible to

the effects of these fibrils forming aggregates (Luk et al., 2009a; Sacino et al., 2013; Volpicelli-

Daley et al., 2011). It is still unknown what compels endogenous α-syn to so readily conform to

the template set out by these aggregates, and it appears that as neurons get older they are less

able to counteract these toxic effects. Select studies have suggested that the NAC regions within

α-syn could mediate this process (Volpicelli-Daley et al., 2011; Waxman et al., 2010). Cells

transduced with PFF devoid of the NAC region failed to induce intracellular aggregation, while

N or C terminal truncated aggregates resulted in inclusion patterns indistinguishable from wild-

type PFF (Volpicelli-Daley et al., 2011; Waxman et al., 2010). A similar outcome was observed

when endogenous α-syn contained point mutations within the NAC region. The V66P, T72P,

and T75P point mutations significantly impeded intracellular inclusion formations following

treatment with exogenous wild-type PFF (Waxman et al., 2010). Therefore, α-syn’s

hydrophobic region appears to be crucial in this aggregation cascade, but so far there is still

insufficient evidence regarding how the NAC region potentially induces this process. It is clear

that the addition of exogenous aggregates can and will seed the formation of intracellular

inclusions, and elucidating the details of this process will be the next step in better

understanding α-syn’s prion-like propagation.

In addition to demonstrating the intraneuronal seeding capabilities of α-syn aggregates, these in

vitro models were also used to monitor interneuronal propagation. Paula Desplats and

colleagues were one of the first to examine the migration of α-syn. In their study two distinct in

vitro cell populations were created in which they first stably expressed myc-tagged α-syn, while

the other was labelled with Qtracker (Desplats et al., 2009). Upon co-incubation of these two

distinct populations, it was discovered that the cells labelled with Qtracker started to

progressively accumulate myc-tagged α-syn. These Qtracker recipient cells displayed

cytoplasmic myc-tagged α-syn inclusions that were Triton X insoluble and stained positively for

ubiquitin and Thio S. Other groups also started to report similar findings. In cells stably

expressing α-syn tagged with either DsRed or AcGFP1, approximately 8-10% of the recipient

cells were positive for α-syn from the donor populations after a 7-day co-incubation period

(Hansen et al., 2011). The authors also noted the appearance of punctate cytoplasmic structures

following the transfer of α-syn. The aggregates usually consisted of a α-syn-GFP core

surrounded by a larger area of endogenous DsRed or vice versa. More recently live cell imaging

29

and flow cytometry have been incorporated to provide a more refined and accurate method of

tracking migration (Reyes et al., 2015). These findings further substantiate the notion that α-syn

can propagate and can induce the recruitment of intracellularly expressed α-syn. Others have

even shown that α-syn aggregates can travel anterogradely from the soma via axons in order to

reach neighbouring cells (Freundt et al., 2012). Microfluidic devices that isolated neuronal cell

bodies from axons allowed Freundt and colleagues to study the migration of fluorescently

labelled α-syn fibrils. When added to compartments containing the cell bodies, these fibrils

traveled at an average velocity of 63 μm/min down the axon to reach naïve cells innervated by

these axons. The authors reported observing punctate α-syn structures within the naïve neurons

4 days post treatment. Interestingly, it was later discovered that direct cell-to-cell contact was

not even necessary in order to facilitate α-syn propagation (Freundt et al., 2012). The incubation

of naïve cells in the conditioned media from the donor cell populations is sufficient to induce

transfer and cellular uptake (Danzer et al., 2012; Danzer et al., 2011; Hansen et al., 2011).

Baseline luminescence was detectable using the PCA in naïve cells that had been incubated in

the conditioned media of cells expressing both the N and C-terminal α-syn-luciferase fragments

(Danzer et al., 2012; Danzer et al., 2011). Thus, studies using these cellular models provided

valuable information regarding the migratory behaviour of α-syn, but they were only the

beginning. In order to truly ascertain if α-syn did possess prion like characteristics, in vivo

animals models, which provide a more realistic neuronal environment, were the next step.

1.5.2. In Vivo Models

In a landmark study, the striatum and cortex of young transgenic mice were inoculated with

brain lysate derived from older cohorts that exhibited extensive Lewy body pathology (Luk,

Kehm, et al., 2012a). Following these injections mice began exhibiting Lewy body-like

inclusions that consisted of hyperphosphorylated α-syn species. Normally this pathology

developed in non-injected transgenic littermates on average after 200 days, while inoculated

animals developed inclusions within 90 days. Similar α-syn pathology was undetectable in age-

matched controls treated with PBS, but more interestingly mice inoculated with lysate from

nonsymptomatic transgenic cohorts failed to exhibit pathology within 90 days. This indicates

that inoculation alone is not the instigating factor. The aggregated α-syn species within the

inoculant are required to cause the accelerated development of inclusions. This experimental

30

paradigm has even been adapted to a primate model (Recasens et al., 2014). Monkeys that

received intranigral or intrastriatal inoculations of PD-derived LB extracts exhibited significant

nigrostriatal neurodegeneration, which was not observed in the control animals injected with

soluble α-syn. These results provide additional evidence that these aggregate forms of α-syn

play an important role in pathology propagation. Studies have even shown that treating

transgenic mice with α-syn specific antibodies helps drastically reduce the transfer of α-syn

aggregates by promoting microglial uptake of these toxic species (Bae et al., 2012). Treated

animals exhibited less pathology and performed better on functional tests. Furthermore, it

appears that the injected pathogenic α-syn species doesn’t have to be produced from a natural

source. Artificially created fibrils can elicit pathology remarkably similar to brain lysates from

diseased animals (Luk et al., 2012; Paumier et al., 2015). Inoculation with preformed fibrils

(PFFs) similarly resulted in wide spread α-syn inclusions that stained positively for ubiquitin,

Thio S, and were detergent insoluble containing high molecular weight α-syn species. Others

have also reported that injection of pathogenic brain lysates or fibrillar species caused similar

propagation and detrimental outcomes. Unilateral intracerebral injections resulted in

intraneuronal deposits that were distributed bilaterally from the olfactory bulb to the spinal cord

(Desplats et al., 2009; Masuda-Suzukake et al., 2013; Mougenot et al., 2012; Peelaerts et al.,

2015). However, regions containing neurons that either projected to or received input from the

inoculation sites developed the most prominent α-syn pathology (Luk et al., 2012a; Masuda-

Suzukake et al., 2013). Even injection into peripheral regions such as the hind limb

intramuscular junction resulted in the formation of α-syn inclusions and motor impairments

within A53T or WT transgenic mice (Sacino et al., 2014). Also, in addition to having

aggregated α-syn species within the initial inoculant, α-syn also needs to be endogenously

expressed in order for pathology induction and propagation (Luk et al., 2012a; Masuda-

Suzukake et al., 2013). α-Syn null mice injected with pathological aggregates failed to exhibit

neuronal lesions regardless of the incubation time (Luk et al., 2012a). It appears that the seeding

capabilities of the injected aggregates are ineffective in the absence of endogenous recruits.

Thus further substantiating the notion that α-syn pathology spreads in a prion-like manner

because both an instigator and an endogenous recipient are required in order to spread the

disease.

These in vitro and in vivo models (summarized in Error! Reference source not found.) have

provided ample evidence that lend support to the Prion Hypothesis. However, even though

31

many of these studies demonstrated this migratory process, they did not address the mechanistic

aspects such as how α-syn might be released, propagated extracellularly, and taken up in to

cells. Therefore the next few sections will examine in greater detail the specific components

involved in this migratory pathway, in order to provide a more comprehensive presentation of

how this disease might spread (see Figure 4.).

1.5.3. α-Syn Release

1.5.3.1. Exocytosis

It has been known for some time now that α-syn can be found in bodily fluids such as plasma,

cerebrospinal fluid, and brain interstitial fluid (Aasly et al., 2014). The α-syn within these fluids

is most likely derived from cells such as erythrocytes and neurons, which express high levels of

α-syn (Emmanouilidou et al., 2011). However, the way in which α-syn translocates from the cell

cytoplasm to the extracellular milieu is still somewhat unclear. One of the first mechanisms

proposed was an exocytic release of this protein. Studies have reported a high level of α-syn

within the media of cells overexpressing this protein (Emmanouilidou et al., 2010). This release

was not the result of cell death or membrane leakage because other overexpressed proteins such

as β-galactosidase were not recovered in the media (Jang et al., 2010; Lee et al., 2005).

Furthermore, when cells were grown at lower a temperature, which has been known to inhibit

exocytosis, the level of extracellular α-syn was significantly diminished (Lee et al.,2005).

Researchers also noted that intracellularly a small portion of α-syn within these overexpressing

cells were localized within vesicles (Jang et al., 2010), a pattern that was not observed for other

overexpressed proteins such as β-galactosidase (Jang et al., 2010; Lee et al., 2013; Lee et al.,

2005). The vesicular translocation and release of α-syn appeared to increase under stressful

conditions that promoted protein misfolding (Jang et al., 2010). Protein aggregation can be

induced by targeting regulatory organelles using reagents such as bafilomycin (lysosomal

inhibitor) and MG132 (proteasomal inhibitor) (Jang et al., 2010; Lee et al., 2013; Lee et al.,

2005; Poehler et al., 2014). Since these organelles maintain homeostasis by degrading misfolded

or aggregated proteins, inhibiting their function causes a build-up of these toxic species. Under

these conditions vesicular translocation and release of monomeric and aggregated α-syn was

augmented (Jang et al., 2010; Lee et al., 2013; Lee et al., 2005). Proteasomal inhibition caused a

32

Table 3. Overview of in vitro and in vivo α-syn propagation studies

Table highlights different in vivo and in vitro studies that have investigated the intercellular

transfer of pathological α-syn (transfer), intracellular aggregation induced by seeding with

extracellular fibrils (seeding) or have examined a combination of both aspects.

In vitro

Models

In vivo

Models

Reference

Transfer Seeding Transfer

X X (Desplats et al., 2009; H. J. Lee et al., 2010)

X X X (Hansen et al., 2011)

X X (Danzer et al., 2011; Sacino et al., 2013)

X (Alvarez-Erviti et al., 2011; Danzer et al., 2012;

Freundt et al., 2012; Kondo et al., 2011; Reyes et al.,

2015)

X (Danzer et al., 2007; Danzer et al., 2009; Luk et al.,

2009b; Nonaka et al., 2010; Volpicelli-Daley et al.,

2011; Waxman & Giasson, 2010)

X (Bae et al., 2012; Kordower et al., 2008; J. Y. Li et al.,

2008; Luk, Kehm, et al., 2012a; Masuda-Suzukake et

al., 2013; Mougenot et al., 2012; Paumier et al., 2015;

Peelaerts et al., 2015; Recasens et al., 2014)

33

Figure 4. Mechanisms of release and uptake of α-syn

Diagram outlines the mechanisms of release (a) and uptake (b) of α-syn, which are outlined in

section 1.5.3 and 1.5.4. a) i) exocytic release of α-syn via a non-classical pathway (section

1.5.3.1) ii) Exosomal release of α-syn by fusion of multivesicular bodies (MVBs) and the

plasma membrane (section 1.5.3.2). b) i) Passive diffusion of α-syn can potentially facilitate

uptake and release of this monomeric protein, but evidence is currently tenuous (section

1.5.4.1). ii) Receptor mediated uptake of α-syn, but the exact identity of this receptor remains

elusive (section 1.5.4.2). iii) Exosomal uptake of α-syn via fusion of extracellular vesicles with

the plasma membrane (section 1.5.4.3). These mechanisms are currently the most investigated

and likely mechanisms of α-syn propagation.

34

7-fold increase in α-syn within the media and a 2-fold increase within the vesicle fraction, while

lysosomal inhibition resulted in a 2-fold increase within the condition media and 5-fold increase

within the vesicle fraction (Jang et al., 2010; Lee et al., 2013; Lee et al., 2005). Other toxic

reagents such as 3-MA (autophagy inhibitor) and rotenone (mitochondrial complex I inhibitor)

elicited similar results (Jang et al., 2010). Mass spectrometry revealed that the extracellular and

vesicular α-syn had three times more modifications than their cytosolic counterpart, which

indicates a potential selectivity of aberrant protein conformations for exocytic release (Jang et

al., 2010; Lee et al., 2013). The exact mechanism involved in the targeted vesicular

translocation or the main pathway involved in the release of these aggregates is still somewhat

ambiguous. Apparently, α-syn aggregates were released via a ER/Golgi independent pathway

because brefeldin A (an inhibitor of ER/ Golgi trafficking) failed to inhibit α-syn release (Jang

et al., 2010). Even though it is not uncommon for certain cytosolic proteins such as interleukin-

1β and high-mobility group protein B1 (HMGB1) to utilize non-classical export pathways, the

lack of a standard release mechanism just raises more questions regarding how α-syn is

trafficked intercellularly (Jang et al., 2010; Lee et al., 2005). Therefore, despite growing

evidence indicating the extracellular release of α-syn, there are still many uncertainties

regarding the steps involved in this process.

1.5.3.2. Exosomal Release

Exosomal release of α-syn is thought to be another mechanism by which this cytosolic protein

can escape into the extracellular space. Exosomes are round cup-shaped structures 40-100 nm in

size, which are generated within the late endosome by inward invagination of the limiting

membrane (reviewed in Nickel, 2003). This process creates a structure called a multivesicular

endosome (MVE), which houses smaller compartments known as intraluminal vesicles (ILVs)

(Simons et al., 2009). These MVEs can serve as storage compartments housing proteins within

their ILVs, which can be re-released into the cytosol, degraded via fusion with the lysosome or

exported into the extracellular space when the MVEs merge with the cell membrane (Simons et

al., 2009). Once released, these ILVs are called exosomes, which are considered to be carriers

that can propagate and facilitate the entry of pathogenic material into neighbouring cells. A

significant portion of the released α-syn was reported to be associated with vesicle like

structures (Emmanouilidou et al., 2010). Ultracentrifugation of the culture media and

35

subsequent immunoblot analyses revealed that these vesicles were positive for the exosome

specific proteins, Flotillin and Alix. Electron micrographs of this same fraction revealed the

structures to be rounded and cup-shaped with a diameter ranging from 50-140 nm. Furthermore,

intracellular calcium levels, a known regulator of exosome-mediated release, strongly

influenced extracellular α-syn levels (Emmanouilidou et al., 2010). Administration of 100 nm of

thapsigargin or ionomycin, reagents that raise intracellular calcium concentrations, caused a

significant increase in secreted α-syn while addition of BAPTA-AM, a cell permeable calcium

chelator, hindered release. However, the most striking results arose from inhibiting the

lysosomal pathway. The authors discovered that inhibiting the lysosomal route in the MVE

pathway using methylamine or chloroquine significantly increased extracellular α-syn within the

exosomal fraction (Emmanouilidou et al., 2010). Others have also reported similar increases in

exosomal release of α-syn due to inhibition of lysosomal function using alternative reagents

such as ammonium chloride or bafilomycin A1(Alvarez-Erviti et al., 2011; Danzer et al., 2012;

Poehler et al., 2014). It is believed that one of the roles of these MVEs is to sequester toxic or

aggregated protein species in order to minimize damage to the cell. These isolated proteins are

then trafficked to the lysosome for degradation in addition to being released extracellularly

(Danzer et al., 2012; Simons et al., 2009). However, inhibiting the function of the lysosome

forced cells to rely more heavily on their release mechanisms, resulting in more ILVs released

extracellularly (Alvarez-Erviti et al., 2011; Danzer et al., 2012; Poehler et al., 2014).

Interestingly, Poehler and colleagues also demonstrated that impeding the autophagy lysosomal

pathway in α-syn overexpressing cells not only promoted secretion, but also caused a transition

in the structure of the intracellular α-syn species. They transformed from large aggregates, to

more toxic oligomeric isoforms, which were secreted in association with exosomes. Danzer and

colleagues previously reported similar results. Their study was one of the first to clearly

demonstrate that these exosomes house oligomeric forms of α-syn (Danzer et al., 2012). Using

the protein fragment complementation assay, which identifies α-syn oligomers, they observed a

strong luciferase signal in their exosomal fraction (Danzer et al., 2012). Furthermore, these

oligomers appeared to be located within the lumen and on the exterior membrane of these

exosomes. Combination of proteases and membrane permeabilizing detergents were required to

completely degrade α-syn within the exosomal fractions. These results indicate that not only are

exosomes carriers for the more toxic α-syn species, but that they can also offer significant

protection while transmitting these isoforms.

36

Figure 5. Exosomal biogenesis

Schematic outlines the maturation process for exosomes and the potential pathways the mature intracellular vesicles can take. The early

endosome, which is tube-like in shape, is located near the outer portion of the cytoplasm. As it matures it becomes more spherical in shape

and moves closer to the nucleus. Inward budding of these mature endosome causes the formation of intraluminal vesicles (ILVs), and the

structures that house these vesicles are called multivesicular body (MVB). MVBs can either fuse with the lysosome causing the hydrolysis of

its contents or it can fuse with the plasma membrane releasing its ILVs, which are now called exosomes. These exosomes can act as carriers

transporting the intracellular contents through the extracellular space.

37

1.5.4. α-Syn Uptake

1.5.4.1. Diffusion

Studies presented in section 1.5.1 demonstrated that α-syn is not primarily confined to the cell

cytoplasm. Following cellular release, aggregated versions of this protein can influence and

trigger toxic cascades in neighbouring cells. However, it is unlikely that these effects are

triggered from its extracellular location. Studies have shown that cells can internalize α-syn, but

the method of entry is dependent on the conformational state of the protein. All three synuclein

proteins (α, β, and γ) can translocate across the plasma membrane, but with varying efficiencies

(Ahn et al., 2006). In α-syn, two distinct regions are crucial for this process. Truncated versions

of α-syn lacking either the N-terminal or the NAC domains exhibited significant deficiencies in

membrane translocation (Ahn et al., 2006). Of the two regions, the N-terminal truncated

isoforms exhibited the greatest extracellular localization. The authors discovered that the amino

acid repeat sequence within this region facilitated the transfer into the cell (Ahn et al., 2006).

Mutants that lacked sequences within this region exhibited translocation deficiencies that

positively correlated with number of repeats that were removed. Wild-type α-syn monomers that

possessed both the N-terminal and NAC regions were able to reach detectable levels within the

cell cytoplasm 5 mins after addition into the culture media. Moreover, the uptake of monomeric

α-syn was unaffected by temperatures as low as 4°C or disruptions to the cell’s microfilament

structures, which should hamper receptor mediate uptake (Ahn et al., 2006). Also, the

transfection of a dynamin mutant, which inhibits endocytosis, due to deficiencies in guanosine-

5'-triphosphate (GTP) hydrolysis, failed to affect cellular entry (Ahn et al., 2006; Lee et al.,

2008b). These internalized monomers were not present in any of the vesicular fractions, and

appear to be able to freely travel between the cell interior and exterior (Lee et al., 2008b). The

level of α-syn within these locations at any given time point was primarily influenced by a

concentration gradient. Taken together these results indicate that the internalization of

monomeric α-syn is similar to diffusion because it is temperature insensitive, occurs very

rapidly via a route independent from normal endocytosis, and is strongly influenced by a α-syn

concentration gradient. However, it should be noted that to-date the work by Ahn et al. 2006 and

Lee et al. 2008 concerning α-syn diffusion has yet to be replicated. Consequently, many are

skeptical regarding the assertion that α-syn can travel unrestricted across the plasma membrane.

38

Therefore, until these claims can be further validated movement via diffusion remains very

unlikely.

1.5.4.2. Endocytosis

Unlike α-syn monomers, larger synuclein structures such as fibrils and oligomers are unable to

access the intracellular environment so easily. These larger α-syn conformations potentially

utilize an endocytic pathway to gain entry into cells. Low temperatures, a classical inhibitor of

endocytosis, significantly diminished the level of extracellular α-syn fibrils that were recovered

from the cell cytoplasm (Lee et al., 2008b). Mutations that inhibit the function of Rab5a and

dynamin, proteins that regulate membrane trafficking in endocytic pathways, induced similar

reductions in α-syn aggregate internalization (Lee et al., 2008a; Sung et al., 2001). Fractionation

studies in these same cells revealed a greater amount of α-syn within the membrane fraction

relative to the cytosolic pool because much of the extracellular aggregates were unable to enter

the cytoplasm, and thus remained attached to the cell membrane (Sung et al., 2001). However,

when cellular uptake did occur, the authors noted that the fibrils were predominantly localized

within the vesicular fraction. These internalized proteins colocalized with markers for the early

and late endosome, indicating that α-syn moved through the endosomal compartments, a

common occurrence for proteins that have been endocytosed. Results from this study even

alluded to the presence of a α-syn receptor, which could potentially facilitate this endocytic

process. The authors discovered that treatment of cells with proteases prior to addition of the α-

syn aggregates drastically reduced uptake of both oligomers and fibrils potentially due to the

loss of an extracellular receptor (Sung et al., 2001). Additional studies with cells that had been

permeabilized, with the detergent digitonin, displayed FITC labelled α-syn fibrils primarily

along the plasma membrane, while nonpermeabilized cells exhibited more cytoplasmic

localization (Lee et al., 2008b). These results not only further support the presence of

extracellular binding regions, but also the role of the intracellular machinery in facilitating

uptake, indicating that aggregate internalization is a cell-mediated process with specialized

components.

More recently, another important component in this endocytic process was examined, which

was the mechanism by which α-syn oligomers or aggregates might escape from the intracellular

39

vesicles following uptake. The authors demonstrated that α-syn exhibited remarkably similar

characteristics to adenovirus protein VI, a non-enveloped virus that forms an amphipathic helix

in order to disrupt endocytic vesicle membranes so that it can gain access to the cytoplasm

(Freeman et al., 2013). Since α-syn has also been known to form helical structures that can

disrupt membranes, the authors proposed that α-syn aggregates could generate analogous

ruptures in endocytic vesicles in order to enter cytoplasm. In order to study the effects of α-syn

aggregates on endocytic vesicles, they utilized fluorescently labelled Galectin-3, a protein that

recognizes and binds β-galactoside sugars, which are normally localized on the external plasma

membrane or the interior wall of intracellular vesicles following endocytosis (Freeman et al.,

2013; Varkey et al., 2010). Galectin-3 relocalization was used to identify ruptured vesicles in

earlier experiments examining the function of adenovirus protein VI (Freeman et al., 2013). In

their current study, the authors discovered that treatment with wild-type α-syn fibrils resulted in

the reorganization of mcherry labelled glaectin-3 (chGal3) from a diffuse cytoplasmic pattern to

punctate structures within the cell interior. The fluorescently labelled α-syn fibrils either

colocalized with the punctate mcherry structures or were located near the periphery. Since this

phenotype was not observed after the addition of monomeric wild-type α-syn, the authors

believed that this reorganization was indicative of endocytic vesicle disruption. Similarly to

adenovirus protein VI, α-syn induced vesicle rupture caused an increase in the level of reactive

oxygen species (ROS) and mitochondrial oxidation relative to untreated cells. However, unlike

adenovirus protein VI, which predominantly induced the rupture of endosomal vesicles, the α-

syn aggregates appeared to target lysosomal compartments. The chGal3 vesicles were positive

for the lysosomal marker lysosomal associated membrane protein-1 (LAMP-1) (Freeman et al.,

2013). Thus, based on this and other studies, endocytic uptake of α-syn aggregates appears to be

a viable mechanism. Not only does the cell possess the machinery to facilitate internalization,

but these aggregates also possess the ability to escape the confines of the endocytic vesicles

following uptake.

1.5.4.3. Exosomal Uptake

In addition to endocytosis, exosome mediated membrane translocation is another proposed route

of entry for α-syn aggregates. These exosomes can act as carriers transporting aggregates from

diseased cells to healthy neighbours. α-Syn oligomers contained within exosomes were able to

translocate the plasma membrane much more readily than those unassociated with these lipid

40

vesicles (Danzer et al., 2012). However, exosomal membrane integrity must be maintained in

order to facilitate this translocation process. Recently a study demonstrated that if the exosomal

membranes were compromised via sonication, which disrupts the lipid bilayer, oligomer uptake

was drastically reduced. Thus, based on these results it appears that functionally intact exosomes

are required to promote the uptake of α-syn oligomers. Interestingly, similar transport

mechanisms have also been noted in other diseases. PrP (SC), the abnormal isoform in prion

disease, has also been shown to utilize exosomes to propagate and gain access to cells. These

exosomes even facilitated the transfer of prion pathology when inoculated into mice (Vella et

al., 2007). In AD, which is characterized by extraneuronal inclusions, amyloid-β (Aβ) the

protein found in these deposits was shown to be cleaved within early endosomes and routed to

MVBs in HeLa and N2a cells (Vella et al., 2007). Subsequently a small portion of the Aβ

peptides were later secreted in association with exosomes (Rajendran et al., 2006). In MS,

extracellular exosomes were reported as carrying high levels of miR-219 micro RNA, which

supposedly promotes the formation and maintenance of compact myelin (Rajendran et al.,

2006). Therefore, exosomes appear to be a very versatile transport system that can not only

ferry proteins such as α-syn across the extracellular environment, but also facilitate intracellular

transport.

41

Chapter 2

Hypothesis and Research Aims

The prion-like spread of pernicious isoforms of α-syn is an idea that has garnered a lot of

attention over the past few years. Numerous in vitro and in vivo models have not only

demonstrated that extracellular α-syn fibrils can induce aggregation of its intracellular

monomeric counterpart, but also that these conformational shifts can be propagated to healthy

neighbouring cells (see section 1.5.1 and 1.5.2). For many of these models, preformed α-syn

fibrils or brain homogenate from animals with α-syn inclusions have been used to induce

pathology. However, more recent evidence has begun to indicate that α-syn oligomers are

potentially a more toxic form of this protein (see section 1.4.6). These α-syn oligomers have

even been reported to be elevated within the CSF of both sporadic PD cases as well as

asymptomatic individuals with the LRRK2 mutation (Aasly et al., 2014). Evidence such as these

suggests that oligomers could play a very important role in PD pathogenesis. However, despite

growing interest in these α-syn species, one aspect still remains understudied and that is their

ability to induce aggregation. Since a fundamental component of the prion hypothesis is the

ability of pathological species to recruit monomeric α-syn into aggregated conformations, it is

important to evaluate the seeding capabilities of these oligomers in order to determine if they

could potentially be involved in this pathway. Moreover, comparatively assessing the seeding

capabilities of α-syn oligomers relative to fibrils might even reveal new information about the

aggregation process. Therefore, further exploring the properties of these α-syn oligomers might

lead to discoveries that can help solve some the mysteries surrounding the mechanisms involved

in PD pathogenesis.

Previous studies have demonstrated that similarly to fibrils α-syn oligomers possess the ability

to seed intracellular aggregation (Danzer et al., 2007; Danzer et al., 2009). Furthermore, in other

diseases such as Alzheimer’s oligomeric versions of the pathogenic protein are taken up more

readily than their fibrillar counterparts (Chafekar et al., 2008). Therefore, based on these results

we hypothesize that α-syn oligomers will induce greater intracellular aggregation than fibrils.

The two main aims of this project are 1) determine how effectively oligomers instigate

aggregation by comparing the seeding capabilities of these smaller α-syn species to different α-

42

syn fibrils, which have been traditionally used in aggregation assays 2) ascertain whether

oligomer-seeded pathology can be propagated intercellularly to neighbouring cells. In order to

accomplish aim 1 this study will employ a cell-based model, which utilizes the luciferase

protein fragment complementation assay (PCA) discussed in section (1.4.6). Since this assay

allows α-syn interactions to be quantified, it will be useful when comparing the aggregation

induction potential of the α-syn oligomers and fibrils. The oligomers used in this study will be

based on the model designed by Winner et al., 2011 (see section 1.4.6), which involved the

insertion of point mutations (E35K or E57K) within the α-syn sequence in order to maintain

oligomeric species by inhibiting the formation of β-sheet secondary structures. The larger fibrils

will be generated from either phosphorylated or non-phosphorylated recombinant wild-type

(WT) or mutant (A30P or A53T) α-syn. Following treatment with these different α-syn species,

cellular fractionation and protein immunoblots will be utilized in addition to the luminescence

assay to characterize and compare the resultant aggregation. Immunocytochemistry will also be

used to further expand and verify the findings from the biochemical and PCA assay. In order to

address aim two, a fluorescent cell-based co-culture model will be established through which the

intercellular effects induced by oligomer-seeded pathology can be monitored. Using this model

the extent of pathology development in untreated cell will be quantified relative to the control

condition.

43

Chapter 3

Material and Methods

3.1. Vector Design

The α-syn-luc constructs, contained within a PCDNA 3 vector, were a kind gift from Dr.

Pamela Mclean (Mayo Clinic, Jacksonville Florida). These constructs were transferred into a

PCMV6A vector by incorporating XhoI and BglII restriction sites up and downstream of the α-

syn-luc sequence respectively using PCR primers created by the ACGT corporation (Toronto,

Canada). The sequences were then cut using the respective enzymes (XhoI and BglII) and

ligated into PCMV6A resulting in vectors that contained α-syn-luc 1 or α-syn-luc 2 (all

reagents were acquired from New England Biolabs Whitby, Canada). In order to create a single

vector, which contained both synuclein luciferase constructs, a PCR fragment of the α-syn-luc 1

sequence from the previous ligation was generated. This new α-syn-luc 1 fragment contained

additional SpeI restriction sites incorporated upstream and downstream of the cytomegalovirus

promoter and poly A tail respectively. The α-syn-luc 1 PCR fragment, and the PCMV6A α-syn-

luc 2 (which had a single SpeI site upstream of α-syn-luc 2) vector were cut using the SpeI

enzyme, and the PCR fragment was ligated into the PCMV6A α-syn-luc 2 vector. This ligation

resulted in a single vector that housed both the synuclein luciferase constructs (all reagents

were obtained from New England Biolabs). Orientation of the new ligated α-syn-luc1 sequence

was validated using primers for regions upstream of the CMV promoter and downstream of the

α-syn-luc 1 sequence. A similar approach was also utilized to create one of the controls for this

experiment, which was a vector that housed both the linker-luc 1 and α-syn-luc2 sequences.

3.2. Cell Culture

H4 neuroglioma cells (ATCC, Manassas, Virginia) were used in all, luciferase dimerization,

exogenous protein aggregation, and immunocytochemistry studies. H4 are neuroglioma cells

derived from the brain tissue of a 37 year old of Caucasian male (Krex et al., 2001).

Immunohistochemical analyses have indicated that these cells stain positively for neuron-

44

specific enolase (NSE), which indicates that they are neuroectodermal in origin (Krex et al.,

2001). However, they fail to exhibit reactivity for synaptophysin indicating that they are not of

neuronal origin (Krex et al., 2001). This cell line was chosen for the following experiments

because they exhibit a rapid doubling time and high transfection efficiency in comparison to the

commonly used SHS5Y cells. Other groups who have used the same cell line in order to

conduct the split luciferase enzyme assay, and therefore have already verified the efficacy of

this system. Furthermore, due to their glial origin, these cells exhibit very low levels of

endogenous α-syn, which makes it easier to differentiate between exogenously added α-syn

constructs and endogenous protein.

H4 neuroglioma cells were maintained on 100 x 20 mm tissue culture dishes (Sarstedt, Nümbrecht,

Germany) in 10 mL of Dulbecco’s modified Eagle medium (DMEM) with 10% fetal bovine serum

(FBS) (Wisent, St. Bruno, Canada). Cells were passaged using 1X trypsin EDTA (Wisent, St.

Bruno, Canada) and 1/5 of the cell pellet was replatted in 9ml of fresh media, and cells were

maintained at 37°C in 5% CO2.

3.3. Stable Cell Line Generation

Once H4 cells reached 80-90% confluence, the synuclein luciferase vectors were transfected

using Lipofectamine 2000 (Life Technologies, Ontario, Canada). A ratio of 2:1 microliters

Lipofectamine to micrograms of plasmid DNA was used respectively. 48 hrs post transfection,

cells were exposed to media that contained 1ug/ml of puromycin (Bioshop, Canada) in order to

begin the selection process, and cells were maintained in media containing this selection factor

for the duration of the experiment. This method generated a mixed population of stable cells that

exhibited robust expression of our fusion proteins for approximately 10 passages. The presence

of the constructs was verified using western blotting and luminescence assays.

3.4. Gaussia Assay

Stable H4 cells containing the synuclein luciferase constructs were collected by adding cold

Phosphate Buffered Saline (PBS) (Wisent, St. Bruno, Canada) to plate and then scrapping off

adherent cells using a cell scraper (Thermo Scientific, Rockford, Illinois). Cell solution was

45

Figure 6. Double promoter vector ligation

Schematic outlines the transfer of α-syn-luc 1 and α-syn-luc 2 from their original PCDNA 3 vector into a single PCMV6A vector. Initially,

both constructs were transferred into a PCMV6A vector. Then primers were generated that allowed the replication of the segment that

contained the PCMV promoter, α-syn-luc 1, and the poly A tail. These primers also incorporated SpeI sites up stream of the promoter and

downstream of the poly A tail. The newly formed PCR product was digested and ligated into an opened PCMV6A that already contained α-

syn-luc 2. This ligation generated a single vector that housed both α-syn-luc 1 and α-syn-luc 2 each with its own promoter and poly A tail.

2

46

collected and spun down at 800 G at 4°C using tabletop centrifuge. Following spin down, the

cell pellet was hypotonically lysed using 10 mM Tris pH 7.5 buffer. The cell lysate was then

aspirated twice through a 27G needle (BD Bioscience, Ontario, Canada) in order to further

promote cell lysis. Lysate was then spun at 800G for 10 min at 4°C on a tabletop centrifuge.

The supernatant was collected and used for subsequent luminescence experiments. A Bradford

protein assay was used to equilibrate the protein concentration across the different conditions to

1ug/ul for all luminescence assays.

3.5. Coelenterazine Preparation

Native coelenterazine (Nanolight Technologies, Arizona, United States) was purchased in a

lyophilized form and diluted to a stock concentration of 1 mM in 100% ethanol (Caledon

Laboratories, Ontario, Canada). This stock solution was then diluted to a working concentration

of 40 μM in 10 mM Tris for all luminescence experiments.

3.6. Luminescence Assay

All luminescence assays were performed on a Spectra Max L luminometer (Molecular devices,

California, United States) programed to read 480 nm wavelengths. 50 μl of cell lysate at a

concentration of 1 μg/μl was aliquoted into 96 well Costar white opaque plates (Fisher

Scientific). Samples were then electronically injected with 50 μl of 40 μM coelenterazine

working solution and an endpoint assay was conducted with an integration time of 5sec.

3.7. Electron Microscopy

The α-syn fibrils were imaged using electron microscopy (EM) in order to characterize their

structure. 10 μl of the α-syn fibrils diluted in PBS were loaded onto a pioloform filmed copper

grid and stained with 10 μl of 1% phosphotungstic acid (Bio Basic, Ontario, Canada) for 2

minutes. The samples were examined and photographed in a Hitachi H7000 transmission

electron microscope at an accelerating voltage of 75Kv.

47

a) b)

(5min) (5min)

Figure 7. Fractionation protocol outline

a) Flow diagram outlines the fractionation protocol that was initially utilize d to isolate soluble,

membrane and insoluble fractions within cell lysate. b) Following optimization, a shortened

protocol was created that allowed the isolation of detergent soluble and insoluble fraction, which

was later used to assess level of α-syn aggregation.

48

3.8. Monomer purification and phosphorylation

E.coli were subcloned with α-syn cDNA (WT or mutant), which was contained within a pET-

28a vector. Following α-syn expression, bacterial pellet was suspended in PBS containing 1mM

phenylmethylsulfonyl fluoride. The bacterial suspension was then sonicated for 30 sec and

boiled for 15 min, followed by ultracentrifugation at 150,000 x g. The supernatant was dialyzed

against 50 mM Tris, pH 8.3, loaded onto a Q-sepharose column and eluted with a 0-500 mM

NaCl step gradient. The eluents were desalted and dialyzed in 5 mM phosphate buffer, pH 7.3.

In order to induce phosphorylation of the ser129 residue, α-syn vector was co-transfected with a

polo-like kinase vector. This kinase has been shown to be very efficient at specifically

phosphorylating the ser129 residue. Presence of phosphorylation was assessed using

immunoblots and HPLC.

3.9. Fibril and Oligomer Treatment

α-Syn fibrils and oligomers were formed by diluting α-syn monomers in PBS and shaking at

200 rpm at 37°C for 10 days. 24 hrs prior to treatment, 600,000 H4 neuroglioma cells stably

expressing α-syn-luc 1 and α-syn-luc 2 were seeded on a 60 mm tissue culture plate (Sarstedt,

Nümbrecht, Germany). Fibrils and oligomers were diluted to a concentration of .5 μg/μl in PBS

and sonicated for 30 sec on ice using a Fisher Scientific Sonic Dismembrator. After sonication,

α-syn fibrils were added to the culture media resulting in a final concentration of either 1μg/ml,

3 μg/ml or 10 μg/ml. 24 hrs after incubation with the α-syn fibrils and oligomers, cell were

collected using 1X trypsin EDTA.

3.10. Cell Death Assay

After fibril treatment cell death was assessed using a Trypan Blue (Gibco, Ontario, Canada) cell

death assay. Cells contained DMEM with 10% FBS were mixed with the Trypan blue solution

in a 1:1 ratio, and the number of cell, which had not taken up the dye were measured using a

hemocytometer.

49

3.11. Fractionation

In order to characterize the level of soluble and insoluble α-syn, cells that were treated were

partitioned into different fractions (see Figure 7.). Following fibril treatment the cell pellet was

subjected to different buffers in order to separate the cytosolic, membrane and insoluble

fractions. Initially, the pellet was hypotonically lysed using swelling buffer (10mM HEPES and

18 mM potassium acetate at pH 7.2 + protease inhibitor cocktail (PIC)). The resulting lysate was

spun down at 14000 rpm at 4°C on a tabletop centrifuge for 5 mins. Both the supernatant and

pellet were collect for separate purposes. The supernatant was spun again at 100,000 G for 15

mins at 4°C using a tabletop ultracentrifuge (Beckman Coulter, Ontario, Canada) in order to

sediment any vesicles or debris. The supernatant from this spin was utilized as the soluble

(cytosolic fraction). The pellet from the earlier spin of the cell lysate was suspended in Tris

lysis buffer (100mM NaCl, 50mM Tris, 1mM EDTA + 1% Trition X v/v + PIC). The solution

was then spun at 14,000 rpm for 5min at 4°C. The supernatant from this spin was collected and

utilized as the membrane fraction while the pellet was considered detergent insoluble and

treated with 8M urea in a 20 mM Tris buffer pH 7.5. This solution was then sonicated for 10 sec

on ice and then spun at 100,000G for 30 min using a tabletop ultra centrifuge. The supernatant

was collected and used as the insoluble fraction. After the initial optimization, all future

protocols utilized only detergent soluble and insoluble fractions (see Figure 7.)

3.12. Western Blotting

Samples were incubated in 1X sample buffer (106 mM Tris HCl, 141 mM Tris Base, 2%

v/vLDS, 10% v/v Glycerol, 0.51 mM Ethylenediaminetetraacetic acid (EDTA), 0.22 mM

SERVA Blue G250, 0.175 mM Phenol Red, pH 8.5) at 70°C for 10min. They were then loaded

onto a 12% Bis Tris gel (self-casting using Biorad casting equipment) and run at 200V for

50min in 3-(N-morpholino)propanesulfonic acid (MOPS) buffer (50 mM MOPS, 50 mM Tris

Base, 0.1% v/v SDS, 1 mM EDTA, pH 7.7). Proteins were then transferred onto a 0.2 μm pore

sized nitrocellulose membrane (Bio Rad, Ontario, Canada) at 55V for 80min in transfer buffer

(25 mM Bicine, 25 mM Bis-Tris, 1 mM EDTA, pH 7.2). After transfer, membranes were

blocked for 60 min with 5% evaporated milk (Bioshop, Canada) in tris buffered saline (0.5 M

NaCl and 0.05 M Tris at pH 7.2) and 1% TWEEN 20 (Bioshop, Canada) (TBST). Membranes

50

were then incubated with primary antibody in 2.5% evaporated milk in Tris buffered saline and

tween (TBST) for 60 min at room temperature or overnight at 4°C. The primary antibodies used

include anti-α-synuclein antibody (monoclonal #42, BD Transduction, 38 Sparks, Maryland),

anti-gaussia antibody (polyclonal from Nanolight Technologies, Arizona, United States), anti-

tubulin antibody (Cell signalling, Massachusetts, United States), anti-actin antibody (Thermo

Scientific, Illinois, United States), and anti N and C-terminal Gaussia antibody (produced by the

Fraser Lab, Tanz Centre, Toronto Canada). Membranes were washed 3 times for 5 min with

TBST, and then incubated for 60 min at room temperature with goat-anti-mouse or goat-anti-

rabbit IgG secondary antibodies (Immunopure, Thermo Scientific, Rockford, Illinois) in 2.5 %

evaporated milk and TBST. Membranes were washed again 3 times for 5 min in TBST before

the addition of Western lighting Plus ECL solution (PerkinElmer, Massachusetts, United

States). Signals were resolved on a Bio Rad ChemiDOc XRS+, and the bands were quantified

using Image Lab 2 software.

3.13. Fluorescent Imaging

5 x 104 H4 neuroglioma cells were seeded onto square coverslips (VWR, Alberta, Canada) and

incubated at 37°C in 5% CO2 12 hrs prior to treatment with synuclein fibrils (for fibril treatment

see section 2.8). 24hrs after exposure to exogenous proteins, these cells were fixed with 4%

paraformaldehyde (PFA) (Bioshop, Canada Inc) for 8 min followed by three 5min washes with

PBS. Coverslips were then incubated in PBS containing 0.1% Triton X100 (Bioshop Canada,

Inc) for 5min, and then treated with .02% Thioflavin S (Sigma-Aldrich, Missouri, United States)

diluted in PBS for 8min. Excess Thioflavin S was removed using two 2 min washes with 50%

ethanol. Coverslips were then blocked for 20 min in PBS containing 0.1% Triton X100, and 5%

goat serum. After blocking, slides were incubated with primary antibody diluted in PBS

containing 5% goat serum for 1hr, followed by three 5 min washes with PBS. Primary

antibodies used include anti-α-synuclein antibody (monoclonal, Life Technologies, Ontario,

Canada), and anti-gaussia antibody (polyclonal from Nanolight Technologies, Arizona, United

States). Coverslips were then incubated with Alexa Fluor 488 goat-anti-mouse or Alexa Fluor

555 goat-anti-rabbit IgG secondary antibodies (Life Technologies, Ontario, Canada) for 40 min

followed by three 5min washes in PBS. Following the final wash step, slides were then

incubated with 4', 6-diamidino-2-phenylindole (DAPI) (Roche, Quebec, Canada) diluted at a

ratio of 1:1000 in PBS for 15 mins. This incubation period was then followed up with three 5

51

min washes in PBS. Coverslips were mounted onto slides using Prolong Diamond antifade

reagent (Life Technologies, Ontario, Canada) and allowed to dry overnight. Slides were imaged

on a Leica TCS SP8 non-resonant confocal microscope.

3.14. Co-seeding

In order to observe the intercellular propagation of α-syn two distinct populations of H4

neuroglioma cells were co-seeded and observed for a time course of 24 and 48 hrs (see Figure

8.). The first population of naïve H4 cells (donor cells) was grown in a six well plate (Sarstedt,

Nümbrecht, Germany) and transfected with either PCDNA3 or α-syn using Lipofectamine 2000

(Life Technologies, Ontario, Canada). 24 hrs post transfection cells were treated with either PBS

or 10 ug/ml of IAPP, E35K or E57K. 24 hrs after treatment with either PBS or exogenous

aggregates, donor cells were collected using 1X trypsin EDTA, treated with Cyto ID (see

section 3.14), and then co-seeded on cover slips with a second population of H4 cells (recipient

cells). Recipient cells were transfected 24 hrs prior to the co-incubation with the α-syn-venus

fusion constructs (α-syn-venus 1 and α-syn-venus 2) acquired from the lab of Dr. Pamela

Mclean (Mayo Clinic, Jacksonville Florida). In order to facilitate intercellular transfer from

donor to recipient population, cells were seeded at a 6:1 ratio of donor to recipient cells

respectively. Following co-seeding, the two distinct populations were co-incubated from 24 and

48 hrs. At the end of each incubation, the coverslips containing the two cells populations were

collected, fixed with PFA (see above), stained with DAPI, and the imaged on a Leica TCS SP8

non-resonant confocal microscope.

3.15. Cyto ID Staining

Cyto ID (Enzo scientific, New York, United States) is a long-term cell tracer kit, which

incorporates a red fluorescent dye into the cell membrane’s lipid bilayer. This dye is non-toxic,

can be used in living cells, and can be sustained for 96 hrs. For the purposes of this experiment

Cyto ID was utilized to differentiate the donor population from the recipient population

following co-seeding.

52

Figure 8. Co-seeding outline

Day 1: donor cells, which contain naïve H4 cells, were transfected with either α-syn or PCDNA

3 constructs. Day 2: these donor cells are then treated with 10 μg/ml of exogenous protein

(E35K, E57K or IAPP). Concurrently, a second distinct population of naïve H4 cells (recipient

cells) were co-transfected with α-syn-venus 1 and α-syn-venus 2 constructs. Day 3: donor cells

were stained with Cyto ID, and then co-cultured with recipient cells a ratio of 6:1, donor to

recipient respectively, on glass coverslips. Day 4: collect 24 hr time point. Day 5: collect 48 hr

time point.

53

Approximately 2 x 106 cells from donor population were collected in a 15 ml conical tube and

spun down at 400 G on a table-top centrifuge (Thermo Scientific, Rockford, Illinois) into a

pellet. The pellet was subsequently washed with a 1X Hanks buffered saline solution (HBSS)

(Enzo scientific, New York, United States). Following the wash step, the cells were once again

pelleted using a 400 G spin and treated with 1ml of 2X labeling buffer solution (Enzo scientific,

New York, United States) for 10 mins. After 10 mins the labeling buffer solution containing the

cells were added to a 2X Cyto ID Red Tracer Dye solution (prepared by adding 2 μl of Cyto ID

Red Tracer Dye to 1ml of 1X labeling buffer). The combined solution was mixed thoroughly

and incubated for 3 mins. The staining process was stopped by adding an equal volume (2 ml) of

stop buffer solution (prepared by adding 2000 μl of FBS to 9.8 ml of 1X HBSS). After stopping

the reaction, solution was spun down at 400 G and the pellet was subsequently washed 3 times

with 10 ml of DMEM with 10% FBS. Once thoroughly washed, stained donor cells were

counted using a hemocytometer and approximately 60 x 104 cells were seeded with recipient

cells.

3.16. Quantification of Punctate Structure Formation

Following co-culture, the formation of punctate structures within the recipient cell population

was quantified and compared between the different control and experimental conditions. For the

quantification process 10 images were taken at random per condition per experiment. Following

image acquisition, the participant was blinded and asked to record the percentage of cells that

possessed green punctate structures relative to the total number of green cells. An average

percentage was obtained from the ten pictures per condition for each experiment. These values

were then compared using a one-way ANOVA for any statistically significant differences

between the conditions.

3.17. Normalization and Statistical Analysis

All immunoblots were quantified using the Bio-Rad image lab 5.1 software. Each immunoblot

was initially normalized to its respective loading control (either actin or tubulin) in order to

account for any anomalies resulting from loading differences. These values were then

54

subsequently normalized to the PBS control for each immunoblot in order to get relative values

that could be compared between experiments. However, in order to normalize the different

values to the PBS control, this condition was artificially set as a 100% and in doing so this

resulted in a standard deviation of zero and no error bars. Due to this alteration, this condition

could not be used in any comparative statistical analysis because values artificially created to

possess zero variability might result in false positive outcomes. Therefore, for all 1-way

ANOVA comparisons the normalized IAPP condition served as the control.

A 1-way ANOVA with a Tukey’s HSD post-hoc test was utilized for the immunoblot and cell

death assay analyses (all statistically analyses were performed on SPSS). All data sets were

assessed to ensure that the requirements of the 1-way ANOVA were met. Normality of the data

was determined using the Shapiro-Wilk test for normality, lack of outliers was verified using

boxplots, and homogeneity of variance was confirmed using the Levene’s test. Only the results

from the he luminescence assay failed to exhibit homogeneity of variance, and thus could not be

interpreted using a standard 1-way ANOVA. Therefore, these results were analysed using the

modified 1-way welch ANOVA, which is designed to take into account lack of homogeneity of

variance.

55

Chapter 4

Results

4.1. Construct Expression

When studying aggregation, being able to monitor and measure a change in the solubility of the

protein of interest is essential. Therefore, the α-syn protein fragment complementation assay

(PCA) utilized in the study by Outeiro et al. 2008 was ideal for these experiments because it

allows α-syn interaction to be directly quantified (see section 1.4.6). In order to ensure the

consistency of this assay, ubiquitous cell expression of the PCA constructs was achieved by

generating H4 cells that stably expressed the fusion proteins. Within these cells the α-syn-luc 1

and α-syn-luc 2 constructs are approximately 35 and 32 kD in size respectively, and surprisingly

α-syn-luc 1 is expressed more highly than α-syn-luc 2 even though both constructs are driven by

identical promoters (Figure 9b). This result was consistent regardless of whether the constructs

were expressed individually or together within H4 cells and similar finding were also observed

using a different antibody (Figure 13a).

In terms of functionality, cells expressing only one of the two constructs exhibited background

levels of luminescence nearly identical to those that contained an empty vector. The recorded

signal ranged between 2.07 to 2.35 x 103 relative luminescence units (RLU) (Figure 9c).

Furthermore, the level of false positive signal due to nonspecific interactions between the two

luciferase halves, determined using the cells that contained linker-luc 1 and α-syn-luc 2, was

two-fold greater than cells containing only a single fusion construct or the empty vector (p <

.05, one way ANOVA with Tukey’s HSD post-test, n=3) (Figure 9c). Nonetheless, the greatest

activity was observed only in cells that contained both full-length constructs (α-syn-luc 1 and α-

syn-luc 2). This subgroup exhibited luminescence activity that was 7-fold greater than the empty

vector control and 3-fold greater than the linker-luciferase control (p < .05, one way ANOVA

with Tukey’s HSD post-test, n=3). Thus, our cells stably expressed both constructs, which were

only functionally active when the full-length α-syn-luc1 and 2 were interacting.

56

Figure 9. Expression and functionality of endogenous α-syn luciferase constructs

a) Schematic of the α-syn luciferase fusion constructs combinations stably expressed by

different H4 cell populations. b) Immunoblot of lysate from H4 cells stably expressing different

combinations of the α-syn luciferase fusion constructs. α-syn-luc1 and α-syn-luc 2 were

approximately 35 and 32 kD respectively, with α-syn-luc1 exhibiting higher expression. Blots

were probed with a polyclonal gaussia antibody. c) Relative luminesce units (RLU) produced

due to α-syn interaction in cells stably expressing different combinations of α-syn luc 1 and α-

syn luc 2. Cells with both full length constructs exhibited significantly greater luminescence

than the control conditions. Data expressed as mean± SEM. 1 way ANOVA, Tukey HSD post

hoc test, n=3.

57

4.2. Characterization of α-Syn Species

The treatments utilized in the study consisted of two distinct populations of α-syn species. The

first were oligomers, which were generated based on the design used in the study by Winner et

al. 2011 (see section 1.4.6). Point mutations (E35K or E57K) were inserted into the α-syn

sequence, which should inhibit β-sheet formation and thereby help maintain α-syn in an

oligomeric state. In addition to these oligomers, larger fibrils were generated from either

recombinant WT α-syn or recombinant α-syn that had the PD mutations (A30P or A53T). Some

of the fibrils were even generated from phosphorylated versions of α-syn by co-expressing

either WT or mutant α-syn with polo-like kinase 2 (PLK2) within E.coli. This enzyme

specifically phosphorylates the Ser129 residue, a modification that has been shown to promote

the fibrillization process. Previous HPLC experiments demonstrated that phosphorylated and

unphosphorylated versions of the protein possess different elution profiles. We confirmed that

there was very little indication of a nonphosphorylated profile within the phosphorylated

synuclein eluant.

Transmission Electron Microscopy (TEM) studies were conducted on the generated fibrils and

oligomers in order to assess the structural characteristics of the different populations. The fibrils

possess long strand-like characteristics, and those made from the α-syn mutants in particular had

denser fibrillar structures (Figure 10). Phosphorylation of the serine 129 residue appeared to

promote fibrillization especially for the A30P and WT α-syn fibrils, which contained more

strand-like structures relative to the non-phosphorylated versions (Figure 10). The E35K and

E57K populations, on the other hand, exhibited a heterogeneous composition of small punctate

circular conformations, similar to the ones reported by Winner et al. 2011, as well as short

fibrillar-like strands.

58

Figure 10. Characterization of α-syn fibrils and oligomers

Transmission Electron Microscopy was used to visualize the structure of the α-syn species used

for the different treatments in this study. The fibrils made from either WT or the mutant

isoforms of α-syn possess long strand-like structures, the mutants in particular exhibited greater

fibrillization. Following phosphorylation, WT and A30P exhibited greater formation of these

thread-like structures. Both the E35K and E57K populations possess a heterogeneous

composition of circular structures as well as short fibrillar strands.

100nm 100nm 100nm

100nm100nm100nm

100nm 100nm

WT A30P A53TN

on

-ph

osp

horyla

ted

P

ho

sph

ory

late

d

Ser

129

E35K E57K

59

4.3. Dose Response with Wild-type α-Syn Fibrils

In order to determine the appropriate concentration that would induce aggregation, α-syn luc 1

and α-syn luc 2 stable cells were initially treated with 1, 3 or 10 μg/ml of wild-type α-syn fibrils

for 24 hrs. The logarithmic dose response was based on concentrations used by other studies

within the literature (Luk et al., 2009b; Volpicelli-Daley et al., 2011). Lysates from treated cells

were separated into three fractions (soluble, membrane, and insoluble) and α-syn redistribution

within these fractions was assessed relative to the vehicle (PBS) treated counterparts (Figure

11). The 1 μg /ml treatment condition did not produce any striking differences in the level of α-

syn-luciferase redistribution relative to the PBS control. An increase in monomeric α-syn was

noted in the total lysate and insoluble fractions for the treated cells; however, this is likely the

breakdown of the exogenously added fibrils under the denaturing condition of the SDS gel. The

3 μg /ml treatment yielded more appreciable differences mainly within the insoluble fraction,

which normally contains aggregated proteins. A faint band was observed around 26-37 kD, the

range for the luciferase constructs. This difference became even more prominent when the fibril

concentration was increased to 10 μg /ml. A large smear within the 26-37 kD range along with

an increased amount of monomeric α-syn and smaller degradation products was observed within

the insoluble fraction, while the control condition remained blank. However, it should be noted

that using the Syn-1 antibody it is not possible to differentiate between α-syn that belongs to the

exogenous treatment versus those generated from the endogenous fusion proteins. Thus, the

smear pattern within the insoluble fraction could possibly belong to exogenous aggregate

fragments that coincidentally migrate between 26-37 kD. Therefore, in addition to establishing

the treatment concentration that will be used for subsequent experiments, these results also

identify an area that needs to be further investigated.

60

Figure 11. α-Syn wild-type fibril dose response

H4 cells expressing the luciferase constructs were treated with increasing concentrations of

wild-type fibrils (fib) (1, 3, 10 μg/ml) in order to determine the concentration range that would

induce the greatest aggregation. Cell lysate from each treatment was sorted into varying

fractions (total lysate, soluble, membrane, and insoluble) and blots were probed with the Syn-1

antibody. The 10 μg/ml treatment condition induced the greatest aggregation as indicated by the

large smear between 26 - 37 kD, which could potentially be the α-syn-luciferase fusion proteins,

while the PBS treated control exhibited no change. An increase in monomeric α-syn as well as

smaller degradation products was also observed, but this can likely be attributed to the

breakdown of the exogenous fibrils on the SDS gel.

μg

μg μg

61

4.4. Cell Viability

Cell death resulting from the WT fibril dose response was determined using a trypan blue

exclusion assay. Cell viability was assessed 24 hrs post-treatment by comparing the percentage

of live cells to those treated with the vehicle (PBS) (Figure 12a). For treated cells, viability

ranged between 85-90%, and even though minor decreases were observed following treatment

with increasing fibril concentrations, these differences were not significant relative to the

control condition (P >.05 one way ANOVA with Tukey’s HSD post test, n=3). Similar results

were also observed following exposure to 10 ug/ml of either the different fibrils or the E35K

and E57K treatments. After the 24 hr exposure, cell viability ranged between 84-94% and

salient differences relative to the PBS control were not observed (P >.05 one way ANOVA with

Tukey’s HSD post test, n=3) (Figure 12b).

4.5. Aggregation Induction Following Treatment with Varying α-Syn

Species

When assessing the level of aggregation induced by either the E35K and E57K treatments or the

fibrils, the subsequent experiments focused on the redistribution of endogenous α-syn-luciferase

fusion proteins, from the soluble to the insoluble fraction. Within the detergent soluble fraction,

α-syn-luc 1 and α-syn-luc 2 protein levels were fairly equivalent across the different treatments

for immunoblots probed with Syn-1 (Figure 13a). Densitometric analyses further confirmed the

lack of any significant differences between the conditions for either α-syn-luc 1 or α-syn-luc 2

(P >.05 one way ANOVA with Tukey’s HSD post test, n=3) (Figure 13b). Values were

normalized to the PBS control, but this condition was excluded for all statistical analyses (see

section 3.15). Islet amyloid polypeptide (IAPP) was used as an alternative control in place of

PBS. IAPP is a pancreatic protein that forms fibrils that possess β-sheet secondary motifs. This

control protein was used to determine if non-synuclein based fibrils would be able to recruit

endogenous monomeric α-syn into aggregated conformations.

62

Figure 12. Cell viability

a) Percentage of live cells following treatment with increasing concentrations of WT α-syn. b)

Cell viability following exposure to different exogenous aggregates. Significant differences

relative to the PBS control weren’t observed following the WT fibril dose response or exposure

to the other α-syn species. Data expressed as mean± SEM, 1 way ANOVA, Tukey HSD post

hoc test, n=3.

63

a)

b) α-syn-luc 1

α-S

yn

-lu

c 1 L

evel

s

64

Figure 13. Analysis of Triton X-100 detergent soluble fraction

a) H4 cells stably expressing the α-syn fusion constructs were subjected to a 24 hr treatment

with PBS, IAPP, α-syn fibrils or the E35K and E57K treatments. Proteins that were solubilized

by Triton X-100 were run on blots probed with the α-syn antibody Syn-1. There appeared to be

little to no difference in the protein levels following treatment with either the oligomers or the

fibrils. b) and c) Densitometric analyses conducted on the α-syn-luc 1 and α-syn-luc 2 bands

from the blots probed with Syn-1. All values were normalized to the PBS control, but this

condition was excluded from all statistical analyses. No significant difference was observed for

either α-syn-luc 1 or α-syn-luc 2. Data expressed as mean± SEM, 1 way ANOVA, Tukey HSD

post hoc test, n=3.

α-syn-luc 2 c)

α-S

yn

-lu

c 2

Lev

els

65

Within the insoluble fraction, more striking differences were observed relative to the IAPP and

PBS control treatments (Figure 14). Aside from the A53T_P condition, significant aggregation

was observed between 30-40 kD, which is the range for the luciferase fusion proteins. However,

it was also noted that naïve H4 cells that did not contain the luciferase fusion proteins also

demonstrated aggregation within this range when treated with A30P fibrils. This outcome is

likely due to fragments from the exogenous protein treatments exhibiting similar mobility as the

luciferase fusion proteins. Using the Syn-1 antibody it is not possible to discriminate α-syn

belonging to the exogenously added aggregates from endogenous α-syn that is part of the

luciferase constructs. Therefore, a more specific detection method is required in order to

accurately assess the effects of the different extracellular treatments on the endogenous α-syn-

luciferase fusion proteins.

In order to better differentiate the aggregated species within the insoluble fraction, specific

antibodies that recognize distinct segments with the N and C-terminal luciferase fragments were

generated (Figure 15a). Each antibody was not only specific, but also exhibited minimal

background and cross-reactivity for the opposite fragment (Figure 15b). When the insoluble

fraction was re-probed with these antibodies clear differences between the treatments was once

again observed, but now the identity of these aggregated species was more clear. For these blots

the E35K, E57K, A30P, A30P_P, A53T, and WT_P treatments initially appeared to induce high

levels of aggregation (Figure 16a). However, upon further verification using densitometric

analyses, only the E35K and E57K treatments induced any significant effects (Figure 16b). For

blots probed with N-terminal luciferase antibody (luciferase 1) the E35K condition was

significantly greater than the control and all the fibril treatments, while E57K was only greater

than the IAPP, WT, WT_P, and A30P_P exogenous treatments (p < .05, one way ANOVA with

Tukey’s HSD post-test, n=3). For blots probed with the C-terminal luciferase antibody

(luciferase 2) only the E35K condition was significant relative to IAPP (p < .05, one way

ANOVA with Tukey’s HSD post-test, n=3). The fractionation and immunoblot studies revealed

that the E35K and E57K treatments could induce significant aggregation.

66

Figure 14. Triton X-100 detergent insoluble fraction

H4 cells stably expressing the α-syn fusion constructs were subjected to a 24 hr treatment with

either PBS or different extracellular forms of α-syn . Proteins from the lysate that were insoluble

to Triton-X were probed with the α-syn antibody Syn-1. Even though significant aggregation

was observed across the different treatments, it was evident that Syn-1 could not differentiate de

novo luciferase fusion protein aggregation from exogenous fibrillar or oligomeric fragments that

migrated within the same range. H4 naïve cells treated with A30P exhibited aggregation in the

same range as cells expressing luciferase fusion proteins that were treated with the different α-

syn species.

67

Figure 15. Luciferase 1 and 2 antibodies

a) Peptide sequences within the N and C-terminal luciferase fragments that were recognized by

luciferase 1 and luciferase 2 respectively. b) A test with the luciferase 1 and luciferase 2

antibodies, which demonstrated specificity, low background, and minimal cross-reactivity.

a)

b)

68

a)

15

20

30

40

50

40

PBSIA

PPE35K

E57KH

4:naiv

e:+A30P

+P +P +P-P -P -P

A30P A53TWT

actin

α-syn-luc:1:

AB::luciferase:1

69

Figure 16. Analysis of Triton-X 100 insoluble fraction using luciferase 1 and 2 antibodies

a) The Triton-X 100 insoluble fraction was re-probed with antibodies against either N or C-

terminal luciferase fragments. WT_P, A30P, E35K and E57K treated cells exhibited the greatest

aggregation for blots probed with both antibodies. b) Densitometric analyses of blots probed

with the luciferase 1 and luciferase 2 antibodies. Only the E35K and E57K treatments exhibited

significantly greater aggregation relative to IAPP and the fibril treatments. Values were

normalized to the PBS control, but this condition was excluded from all statistical analyses.

Data expressed as mean± SEM. 1 way ANOVA, Tukey HSD post hoc test, n=3.

b)

70

4.6. Luminescence Assay

Luminescence resulting from α-syn interaction was another way in which this study attempted

to directly quantify and compare α-syn aggregation induced by the different treatments.

However, despite repeated trials there was some variability between experiments for certain

treatments, and significant patterns were not observed across the different conditions (Figure

17). The luminescence assay was unable to recapitulate the findings observed in the biochemical

studies. The E35K and E57K treatments did not induce any significant differences relative to

IAPP or the fibrils (P > .05 one way Welch ANOVA, n= 3). All luminescence values were

normalized to the PBS control, but this treatment condition was excluded from all statistically

analyses.

4.7. Fluorescent Imaging of E35K and E57K Treated Cells

Immunocytochemistry provided the opportunity to verify E35K and E57K induced aggregation

from a different perspective. The α-syn-luciferase expressed within H4 cells was detected using

a polyclonal gaussia antibody, while a Thioflavin S (Thio S) dye was used to detect the presence

of β-sheets. These are secondary structural conformations normally adopted by aggregated

version of α-syn. Co-compartmentalization (depicted in yellow) indicates areas where the α-syn-

luciferase fusion proteins have either formed or been incorporated into aggregates that possess

β-sheet structure. The study revealed that the cytoplasmic distribution of α-syn-luc 1 and 2 was

strongly dictated by the exogenous protein treatment (Figure 18). PBS treated cells possessed a

diffuse α-syn-luciferase signal and did not exhibit any significant β-sheet motifs, which was

denoted by a weak Thio S stain (Figure 18). Addition of IAPP, a known β-sheet forming

protein, increased the intensity of the Thio S stain. However, there was no co-

compartmentalization of Thio S positive structures and the α-syn-luciferase constructs, which

remained diffuse throughout the cytoplasm (Figure 18). Only the E35K and E57K treatments

induced significant changes. Incrementally elevating the dose of E35K not only increased β-

sheet structure formation, but also prompted the redistribution of the α-syn-luciferase constructs

to more punctate formations, which co-compartmentalized with the structures positively stained

with Thio S (indicated by the arrows).

71

Figure 17. Quantification of aggregation using luminescence

Bioluminescent signal produced due to α-syn interaction was compared following a 24 hr

exposure to varying exogenous aggregates at a concentration of 10 μg/ml. Certain conditions

exhibited high variability between experiments, and no significant difference was observed

across the treatments. Values were normalized to the PBS control, but this condition was

excluded from all statistical analyses. Data expressed as mean± SEM, 1-way Welch ANOVA,

n= 3.

72

73

Figure 18. Changes in distribution of α-syn following exposure to E35K or E57K

H4 cells were treated with PBS, 10 μg/ml of IAPP or one of the oligomers at increasing

concentrations. α-Syn-luc1 and α-syn-luc 2 (red) were labelled using a polyclonal gaussia

antibody and aggregation was detected using the thioflavin S dye (Thio S) (green). Co-

compartmentalization of α-syn-luc1 and 2 with thioflavin positive structures is indicated in

yellow. Only treatment with 10 μg/ml of either E35K or E57K yielded any appreciable changes.

The α-syn-luciferase constructs were redistributed into punctate structures (indicated by the

arrows), which co-compartmentalized with Thio S positive structures. A magnified image of one

of the cells from the 10 μg/ml treatment condition for both E35K and E57K shows that these

yellow punctate structures are distributed throughout the cytoplasm and that there is not 100%

co-compartmentalization between Thio S and the luciferase fusion proteins.

74

Closer examination of these E35K treated cells at a higher magnification also revealed the

presence of numerous Thio S positive structures that did not co-compartmentalize with the

luciferase fragments. Since these β-sheet conformations did not seem to originate from the

fusion proteins, the other likely source would be the E35K treatments. However, since

oligomers normally lack β-sheet conformations, these findings indicate that the E35K and E57K

treatments likely contain a mixture of oligomers and fibrils. Assessment of the E57K treatment

also yielded similar results.

4.8. Fluorescent Protein Fragment Complementation

An alternative protein fragment complementation assay, which utilized fluorescence, provided

the opportunity to further validate the immunocytochemical results obtained with the luciferase

constructs. For these experiments cells were transfected with different combinations of the α-

syn-venus constructs (Figure 19a). In terms of fluorescent signal, only cells co-transfected with

both α-syn-venus 1 and α-syn-venus 2 yielded any activity, which was diffuse throughout the

cell cytoplasm (Figure 19b). Since transfection was utilized in order to incorporate the α-syn-

venus constructs, ubiquitous expression of the fusion proteins was not observed. Nevertheless, a

consistent transfection efficiency of 25% was observed across repeated trials.

When cells containing both fluorescent fusion proteins were exposed to 10 ug/ml of either E35K

or E57K, a significant redistribution to more punctate structures was observed (Figure 20). This

pattern closely mirrored the results observed with the luciferase counterparts. However, with

these fluorescent constructs the redistribution was much more prominent, and both the E35K

and E57K treated cells exhibit identical phenotypes.

75

Figure 19. Fluorescent protein fragment complementation assay

a) α-Syn-venus fusion protein combinations within H4 cells. b) Confocal images depicting the

level of fluorescent activity for each of the different transfection combinations. Only cells co-

transfected with α-syn venus 1 and α-syn venus 2 exhibited any fluorescent activity.

α-syn-venus 1 α-syn-venus 2

α-syn-venus 1+

α-syn-venus 2

Linker-venus 1+

α-syn-venus 2

Venus 1 α-syn Venus 2α-syn

Venus 1

Venus 2 α-synVenus 2 α-syn

Venus 1 α-syn

Vector only α-syn-venus 1 α-syn-venus 2

α-syn-venus 1 + α-syn-venus 2

linker-venus 1 + α-syn-venus 2

a)

b)

76

Figure 20. E35K and E57K induced redistribution of α-syn venus proteins

Significant redistribution of the of the α-syn venus constructs, from diffuse to more punctate

structures, was observed in cells expressing the a-syn venus 1 and a-syn venus 2 constructs

following a 24 hr treatment with 10 μg/ml of either E35K or E57K.

77

4.9. Propagation of E35K and E57K Induced Morphological Changes

Due to the fluorescent PCA’s ability to clearly depict changes in α-syn distribution, this assay

was used in the co-culture study, which assessed whether E35K and E57K induced changes

could be propagated intercellularly. For this experiment, cells transiently transfected with the

fluorescent fusion proteins (recipient population) were co-incubated with cells transfected with

α-syn that had been previously treated with IAPP, E35K or E57K (donor population). The two

distinct populations were co-incubated in order to determine if the pathology induced within the

donor cells could be propagated to the untreated recipient cell population. Cells were co-

incubated at a 6:1 ratio of donor cells (labelled in red) to recipient cells (labelled in green)

respectively for either 24 or 48 hrs.

To account for any effects resulting primarily from the co-incubation of two distinct

populations, PBS treated naïve H4 cells were co-cultured with cells expressing the fluorescent

constructs. We discovered negligible redistribution of the α-syn-venus constructs during the 24

and 48 hr time points, indicating that co-incubation alone had minimal effect (Figure 21).

Furthermore, treatment of H4 naïve cells that possess minimal levels of endogenous α-syn with

the oligomer prone mutants, prior to co-culture, was also unable to induce changes within the

recipient population. These results exclude the possibility of carry-over of exogenous

recombinant α-syn from recipient to donor cells. We then proceeded to determine if exposure to

non-synuclein based fibrils would induce morphological changes that can be propagated to

neighbouring cell populations. Since the presence of endogenous α-syn is required for

aggregation induction and propagation, H4 cells were initially transfected with α-syn followed

by treatment with 10 μg/ml of IAPP. These cells were then co-incubated with the recipient cell

population. This treatment resulted in mild punctate structures formation after 24 hrs; however,

these inclusions did not persist at the 48 hr time point (Figure 21).

Noticeable changes only occurred after α-syn overexpressing cells were treated with 10 μg/ml of

either E35K or E57K. After 24 hrs punctate structures were distributed around the nucleus and

throughout the cytoplasm (indicated by white arrows) for both treatments (Figure 21). This

phenotype became significantly more prominent after the 48 hr co-incubation period where

large distinct punctate structures were observed (indicated by white arrows) especially for the

78

E35K treatment, which appeared to possess slightly larger aggregates (Figure 21). Also, the

diffuse cytoplasmic distribution noted at 24 hrs was almost nonexistent at the 48 hr time point,

indicating significant recruitment of the α-syn-venus fusion proteins. For the E35K and E57K

conditions after 24 hrs, 36 and 35% of the recipient population displayed this phenotype

respectively (Figure 22). This number increased to 55% for E35K and 58% for E57K after the

48 hr co-incubation period (Figure 22). Furthermore, it was confirmed that this phenomenon

was not a consequence of cell death because relative to the controls, the experimental conditions

did not exhibit any significant changes in viability during the 24 or 48 hr co-incubation period

(P > .05 one way ANOVA with Tukey HSD post test, n= 3) (Figure 23).

79

80

81

Figure 21. Propagation of E35K and E57K induced morphological changes

Two distinct populations of cells were co-incubated in order to determine if E35K or E57K

induced phenotype can be propagated intercellularly. Donor cells stained with the dye Cyto ID

(in red) were either untrasnfected or transfected with PCDNA or wild-type α-syn. These cells

were then treated with either a vehicle (PBS) or 10 μg/ml of an exogenous aggregant (IAPP,

E35K or E57K) prior to being co-incubated with the recipient cells (labelled in green), which

contain cells co-transfected with α-syn-venus 1 and α-syn-venus 2. Co-incubation periods were

either 24 or 48 hrs. Punctate structure formation was only observed when cell containing the α-

syn-venus constructs were co-incubated with cell that had been transfected with α-syn and

treated with either E35K or E57K. The phenotype worsened as the co-incubation time was

extended. The diffuse cytoplasmic distribution of α-syn-venus 1 and α-syn-venus 2 was almost

nonexistent at the 48 hr time point.

82

Figure 22. Percentage of recipient cells with punctate morphology

a) After 24 hrs, recipients cells co-incubated with E35K or E57K treated donor cells exhibited

significant punctate morphology (36% ± 2%) and (35% ± 4%) respectively compared to the

PCDNA (13%± 4%) ,IAPP (19%± 4%) , E35K(16%± 5%), and E57K(20%± 5%) controls. b)

At 48 hrs, the E35K and E57K conditions possessed (55% ± 5%) and (58% ± 5%) punctate

morphology in recipient cells respectively, while the empty vector (12%± 2%), IAPP (15%±

3%) , E35K(21%± 5%), and E57K(23%± 4%) controls remained relatively constant. Data

expressed as mean± SEM. 1 way ANOVA, Tukey HSD post hoc test, n=3.

83

Figure 23. Cell viability following co-seeding

Cell viability was assessed following co-seeding of α-syn-venus construct expressing cells and

donor population . No significant difference was observed between the conditions. Data

expressed as mean± SEM, 1 way ANOVA, Tukey HSD post hoc test, n=3.

84

Chapter 5

Discussion

5.1. Study Aims

Our study focused on creating an aggregation model using oligomers that faithfully

recapitulated the stages involved in the prion-like propagation of α-syn pathology. This involved

not only assessing how effectively α-syn oligomers induce aggregation relative to fibrils, but

also ascertaining if oligomer-induced intracellular changes can be propagated to neighbouring

cells. This section will examine and interpret both the positive and negative results in addition to

discussing how these findings align with the current body of knowledge.

5.2. Fibril Aggregation

5.2.1. Soluble Fraction Analysis

Fractionation was used to evaluate the redistribution of the α-syn-luciferase fusion proteins from

the detergent soluble to the detergent insoluble fraction, a customary transition when proteins

aggregate. For our study, analysis of the detergent soluble fraction revealed negligible

differences following treatment with either the fibrils or oligomers relative to the control

condition (Figure 13). This lack of change was initially surprising given that protein

accumulation was observed within the insoluble fraction (Figure 16). Customarily, aggregation

of the α-syn-luciferase proteins should be accompanied by a subsequent decrease within the

respective soluble fraction because these fusion proteins are being removed from the soluble

pool. Interestingly, we discovered that other in vitro aggregation studies also reported similar

findings (Luk et al., 2009a; Sacino et al., 2013). In these studies little to no change occurred

within the soluble fraction following a 24 hr treatment with either WT or mutant α-syn fibrils,

even though significant protein accumulation was observed within insoluble fraction. Only fibril

incubation periods that ranged from 5 days to 2 weeks managed to induce any noticeable

reductions within the detergent soluble fraction (Volpicelli-Daley et al., 2011).

85

The lack of change within the soluble fraction for our study and those done by others could be

attributed to the shorter incubation time (Luk et al., 2009a; Sacino et al., 2013). Since α-syn is a

soluble protein, a large quantity is normally present within the soluble fraction. A 24 hr

exposure to either fibrils or oligomers might only recruit a small portion of the soluble α-syn-

luciferase proteins. This relative change is very small compared to the overall amount that

remains within the soluble fraction, and thus the redistribution is not as noticeable (Figure 13).

Unfortunately, for our study, extended incubation periods could not be used because prolonged

exposure to certain α-syn species such as the E35K and E57K isoforms caused a drastic increase

in cell death. A 24 hr incubation period was the most optimal time course when using the lysine

mutants. Consequently, in order to perform an unbiased comparison, the incubation period for

the other extracellular treatments was also maintained at 24 hrs. It is possible that significant

recruitment of soluble α-syn species might not occur in such a short time frame.

The lack of change within the soluble fraction could also be due to the initiation of

compensatory mechanisms within the cell, which maintained a constant level of the α-syn-

luciferase protein. It has been reported previously that upregulation of α-syn was observed

following exposure to agents that promote aggregation or other changes that induce cytotoxicity

(Manning-Bog et al., 2002; Quilty et al., 2006; Vila et al., 2000). Therefore, a similar process

could have also occurred within our study in response to the effects of the extracellular

treatments. It is possible that an increased synthesis of α-syn-luciferase was triggered in order to

compensate for the loss of the fusion proteins that were recruited into aggregates. Consequently,

this increased α-syn-luciferase expression could effectively mask the redistribution of the fusion

proteins into the insoluble pool. The induction of this process could also explain the findings

observed in our study, in which α-syn aggregation was observed, but not the concurrent

decrease within the soluble fraction.

5.2.2. Insoluble Fraction Analysis

In contrast to the soluble fraction, changes within the insoluble pool were much more noticeable

(Figure 14). Since this fraction normally possesses very low protein levels, it is understandable

that even small increases caused by the transition of α-syn luc 1 or α-syn luc 2 from soluble to

insoluble, are much more apparent. Immunoblots with cell lysate from the insoluble fraction

86

were initially probed with the α-syn antibody Syn-1, which consistently detected a range of

bands increasing in size for fibril and oligomer treated cells, while the PBS and IAPP treated

population remained blank (Figure 14). At first, these bands were thought to be endogenous α-

syn that had been recruited into increasingly larger multimeric structures due to the seeding

effects of the extracellular treatments. However, upon further investigation this conclusion

seemed very unlikely because even naïve H4 cells, which possess negligible amounts of

endogenous α-syn, exhibited a similar pattern of equal intensity when treated with A30P fibrils

(Figure 14). Therefore, it is unlikely that the pattern observed following treatment with the other

α-syn aggregates is due to the recruit of endogenous α-syn.

Alternatively, these α-syn bands could be fragments that broke off from the extracellular

treatments. This breakdown could be a consequence of the sonication step that was

incorporated into the experimental protocol. For our study, a sonication protocol similar to ones

employed in other aggregation experiments, was utilized in order facilitate the uptake of large α-

syn species (Luk et al., 2009a; Volpicelli-Daley et al., 2011). This disruption process could have

created smaller insoluble by-products from the larger α-syn aggregates, which were

subsequently internalized, and these fragments could have created the banding pattern. Thus, the

findings observed within the insoluble fraction are likely a consequence of the experimental

protocol. Unfortunately, since some of these fragments also exhibited the same mobility as the

α-syn-luciferase proteins, the Syn-1 antibody could not be used to distinguish exogenous α-syn

belonging to the fibrils or oligomers from the endogenous α-syn attached to the luciferase

fragments. Consequently, this prompted the creation of more specific antibodies that would

allow us to identify the α-syn-luciferase fusion proteins.

The luciferase 1 and 2 antibodies were significantly better at discriminating the endogenous α-

syn-luciferase constructs from the extracellular treatments and possessed lower background than

the commercially available polyclonal gaussia luciferase antibody. Using these antibodies we

discovered that cells treated with PBS did not exhibit any significant aggregation. Similar results

were also observed for cells exposed to IAPP, a pancreatic protein that aggregates into insoluble

amyloid fibrils (Figure 16). We discovered that despite possessing secondary structural

conformations similar to α-syn fibrils, IAPP was unable to induce aggregation. This inability

could stem from differences in the primary sequence of the protein, which does not share any

87

homology with α-syn. This intrinsic difference could affect IAPP’s affinity for α-syn and

subsequently its ability to recruit endogenous monomers. Other studies have argued the

importance of primary sequence homology in order to effectively induce seeding (Han et al.,

1995; Jarrett & Lansbury, 1992). Therefore, our findings in addition to reports from other

studies suggests that seeding is a chemically discriminating event and that differences in

primary sequence homology can affect the efficiency of the process. This fundamental

difference could have impeded IAPP’s ability to recruit intracellular α-syn, but more

importantly these results indicate that the aggregation observed in our study is specifically

attributable to extracellular α-syn treatments.

Unlike IAPP, the WT_P, A30P, A30P_P, and A53T fibril treatments initially appeared to induce

significant aggregation (Figure 16). Cells exposed to these fibrils had relatively intense bands

within the insoluble fraction that corresponded to either α-syn-luciferase 1 or α-syn-luciferase 2.

However, densitometric analyses later revealed the absence of any significant differences,

especially relative to the control condition (Figure 16). These results were surprising because

numerous studies within the field have validated the aggregation inducing potential of α-syn

fibrils (Freundt et al., 2012; Luk, Kehm, et al., 2012a; Luk et al., 2009a; Peelaerts et al., 2015;

Sacino et al., 2013; Volpicelli-Daley et al., 2011; Waxman & Giasson, 2010). In vitro

experiments have demonstrated that different types of fibrils either composed of WT or mutant

α-syn are able to cause a significant accumulation of protein within the insoluble fraction, and

that this phenotype progressively worsens as the incubation time is extended. Furthermore,

fibrils assembled from mutant versions of α-syn such as A53T or E46K can induce aggregation

much more rapidly and extensively than their WT counterparts (Sacino et al., 2013). Thus, due

to the large body of evidence documenting the proficiency of synthetically produced fibrils in

inducing aggregation, it was surprising that our study was unable to recapitulate these results

(Freundt et al., 2012; Luk, Kehm, et al., 2012a; Luk et al., 2009a; Sacino et al., 2013; Volpicelli-

Daley et al., 2011; Waxman & Giasson, 2010).

This discrepancy could be associated with the 24 hr incubation period, which might not have

been a sufficient amount of time for the preformed α-syn fibrils to effectively induce

intracellular aggregation. Previous studies that employed extracellular fibril treatments utilized

varying exposure times that range from 24 hrs to 2 weeks (Luk et al., 2009a; Sacino et al., 2013;

88

Volpicelli-Daley et al., 2011; Waxman & Giasson, 2010). Experiments that used a 24 hr

incubation period, similar to our study, also utilized chemical reagents that assisted with the

uptake of the extracellular fibrils. One study used Bioporter, this reagent captures extracellular

proteins within a lipid formation, and then this lipid-protein complex is transported into cells

(Luk et al., 2009a). Our study did not utilize agents such as these in order to assess the seeding

potential of α-syn fibrils and oligomers under unassisted conditions, which better recapitulates

the processes that occur in vivo. Therefore, in the absence of this chemical agent the α-syn

fibrils, which are significantly large, might not be able to translocate across the plasma

membrane as quickly, and this could have hindered aggregation induction. Extended incubation

periods might offset this issue, but as mentioned previously prolonged exposure was not feasible

for our study. Consequently, this shorter incubation period could be the main reason we failed to

observe any significant results not only between the control condition and the fibrils, but also

between the different fibril treatments themselves. Previous in vitro studies have shown that

certain mutations such as A53T bestow much faster aggregation kinetics, and that these fibrils

can recruit endogenous α-syn and propagate the pathology more rapidly than their WT

counterparts (Conway et al., 1998; J. Li et al., 2001; Narhi et al., 1999; Sacino et al., 2013). The

inability of our study to recapitulate these findings could once again be attributed to the 24 hr

incubation period, which was likely an insufficient amount time for the A53T fibrils to enter

cells and exert their full effect.

5.3. E35K and E57K Induced Aggregation

Cells treated with the E35K and E57K exhibited significant aggregation not only relative to the

IAPP treatment condition, but also in comparison to the phosphorylated and unphosphorylated

WT and mutant fibrils (Figure 16). Intracellular aggregation was heavily dependent on

extracellular treatment concentration, and the inclusions that were formed also possessed β-

sheet structural conformations like the aggregated proteins found within Lewy bodies (Figure

18). Our results closely align with the growing evidence regarding the toxicity and aggregation

capabilities of α-syn oligomers. Whether it is increased cytotoxicity due to alternations in

calcium homeostasis, impairments to searching behaviour in C.elegans or reductions in the

climbing response in Drosophila, α-syn oligomers have been demonstrated to be just as, if not

more, potent than their fibrillar counterparts (Karpinar et al., 2009; Winner et al., 2011). The

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principal aim of our study was to examine the aggregation induction potential of α-syn

oligomers.

We initially hypothesized that α-syn oligomers would be significantly better than fibrils at

inducing aggregation of intracellular α-syn. The results from our study support this statement to

a certain extent. Even though the E35K and E57K treatments were able to induce significantly

greater aggregation than the fibrils, these extracellular treatments were not solely composed on

oligomers. Both the E35K and E57K solutions contained a mixture of oligomeric and fibrillar

forms of the protein. This issue was initially identified when the different populations were

analysed using EM. The E35K and E57K treatments contained structures that resembled fibrillar

conformations. The existence of fibrils within our E35K and E57K treatments was further

confirmed when we noted the presence of Thio S positive structures that did not co-

compartmentalize with the luciferase fragments. Since these β-sheet conformations did not seem

to originate intracellularly, the only other likely source would be our extracellular treatments.

Based on these initial findings, it is very likely that the E35K and E57K treatments contain a

heterogeneous mixture of oligomers and fibrils. Initial attempts at separating these different

conformations using size exclusion were unsuccessful, indicating that a majority of the species

within the E35K and E57K treatments are similar in size.

Despite this mixed population, we argue that the aggregation observed in our study was

primarily due to the oligomers. The idea that oligomeric α-syn can seed aggregation is not a

foreign concept (Danzer et al., 2007; Danzer et al., 2009; Illes-Toth et al., 2015). Studies have

demonstrated that α-syn can form heterogeneous populations of oligomers some of which can

seed aggregation. This seeding ability is similar to that of fibrils, but there is one main

difference. The fibrils, in the absence of membrane transduction agents, require incubation

periods longer than 24 hrs (Luk et al., 2009a; Sacino et al., 2013; Volpicelli-Daley et al., 2011).

Even in our study, treatment with α-syn fibrils did not cause significant aggregation within 24

hrs. Therefore, it is unlikely that the fibril population within the E35K and E57K solutions

facilitated the aggregation that was observed. However, unlike fibrils, oligomers-induced

aggregation has been reported to occur within a 24 hr period (Danzer et al., 2007). Even though

we utilized a different form of oligomer stabilization than the study by Danzer et al. 2007, we

observed comparable seeding effects within the same timeframe in our study. Aggregation was

initially assessed biochemically and then further verified using immunocytochemistry. We

90

observed that the E35K and E57K treatments induced noticeable changes within the cytoplasmic

distribution of α-syn similarly to the oligomers used in the study by Danzer et al. 2007. Thus,

we argue that despite the heterogeneous nature of our E35K and E57K treatments, it is the

oligomers that are responsible for the intracellular changes that were observed in our study.

The underlying mechanisms that allow oligomers to facilitate aggregation more rapidly is still

unclear, but we speculate that it could be related to the selective uptake of oligomers relative to

the fibrils. Differences in accessibility to the cytoplasm could significantly influence the speed

of intracellular aggregation. Previously it has been shown that uptake of particles into

mammalian cells is size and shape dependent (Chithrani et al., 2006). Consequently, the smaller

structure of these oligomers might facilitate cellular entry. For example, it has already been

shown that oligomeric Aβ1-42 can be internalized much more readily than fibrillar isoforms via

an endocytic mechanism (Chafekar et al., 2008). Therefore, it is reasonable to assume that a

similar process might also occur with α-syn oligomers. In addition, previous experiments have

also alluded to the presence of α-syn membrane receptors, which these oligomers might utilize

to enter cells (Lee et al., 2008b). However, further research examining the uptake of α-syn

oligomers relative to fibrils is needed in order to verify these ideas. Our study has established

that the underlying process is possible, and the next step involves identify the cellular

mechanism involved.

It should also be noted that despite the significant aggregation observed following treatment

with either E35K or E57K, some modest variability was observed especially with respect to the

luciferase 1 and 2 antibodies. These antibodies exhibited variances in the level of aggregation

that was detected. Densitometric analysis of blots probed with luciferase 1 revealed that both

E35K and E57K treatments induced significant aggregation relative to a majority of the other

conditions, while blots probed with luciferase 2 only exhibited a single statistically significant

outcome. It is possible that the luciferase fragments could have affected the half-life of the

fusion proteins. α-Syn attached to the N-terminal luciferase fragment (luc 1) could possess

greater stability and resistance to degradation. A similar phenomenon has been observed with

the α-syn-GFP fusion proteins where one fragments possessed a slower turnover rate relative to

the other (Outeiro et al., 2008). If α-syn-luc 1 possesses a longer half-life, a greater portion of

this particular fragment would remain within the cytoplasm and is more likely to aggregate. This

could potentially explain the discrepancy observed within the insoluble fraction. Alternatively,

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these discrepancies between the immunoblots could be a detection issue, which is associated

with the ability of the luciferase 1 and 2 antibodies to recognize their specific epitopes. The

epitopes for the luciferase 2 antibody could have become masked or denatured during

processing of the insoluble fraction. For our study, the insoluble fraction was subjected to 8 M

urea and sonication in order to promote solubilization, and this process could have denatured or

negatively affected the luciferase 2 antibody binding regions. Consequently, a lower number of

viable epitopes could affect detection efficiency resulting in the discrepancies that were

observed. It is also possible that the antibodies themselves possess intrinsic differences in either

structure or conformation, which could affect their ability to bind to their region of interest.

However, regardless of the modest variability observed using these antibodies, it is still clear

that both the E35K and E57K treatments can induce significantly greater aggregation than the

other treatments.

5.4. Luciferase PCA

In contrast to the biochemical and fluorescent studies, quantification of intracellular aggregation

using luminescence yielded some counterintuitive results. Using the luciferase PCA salient

differences were not observed relative to the control condition following treatment with the

different α-syn species (Figure 17). Since positive results had already been observed using

fractionation and immunoblot analysis, these findings were unexpected. The luciferase PCA was

employed in our study due to its sensitivity and ability to detect protein interactions. This assay

has been used in numerous different in vivo and in vitro systems to study processes such as the

twin arginine translocation pathway in bacteria, the activation of G-protein-coupled receptors,

and the interaction of mitogen-activated protein kinases (Michnick et al., 2006; Zhao et al.,

2008). More recently, it has been used to detect α-syn within exosomes, the extracellular media,

and even to monitor intercellular propagation of α-syn (Danzer et al., 2012; Danzer et al., 2011;

Outeiro et al., 2008; Tetzlaff et al., 2008). Thus, it was unusual that this well characterized assay

did not behave optimally. Since much of our data challenges the results from the PCA, it is very

likely that this assay was unable to accurately assess the changes induced by aggregation.

Therefore, the following section will examine potential factors that could have compromised the

functionality of this assay. A deficiency in construct expression could significantly interfere

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Figure 24. Conformational Specificity required for the Luciferase PCA

In order to obtain the maximum bioluminescent signal, the luciferase fragments must be brought

together in the appropriate conformation (a). Interactions that do not satisfy this specific

conformational requirement (b or c) will result in little to no bioluminescent signal. Since

aggregation is not a process that selectively facilitates specific interactions, this might explain

why the assay was unable to accurately detect inclusion formation.

a) b)

c)

α-syn luc 1

α-syn luc 2

α-syn

α-syn

luc 1

luc 2

α-syn

α-syn

luc 1

luc 2

93

with performance of the PCA. When the α-syn-luciferase stable cells were originally assessed,

unequal levels of the fusion proteins were observed. α-Syn-luc 2 appeared to exhibit reduced

expression relative to α-syn-luc 1. Since the PCA will only produce a signal when the

complementary luciferase halves are brought together, reduced expression of one of the

fragments could seriously affect the functionality of the assay. This disparity is unlikely to stem

from issues with translation or protein processing because previous experiments that have

employed the same luciferase PCA, within an identical cell line, did not report any variances in

the expression of the complementary fusion proteins (Danzer et al., 2012; Danzer et al., 2011;

Tetzlaff et al., 2008). In addition, for our study extra measures were taken to prevent these

issues. Both fusion proteins were housed within the same vector and each one was provided

with its own promoter and polyadenylation sequence. This setup not only ensured that a cell that

contained the vector possessed both α-syn-luciferase halves, but also that each fragment had a

very good chance of being expressed. We also observed that individual expression of the fusion

proteins, within the same vector, resulted in a similar expression pattern, and this further

substantiates the notion that the variance between α-syn-luc 1 and α-syn-luc 2 is not an

expression issue. Consequently, the disparity in the level of α-syn-luc 1 and α-syn-luc 2 could

be due to differences in turnover of the α-syn-luc 2 fusion protein relative to α-syn-luc 1. The

presence of the N-terminal luciferase fragment (luc 1) might afford greater stability and even

prevent the α-syn-luc 1 fusion protein from experiencing the same level of degradation, and

could possibly explain the variance. This hypothesis could be tested using a cycloheximide

chase assay. Following the addition of cycloheximide, the levels of either α-syn-luc 1 or α-syn-

luc 2 can be assessed over a given time frame in order to determine if incorporation of luciferase

fragments affects the half-life of the protein.

An alternative explanation is that the assay could not function properly due to irregularities in

fusion protein interaction. One of the shortcomings of PCA is the conformational specificity that

is required. In order to obtain a bioluminescent signal, the α-syn attached to the luciferase

fragments needs to interact in a manner that brings the luc 1 (aa 1-193) and luc 2 (aa 94-185)

fragments together properly in order to reconstitute the enzyme (Remy & Michnick, 2006)

(Figure 24). If this precise interaction is not facilitated, the signal will either be significantly

reduced or not emitted. For example, if the α-syn attached to the luciferase fragments were to

interact in an antiparallel fashion or if there was only partial α-syn-α-syn interaction, this would

94

hinder the alignment of the luciferase fragments (Figure 24). It is very unlikely that during

aggregation the soluble fusion proteins would always be recruited in a manner that would

favour reconstitution of the luciferase enzyme, and therefore using this assay as a quantitative

measure of aggregation becomes increasingly difficult. In our case, it is very likely that a

majority of the fusion proteins recruited into aggregates are trapped in unsuitable conformations

for bioluminescence. Consequently, much of the signal that is produced would come from the α-

syn-luciferase proteins that remain soluble. However, if the soluble protein levels remain

constant due to increased synthesis of α-syn-luciferase 1 and 2, changes induced by the

extracellular α-syn species would be really hard to gauge. Since the luciferase PCA was

originally designed to measure subtle changes in protein self-assembly, it might not perform

optimally under condition in which large scale aggregation is induced.

In addition to this explanation, another likely possibility is that treatment with extracellular

oligomers or fibrils does not affect total luminescence, only the intracellular distribution of the

preassembled fusion proteins that are emitting the signal. This hypothesis assumes that the

extracellular treatments will not significantly impact total bioluminescent activity. This could

explain the nearly identical luminescence signal observed between the control and experimental

conditions for our experiments (Figure 17). However, even though the extracellular α-syn fibrils

and oligomers might not significantly affect total luminescence, they can still cause the

recruitment of the preassembled fusion proteins into larger inclusions. These proteins will

continue to produce a luminescent signal, but now they will do so while incorporated into larger

aggregates. This change induced by the extracellular treatments will not affect total

luminescence, only the cytoplasmic distribution of the preassembled proteins that produce the

signal. Since the luciferase PCA was designed to quantify total luminescence, it will not be able

to accurately gauge these changes because total luminescent activity remains constant even

though the fusion protein subcellular localization has changed.

This hypothesis is further validated by results from the fluorescent PCA, which behaves

similarly to the luciferase PCA. A preliminary quantification of the fluorescence emitted by

cells expressing α-syn-venus 1 and α-syn-venus 2, following treatment with either E35K or

E57K, revealed negligible difference relative to the PBS control (Figure 25). However, when the

effects of the treatments were assessed using confocal microscopy, significant differences were

noted. The cytoplasmic distribution of the α-syn-venus constructs changed from diffuse to

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punctate following treatment with either E35K or E57K (Figure 20). This discrepancy between

the quantitative and qualitative measures further indicates that the emitted signal does not

accurately portray the changes that have occurred (Figure 25). Since the quantitative component

of the assay was designed to measure total fluorescence, it would not be able to detect changes

in the distribution of the preassembled fusion protein, which can be easily observed visually.

Thus, due to the evidence provided by the fluorescent PCA, we are more inclined to support this

hypothesis. The inability of the PCA to accurately report aggregation is most likely related to

the design of the assay, which only allows assessment of total luminescence or fluorescence,

processes that might not be affected by the extracellular treatments.

5.5. Intercellular Propagation

Intercellular propagation is a fundamental component of the prion hypothesis, and this section

examines the model created in our study, which depicts the propagation of E35K and E57K

induced morphological changes. The spread of pathology due to the transfer of toxic species is

now a basic component of disease progression for many different neurodegenerative disorders.

For PD, α-syn’s migratory capabilities garnered a lot of attention following the clinical studies

conducted by Braak et al. (2003), Li et al. (2008), and Kordower et al. (2008). These authors

surmised that PD pathology could be spread from one cell to another due to the propagation of

toxic isoforms of α-syn. Their suspicions were later confirmed by different in vitro and in vivo

models, which not only demonstrated that α-syn from donor cells can enter and colocalize with

α-syn from the recipient population, but also that this process occurs independent of cell-to-cell

contact (Desplats et al., 2009; Freundt et al., 2012; Hansen et al., 2011; Luk et al., 2009a;

Masuda-Suzukake et al., 2013). Conditioned media containing secreted forms of α-syn is a

sufficient conduit that can transfer the protein into recipient cells. However, despite the wealth

of information gained from these studies, many of them did not take into consideration the role

of oligomeric species in pathology induction and propagation. Since oligomeric versions of α-

syn are now considered to be a more toxic isoform, it is important to establish their role within

this process.

96

0

1

2

3

4

5

6

RF

U x

10^

5

max

Total fluorescence

Total fluorescence

oligomers

Pre-treatment

Post-treatment

punctate structures

Figure 25. Reorganization of the fluorescent PCA fragments

a)Schematic outlines the potential effects of the E35K and E57K treatments on the intracellular

distribution of α-syn-venus fusion proteins. Due to high baseline activity during pre-treatment

conditions, the extracellular α-syn treatments do not significantly affect total fluorescence,

which is already close to the maximum level. These treatments primarily cause the redistribution

of preassembled fusion proteins. These fused proteins will continue to produce a fluorescent

signal, but now they will do so while incorporated into larger inclusions. Total fluorescence

remains constant, while cytoplasmic α-syn-venus distribution has shifted. b) Preliminary

fluorescent quantification data indicated negligible differences between the control (VS1 +VS2)

and the experimental [(VS1 +VS2) +E35K or (VS1 +VS2) +E57K] conditions.

a)

b)

VS1= α-syn-venus 1

VS2= α-syn-venus 2

97

Therefore, our study used the E35K and E57K treatments to promote α-syn pathology

propagation from donor to recipient cells. In our model, cells previously exposed to E35K,

E57K or IAPP fibrils (donor cells) were co-incubated, for either 24 or 48 hrs, with untreated

cells in order to determine if changes within the donor population can be transmitted to the

untreated (recipient) population. Donor cells were labelled with the dye Cyto ID, while recipient

cells contained the fluorescent α-syn-venus 1 and α-syn-venus 2 constructs. Changes within the

distribution of the α-syn-venus constructs were used to determine if propagation had occurred.

The results from our propagation study demonstrate that the changes induced by either E35K or

E57K can be potentially transferred to neighbouring cells. With our model, punctate structures

only appeared within recipient cell populations that had been co-incubated with donor cells

exposed to either E35K or E57K (Figure 21). Furthermore, a significant phenotype was not

observed when recipient cells were co-incubated with donor cells treated with IAPP or cells that

were treated with the oligomer prone mutants but were devoid of endogenous α-syn. As

mentioned previously, the propagation of pathological conformations of α-syn is an area of

growing interest. Numerous studies have reported on the release, spread, and uptake of

aggregated forms of α-syn (Chang et al., 2013; Danzer et al., 2012; Emmanouilidou et al., 2010;

Jang et al., 2010; Lee et al., 2005; Lee et al., 2008b; Sung et al., 2001). These models have

monitored a variety of α-syn species in terms of propagation as well as aggregation induction.

Within our model we observed that approximately 35-36% of recipient cells displayed punctate

structures when incubated with either E35K or E57K treated donor cells that possessed

significant endogenous α-syn. This number rose to 55% for E35K and 58% for E57K treated

donor cells when the co-incubation period was extended from 24 to 48 hrs. These changes were

not only significant relative to the control conditions, but the increase in the percentage of

affected recipient cells indicates a potential spreading process.

In other in vitro propagation models, between 4 - 47% of the recipient population exhibited

changes following co-culture (Desplats et al., 2009; Freundt et al., 2012; Hansen et al., 2011;

Reyes et al., 2015). In certain studies, such as the one by Desplats et al. 2009, the level of α-syn

within the donor and recipient cell population was significantly overexpressed. Intake of

extracellular α-syn by recipient cells that already express above average levels of the protein

could overwhelm regulatory organelles such as the lysosome. Impediments in clearance and

98

degradation of these excess proteins could additionally facilitate aggregation, which might

explain the rapid changes observed in this study. Despite observing extensive pathology within

the recipient population, the authors also reported that extended incubation periods triggered

significant cell death. For our study, lower levels of expression, which might have hindered

pathology propagation, were employed in order to minimize cell death and establish an

experimental paradigm that can be applied to extended incubation periods. Our model provides

the opportunity to study the early stages of α-syn propagation prior to the commencement of

significant toxicity. This is an important stage in pathology development, and understanding this

phase will be critical when creating therapeutics that either impede or prevent the pathology

from becoming too aggressive.

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Chapter 6

Conclusion

Our study demonstrated that treatment with the oligomer prone mutants, either E35K or E57K,

induced significant intracellular aggregation, relative to the control and fibril treatments.

Following exposure to these lysine mutants the intracellular distribution of the fusion proteins

became more punctate in nature, and some of these structures possessed β-sheet structural

motifs similar to that of aggregated amyloidogenic proteins. Based on these results it appears

that the oligomer prone mutants can exert influences on the intracellular fusion proteins much

more rapidly than some of the fibril treatments possibly due to differences in accessibility to the

intracellular environment. Since oligomers are typically smaller than fibrils they might be taken

up more readily. Our study highlights another potential pathway through which oligomeric

isoforms of α-syn can influence the intracellular environment. These findings further stress the

importance of elucidating the mechanisms responsible for α-syn cellular entry. Understanding

these pathways will allow us to develop therapeutics that specifically target the mechanisms

responsible for the uptake of these pathogenic isoforms.

The results from our study also suggest that the changes induced by these oligomer prone

mutants can be propagated to neighbouring untreated cells. Using our in vitro co-culture model

we observed the formation of punctate structures within untreated (recipient) cells following co-

incubation with the treated (donor) population. However, this phenotype was only observed

when these recipient cells were co-incubated with a donor population that not only possessed

significant levels of endogenous α-syn, but had also been exposed to the oligomer prone

mutants. The results from this model nicely recapitulate the findings from previous studies,

which have demonstrated that in addition to the pathological instigator, endogenous α-syn also

needs to be present in order to facilitate changes within the untreated population. Furthermore,

the percentage of affected recipient cells increased as the co-incubation time was extended,

which further suggests a prion-like propagation mechanism for the phenotype observed in our

co-culture study. Even though further experiments are required to verify this idea, establishing a

functional model provides the opportunity to address numerous other avenues such as release

and uptake pathways, which are an area of growing interest.

100

Also, while assessing aggregate propagation we gained further insight regarding the utility and

limitations of the luciferase PCA, one of the main assays we incorporated into our study in order

to quantify α-syn interaction. In our hands this assay did not recapitulate the results observed in

our biochemical and immunocytochemical experiments. We believe the PCA’s inability to

detect the changes induced by our extracellular treatments is due to limitations in the assay,

which only allows detection of total luminescence. Since our treatments appear to cause the

redistribution of the pre-assembled fusion proteins, a process that likely does not affect total

bioluminescent activity, the results reported by the luciferase PCA might not be an accurate

representation of the subcellular changes that have taken place. Since PCA based methods are

steadily gaining popularity, it is important to understand their limitations especially when

quantifying self-assembly of amyloidogenic proteins.

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Chapter 7

Future Directions

In order to improve this study, alternative separation techniques should be explored into order to

better isolate the oligomeric and fibrillar species for both the E35K and E57K treatments. Since

oligomers and fibrils typically vary in size and mass, ultracentrifugation could potentially be

used to separate the two populations. Previously 16,000 G spins for 5-10 mins have been shown

to be sufficient at pelleting insoluble fibrils (Conway et al., 1998). Ultracentrifugation can be

used in conjunction with Thioflavin T assays, which can be used to assess the level of β-sheet

positive structures within the pellet and supernatant. Immunoblot analysis using the A11

antibody, which specifically targets oligomers, can also be used to identify the level of

oligomeric species within the two fractions. (Winner et al., 2011). These different assessment

methods can help determine the quality of separation of the two different α-syn conformations.

Once the fibrillar and oligomeric isoforms have been isolated, the aggregation assay can be

rerun in order to determine, which species was responsible from the aggregation that was

observed in our study.

In addition to better separation techniques our co-culture model can also be improved by

incorporating stable recipient and donor cells lines into the system. Currently, the two distinct

populations are transfected with the appropriate vectors needed to induce expression of α-syn

and the α-syn-venus fusion proteins. However, transfected elements are not as conducive to

extended incubation periods, and transfection efficiency sometimes exhibits inter-experimental

variability. Therefore, creating stable cells lines will improve the replicability of our results in

addition to increasing the utility of this model. We can extend the incubation period and

examine the progressive development of punctate structures within the recipient population. Our

stable co-culture model can also be integrated with a live cell imaging system to continually

monitor changes within the recipient populations. Reagents such as the Cyto ID dye as well as

the α-syn-venus constructs are nontoxic and amenable to longer incubation periods. Rather than

acquiring an image that is representative of only a specific time point within this process,

incorporating a live cell imaging system into our model will allow uninterrupted observation of

the changes that occur.

102

Our improved oligomer induced α-syn propagation system provides the opportunity to not only

evaluate aggregation from a different perspective, but also examine in detail the different

processes involved in the migration of α-syn. Furthermore, using an in vitro system, we have the

ability to isolate and manipulate individual components in order to ascertain the mechanisms

involved within the overall process. Either through the co-culture system or by treating cells

with conditioned media from different cell populations, processes such as uptake, propagation,

and release can be further studied, and incorporating live cell imaging provides the added

benefit of viewing this process in its entirety.

One of the first processes we will examine will be α-syn release, which has been attributed to

two main pathways, either exocytosis or exosomal release (Danzer et al., 2012; Emmanouilidou

et al., 2010; Jang et al., 2010; H. J. Lee et al., 2013; H. J. Lee et al., 2005). Unfortunately, there

is still not a consensus regarding the predominant pathway utilized to transport α-syn. The co-

culture system designed in our study is a good tool to address uncertainties such as these and

others regarding α-syn propagation. Analyzing the conditioned media from the 24 and 48 hr co-

cultures can help determine the main release pathway. α-Syn released via exocytosis will

predominantly appear as free floating protein within the media, while those released via an

exosomal mechanism will be found in association with vesicle-like structures.

Ultracentrifugation can help separate these distinct fractions, which can be later analysed to

determine the relative amounts of α-syn. We have already developed and ultracentrifugation

protocol for the exosomal fraction and our preliminary attempts were successful (Figure 26). We

were able to isolate exosomes, which were identified using antibodies against flottilin, a

membrane protein often enriched within the exosomal fraction. This approach can be used in

conjunction with electron microscopy to verify the presence of exosomes. Isolation and analysis

of extracellular α-syn using ultracentrifugation and biochemical assays respectively will provide

some insight regarding the release pathway that is preferentially utilized by the protein. This

pathway can then be further studied by using chemical agents that preferentially target its

processes.

Brefeldin A or BAPTA AM are chemical agents which have been shown to significantly affect

exocytosis, the former inhibits ER to golgi protein transport by impeding formation of transport

103

flotillin

Figure 26. Exosome Isolation protocol

a) Schematic outlines the optimized ultracentrifugation protocol that ensures the highest level

of purity and the maximum yield when isolating exosomes from the culture media. b)

Immunoblot probing the different fraction from the exosome isolation protocol. Exosomes

were identified using flotillin, a membrane protein enriched within these vesicles. The

ultracentrifugation protocol was able to clearly isolate exosomes, which otherwise are only

present within either live or dead cells.

Fractions:

1- dead cells

2- cell debris

3- Supernatant

4- PBS wash

104

vesicles, while the latter sequesters intracellular calcium, which is essential for exocytosis

(Emmanouilidou et al., 2010). Treating the donor cells with either one of these reagents should

significantly impair exocytosis, and the media can be subsequently analysed to determine the

relative change in α-syn release. Furthermore, by exposing the recipient cells to the conditioned

media from the chemically treated donor population, changes in aggregation induction within

the recipient cells can also be monitored. This system provides the dual benefit of assessing

both the immediate and downstream changes. A similar approach can also be utilized for

exosomal release. Since the exosomal and autophagy pathway share common mechanisms,

treatments that affect autophagy also strongly influence exosome formation and release (Danzer

et al., 2012; Simons & Raposo, 2009). The reagent rapamcyin has been previously utilized to

inhibit exosomal release by promoting the degradation of these vesicles within the lysosome.

Once again, changes in extracellular α-syn levels as well as shifts in aggregation induction

within the recipient cell population can be assessed.

In addition to release, the co-culture model can also be used to study pathways involved with

aggregate uptake, which might be endocytic in nature. This pathway in particular is susceptible

to manipulation using the dynamin K44A mutant (Lee et al., 2008b). When transfected, this

mutant has been shown to inhibit receptor-mediated endocytosis by preventing the scission of

nascent vesicles from the membrane. Recipient cells transfected with these constructs should be

unable to undergo endocytosis. Previous studies have shown that this effectively prevents entry

of extracellular aggregants (H. J. Lee et al., 2008b). However, many of these studies utilized

synthetically formed fibrils, and it would be interesting to determine if the oligomer-induced

species released from the donor cells utilize a similar entry mechanism. In addition to the

dynamin K44A mutant, the chemical agent Dynasore, which is a GTPase inhibitor that blocks

dynamin-dependent endocytosis could also be used to arrest the uptake of extracellular

aggregates. The recipient cells can be incubated in the conditioned media from the donor cell

population to which 50-80 μM of Dynasore can be added. This approach ensures that

Dynasore’s effects will primarily affect the recipient cell population. Thus, the utility of the in

vitro propagation model is not only the flexibility to target and manipulate individual

components within the system, but also the ability to monitor the immediate and downstream

changes that result from these manipulations.

105

In addition to monitoring the changes that occur within the cells, another interesting avenue that

has yet to be fully explored is the isolation of pathological α-syn species that are propagated

intercellularly. In order to accomplish this feat, a combination of different approaches will be

utilized. Firstly the conditioned media from the donor cell population, which most likely

contains the aforementioned pathological α-syn species, will be treated with biotinylated

polyclonal antibodies that target a wide range of epitopes within α-syn. Since the exact

conformation of this release species is still unclear, a polyclonal antibody has the best chance of

binding these released α-syn isoforms. Following incubation with the antibody, the condition

media will be run through a streptavidin resin column. Due to the strong and specific interaction

between biotin and streptavidin, the antibody-antigen complex will most likely bind to the

column and become isolated from the rest of the media. The antibody bound α-syn isoforms can

later be eluted from the column using chaotropic agents such as thiocyanate (SCN−),

trifluoroacetate (CF3 COO−). These agents will help dissociate the antibody-antigen complex

without significantly denaturing or damaging the protein bound to the antibody. The eluted

solution can then be subjected to size exclusion chromatography in order to separate species of

different sizes. A portion of the different fractions can be added to separate populations of

recipient cells in order to identify the fraction that contains pathological α-syn isoform that is

able to induce punctate structure formation. The α-syn within this fraction can then be further

analysed using electron microscopy and circular dichroism to identify any significant

conformations that might be present. Using this isolation procedure not only can the

pathological isoforms of α-syn be further studied, but therapeutics can be designed that

specifically target these isoforms.

106

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