The Interplay Between Lewy Body-Like Alpha-Synuclein Aggregates Nd Protein Degradation Pathways in Cell-Based Model of Parkinson's Disease

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Publicly Accessible Penn Dissertations

2013

The Interplay Between Lewy Body-Like Alpha-Synuclein Aggregates nd Protein Degradation Pathways in Cell-Based Model of Parkinson's Disease

Selcuk Aski Tanik University of Pennsylvania, [email protected]

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Recommended Citation Tanik, Selcuk Aski, "The Interplay Between Lewy Body-Like Alpha-Synuclein Aggregates nd Protein Degradation Pathways in Cell-Based Model of Parkinson's Disease" (2013). Publicly Accessible Penn Dissertations. 705. https://repository.upenn.edu/edissertations/705

This paper is posted at ScholarlyCommons. https://repository.upenn.edu/edissertations/705 For more information, please contact [email protected]. The Interplay Between Lewy Body-Like Alpha-Synuclein Aggregates nd Protein Degradation Pathways in Cell-Based Model of Parkinson's Disease

Abstract Cytoplasmic alpha-synuclein (a-syn) aggregates, including Lewy bodies (LBs), are pathological hallmarks of a number of neurodegenerative diseases, most notably Parkinson's disease (PD). Activation of intracellular protein degradation pathways (Pdps) to eliminate these aggregates has been proposed as a therapeutic approach for PD and other synucleinopathies, but the interplay between LB-like a-syn aggregates and Pdps is not completely understood. Here, we investigate this interplay by utilizing a recently developed cellular model in which intracellular LB-like a-syn inclusions accumulate after delivery of pre-formed a-syn fibrils (Pffs) into a-syn-expressing HEK293 cells or cultured primary neurons. This thesis describes the interplay between LB-like aggregates and Pdps, as well as the examination and optimization of the cellular model for this study. We demonstrate that the efficiency of the model can be greatly improved by use of mutant a-syn expressing cells and truncated Pffs. We also show that only a minute amount of Pffs that gain access to cells, and the LB-like aggregates are primarily composed of endogenous a-syn, thereby rendering levels of insoluble endogenous a-syn an excellent read-out for our study. Utilizing the optimized model, we show that a-syn inclusions cannot be effectively degraded by Pdps, even though they colocalize with essential components of these pathways. Furthermore, once formed, a-syn aggregates persist even after reduction of soluble a-syn levels, suggesting that pathological a-syn inclusions are refractory to clearance. Importantly, while proteasome function appears unaltered, we find that a-syn aggregates impair macroautophagy by reducing autophagosome clearance, which may play a role in the increased cell death observed in aggregate-bearing cells. Our results indicate that, after accumulation LB-like of a-syn aggregates, activation of Pdps may not effectively clear pathology or enhance cell survival. However, treatments that could prevent the inhibitory effect of a-syn aggregates on autophagy may be beneficial. eW believe this work improves our understanding of the role of a-syn aggregates in synucleinopathies, and it may contribute to the efforts to develop potential therapies for these devastating diseases.

Degree Type Dissertation

Degree Name Doctor of Philosophy (PhD)

Graduate Group Biology

First Advisor Virginia M. Lee

Keywords autophagy, degradation, Lewy body, Parkinson's disease, preformed fibril, synuclein

Subject Categories Cell Biology | Neuroscience and Neurobiology

This dissertation is available at ScholarlyCommons: https://repository.upenn.edu/edissertations/705 THE INTERPLAY BETWEEN LEWY BODY-LIKE ALPHA-SYNUCLEIN AGGREGATES AND

PROTEIN DEGRADATION PATHWAYS IN A CELL-BASED MODEL OF PARKINSON’S

DISEASE

Selcuk Aski Tanik

A DISSERTATION

Biology

Presented to the Faculties of the University of Pennsylvania

in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

2013

Supervisor of Dissertation

______

Virginia M-Y Lee, John H. Ware 3rd Endowed Professor in Alzheimer's Research

Graduate Group Chairperson

______

Doris Wagner, Associate Professor in Biology

Dissertation Committee

Nancy Bonini, Professor of Biology James Shorter, Assistant Professor of

Biochemistry and Biophysics

Harry Ischiropoulos, Research Professor of Deijan Ren, Associate Professor of Biology

Pediatrics

I dedicate this dissertation to my beloved parents: particularly my dear mother who suffers from Parkinson’s disease; my dear father, who has always been an inspiration

to me; and finally, to my dear wife who exhibited remarkable patience and support

during many years this study kept us apart.

ACKNOWLEDGMENTS

I thank Drs. Virginia Lee and John Trojanowski for the privilege of completing my dissertation studies in CNDR, under their supervision. I also thank my thesis committee

chair Dr. Nancy Bonini, and committee members Drs. James Shorter, Deijan Ren, and

Harry Ischiropoulos for their support and guidance. I am very grateful to Dr. Kurt

Brunden for his help in completion of the project. I thank Dawn-Marie Riddle, and Dr.

Mecky Pohlschroedinger for their valuable support. Additionally, I would like to thank

Christine Schultheiss for her for her hard work and patience. I also thank Drs. Nesrin

Ozoren, Todd Cohen, Youngshim Lim, Laura Volpicelli-Daley and Scott Pesiridis for

their help and Dr. Kelvin Luk for his contributions to my study. Finally, I thank many dear friends of mine among past and present employees of CNDR, and my class-mates in

graduate school, for their friendship and support.

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ABSTRACT

THE INTERPLAY BETWEEN LEWY BODY-LIKE ALPHA-SYNUCLEIN

AGGREGATES AND PROTEIN DEGRADATION PATHWAYS IN A CELL-

BASED MODEL OF PARKINSON’S DISEASE

Selcuk Aski Tanik

Virginia M.-Y. Lee

Cytoplasmic alpha-synuclein (α-syn) aggregates, including Lewy bodies (LBs), are pathological hallmarks of a number of neurodegenerative diseases, most notably Parkinson‟s disease (PD).

Activation of intracellular protein degradation pathways (Pdps) to eliminate these aggregates has been proposed as a therapeutic approach for PD and other synucleinopathies, but the interplay between LB-like α-syn aggregates and Pdps is not completely understood. Here, we investigate this interplay by utilizing a recently developed cellular model in which intracellular LB-like α-syn inclusions accumulate after delivery of pre-formed α-syn fibrils (Pffs) into α-syn-expressing

HEK293 cells or cultured primary neurons. This thesis describes the interplay between LB-like aggregates and Pdps, as well as the examination and optimization of the cellular model for this study. We demonstrate that the efficiency of the model can be greatly improved by use of mutant

α-syn expressing cells and truncated Pffs. We also show that only a minute amount of Pffs that gain access to cells, and the LB-like aggregates are primarily composed of endogenous α-syn, thereby rendering levels of insoluble endogenous α-syn an excellent read-out for our study.

Utilizing the optimized model, we show that α-syn inclusions cannot be effectively degraded by

Pdps, even though they colocalize with essential components of these pathways. Furthermore, once formed, α-syn aggregates persist even after reduction of soluble α-syn levels, suggesting that pathologic α-syn inclusions are refractory to clearance. Importantly, while proteasome function appears unaltered, we find that α-syn aggregates impair macroautophagy by reducing autophagosome clearance, which may play a role in the increased cell death observed in aggregate-bearing cells. Our results indicate that, after accumulation LB-like of α-syn aggregates, activation of Pdps may not effectively clear pathology or enhance cell survival. However, treatments that could prevent the inhibitory effect of α-syn aggregates on autophagy may be beneficial. We believe this work improves our understanding of the role of α-syn aggregates in synucleinopathies, and it may contribute to the efforts to develop potential therapies for these devastating diseases.

TABLE OF CONTENTS

ACKNOWLEDGMENTS ...... III

ABSTRACT ...... IV

LIST OF ILLUSTRATIONS ...... IX

LIST OF ABBREVIATIONS ...... X

CHAPTER 1: GENERAL INTRODUCTION ...... 1

1.1 Parkinson’s disease and other synucleinopathies ...... 1

1.2 α-Synuclein ...... 4

1.3 Role of α-syn in pathogenesis ...... 8

1.4 Protein degradation pathways ...... 10

1.5 Pdps and neurodegeneration ...... 15

1.6 Pdps and α-syn degradation ...... 16

1.7 Models of synucleinopathies ...... 16

1.8 Proposed therapeutic approaches for synucleinopathies ...... 21

CHAPTER 2: OPTIMIZATION OF PFF TRANSDUCTION IN HEK293 CELLS 25

2.1 Abstract ...... 25

2.2 Introduction ...... 26

2.3 Results ...... 27 Improvement of Pff transduction efficiency by use of truncated Pffs and A53T mutant α-syn expression ...... 27 Expression of PD-linked A30P or E46K mutant α-syn does not improve aggregate formation...... 28 Transduction without reagent is feasible, but inefficient in HEK293 A53T α-syn cells ...... 31 vi

2.4 Discussion...... 33

CHAPTER 3: DEGRADATION OF EXOGENOUS PFFS IN NON-NEURONAL CELLS ...... 35

3.1 Abstract ...... 35

3.2 Introduction ...... 36

3.3 Results: ...... 37 Pffs are cleared from cells and this clearance is slowed by lysosomal inhibitors ...... 37 Pffs do not significantly colocalize with early endosomes or lysosomes ...... 37 A small proportion of added Pffs gain entry to the cells ...... 39

3.4 Discussion...... 43

3.5 Materials and Methods for Chapters 2 and 3 ...... 45 Mammalian Cell Cultures ...... 45 Recombinant α-syn proteins and generation of α-syn Pffs ...... 46 Pff Transduction ...... 46 Drug Treatments ...... 47 Plasmids and Transfections ...... 48 Immunofluorescence ...... 48 Immunoblotting ...... 49 Quantifications, Image Processing, Statistics ...... 50

CHAPTER 4: LEWY BODY-LIKE Α-SYNUCLEIN AGGREGATES RESIST DEGRADATION AND IMPAIR MACROAUTOPHAGY ...... 51

4.1 Abstract ...... 51

4.2 Introduction ...... 51

4.3 Results ...... 54 Insoluble α-syn aggregates are not degraded by autophagy and their clearance is severely impaired...... 57 Typical autophagy substrates accumulate in α-syn aggregate-bearing cells...... 63 α-Syn aggregates also interact with autophagy and proteasome components in neurons but are resistant to degradation...... 63 Autophagy function is impaired in -syn aggregate-bearing cells...... 67

vii

Autophagosome clearance is impaired in -syn aggregate-bearing cells...... 70 Lysosome function appears normal in -syn aggregate-bearing cells...... 71 Autophagy activation enhances cellular toxicity in -syn aggregate-bearing cells...... 79

4.4 Discussion...... 81

4.5 Materials and Methods ...... 85 Mammalian and Primary Neuronal Cell Cultures ...... 85 Recombinant α-syn proteins and generation of α-syn Pffs ...... 87 Pff Transduction ...... 87 Drug Treatments ...... 88 Plasmids and Transfections ...... 89 Immunofluorescence ...... 90 Immunoblotting ...... 90 Immuno Electron Microscopy ...... 91 Quantifications, Image Processing, Statistics ...... 92

4.6 Supplementary Materials and Methods...... 92 Drug Treatments, Plasmids and Transfections ...... 92 Toxicity Assays ...... 93

CHAPTER 5: GENERAL CONCLUSIONS AND DISCUSSION ...... 95

5.1 Possible mechanisms of α-syn aggregate mediated impairment of autophagy ... 95

5.2 LB-like seeded aggregates and non-fibrillar α-syn inclusions ...... 97

5.3. Insights into the therapeutic implications of our study ...... 98

5.4 Insights into the possible interplay between LBs, autophagy and mitochondrial function in neurodegeneration ...... 99

5.5 Concluding remarks ...... 104

BIBLIOGRAPHY ...... 105

viii

LIST OF ILLUSTRATIONS

Figure 1-1: Degeneration of Substantia Nigra in PD ...... 2 Figure 1-2: Simplified schematic diagram of mammalian autophagy ...... 12 Figure 1-3: Pre-formed fibril transduction model of α-syn aggregation ...... 20 Figure 2-1 Transduction efficiency is improved by mutant a-syn expression and use of truncated Pffs ...... 29 Figure 2-2: E46K and A30P mutant α-syn expression does not increase aggregate formation ...... 30 Figure 2-3: Transduction without reagent is feasible, but inefficient in HEK293 A53T α- syn cells ...... 32 Figure 3-1: Transduced Pffs are cleared from cells and this clearance is slowed by lysosomal inhibitors ...... 38 Figure 3-2: Majority of transduced Pffs are not localized into early endosomes or lysosomes ...... 40 Figure 3-3: A minute amount of Pff gain entry to cells ...... 41 Figure 3-S1: Extracellular Pffs are refractory to removal by washing or trypsinization: 44 Figure 4-1: Co-localization of protein degradation pathway components with α-syn aggregates ...... 56 Figure 4-2: Autophagy does not degrade α-syn aggregates and clearance of aggregated α- syn is severely impaired ...... 58 Figure 4-S1: Modulation of autophagy and α-syn aggregates ...... 59 Figure 4-S2: The UPS does not effectively clear α-syn aggregates...... 62 Figure 4-3 Typical autophagy substrates accumulate in α-syn aggregate-bearing cells. . 64 Figure 4-4: α-Syn aggregates interact with protein degradation pathway components but cannot be effectively degraded in neurons...... 66 Figure 4-5: Canonical autophagy substrates accumulate in neurons with α-syn accumulations...... 68 Figure 4-6: Autophagy function is reduced in α-syn aggregate bearing cells...... 69 Figure 4-7: Autophagosome clearance is impaired in α-syn aggregate-bearing cells...... 72 Figure 4-8: Lysosome morphology, but not catalytic function, is affected in Pff-td cells...... 74 Figure 4-S3: Effect of Pff-td on Lysosomes ...... 76 Figure 4-9: Autophagosomes associated with p-α-syn aggregates are impaired in maturation ...... 77 Figure 4-S4: Treatments that activate autophagy increase the toxicity of α-syn aggregates.: ...... 80 Table 1: Antibodies used in this study: ...... 94 Figure 5-1: A hypothetical model of neurodegeneration in PD ...... 103

LIST OF ABBREVIATIONS 3-MA 3-methyladenine AD Alzheimer's disease α-syn Alpha-synuclein ALP Autophagy-Lysosome pathway CMA Chaperone mediated autophagy Cq Chloroquine DLB Dementia with Lewy bodies EM Electron microscopy HMW High molecular weight Htt Q72 Huntingtin exon1 fragment with 72 glutamine repeats IB Immunoblot IF Immunofluorescence KO Knockout LB Lewy body LN Lewy neurite MSA Multiple system atrophy NAC Non-amyloid component PD Parkinson's disease PBS Phosphate buffered saline PBS/Pff-td Transduced with PBS or Pff, respectively Pdp Protein degradation pathway Pff Pre-formed alpha-synuclein fibrils Rap Rapamycin SDS Sodium dodecyl sulphate SNc Substantia nigra pars compacta TX Triton X-100 Ub Ubiquitin UPS Ubiquitin proteasome system Wt Wild-type P-α-syn Phospho-alpha-synuclein

Chapter 1: General Introduction

1.1 Parkinson’s disease and other synucleinopathies

Parkinson‟s disease (PD) is the most common neurodegenerative movement disorder and it is the second most common neurodegenerative disorder after Alzheimer‟s disease (AD) (de Lau & Breteler, 2006). It is estimated that over 10 million people are impaired by PD, and this number is expected to significantly increase with aging population (de Lau & Breteler, 2006). Motor symptoms of PD include resting tremor, muscle rigidity and slowness of movement (bradykinesia). These most prominent symptoms of PD are caused by loss of dopaminergic neurons in substantia nigra pars compacta (SNc) and resulting brain dopamine insufficiency (Figure 1-1). Currently, PD patients are routinely treated by medications that exert their effects by increasing dopamine levels in brain. However, this approach only alleviates the motor symptoms, and progressive neurodegeneration typically decreases responsiveness to such therapies and eventually the symptoms worsen, in some extreme cases to the extent of complete loss of movement (akinesia). Dopamine replacement therapies are also ineffective against dementia that appear later in the course of disease in approximately 30% of patients

(Aarsland & Kurz, 2010).

Dementia with Lewy bodies (DLB) is the second most common cause of dementia after AD (Goedert et al, 2001), and is characterized by dementia, hallucinations

Figure 0-1-1: Degeneration of Substantia Nigra in PD

Figure 1-1: Degeneration of Substantia Nigra in PD (A) Images of midbrain sections of a neurologically healthy control (left), and a PD patient (right). In PD midbrain section, pigmented neurons of substantia nigra (black arrowheads) are severely reduced as a result of neurodegeneration. (B) Immunohistochemistry images of midbrain tissue stained with an α-syn antibody. At the left, a pigmented dopaminergic neuron with normal appearance is shown. At the right, a LB in a pigmented cell, and a proximal Lewy neurite (LN) are shown. Images kindly provided by Mr. John Robinson (Center for Neurodegenerative Disease Research, PA). Scale bar: 50µm.

as well as parkinsonian symptoms such as bradykinesia and rigidity. Multiple system atrophy (MSA) is a terminal neurodegenerative disorder, causing autonomic nervous impairment, parkinsonism and cognitive dysfunction. Individuals diagnosed with MSA have an average life span of 8 years only (Bower et al, 1997). As in PD, there are no treatments available to slow down or stop the progression of DLB and MSA.

The diseases mentioned above, as well as several others, such as: a subtype of

AD (Lewy body variant of Alzheimer‟s disease) and neurodegeneration with brain iron accumulation type 1 (NBIΑ-1), have one major commonality: the presence of insoluble inclusions composed of the protein alpha-synuclein (α-syn) (Goedert, 2001; Neumann et al, 2000). The pathological hallmarks of PD are Lewy bodies (LBs) and Lewy neurites

(LNs), which are fibrillar α-syn inclusions found in neuronal soma and neuritic processes respectively (Spillantini et al, 1997). The presence of LBs and LNs is correlated to functional deficits and cell loss observed in synucleinopathies: in PD LBs and LNs are common in brainstem particularly in SNc, in DLB, LBs are found in cortical areas and in

MSA, α-syn accumulations, namely glial cytoplasmic inclusions (GCIs) are widely distributed among oligodendrocytes in affected regions (Spillantini, 1999). The presence of these inclusions in all these diseases and the connection between their predilection sites and disease symptoms strongly implicate α-syn aggregation with disease pathogenesis. Moreover, mutations in α-syn, namely A53T, E46K and A30P or α-syn gene locus multiplications, that may directly or indirectly promote α-syn aggregation cause familial PD (Kruger et al, 1998; Polymeropoulos et al, 1997; Singleton et al, 2003;

Zarranz et al, 2004)., supporting that α-syn aggregation is a causative factor in disease

3 pathogenesis Therefore, these neurodegenerative diseases are collectively referred as

“synucleinopathies” (Goedert & Spillantini, 1998).

1.2 α-Synuclein

α-Synuclein is a 140 amino acid protein highly expressed in central nervous system neurons, such that it constitutes 1% of total soluble brain protein (Iwai et al,

1995). It can also be found at high levels in other tissues such as bone marrow and erythrocytes (Scherzer et al, 2008). The protein includes a N-terminal region composed of KTGEGV repeat sequences (Maroteaux et al, 1988), a hydrophobic central region

(Ueda et al, 1993; Weinreb et al, 1996), and a negatively charged C-terminal region

(Crowther et al, 1998).

Recombinant α-Syn can form fibrillar aggregates in vitro, in a nucleation and concentration dependent manner (Conway et al, 1998; Wood et al, 1999). These 10-19nm wide fibrils are typical amyloid: insoluble, protease K resistant and rich in antiparallel β- pleated sheet structure (Conway et al, 1998; Conway et al, 2000; Giasson et al, 1999).

Fibrils made of purified recombinant α-syn are both ultrastructurally and biochemically very similar to the α-syn immunoreactive fibrils found in LBs (Conway et al, 2000;

Giasson et al, 1999), which may indicate that LB formation is a process primarily driven by α-syn fibrillization. It has been shown that PD-linked α-syn mutations A53T and

E46K increase the rate of fibrillization in vitro (Choi et al, 2004; Conway et al, 1998). In addition, α-syn locus multiplications found in familial cases of PD, may increase

4 intracellular α-syn concentration and thus the rate of fibrillization. The PD-linked A30P mutation does not affect the rate of fibrillization in vitro (Conway et al, 1998).

The central, hydrophobic region of α-syn is initially referred as Non-amyloid component (NAC) domain, since it was first purified along with amyloid plaques from brains of AD patients. However, more recent studies demonstrated that α-syn does not co-aggregate with amyloid-β in extracellular space (Bayer et al, 1999), and the co- purification of NAC domain was likely due to a contamination by intracellular LBs, which are common in AD stricken brain tissues. NAC region spans the aminoacids 71-82 and gives α-syn the ability to form amyloid fibrils (Giasson et al, 2001). This region can fibrillize by itself at a much faster rate indicating that the remainder of the protein does not significantly contribute to fibrillization process (Giasson et al, 2001).

N-terminus of α-syn is believed to be involved in membrane interaction and studies on α-syn crystal structure showed that this region, which is natively unfolded, assumes an alpha-helical conformation when bound to lipid membranes (Davidson et al,

1998; Mihajlovic & Lazaridis, 2008). Although PD-linked α-syn mutation A53T is not shown to affect lipid binding, other PD-linked mutations modify this interaction in opposite ways, such that A30P mutation reduces lipid binding, whereas E46K promotes it

(Choi et al, 2004; Perrin et al, 2000). These conflicting results may suggest that lipid binding ability of α-syn does not play a direct or pivotal role in α-syn pathogenesis.

C-terminus of α-syn is highly acidic due to the abundance of glutamate residues and it is a evolutionarily less conserved part of the protein (Maroteaux et al, 1988). C-

5 terminally truncated α-syn aggregates faster in vitro (Murray et al, 2003) and truncated α- syn can be found in LBs, however it not clear if this modification occurs after aggregation, post-translationally before aggregation, or if these truncated α-syn species are alternative splicing variants (McLean et al, 2012).

α-Syn has two homologs in vertebrates: β- and ɣ-synuclein (Ninkina et al, 1998;

Tobe et al, 1992). β-synuclein is highly homologous to α-syn but lacks the majority of the

NAC domain, hence the ability to form amyloid fibrils. β-synuclein is not observed in any type of α-syn inclusions and believed to have an inhibitory effect on α-syn aggregation (Hashimoto et al, 2001). Mutations in β-syn gene have been linked to familial form of DLB (Ohtake et al, 2004). ɣ-Synuclein exhibits less homology to other synucleins, and there is no evidence indicating that it plays a role in synucleinopathies.

However, it has shown to be overexpressed in breast tumors and therefore can be used as a marker for tumor progression (Bruening et al, 2000).

In neurons, α-syn is localized to synapses, and found in close proximity to synaptic vesicles (Iwai et al, 1995). α-Syn is delivered to synapses later in development

(in about 18 weeks of gestation, (Galvin et al, 2001b)) and genetic ablation of α-syn does not impair synaptogenesis indicating that this protein does not play a vital role in formation of synapses (Abeliovich et al, 2000).

α-Syn knockout (KO) animals show only minor phenotypic changes (Abeliovich et al, 2000), and comparable results were obtained from α-syn/β-syn double KO studies suggesting that the failure to detect dramatic changes is not due to compensatory effects

6 of the other neural synuclein protein (Chandra et al, 2004). Similarly, overexpression of

α-syn in murine models did not cause overt physiological changes, apart from secondary effects of synaptic or neuronal degeneration accompanied by α-syn aggregation (Giasson et al, 2002). Therefore, there is no consensus on physiological function of α-syn.

Nevertheless; these studies provided evidence on possible functions of α-syn.

Most α-syn ablation studies showed effects, albeit subtle, in synaptic function.

Since α-syn binds membranes, and found in close association with synaptic vesicles, it was proposed that it may regulate synaptic transmission. Indeed α-syn KO mice have shown to possess a reduced pool of reserve synaptic vesicles (Cabin et al, 2002) and α- syn overexpression in PC12 and chromaffin cells results in a limited increase in docked synaptic vesicles suggesting that a late step in exocytosis is affected (Larsen et al, 2006).

Perhaps the most convincing evidence about α-syn function was presented in a study from α-syn/β-syn /ɣ-syn triple KO mice, which suggested that α-syn acts as a synaptic chaperone assisting presynaptic SNARE complex assembly (Greten-Harrison et al,

2010). This finding is supported by prior evidence indicating that α-syn has homology to chaperone proteins, have in vitro chaperone activity and can rescue neurodegeneration observed in mice lacking chaperone-like CSPα protein (Chandra et al, 2005).

Alternatively, it has been proposed that α-syn may regulate dopamine transporter

(DAT) delivery to plasma membrane, inhibit tyrosine hydroxilase (TH) (Lee et al, 2001;

Perez et al, 2002; Wersinger & Sidhu, 2003), and α-syn phosphorylation, which is mostly observed under pathological conditions, upregulates TH activity (Wu et al, 2011). Thus,

α-syn may play a role in determining intracellular dopamine levels but the physiological and pathological relevance of this is yet to be determined.

Moreover, studies indicate that α-syn may regulate activities of other enzymes.

Synuclein family members are potent inhibitors of phospholipase D2 in vitro (Jenco et al,

1998; Payton et al, 2004), and they may also inhibit protein kinase C (Narayanan et al,

2005; Ostrerova et al, 1999). However, more recent studies failed to demonstrate biological relevance of these findings and link them to the role of α-syn in pathogenesis.

1.3 Role of α-syn in pathogenesis

Similar to its normal function, the role of α-syn in pathogenesis is not completely understood. However, evidence suggests that α-syn aggregation plays a significant role in pathogenesis. Firstly, α-syn aggregates are found in all synucleinopathies and they are enriched in regions in brain regions associated with symptoms (Trojanowski & Lee,

2001). Secondly, two of three PD-linked α-syn mutations A53T and E46K increase α-syn aggregation in vitro (Choi et al, 2004; Conway et al, 1998), and the remaining mutation,

A30P, may indirectly promote aggregation by reducing lipid bound α-syn and increasing the concentration of free α-syn that can readily aggregate (Perrin et al, 2000). Moreover, in murine models of synucleinopathies overt phenotype and cell loss are typically observed after formation of insoluble α-syn aggregates (Giasson et al, 2002; Luk et al,

2012; Trojanowski & Lee, 1999), although more subtle phenotypic changes may be observed prior to initiation of neurodegeneration and LB-like α-syn aggregate formation

(Hashimoto et al, 2003). These aggregates may impair cellular function and cause cell

8 death through a wide variety of mechanisms, they could sequester important cellular proteins or vesicles, disrupt cytoskeleton, impair protein trafficking or other vital cellular processes. It is unlikely that aggregation of α-syn confers its toxicity through sequestering soluble α-syn thereby preventing its normal function, since none of α-syn KO models exhibited PD-like phenotype.

An alternative, but not mutually exclusive, hypothesis is that soluble α-syn oligomers or protofibrils are toxic to cells. Tubular, annular or spherical multimeric α-syn structures have been observed in vitro (Ding et al, 2002; Volles et al, 2001) and it has been suggested that annular α-syn structures may act as pores in membranes, such as mitochondrial membrane, causing mitochondrial dysfunction, oxidative stress and apoptosis via cytochrome c release. (Volles et al, 2001). Still, although oxidative stress and mitochondrial dysfunction are very strongly implicated in pathogenesis of synucleinopathies, particularly PD (Jenner & Olanow, 1996; Vila et al, 2008), the evidence implicating pore-like oligomeric α-syn structures with these defects is sparse and such structures are yet to be detected in cells, or in vivo. Apart from that, the results of several cell culture studies indicate that α-syn oligomers may also block protein degradation by lysosomes and proteasome (Cuervo et al, 2004; Martinez-Vicente et al,

2008; Snyder et al, 2003), which may indirectly cause toxicity.

Additionally, the extent of post translational α-syn modifications are shown to be dramatically increased in synucleinopathies and it was suggested that they may play a role in pathogenesis. While α-syn is scarcely phosphorylated at S129 in neurons, LBs and other α-syn aggregates are heavily phosphorylated. In addition, α-syn ubiquitination is 9 only observed in the presence of α-syn aggregation (Fujiwara et al, 2002; Kuzuhara et al,

1988). Nitrated α-syn is found in LBs and other α-syn aggregates, whereas it cannot be detected in healthy cells (Paxinou et al, 2001). None of these modifications are required for α-syn aggregation in vitro and in vivo (Michell et al, 2007; Sampathu et al, 2003), and although the evidence about the effects of these modifications on kinetics of α-syn aggregation is somewhat conflicting (Lee et al, 2008; Meier et al, 2012; Souza et al,

2000; Yamin et al, 2003), the trend is that they increase the rate of aggregate formation or stabilize the aggregates. Although it is possible that α-syn is more commonly modified in disease state, it is also very likely that newly synthesized α-syn is continuously modified but these modified forms are rapidly cleared and not readily detected unless they gain resistance to clearance by being recruited to pathological aggregates.

1.4 Protein degradation pathways

Protein degradation pathways (Pdps) are cellular mechanisms evolved to identify, and hydrolyze proteins that are needed to be removed from cells. Pdps function in post- translational regulation of protein levels, removal of damaged, misfolded, or aggregated proteins, as well as disassembly of proteins to acquire amino acids during starvation.

There are two major Pdps in eukaryotes that take part in degradation of intracellular proteins. Ubiquitin proteasome system (UPS) and autophagy-lysosome pathway (ALP)

(Rubinsztein, 2006). UPS is typically involved in degradation of short lived, cytoplasmic or nuclear proteins. In UPS, target protein is tagged by a chain of small proteins called ubiquitin. The ubiquitin chain is attached to lysine residue on the target protein via an isopeptide bond, and it is recognized by the 19S regulatory subunit of the proteasome. 10

This 19S proteasome takes part in removal of ubiquitin tag and unfolds the target protein, so that it can access the active sites of the proteases buried in barrel shaped 20S proteasome catalytic subunit (Bedford et al, 2010). Localization of active sites of the proteasome inside a barrel shaped protein which can be accessed only through a narrow channel, minimizes the possibility of uncontrolled degradation of surrounding proteins by the proteasome, however it creates a restriction such that proteins needs to be unfolded before degradation. This restriction creates two problems, firstly large aggregates cannot be directly degraded by proteasome, secondly misfolded proteins or oligomers which are resistant to unfolding can block the entry to proteasome and inhibit its function.

Chaperone mediated autophagy (CMA), microautophagy, and macroautophagy are three constituents of ALP and their commonality is that they involve translocation of cytoplasmic substrates to lysosomes or autophagolysosomes for degradation. In CMA, cytoplasmic proteins that carry the KFERQ motif are recognized by the chaperone Hsp70 and translocated to lysosomes through Lamp2a channels (Cuervo, 2010). Similar to UPS, this pathway necessitates unfolding of proteins and can be blocked by protein aggregates.

Approximately 30% of cytoplasmic proteins contain KFERQ motif and believed to be degraded by CMA (Cuervo, 2010). Microautophagy is the least understood and studied type of autophagy and is believed to occur by engulfment of small portions of cytoplasm directly by lysosomes. Neither its exact function, nor the extent of its role in total cellular protein degradation is clearly understood (Santambrogio & Cuervo, 2011).

Macroautophagy is the most prominent type of autophagy and therefore usually simply referred as autophagy. In autophagy, a number of proteins orchestrate formation of a

Figure 1-0-2: Simplified schematic diagram of mammalian autophagy

Figure 1-2: Simplified schematic diagram of mammalian autophagy With the activation of autophagy, pre-autophagosomal structure (PAS) is formed by assembly of autophagy related atg proteins. PAS primers formation of cup shaped

membranous phagophore, which is characterized by the enrichment of membrane bound LC3-II. While phagophore expands by recruitment of additional lipids, it engulfs the target it recognizes via adaptor proteins, such as p62 that binds ubiquitin and LC3. With the complete engulfment of the substrate, a closed, double membrane structure called autophagosome is formed. While autophagosomes are transported to lysosome rich perinuclear areas on microtubule tracks, they fuse with endosomes such as CD63 containing multivesicular bodies, forming amphisomes. Amphisomes eventually fuse with lysosomes that are decorated with Lamp1. This fusion results in formation of autophagolysosomes, in which lysosomal proteases degrade the autophagy substrates, p62 and LC3 located in the inner membrane of the autophagosome.

12 membranous body called pre-autophagosomal structure (PAS). PAS formation is followed by extension of lipid core to form a cup shaped membranous structure called the phagophore. The origin of the lipids that form this structure is debated but it is believed that endoplasmic reticulum is the major source, while golgi, plasma membrane, mitochondrial membrane and free lipids in cytoplasm may be contributing (Axe et al,

2008). Phagopore, which can be identified by phosphoethanolamine conjugated microtubule binding protein light chain 3–II (LC3-II) which decorates it, expands by progressive recruitment of additional lipids. Although it was initially thought that autophagy is an unspecific process, now significant evidence supports that prior to and during phagopore formation, adaptor proteins bind autophagy substrates and LC3-II, thereby recruiting newly forming autophagosome to the substrate to be degraded. One of the best examples of such adaptors is, the protein SQSTM-1/p62 which has both ubiquitin and LC3 binding sites therefore can tag ubiquitinated protein aggregates for autophagic degradation (Bjorkoy et al, 2005). Interestingly, PD-linked genes parkin and Pink-1 havealso shown to play a role in specific degradation of damaged mitochondria via autophagy (Narendra et al, 2008; Vives-Bauza et al, 2010), a process referred as mitophagy. When both ends of phagophore fuse, engulfing the substrate, a double- membrane autophagosome forms. Autophagosomes are transported to lysosomes in a microtubule dependent manner and during this process, they can also fuse with other cytoplasmic vesicles such as late endosomes, or multivesicular bodies, to form amphisomes (Eskelinen et al, 2005). The events that take place between autophagosome formation and autophagosome/lysosome fusion are called autophagosome maturation.

Autophagosome/lysosome fusion results in formation of autophagolysosomes and 13 proteolytic degradation of the substrate, as well as LC3-II and p62 that are located in the inner membrane of the autophagolysosome. This event results in release of degradation products for use as building blocks, and recycling of undegraded components such as proteins located in outer membrane of autophagosomes, and proteolytic lysosomal enzymes, consequently leading to clearance of autophagosomes from cytoplasm.

Autophagy, particularly during PAS and autophagosome formation steps, is orchestrated by a family of autophagy related proteins. In addition to the proteins that play a role in relaying initiation signals to autophagic machinery, such as ULK1 and beclin-1, proteins that are vital in autophagic process such as atg5, atg7 are often genetically ablated to eliminate autophagic activity in vitro or in vivo (Hara et al, 2006; Komatsu et al, 2006).

Autophagy is a tightly regulated process, particularly mTOR pathway is a major regulator of autophagy. Reduction in availability of amino acids, glucose, ATP/AMP ratio or growth factors inhibit mTOR activity, thereby activating autophagy. Among these factors, amino acid starvation or direct pharmaceutical inhibition of mTOR with drugs, such as rapamycin, are the most commonly used methods to activate autophagy

(Ravikumar et al, 2010). Autophagy can also be regulated through mTOR independent mechanisms, such as calpain activity, Ca+2 levels, and IP3 levels (Harr & Distelhorst,

2010). Furthermore, it has also been shown that proteolytic stress caused by proteasome dysfunction, oxidative stress, aging, certain disaccharides and spingolipids are among those that can affect autophagic activity (Ravikumar et al, 2010).

1.5 Pdps and neurodegeneration

There are several lines of evidence implicating Pdps with neurodegerative diseases. Firstly, majority of these diseases exhibit Pdp pathology such as accumulation of autophagic vacuoles in AD and PD (Boland et al, 2008; Higashi et al, 2011).

Furthermore, inhibition of these pathways via pharmaceutical means or genetic ablation of important Pdp genes are neurotoxic, suggesting that Pdp inhibition may be a mechanism through which primary defects indirectly lead to cell death in neurodegenerative diseases. Virtually all neurodegenerative diseases involve aggregation of disease specific proteins, such as α-syn in synucleinopathies, tau/amyloid-β in AD, and polyglutamine expanded huntingtin in huntington disease. Thus, impaired Pdp function may also be an initiating factor in pathogenesis since it has been shown that in the absence of Pdp function, cells can form aggregates similar to the ones observed in neurodegenerative diseases. However, it is not clear why different proteins aggregate only in certain brain regions in different neurodegenerative diseases if Pdp impairment played a strictly causative role in the pathogenesis. Therefore it is still debated whether impairment of Pdp function is mostly a result of disease pathology, or it plays a significant role in the pathogenesis.

As aforementioned, neurodegenerative diseases involve protein inclusions and autophagy was shown to be able to degrade inclusions of some of such proteins including huntingtin. Since as in PD, there is evidence that protein inclusions contribute to pathology also in other neurodegenerative diseases, activation of autophagy was offered as a therapeutic approach for such diseases. This topic will be discussed in section 1.8. 15

1.6 Pdps and α-syn degradation

As its function and role in pathology, the degradation of α-syn was also a topic that has not been completely clarified. However, recent in vivo and in vitro studies shed light on this matter and it is now widely accepted that natively unfolded α-syn, particularly when phosphorylated, is mainly degraded by the proteasome in a ubiquitin independent manner (Ebrahimi-Fakhari et al, 2011; Machiya et al, 2010). However, when

α-syn levels increase and α-syn forms soluble oligomers that cannot access catalytic sites of the proteasome because of their size and inability to be unfolded, autophagy plays a more important role in their degradation. α-Syn has a CMA motif and can be degraded by this pathway, but similar to proteasome, CMA fails to degrade α-syn when it forms stable oligomers (Cuervo et al, 2004).

Unlike soluble α-syn species, degradation of insoluble α-syn aggregates is not well documented. This is mainly due to lack of in vitro models which faithfully recapitulate formation of highly insoluble pathological α-syn aggregates. Although a number of studies indicate that autophagy can clear α-syn aggregates (Spencer et al,

2009; Wong et al, 2008)and elsewhere), the structures identified as α-syn aggregates in these studies are often fundamentally different than bona fide α-syn aggregates observed in disease stricken brain tissues. This topic will be further discussed in section 1.7.

1.7 Models of synucleinopathies

Due to prominence of α-syn aggregates in synucleinopathies, a large number of in vitro and in vivo models of α-syn aggregation were developed in the last decade. Unlike

16 many aggregate prone proteins, such as poly glutamine expanded Htt, α-syn does not readily aggregate when overexpressed. Particularly in cell based in vitro models, α-syn remains soluble even at very high expression levels and additional manipulations such as exposure to oxidative, nitrative or proteolytic stress are required to induce α-syn aggregation (Lee et al, 2002; McLean et al, 2001; Paxinou et al, 2001). Apart from having a relatively low efficiency, this approach has another limitation: since the manipulations that are needed to generate α-syn inclusions affect Pdp function, such models are not ideal to study degradation of these aggregates by Pdps, or their effect on

Pdp function. Co-overexpression of α-syn with snyphillin-1, which is shown to colocalize with LBs, results in formation of inclusions that contain α-syn in a small percentage of the cells (Engelender et al, 1999). In addition, overexpression of α-syn tagged with GFP also causes minor α-syn aggregation (McLean et al, 2001). The main caveat with these two models is the possibility that α-syn is “packaged” amorphously into the inclusions whose formation is driven by external factors, such as GFP aggregation and aggresome response (Kopito, 2000). On the other hand, purified α-syn can form aggregates that share similar ultrastructure with LBs and LNs, indicating that formation of these highly organized aggregates is likely driven by α-syn itself.

Although insoluble α-syn aggregates are observed in in vivo models of α-syn overexpression, there is evidence suggesting these models may not accurately recapitulate human disease. First of all, substantia nigra pars compacta (SNc), which is a predilection site of the aggregates in PD brain, is often resistant to aggregate formation and cell loss, whereas many regions that are not particularly affected in human disease, such as spinal cord, show significant pathology (Giasson et al, 2002). Secondly, in 17 several studies, the α-syn immunoreactive structures identified as aggregates are small puncta, which are neither fibrillar nor phosphorylated and there is no convincing biochemical data showing that they are insoluble. It may be highly misleading to draw conclusions about potential pathways that can clear pathological LB-like α-syn aggregates based on such models.

Although α-syn overexpression can cause PD in humans, attempting to replicate disease conditions in mice with α-syn overexpression may cause artifacts such as formation of amorphous inclusions before LB-like, amyloid aggregates can form. Indeed, such a study describes the α-syn immunopositive structures as: “less circumscribed, are present in the nucleus, and lack fibrillar components” moreover, there were no data shown about the solubility or phosphorylation state of these accumulations, indicating that these structures do not represent pathological LBs (Masliah et al, 2000). In addition, an inducible α-syn mouse model from our lab (Lim et al, 2011) showed that younger mice (12 month-old) had small, partially phosphorylated, soluble α-syn immunoreactive puncta, that can be cleared from certain brain regions after cessation of α-syn expression.

On the other hand, aged mice (20 month-old) had larger, insoluble, highly phosphorylated α-syn aggregates that are much more similar to LBs. Furthermore, α-syn puncta in younger mice were not associated with cell loss, whereas aged mice had overt cell loss on predilection sites of LB-like α-syn aggregates (Lim et al, 2011). These data indicate that α-syn overexpression may initially cause formation of α-syn immunoreactive structures which are fundamentally distinct from LBs, while formation of α-syn accumulations that could be used as LB-models necessitates more time. It could be speculated that this “lag period”, is the underlying reason for the difficulty of 18 generating cell-based models of LB formation. Elimination of this lag period would enable LB-like aggregate formation in cell culture, without generation of other types of, and potentially misleading α-syn accumulations.

Recently, we have shown that intracellular, LB-like α-syn aggregates can be efficiently “seeded” by introduction of exogenous α-syn fibrils (Pffs) to cultured cells that express α-syn, including primary neurons (Figure 1-3) (Luk et al, 2009; Volpicelli-

Daley et al, 2011). This method is extremely efficient such that more than 80% of the cells in the culture eventually form α-syn aggregates, more importantly; the seeded aggregates that form after transduction are remarkably similar to LBs and LNs in many aspects such as of their biochemical properties, ultrastructure, and post-translational modifications. Moreover, introduction of Pffs per se, is not toxic and does not affect function of Pdps, rendering these models ideal to study the effect of α-syn aggregates on

Pdps and cell viability, as well as the potential Pdps that can degrade them. Our cellular models of α-syn seeding and recruitment of endogenous α-syn to yield LB-like aggregates may replicate key aspects of synucleinopathies, as several recent lines of evidence suggest that the spreading of amyloid protein pathology, such as seen with α- syn in PD, may result from the neuronal release and uptake of extracellular seeds (Angot et al, 2010; Brundin et al, 2010; Luk et al, 2012; Soto, 2012). Indeed, these recent findings help explain prior observations about linear, non-random, spread of α-syn pathology (Braak et al, 2003) , and the appearance of LBs in neuronal grafts implanted into the brains of PD patients (Kordower et al, 2008).

There are also models of synucleinopathies which are not dependent of α-syn aggregate formation: Since the most prominent pathology, and the underlying reason of 19

Figure 0-3: Pre-formed fibril transduction model of α-syn aggregation

Figure 1-3: Pre-formed fibril transduction model of α-syn aggregation (A) A schematic representation of pre-formed fibril (Pff) transduction method. Recombinant α-Syn (shown in red) is allowed to form fibrils, which are sonicated to form Pff fragments. Fragmented Pffs (in neurons) or Pff-lipid complex generated by incubation of fragmented Pffs with lipid-based tranduction reagent (in non-neuronal cells), are added to cell media. Both in neuronal and non- neuronal cells, transduced Pffs gain entry to cell and recruit endogenously expressed α-syn (shown in green) leading to formation of LB and LN-like α-syn aggregates. (B) Phospho-α-syn (p-α-syn, green) IF images showing: (Left) A high magnification view of a Pff-td HeLa cell with a LB-like aggregate. Note the dense core, and radiating filamentous structures. (Middle, right) Pff-td neurons showing both LB-like somatic aggregates and LN-like neuritic aggregates.

motor symptoms is the loss of dopaminergic neurons in substantia nigra in PD, toxin- mediated ablation of these cells has been used to generate α-syn independent models of

PD in several organisms ranging from flies to non-human primates. The most common toxins used are the by-product of an illicit drug synthesis attempt MPTP, the herbicide paraquat, and the natural insecticide rotenone. All these toxins are associated with increased risk of PD, in humans (Gorell et al, 1998; Langston et al, 1999; Liou et al,

1997) and inhibit mitochondrial complex I thereby causing selective degeneration of nigrostriatal dopaminergic neurons (Betarbet et al, 2000). Although some studies report that paraquat and rotenone increase α-syn expression induce formation of α-syn aggregates (Betarbet et al, 2000; Manning-Bog et al, 2002). MPTP monkey model, which is the gold-standard of toxin-based models, and brains of MPTP-intoxicated patients do not have α-syn aggregates (Forno et al, 1993) which suggest that mitochondrial dysfunction/oxidative stress mediated dopaminergic cell loss may be downstream of α- syn aggregate formation.

1.8 Proposed therapeutic approaches for synucleinopathies

Currently, only therapies available for synucleinopathies are the ones aiming to alleviate parkinsonian motor symptoms of these diseases. Although these symptomatic treatments such as levodopa, MAO inhibitors or deep brain stimulation are very helpful particularly in the earlier stages of the disease, they tend to be less effective at more advanced stages and they do not improve common non-motor symptoms such as cognitive decline observed particularly in DLB. In addition, such therapies do not slow down autonomic dysfunction in MSA which renders the disease lethal. Therefore, a 21 therapeutic strategy which would stop or slow down the disease progression is required.

Stem cell based therapies are promising (Schwarz & Schwarz, 2010), but are far from ready to be employed for routine clinical use. Furthermore, transmissible nature of α-syn pathology may dramatically limit the success rate of such therapies.

For reasons we discussed before, reduction of α-syn aggregation is one of the most commonly studied potential therapeutic approach. There are multiple ways to elicit a decrease in α-syn aggregates: reduction of α-syn synthesis, inhibition of α-syn aggregation, or induction of clearance of α-syn aggregates. α-Syn expression may be reduced by delivery of anti-α-syn siRNAs or activation of Pdps that degrade α-syn.

Although there is no convincing human data on this subject, based on the findings on mouse models (Abeliovich et al, 2000), it is plausible to expect that a decrease in α-syn levels will not have a detrimental effect on human brain function. However, this approach is primarily preventative and although it may have significant benefits before formation of the aggregates, its effectiveness may be limited after the symptoms of the disease manifest. Inhibitors of α-syn aggregation are also investigated as potential treatment options (El-Agnaf et al, 2004), however, unlike inhibiting an enzymatic function, inhibiting interaction of large protein surfaces (for α-syn, hydrophobic NAC region) has proven to be very challenging. Because of the nature of the likely mechanism of inhibition, i.e. binding NAC region and blocking further α-syn binding by steric hindrance, such inhibitors should be relatively large, and be used at near-equimolar ratios with α-syn, which makes them poor drug candidates. It was suggested that β-synuclein inhibits α-syn aggregation (Hashimoto et al, 2001), but the evidence is not compelling

22 enough to render β-syn expression a viable therapeutic approach, particularly considering the technical challenges of gene therapy. If possible, removal of pre-existing aggregates could perhaps be the most practical therapy for synucleinopathies, since it would not necessitate diagnosis of the disease before manifestation of clinical symptoms and it could be beneficial to the patients at more advanced stages of the disease. A small number of studies claim benefit of immunotherapy in mouse α-syn overexpression models, however, these studies were based on the aggregation model we previously discussed (Masliah et al, 2000), i.e. the α-syn accumulations observed in this model were vastly different from LBs or LNs. Moreover, while the first study based on this model claimed that immunotherapy clears intracellular α-syn accumulations (Masliah et al,

2005) the second study based on the same mouse model, but carried out after the advent of α-syn transmission evidence (Bae et al, 2012) attributed the effect of the immunotherapy to clearance of extracellular α-syn seeds. Perhaps, this example may emphasize the vital importance of accurate models of synucleinopathies. Finally, possibly the most discussed approach to remove α-syn aggregates is activation of autophagy, particularly since it has been shown that aggregates of several neurodegenerative disease related proteins can be cleared via this pathway (Bjorkoy et al, 2005; Williams et al,

2006). However, as discussed before, α-syn aggregation models used in such studies may not fully recapitulate α-syn aggregation in synucleinopathies and the limited effect observed in a number of such studies may reflect indirect reduction of aggregation due to reduction of soluble α-syn, and/or degradation of non-fibrillar α-syn deposits which are not representative of LBs or LNs. It is very important to note that the study that provides the bulk of the in vivo evidence on the role of autophagy in degradation of α-syn 23

“aggregates” is based on the controversial α-syn accumulation model that was used for the aforementioned α-syn immunotherapy studies (Spencer et al, 2009). Although activation of autophagy may appear to be a plausible approach, recent findings showing that such manipulations in cells with impaired autophagosome clearance may be detrimental (Boland et al, 2008; Wong & Cuervo, 2010; Zhang et al, 2011) emphasizes the fact that this strategy may lead to undesirable outcomes, and before offering it as a viable approach one must be precautious enough to investigate if these aggregates can indeed be cleared by autophagy and if they affect autophagy function using the most accurate α-syn aggregation models available.

Chapter 2: Optimization of Pff Transduction in HEK293 Cells

2.1 Abstract

Faithful models of α-syn aggregation are indispensible to study LBs, and their role in pathogenesis. We previously introduced a model of α-syn aggregation in which intracellular LB-like α-syn inclusions accumulate after cationic-lipid assisted internalization of pre-formed α-syn fibrils into α-syn-expressing cells. However, a limitation of the model was that only a minority of the treated cells developed aggregates, decreasing the sensitivity of the biochemical analyses that aim to identify the cellular impact of α-syn aggregates. Here, we demonstrate that the model can be significantly improved by use of truncated Pffs and mutant α-syn expression. We show that, when seeded with c-terminally truncated Pffs, A53T mutant α-syn has a higher propensity to form aggregates compared to both wild-type and other PD-linked α-syn mutants, such that almost 80% of cells formed aggregates after transduction of A53T mutant a-syn expressing cells. However, even though the amount of aggregated α-syn is dramatically increased in the optimized system, cationic-lipid reagent is still needed to attain sufficient

α-syn aggregation. We believe that our findings rendered the Pff-transduction model a better in vitro tool to study synucleinopathies, and we hope that it will be helpful in our efforts to gain an insight into the role of α-syn aggregates in neurodegenerative disease.

2.2 Introduction

To understand the mechanism of LB formation and its possible role in pathogenesis, numerous in vitro models of α-syn aggregation has been generated.

However, it has not been possible to model LB formation in various cell lines by simple

α-syn overexpression and this may due in part to the fact that α-syn is highly soluble and exist normally as an unstructured protein. Through use of different methods including co- overexpression with other proteins, exposure to oxidative stress, nitrative insult (McLean et al.,2001 ; Paxinou et al., 2001 ; Lee et al., 2002 and elsewhere), several other groups succeeded in generation of α-syn aggregates only in a small percentage of cells. In addition to their scarcity, those aggregates did not show important characteristics of LB, such as, containing fibrillar α-syn, being ubiquitinated (Kuzuhara et al., 1988) and phosphorylated (Fujiwara et al., 2002). The need for a more efficient model that would recapitulate α-syn aggregate formation in vitro was met when we developed a novel method for introducing preformed α-syn fibrils (Pffs) into cells. α-Syn readily fibrillizes in test tube (Conway et al., 1998 ; Giasson et al., 1999) and these amyloid Pffs can be delivered to different cell types through a liposome mediated protein transduction method. Once they gain entry inside cells, Pffs are able to recruit endogenous α-syn into

LB-like aggregates. This system creates unique opportunity to investigate whether or not intracellular α-syn aggregates affect cellular function. However, this model was limited in the sense that only 15-20% of transduced cells form large (>5µm in diameter) inclusions

(Luk et al., 2009). A higher efficiency was required to biochemically determine the functional consequences of the inclusions on the cell and to test how pharmacological

26 manipulations can impact aggregate formation. We hypothesized that inclusion formation would be accelerated by use of disease associated mutant α-syn in our model.

2.3 Results

Improvement of Pff transduction efficiency by use of truncated Pffs and A53T mutant α-syn expression

We have previously shown that when transduced with wild-type (wt) Pffs, approximately 40% of the cells that express wt α-syn form phosphorylated, insoluble α- syn aggregates and only 15-20% of these cells form large (>5um in diameter) inclusions

(Luk et al., 2009). However, a higher efficiency was required to biochemically determine the functional consequences of the inclusions on the cell and determine how pharmacological manipulations can impact aggregate formation. We hypothesized that inclusion formation would be accelerated by transducing cells that express A53T mutant

α-syn, that is linked to familial PD and has accelerated fibrillization in vitro (Conway et al, 1998). We also wanted to test if Pffs generated from c-terminally truncated α-syn

(amino acids 1-120), which has been shown to fibrillize faster in vitro (Murray et al,

2003), will result in increased transduction efficiency. To test these possibilities, HEK293 cell lines that express comparable levels of wt and A53T mutant α-syn were transduced with wt Pffs or 1-120 truncated Pffs and inclusion formation was visualized using mAB

81A which detects phosphorylated, aggregated forms of α-syn, including LBs in human brain tissue. As expected, we observed that cells expressing A53T mutant α-syn generate more pathology when transduced with both wt and truncated Pffs. Furthermore, truncated

Pffs seed more efficiently in both wt α-syn and A53T α-syn expressing cells (Figure 2-

1A, B). Overall, the 4.7-fold increase in transduction efficiency (16.7%±3.5% vs 78.5%

±0.7%) was elicited using cells expressing A53T α-syn and 1-120 truncated Pffs for transduction. Our observations by IF was confirmed by immunoblotting, indeed, when 1-

120 and wt Pffs were compared in wt α-syn expressing cells, 1-120 truncated Pffs seeded dramatically more pathology (Figure 2-1C) and when 1-120 Pffs were used to transduce wt or A53T α-syn expressing cells, A53T expressing cells produced significantly higher pathology (Figure 2-1D) confirming our IF data. In particular when comparing wt and

A53T cells, the differences in pathology were even more pronounced by IB supporting the IF observation that the aggregates are not only more frequent in A53T cells, but also larger in size.

Our significant improvement on transduction efficiency enabled us to monitor previously unidentified effects of the α-syn inclusions, such as cytotoxicity and impairment of autophagic maturation as discussed in Chapter 4.

Expression of PD-linked A30P or E46K mutant α-syn does not improve aggregate formation.

Since PD associated A53T mutant α-syn produced more aggregates, we wanted to test if expression of other PD associated mutants such as A30P and E46K (Kruger et al,

1998; Zarranz et al, 2004) would cause an increase in aggregate formation. We

Figure 4 Transduction efficiency is improved by mutant a-syn expression and use oftruncated Pffs

Figure 2-1: Transduction efficiency is improved by mutant a-syn expression and use of truncated Pffs:(A) Immunofluorescence (IF) pictures showing HEK293 cells stably expressing wild-type (wt) (left) or A53T (right) a-syn that were transduced with wt (top) or 1-120 truncated (bottom) a-syn Pffs, which are both myc tagged at c-terminus. Cells fixed after 48h of starvation and a-syn aggregates (red), Pffs (green), and nuclei (blue) were visualized with phospho-a-syn (p-a-syn), myc antibodies and DAPI respectively. 1-120 Pff transduced A53T a-syn cells generated the highest amount of pathology. (B) Quantification of IF data, n>400 cells for each condition. (C) Immunoblot with p-a-syn, n-terminal a-syn (SNL4), myc, p62 and a-tub as loading control. When transduced with comparable amounts of Pffs, 1-120 transduced wt a-syn expressing cells generate significantly more pathology compared to wt a-syn transduced wt a-syn expressing cells. (D) IB with p-a-syn, c-terminal a-syn (Syn211), p62 and a-tub. A53T a-syn expressing cells generate more pathology when compared to wt a-syn cells. Together, these data indicate that use of A53T a-syn cells, and 1-120 truncated Pffs dramatically increases a-syn pathology [data from a minimum of 3 independent experiments, error bars +SEM.**denotes p<0.005 according to student‟s t test. Scale bars 10um].

Figure 2-0-5: E46K and A30P mutant α-syn expression does not increase aggregate formation

Figure 2-2: E46K and A30P mutant α-syn expression does not increase aggregate formation (A,B) Naive HEK293 cells transiently transfected with wt and A53T, E46K, A30P mutant α-syn and transduced with 1-120 α-syn Pffs. (A) IF pictures showing p-α-syn aggregates (red). (B) IB picture showing α-syn

expression and aggregated/ phosphorylated α-syn levels. Both IF and IB data indicates that A53T mutant α-syn has the highest preponderance to form aggregates and other α-syn mutants do not increase aggregation in this experimental setting.

30 transiently transfected naïve HEK293 cells, with wt, A53T, A30P and E46K mutant α- syn and transduced the cells with 1-120 Pffs. We observed that under the conditions we tested, A53T mutant α-syn was the most efficient in aggregate formation, whereas A30P and E46K mutant did not show increased efficiency over wt α-syn, in fact they trended towards decreased efficiency (Figure 2-2A, B).

Transduction without reagent is feasible, but inefficient in HEK293 A53T α-syn cells

Previously, it had been shown that p-α-syn aggregate seeding through Pff transduction is not feasible without use of the cationic reagent Bioporter, if wt α-syn Pffs and wt α-syn expressing cells are used (data not shown). Because of the dramatic increase elicited with A53T α-syn expression and transduction with 1-120 Pffs, we wanted to test if with this new combination, Pff transduction would be sufficiently efficient without use of Bioporter reagent. When we directly added sonicated 1-120 Pffs into culture media, we indeed saw a number of cells forming α-syn aggregates; however, the efficiency was dramatically lower compared to Bioporter mediated transduction (Figure 2-3A). When we treated the cells with chloroquine, a lysosome inhibitor which is also used to increase transfection efficiency (Luthman & Magnusson, 1983), we indeed saw an increase in number of cells with aggregates only in no-reagent Pff transduced cells supporting that unless Bioporter is used, only minute amounts of endocytosed Pffs can escape lysosomal compartments prior to degradation (Figure 2-3A). Indeed, we observed a dramatic increase in Pffs retained in cells upon chloroquine treatment.

Figure 2-0-6: Transduction without reagent is feasible, but inefficient in HEK293 A53T α-syn cells

Figure 2-3: Transduction without reagent is feasible, but inefficient in HEK293 A53T α-syn cells (A) IF pictures showing HEK293 A53T α-syn cells transduced with 1-120 Pffs with or without use of Bioporter. Approximately 20% of cells formed small p-α-syn aggregates in no reagent Pff-td cells. With chloroquine treatment (+Cq), approximately 40% of cells had p-α-syn aggregates and few cells had larger aggregates, comparable to the ones in Bioporter Pff-td cells in size. No aggregates were observed in PBS transduced cells with or without chloroquine. (B) IB with α-syn and p-α-syn showing that without Bioporter (No BioP), aggregate formation is minor. Ws indicates 24h incubation with Pffs followed by wash with media, 1x 48h incubation with regular amount of Pffs, 2x with double amount of Pffs, and 1xCq 20uM Cq treatment during transduction. (C) IF pictures showing naïve HEK293 cells transduced with Pffs without Bioporter. 24h after transduction, there are very few Pffs retained in cells unless the cells are treated

with chloroquine which dramatically enhances retention of Pffs

2.4 Discussion

Here, we aimed to optimize Pff transduction system to increase its usability and efficacy as an α-syn aggregation model. We have found that if A53T α-syn expressing cells are used and they are transduced with 1-120 Pffs, the efficiency of aggregate formation is dramatically increased. It is likely that more positively charged 1-120 Pffs are internalized by the cells more readily, increasing transduction efficiency.

Speculatively, this increased internalization is enabled by their enhanced interaction with the reagent, however, since they seed better also in the absence of reagent and deletion of negatively charged c-terminus is not expected to increase interaction with a cationic reagent, this possibility is less likely. Another possible explanation is that being able to fibrillize at an increased rate, 1-120 Pffs seed faster once they are in cytoplasm, but transduction with A53T α-syn Pffs was not more efficient than wt α-syn Pff (data not shown), which does not support this hypothesis.

We observed that the aggregates in A53T α-syn expressing cells are not only more frequent, but also larger (Figure 1A), suggesting that A53T α-syn is recruited faster to the seeds due to its higher aggregation propensity. The evidence suggests that A30P mutant may form soluble oligomers which inhibit its further fibrillization (Conway et al,

2000; Lemkau et al, 2012). It is also possible that the type of seed is important in fibrillization, thus A30P or E46K α-syn expression might have caused increased aggregate formation in the cells transduced with the same type of mutant α-syn Pffs.

However, the focus of this study was to increase the efficiency of the system to a

33 satisfactory level and having achieved our goal with A53T α-syn/1-120 Pff combination, we did not test the efficiency of other combinations.

Although the effort of rendering expensive Bioporter reagent obsolete was not successful, the related experiments were informative. As a proof of concept, we showed the Bioporter merely facilitates transduction, and if Pffs gain entry to cell by other means, they can also seed aggregates. Our preliminary results suggest that Pffs are degraded in lysosomes, thus upon chloroquine treatment, transduction efficiency is increased in no- reagent Pff transduced cells, where the amount of retained Pffs are the limiting factor. We will further investigate the effect of lysosomal inhibition on Pff transduction in Chapter 3.

Chapter 3: Degradation of exogenous Pffs in non-neuronal cells

3.1 Abstract

Faithful models of α-syn aggregation are indispensible to study LBs and their role in pathogenesis. We introduced a model of α-syn aggregation in which intracellular LB- like α-syn inclusions accumulate after cationic-lipid assisted internalization of pre-formed

α-syn fibrils (Pffs) into α-syn-expressing cells. However, the fate of the Pffs that are introduced to cells was previously not well documented. Here, using cells that do not express α-syn, and therefore do not form LB-like aggregates, we investigated the mechanism through which Pffs gain access into cells. We show that Pffs are likely endocytosed and therefore lysosomal inhibition reduces intracellular clearance of the

Pffs. However, we also show that only a minority of the added Pffs gain access to cells, while the rest remain closely attached to the cell surface. Our data indicates that, at the present, our-cell based model may not be ideal to study the mechanisms of Pff internalization or degradation due to the large amount of extracellular Pffs. However, our results confirm that insoluble endogenous α -syn is excellent marker for pathology, since

LB-like aggregates are largely made of endogenous α-syn, and include only trace amounts of the added Pffs. We believe that the better understanding of the model we provided here will help interpret future results more accurately.

3.2 Introduction

In our α-syn aggregation model, we introduce exogenous α-syn Pff seeds to cultured cells, and once these seeds gain entry to cytoplasm, they recruit endogenous α- syn and induce formation of LB-like aggregates. We have previously showed that if these

Pffs are directly introduced to cells, they are internalized and degraded, and this process can be slowed by lysosomal inhibitors (Figure 2-2, 2-3), suggesting that Pffs are endocytosed and go through heterophagic pathway. However, it remained unknown if

Pffs are internalized with a similar mechanism when Bioporter reagent is used, which is important because this reagent vital in increasing transduction efficiency and therefore indispensible for our further studies in HEK293 model. Furthermore, it was also unclear if these Pffs can be recognized, and removed from cytoplasm by cellular degradation pathways. Elucidation of cellular mechanisms that can clear α-syn Pff before they seed α- syn aggregates, may significantly contribute to the efforts of developing therapies against

PD and related neurodegenerative diseases.

Here, we showed that also when Bioporter is used, use of lysosomal inhibitors increase retention of transduced Pffs, however they do not stop their eventual elimination at our experimental setting. With further investigation, we discovered that with or without use of Bioporter, the majority of the Pffs remains extracellular, and therefore amount of

Pffs by detected by immunoblotting is mainly constituted by extracellular Pffs. We concluded that due to the abundance of the extracellular Pffs, our model is not suitable to study degradation of internalized Pffs with our current set of tools, and the focus should

36 be on the degradation of LB-like α-syn aggregates that are almost exclusively comprised of endogenous α-syn.

3.3 Results:

Pffs are cleared from cells and this clearance is slowed by lysosomal inhibitors

As before, we conducted our Pff degradation studies in naïve cells to refrain from any interference by the recruited endogenous α-syn. When naive HEK293 or HeLa cells

(HeLa results shown, HEK293 results are comparable) are transduced with Pffs (using

Bioporter, unless otherwise indicated), a proportion of Pffs remained associated with the cells 48h after transduction, and this amount was greatly increased when cells were treated with NH4Cl, a lysosomal inhibitor, during the course of the experiment (Figure 3-

1A). Similarly, IB showed that transduced Pffs rapidly decreased in time, and this decrease was slowed, but not completely inhibited by NH4Cl (Figure 3-1B,C). We also observed an identical effect with chloroquine, another lysosomal inhibitor (data not shown).

Pffs do not significantly colocalize with early endosomes or lysosomes

Our results thus far suggest that the Pffs are internalized by the cells through endocytosis. Endocytosed material first goes into early endosomes, which consequently mature and fuse with lysosomes (Eskelinen, 2005). Thus, we expected to see a

Figure 3-0-1: Transduced Pffs are cleared from cells and this clearance is slowed by lysosomal inhibitors

Figure 3-1: Transduced Pffs are cleared from cells and this clearance is slowed by lysosomal inhibitors (A) IF pictures showing Pff-td (with Bioporter) naive Hela cells treated with vehicle(left) or 25mM NH4Cl (right), 4 hours after transduction until cells are scraped (48h after transduction). The amount of retained Pffs (red) was significantly higher in NH4Cl treated cells. (B) IB for α-syn and GAPDH as a loading control. (C) Quantification of IBs from 3 independent experiments. Transduced Pffs were reduced with time and this reduction was slowed down by lysosomal inhibitor NH4Cl. [Error bars indicate SEM. Data from a minimum of 3 independent experiments].

significant subpopulation of Pff colocalize with these structures. IF using EEA1 and

Lamp1 as markers of early endosomes and lysosomes, respectively, showed that Pffs only very rarely (<1%) colocalized with any of these two endosomal markers, even when lysosomal degradation was inhibited (Figure 3-2,data not shown). Considering that Pffs per se, do not colocalize with any of the cytoplasmic markers we tested thus far (p62,

LC3, 20S proteasome; data not shown) and do not undergo phosphorylation or ubiquitination, this finding suggests that the majority of Pff are either inertly localized in cytoplasm or localized extracellularly. In addition, this observation refuted the hypothesis that the majority of the Pffs that do not colocalize with p-α-syn aggregates are trapped in endocytic compartments.

A small proportion of added Pffs gain entry to the cells

To investigate the possibility that the majority of the Pffs remain extracellular, we used confocal microscopy. Lectin-PHA was used to visualize plasma membrane. As, suspected, we observed that the majority of the Pffs appear to be extracellular, although we were also be able to detect occasional intracellular Pffs (Figure 3-3A). Similar results were obtained both in reagent assisted transduced cells and reagent free transduced cells indicating that this phenomenon is not an artifact caused by the transduction reagent (data not shown). Although confocal microscopy is a useful method to investigate a small number of cells at a time, a method which would enable us detect intracellular Pffs more easily, and make semi-quantitative assessments about relative amounts of intracellular and extracellular Pffs was needed. Therefore we devised a method, namely in-out assay, to easily differentiate intracellular and extracellular Pffs. The assay is a two step IF 39

Figure 3-0-2: Majority of transduced Pffs are not localized into early endosomes or lysosomes

Figure 3-2: Majority of transduced Pffs are not localized into early endosomes or lysosomes (A-C) IF pictures showing Pff transduced naive HEK293 cells (A) Pffs only rarely colocalize with early endosomal marker EEA1. (B) Neither in the absence nor in the presence of lysosome inhibitor choloroquine (below, +Cq), Pffs do not colocalize with late endosomes/lysosomes decorated with Lamp1. (C) A magnified view of a chloroquine treated cell, showing lack of significant Lamp1/Pff colocalization.

Figure 3-0-3: A minute amount of Pff gain entry to cells

Figure 3-3: A minute amount of Pff gain entry to cells (A) Confocal-IF images showing Pff-td naive Hela cells incubated with lectin PHΑ-L alexa 488 (green) to delineate plasma membrane. On the left image of a representative field, on the right enlarged view of the corresponding z-stack (top) and 3 more z-stacks obtained from different fields. Majority of the Pffs (red) appeared outside the cell, but closely associated with the plasma membrane, although intracellular Pffs were also detected (white arrows). (B) Pff in-out assay. Cells were transduced with Pffs, 24h after transduction, they were incubated with α-syn antibody LB509 prior to fixation and permeabilization (indicated by “(-)‟). Following fixation and permeabilization (indicated by „(+)”) and incubated with another α-syn antibody SNL4. Controls included staining with both antibodies after fixation and permeabilization (bottom right) and staining with α-tubulin before and after fixation and permeabilization (bottom left). Intracellular Pffs that appear green (white arrows, arrowheads in the insets), were visible but much less common than

extracellular Pffs that appear yellow. In both (A) and (B) nuclei stained with DAPI (blue).

procedure: the first step is to stain the cells with an α-syn antibody, while they are still alive and therefore their membrane is intact. 1 hour incubation to stain extracellular Pffs is done at 4oC to inhibit possible antibody endocytosis and followed by wash steps, fixation and the permeabilization of the cells. After permeabilization, a second α-syn antibody, which will not compete with the first one for Pff binding (antibodies raised against opposite termini are to be chosen), is used to stain both extracellular and intracellular Pffs, and depending on the 2ndry antibodies are used at the end (typically red for the first and green for the second antibody), intracellular Pffs will be stained green only, whereas, being accessible to both antibodies, extracellular Pffs will appear yellow.

Controls include use of both antibodies after permeabilization, use of α-tub antibody before and after permeabilization and switching antibody order. After confirmation of its feasibility in this study, this method has been used in other studies for the same purpose

(Guo & Lee, 2011; Volpicelli-Daley et al, 2011).

Using this new method, we confirmed that the vast majority of Pffs are extracellular (Figure 3-3B). Interestingly, we observed an increase in both extracellular and intracellular Pffs upon lysosomal inhibitor treatment, although the increase in intracellular Pffs appeared to be more dramatic (Figure 3-3B). In support of our previous results, this finding suggests that only a small fraction of Pffs are endocytosed and undergo lysosomal degradation, whereas the vast majority remains extracellular.

3.4 Discussion

Here we showed that it is likely that Pffs can be endocytosed and degraded in lysosomes, however the majority of Pffs does not gain access to cells. These extracellular

Pffs are very closely associated with cells, as rigorous washing has no effect on this association and extensive trypsinization (1 hour) causes only a minor reduction in insoluble Pffs associated with cells (Figure 3-S1). We also observed that lysosomal inhibitors decrease elimination of extracellular Pffs. The underlying mechanism of this phenomenon is not clear, however, it can be speculated that since lysosome inhibitors may disrupt receptor uptake and recycling (Tietze et al, 1980), they may reduce secretion of extracellular proteases, such as matrix metalloproteases, which could degrade Pffs.

Regardless, we concluded that under current circumstances, our model is not suitable to study intracellular degradation of Pffs, thus we focused our efforts to study p-α-syn aggregates that are generated by recruitment of endogenous α-syn and therefore unequivocally intracellular. Since most of these LB-like aggregates only include minute amounts of extracellular seeds, Pffs can be considered as a catalyst that does not play a role in intracellular α-syn levels. The possible impact of extracellular Pffs, should not be disregarded and can be tested by use of naïve or α-syn KO cells that do not form aggregates. However, thus far, we did not observe any significant effect of Pff transduction of cells that do not form LB-like aggregates.

In conclusion, our preliminary studies on transduced Pffs indicated that only minute amounts of them gain access to cytoplasm and seed aggregates, whereas the majority 43

Figure 0-4: Extracellular Pffs are refractory to removal by washing or trypsinization:

Figure 3-S1: Extracellular Pffs are refractory to removal by washing or trypsinization IB for a-syn showing the amount of Pffs associated with naive HEK293 cells after various treatments. T- Triton X soluble ; S- SDS Soluble ;Input – Sonicated Pffs; Tp- Trypsinized; Tp+BioP – Bioporter/Pff Complex trypsinized; nxW – Cells washed n times with PBS before scraping; Tp- Cells are trypsinized, washed before scraping.

remains outside without any significant effect on the cell. Therefore, we focused our efforts to study LB-like intracellular α-syn aggregates alone.

3.5 Materials and Methods for Chapters 2 and 3

Mammalian Cell Cultures

HEK293 cells (QBiogene, CA) stably expressing wt or A53T α-syn were generated as previously described (Luk et al., 2009). Cells were maintained in complete media

[DMEM (Gibco, CA), supplemented with 10% FBS, penicillin/streptomycin and L- glutamine], and 500 µg/ml, 100 µg/ml or no G418 (Gibco, CA) was added to media for wt α-syn, A53T α-syn and naïve cells respectively. The day before the experiment,

HEK293 cells were plated at a density of 60,000-75,000 cells/well on poly-D-lysine

(PDL, 0.1 µg/ml in dH2O) and swine gelatin (0.1% w/v in dH2O) coated 12 mm coverslips, and 150,000-200,000 cells/well PDL coated on 12-well plates (Thermo-Fisher

Scientific, MA).

Hela T-Rex (Invitrogen, CA) A53T α-syn cells were generated as per manufacturer‟s instructions. Briefly, A53T α-syn cDNA in the pCDNA 5TO vector (Invitrogen, CA), was transfected into Hela T-Rex cells and hygromycin B (Thermo-Fisher Scientific, MA) resistant cells that inducibly express A53T α-syn were screened by light microscopy and

IB. Hela T-Rex A53T α-syn cells were maintained in Tet-Free complete medium

(DMEM, supplemented with 10% Tet-Screened FBS penicillin/streptomycin, L- glutamine) including 100 µg/ul G418 and 100 µg/ul hygromycin B. The day before the

45 experiment, Hela T-Rex cells were plated at a density of 25,000-30,000 cells/well on

PDL coated coverslips, and 50,000-60,000 cells/well on 12-well plates. 1 µg/ml dox was added at the time of plating to induce α-syn expression.

Recombinant α-syn proteins and generation of α-syn Pffs

Recombinant wt and C-terminally truncated α-syn (remaining amino acids: 1-120) with and without a C-terminal myc-tag, were expressed in Escherichia coli (E. coli) and were purified as previously described (Giasson et al, 2001). Shortly, E. Coli BL21-RIL cells expressing α-syn from the pRK172 expression vector were lysed using glass mortar and pestle, cell lysate was cleared through heat denaturation followed by centrifugation, and

α-syn was purified from cleared lysate by gel filtration and subsequent ion exchange chromatography.

α-Syn Pffs were generated by incubating purified α-syn (5 mg/mL in PBS) at 37°C with constant agitation for 5 days. Electron microscopy, Thioflavin S, and protein sedimentation assays confirmed generation of insoluble amyloid fibrils at the end of incubation (Giasson et al., 2001). Generated Pffs were aliquoted and stored at −80°C.

Pff Transduction

Protein transduction is defined as receptor and energy independent translocation of proteins into cells. To deliver Pffs into cells, we decided to use a liposome mediated transduction method since it is more efficient and, less injurious to cells compared to other available methods. Pffs that are used for transduction are generated by incubation of

46 purified recombinant α-syn, at 5mg/ml concentration for 24 hours at 37oC on an orbital shaker set to 1000rpm. Obtained fibrils are diluted to 1 to 50 in PBS, sonicated thoroughly with a probe sonicator and incubated 10 minutes with the transduction reagent

(BioPORTER® QuikEase™ : Sigmα-Aldrich, Missouri). For monomer transductions,

100ug/ml α-syn in PBS, and for mock transductions only PBS is used. After incubation,

Pff-Reagent mixture is mixed with warm OMEM + 1X Penn/Strep and incubated for another 5 minutes. Cells that were seeded to 6 or 24 well plates the day before transduction to reach confluence, are washed with OMEM + 1X Penn/Strep once, and incubated with the media containing Pff-Reagent mixture for 4-24 hours. Subsequently, the medium is changed to serum starvation medium (complete medium used to grow the specific cell line but with 0.5% instead of 10% FBS ; i.e. DMEM, 0.5% FBS, 1X

Penn/Strep and 1X L-Glutamine for Hela cells) and the transduced cells are incubated overnight. After overnight serum starvation, the cells are either scraped for biochemical studies or trypsinized and transferred to poly-D-lysine (PDL) coated coverslips for immunocytochemistry (ICC).

Drug Treatments

5 M ammonium chloride (NH4Cl, Thermo-Fisher Scientific, MA), 200 mM chloroquine

(Sigmα-Aldrich, MO) and 100 mM 3-methyladenine (Sigmα-Aldrich, MO) solutions were prepared in dH2O immediately before each experiment. Working concentrations of each drug are described in the figure legends.

Plasmids and Transfections

A53T a-syn pcDNA 5TO and wt a-syn pcDNA 5TO were generated using the inserts from available pcDNA3 a-syn constructs according to pcDNA 5TO cloning instructions

(Invitrogen, CA). a-Syn mutant A30P and E46K had been cloned into pCDNA 3 vector prior to this study and used here without any modification.HEK293 transfections were carried out using Fugene HD transfection reagent (Promega, WI) as per manufacturer‟s instructions.

Immunofluorescence

Cells were fixed with 4% paraformaldehyde in PBS followed by permeabilization with

0.1% TX. After blocking with Bovine serum albumin (BSA, 3% w/v in PBS), cells were incubated in primary antibodies (Table 1) followed by Alexa fluor-conjugated secondary antibodies (Invitrogen, CA). Cells on coverslips were incubated with DAPI and mounted on glass slides in Fluoromount-G (SouthernBiotech, AL). For neurons, 4% w/v sucrose included in the fixation solution. Images were acquired using an Olympus BX 51 microscope equipped with a digital camera DP71 and DP manager (Olympus, PA), except for Figures 6A and 8A, where images are acquired using Nikon Eclipse TE2000-E microscope equipped with CoolSnap HQ camera (Roper Scientific) and NIS Elements

(Nikon Inc, NY).

Immunoblotting

For sequential extractions, cells were scraped into 1% TX in Tris-buffered saline (TBS)

(50 mM Tris, 150 mM NaCl, pH 7.4) and protease/phosphatase inhibitor cocktail at 4°C.

Lysates were sonicated and centrifuged at 100,000xg for 30 min. The pellet was washed and suspended in 2% SDS in TBS. For total protein extractions, cells were scraped and sonicated directly in 2% SDS in TBS. Protein concentrations were estimated by BCA assay (Thermo-Fisher Scientific, MA) and equal amounts of protein were loaded on gels for each sample unless indicated otherwise. For sequential extractions, protein concentrations of TX fractions were used to calculate the volume of lysate to be loaded.

For a given sample, TX and SDS fractions were loaded at a volume ratio of 1:2, to better visualize the less abundant SDS-soluble proteins. This difference in loading amounts was considered when TX and SDS fractions were compared in the quantification panels.

Samples were run on 5-20% gradient SDS-Polyacrylamide gels, and proteins were transferred to nitrocellulose or PVDF membranes for IB. 5% fat-free milk in TBS was used for blocking, except for p-α-syn IB, where 7.5% BSA in TBS was used. For dot blot assays, samples were diluted 1:10 in TBS and loaded to Bio-Dot SF Microfiltration

Apparatus (BioRad, CA). Proteins captured by nitrocellulose membrane were visualized by IB. The primary antibodies used in this study are listed in Table 1. Immunoreactivity was detected via chemoilluminescence by use of HRP conjugated secondary antibodies

(Jackson Immunoresearch, PA), WesternLightning ECL substrate (Perkin Elmer, MA) and LAS-3000 imager (Fuji; Tokyo, Japan), or by use of IRDye secondary antibodies and

ODY-2816 Imager (Li-Cor Biosciences, NE).

Quantifications, Image Processing, Statistics

IB signals were quantified by Multi Gauge V2.3 (Fujifilm; Tokyo, Japan) or Image

Studio V2.0 (Li-Cor Biosciences, NE). Acquired images processed with Adobe

Photoshop CS2 (Adobe Systems Inc., CA) to facilitate visualization where needed. The adjustments applied equally across the entire image and to controls. All experiments were repeated a minimum of 3 times. The number following “±” sign indicates SEM.

Chapter 4: Lewy body-like α-synuclein aggregates resist degradation and impair macroautophagy

4.1 Abstract

Cytoplasmic alpha-synuclein (α-syn) aggregates, referred to as Lewy bodies

(LBs), are pathological hallmarks of a number of neurodegenerative diseases, most notably Parkinson‟s disease (PD). Activation of macroautophagy is suggested to help degradation of certain proteinaceous inclusions, but it is unclear if this pathway is capable of degrading -syn aggregates. Here, we examined this issue by utilizing cellular models in which intracellular LB-like α-syn inclusions accumulate after internalization of pre- formed α-syn fibrils into α-syn-expressing HEK293 cells or cultured primary neurons.

We demonstrate that α-syn inclusions cannot be effectively degraded, even though they co-localize with essential components of both the autophagic and proteasomal protein degradation pathways. The α-syn aggregates persist even after soluble α-syn levels have been substantially reduced, suggesting that once formed, the α-syn inclusions are refractory to clearance. Importantly, we also find that α-syn aggregates impair overall macroautophagy by reducing autophagosome clearance, which may contribute to the increased cell death that is observed in aggregate-bearing cells.

4.2 Introduction

Synucleinopathies are a family of neurodegenerative diseases characterized by cytoplasmic aggregates of the protein alpha-synuclein (α-syn), and include Parkinson‟s disease (PD), PD with dementia (PDD), dementia with Lewy bodies (DLB), and multiple 51 system atrophy (MSA) (Galvin et al, 2001a; Spillantini, 1999). In PD, PDD and DLB, α- syn forms highly insoluble, fibrillar aggregates called Lewy bodies (LBs) and Lewy neurites (LNs) which are located in the neuronal cytoplasm or neuritic processes, respectively. These inclusions are rich in phosphorylated α-syn (p-α-syn), are often ubiquitinated (Fujiwara et al, 2002; Kuzuhara et al, 1988) and are widely distributed in the central nervous system (CNS), where they are associated with neuron loss

(Wakabayashi et al, 2007). Moreover, α-syn mutations and α-syn gene duplication/triplication are linked to familial forms of PD (Kruger et al, 1998;

Polymeropoulos et al, 1997; Singleton et al, 2003; Zarranz et al, 2004). Collectively, these findings implicate α-syn aggregates in the onset and progression of disease. Thus, it is important to gain a better understanding of whether -syn inclusions can be effectively degraded by cells, and whether these aggregates might negatively affect key cellular pathways.

The removal of protein aggregates, as well as misfolded, damaged and regulated proteins, occurs via two main degradation pathways; the ubiquitin- proteasome system

(UPS) and the autophagy-lysosomal pathway (ALP) (reviewed by(Rubinsztein, 2006).

Macroautophagy, a prominent constituent of the ALP, is responsible for removal of both soluble and aggregated proteins, as well as damaged organelles, via their sequestration into double-membrane vesicles called autophagosomes, followed by their degradation by catalytic enzymes after autophagosome/lysosome fusion (Mortimore et al, 1996; Seglen et al, 1996). Macroautophagy (hereafter, autophagy) has been implicated in bulk degradation of pathologic aggregates, such as the poly-glutamine-expanded huntingtin

52 inclusions observed in Huntington‟s disease (Ravikumar et al, 2002). ALP is a vital cellular process and impairment of the ALP can lead to neurodegeneration, as demonstrated in animal models that are deficient in genes coding for autophagy proteins in neuronal tissue (Hara et al, 2006; Komatsu et al, 2006), and in human neurodegenerative lysosomal storage diseases (Futerman & van Meer, 2004; Nixon et al,

2008).

Both the UPS and ALP have been implicated in the degradation of soluble, non- aggregated α-syn (Cuervo et al, 2004; Ebrahimi-Fakhari et al, 2011; Webb et al, 2003), and autophagy is believed to be the pathway responsible for degradation of soluble oligomeric α-syn species (Ebrahimi-Fakhari et al, 2011). However, it is not clear if autophagy plays a role in the clearance of LB-like α-syn aggregates, as a heretofore lack of in vitro models that faithfully recapitulate α-syn aggregation has made it difficult to address this question. Unlike many aggregate-prone proteins, simple overexpression of α- syn does not lead readily to the formation of insoluble inclusions. Therefore, additional manipulations have been used to generate α-syn aggregates in cultured cells, including co-expression of proteins such as synphillin-1 (Engelender et al, 1999) and exposure to proteolytic inhibitors, oxidative stress, or nitrative insult (Lee et al, 2002; McLean et al,

2001; Paxinou et al, 2001), which likely directly or indirectly affect protein degradation pathways. However, the -syn aggregates formed in these various cellular paradigms typically fail to exhibit several important characteristics of LBs, including ubiquitination and the presence of insoluble phosphorylated α-syn. To better model LB-like inclusions in cultured cells, we recently developed models of α-syn aggregation in which the

53 introduction of pre-formed α-syn fibrils (Pffs) into cells, including primary neurons from wildtype (wt) non-transgenic mice, results in recruitment of endogenously expressed α- syn and formation of insoluble α-syn aggregates resembling LBs and LNs (Luk et al,

2009; Volpicelli-Daley et al, 2011). The ability to generate α-syn aggregates efficiently, without using any treatment that directly perturbs protein degradative function, provides model systems to investigate the possible interplay between autophagy and α-syn aggregates.

Here, we have investigated whether Pff-seeded, LB-like α-syn aggregates can be removed by autophagy. We observed that pathologic α-syn aggregates cannot be effectively eliminated and that they cause a partial impairment of autophagosome clearance that could contribute to the decreased viability observed in cells harboring - syn deposits. Thus, our findings provide novel insights into the effects of α-syn aggregates on cellular metabolism and viability, and may have implications regarding potential therapeutic strategies for synucleinopathies.

4.3 Results

Insoluble α-syn aggregates interact with protein degradation machinery.

The HEK293 model of LB-like -syn inclusions, in which -syn-expressing cells are transduced with -syn Pffs to induce aggregate formation (Luk et al, 2009), provides a cellular system to evaluate whether α-syn aggregates undergo proteolytic degradation.

Utilizing this model, we first assessed by immunofluorescence (IF) the localizations of several proteins involved in the ALP (Figure 4-1). These include p62, an adaptor protein that recruits autophagy machinery to protein aggregates (Bjorkoy et al, 2006); LC3, a protein specifically localized to autophagosome membranes when in its lipidated form

(LC3-II) (Kabeya et al, 2000); and Lamp1, a marker of lysosomes and autophagolysosomes (Fukuda, 1991). In control PBS-transduced (PBS-Td) cells without

α-syn aggregates, p62 and LC3 staining was faint and punctuate, and spread across the cytoplasm (Figure 4-1). In contrast, in Pff-transduced (Pff-td) cells bearing α-syn aggregates, p62 and LC3 staining was largely redistributed (Figure 4-1), such that the majority of the p-α-syn aggregates showed co-localization with accumulations of these proteins (97.4% ± 1.3% for p62 and 77.8% ± 5.5% for LC3). Autophagy substrates should progressively localize to autophagolysosomes, which are the product of autophagosome-lysosome fusion and where substrates are normally degraded by lysosomal proteases (Xie & Klionsky, 2007). In contrast with p62 and LC3, the lysosome/autophagolysosome marker Lamp1 did not show appreciable colocalization with α-syn aggregates (10.2% ± 6.0%; Figure 4-1). These data suggest that the α-syn inclusions are associated with the early autophagy machinery, but may not be delivered to autophagolysosomes.

In addition to co-localization with p62 and LC3, the -syn aggregates in Pff-td cells were also observed to be associated with the 20S proteasome (Figure 4-1). It was previously demonstrated that most α-syn inclusions in the HEK293 model are

Figure 4-0-1: Co-localization of protein degradation pathway components with α-syn aggregates

Figure 4-1: Co-localization of protein degradation pathway components with α- syn aggregates . Immunofluorescence (IF) images of HEK293 α-syn cells 24 h after Pff transduction, stained with p-α-syn antibody (red), DAPI (blue), or antibodies to p62, LC3, Lamp1, and 20S Proteasome (green). Dispersed and punctate cytoplasmic staining of p62 and LC3 was observed in PBS-td cells. In Pff-td cells, p-α-syn aggregates showed strong co-localization with p62 and LC3. Similarly, there was diffuse staining of the 20S proteasome in the PBS-td cells, whereas Pff- td cells had cytoplasmic 20S proteasome staining that largely co-localized with α- syn aggregates. In contrast, Lamp1-positive vesicles were observed in both PBS-td and PFf-td cells, and these did not typically co-localize with p-α-syn aggregates. [Scale bars = 20µm].

56 ubiquitinated and that they co-localize with the molecular chaperone, Hsp70 (Luk et al,

2009). These findings suggest that proteasome components, as well as those from the

ALP, are recruited to the -syn aggregates, likely in an attempt to degrade the inclusions.

Alternatively, these proteasome and autophagy constituents may interact aberrantly with aggregates, an event that could affect their function.

Insoluble α-syn aggregates are not degraded by autophagy and their clearance is severely impaired.

To elucidate if α-syn aggregates are degraded by autophagy, we evaluated whether autophagy inhibition or activation affected the levels of p-α-syn. HEK293 cells stably expressing α-syn (HEK293 α-syn) were treated for 24 h with either an inhibitor of autophagosome formation, 3-methyadenine (3-MA) (Kabeya et al, 2000; Seglen &

Bohley, 1992), or an activator of autophagy, rapamycin (Rap). Drug treatment was initiated 4 h after PBS or Pff transduction, a time when only minimal α-syn aggregation has occurred. Immunoblotting revealed that neither 3-MA nor Rap caused a significant change in p--syn levels, although rapamycin treatment trended towards an increase of p-

α-syn (Figure 4-2A,B). Examination of parallel cultures 4 h after drug addition, a time which allows for the detection of cell-associated myc-tagged Pffs, revealed that neither drug affected cellular Pff levels (Figure 4-S1A). The effectiveness of the drug treatments is seen in an LC3 immunoblot, with the expected decrease and increase of LC3-II formation after 3-MA and Rap treatment, respectively (Figure 4-S1A). Likewise, there were corresponding drug-induced changes in the abundance of LC3-positive puncta as visualized by IF (Figure 4-S1B). We also noted that there was a significant decrease in 57

Figure 4-0-2: Autophagy does not degrade α-syn aggregates and clearance of aggregated α-syn is severely impaired

Figure 4-2: Autophagy does not degrade α-syn aggregates and clearance of aggregated α-syn is severely impaired. After 4 h Pff transduction, HEK293 α-syn cells were treated with the 3-MA (10 mM) or rapamycin (Rap, 0.2 µM) in starvation media for 24 h. (A) Representative IB of p-α-syn and α-tubulin (α-tub), and (B) quantification of IBs from 3 independent experiments. “0h” refers to cells that were harvested prior to drug treatment, and “(-)” refers to vehicle-treated cells. (C,D,E) Hela T-Rex cells that express α-syn under the control of a doxycyline (dox)-inducible promoter were induced with dox for 16 h. After removal of dox, cells were transduced for 4 h and then incubated in dox-free starvation media until they were sequentially extracted. (C) Representative IB for endogenous α-syn, p- α-syn and α-tub. Equal volumes of cell extract were loaded. (D) Quantification of IBs from 3 independent experiments. The α-syn levels in TX fractions at 0 h after transduction were used to normalize α-syn levels at other time points. (E) Graph showing total (TX+SDS) α-syn levels. In PBS-td cells, α-syn was rapidly cleared from cells, whereas in Pff-td cells, no net α-syn reduction was observed. Error bars ±SEM. One-way ANOVA with Tukey‟s post-hoc analysis was used to test for statistically significant differences between treatments. NS and ** indicates p>0.05 and p<0.01, respectively. Scale bars = 20 µm.

Figure 0-3: Modulation of autophagy and α-syn aggregates

Figure 4-S1: Modulation of autophagy and α-syn aggregates. (A) 4 h after Pff transduction, cells were treated with vehicle (-),10 mM 3-MA, or 0.2 µM rapamycin (Rap), for an additional 4h until they were harvested in 2% SDS. Representative IB showing

decreased and increased LC3-II levels with 3-MA and Rap, respectively. The levels of myc- tagged Pffs were not significantly affected. (B) IF images of Pff-td cells treated with vehicle (-), 3-MA, or Rapamycin. Abundance of LC3 (red) puncta were greatly reduced with 3-MA treatment and increased with Rap treatment. The abundance of p-α-syn (green) aggregates was not significantly affected by the treatments, however, they did not colocalize with LC3 in 3-MA treated cells, where LC3 is mostly cytoplasmic. (C,D) 4 h after Pff transduction, HEK293 α-syn cells were maintained in nutrient rich media for 20 h, followed by rapamycin treatment for 72 h in the absence or presence of 15 µM Cq in nutrient–rich conditions. (C) IB of p-α-syn, p62, LC3 and the loading control, α-tub. (D) Quantification of p-α-syn levels in IBs from 3 independent experiments, with data normalized to the mean p-α-syn level from vehicle-treated, Pff-td cells. (E,F) HEK293 α-syn cells were transfected with the dominant negative autophagy inhibitor, mutant ATG4BC74A-strawberry, or with empty vector (-). Cells were transduced with Pff for 24 h, treated with 30 µM Cq for an additional 24 h, and then harvested 72 h after transfection. (E) IB of p-α-syn, p62, LC3 and the loading control, α-tub. (F) Quantification of p-α-syn levels from 3 independent experiments, with data normalized to the mean p-α-syn level from empty vector transfected Pff-td cells. [Error bars ±SEM. Student‟s paired t test was used to test for statistically significant differences. NS and * indicate p>0.05 and p<0.05 respectively. Scale bar = 20µm].

59 the amount of LC3 that was recruited to the aggregates when the cells were treated with

3-MA (Figure 4-S1B), confirming that the α-syn inclusions normally associate with autophagosome-conjugated LC3-II and not with cytoplasmic forms of LC3 that are increased after 3-MA treatment.

Although Rap treatment of the HEK293 -syn cells resulted in an enhancement of autophagy (Figure S1A,B) Rap-induced autophagy activation may have been attenuated because of prior activation of autophagy caused by serum deprivation during transduction

(Mortimore & Poso, 1987). To eliminate this possibility, HEK293 -syn cells were treated with Rap for 72 h under nutrient-rich conditions, and Rap did not significantly affect p-α-syn levels in the Pff-td cells, although the observed increase of LC3-II confirmed activation of autophagy (Figure S1C). Similarly, p-α-syn levels were not affected by treatment with the lysosome inhibitor, chloroquine (Cq) (Figure S1C,D), even though this drug clearly affected LC3-II clearance. Importantly, inhibition of autophagy by expression of a dominant negative ATG4B mutant, prevents lipidation of LC3 to LC3-

II, thereby inhibiting autophagosome closure and autophagy (Fujita et al, 2008)) did not lead to an elevation of p--syn (Figure S1E,F), although it resulted in significantly decreased total and LC3-I normalized LC3-II levels in ATG4B transfected cells

(2.34±0.30 vs 0.63±0.05 in Pff-td mock and ATG4B transfected cells, respectively. p<0.01). These data thus confirm that seeded α-syn aggregates in HEK293 cells are not appreciably degraded by autophagy.

As the 20S proteasome subunit was found associated with p--syn aggregates

(Figure 1), we also queried whether inhibition of proteasome function would affect the 60 levels of -syn aggregates. Addition of the proteasome inhibitor, clasto-lactacystin-betα- lactone (c-lac) did not increase Triton-X insoluble p-α-syn or -syn in the Pff-td cells

(Figure 4-S2Α-C), although c-lac treatment led to a significant increase in high molecular weight ubiquitinated proteins (HMW Ub; Figure 4-S2A,D), thereby demonstrating inhibition of proteasomal degradation. These results suggest that α-syn aggregates are not cleared effectively by either autophagic or proteasomal pathways. To further confirm this conclusion, another α-syn expression model was utilized in which α-syn is under the control of a doxycyline (dox)-inducible promoter. After growth for 16 h in dox- containing medium to induce α-syn expression, the cells were transduced with -syn Pffs and subsequently grown for varying lengths of time in dox-free medium to suppress α- syn synthesis. The cells underwent sequential extractions and samples were analyzed by immunoblotting, using a C-terminal α-syn antibody that recognizes endogenous -syn and not the C-terminal truncated 1-120myc -syn Pffs. In PBS-td cells, Triton-X100

(TX) soluble α-syn was rapidly degraded, with the amount of α-syn remaining 72 h after dox removal being 29.9±5.1% of the amount observed immediately (0 h) after transduction (Figure 4-2C,D). In Pff-td cells, TX-soluble α-syn decreased at a faster rate over time, concomitant with a progressive increase in TX-insoluble/SDS-soluble α-syn levels (Figure 4-2C,D). However, no net degradation of total α-syn was detected in the

Pff-td cells 72 h after dox removal, while there was a substantial decrease of total α-syn in the PBS-treated cells (Figure 4-2D,E). Taken together, these data indicate that α-syn aggregates are not effectively degraded by any cellular mechanism in the time-frame of this study, including but not limited to the UPS and ALP.

Figure 0-4: The UPS does not effectively clear α-syn aggregates.

Figure 4-S2: The UPS does not effectively clear α-syn aggregates. HEK293 α-syn cells were transduced with Pffs or PBS and 24 h later were treated with the proteasome inhibitor clasto-lactacystin-beta-lactone (c-lac, 10 µM) for an additional 24 h. The cells were sequentially extracted in 1% TX, followed by 2% SDS. (A) Representative IBs for p-α-syn, α-syn, ubiquitin, and the loading control, GAPDH, with (B,C,D) quantification of IBs from 3 independent experiments. C-lac treatment significantly increased high molecular weight ubiquitinated (HMW ub) protein levels. However, it did not increase insoluble p-α-syn or α-syn, demonstrating that proteasome activity does not reduce α-syn pathology. Pff transduction caused an increase in HMW ub protein levels, which may indicate that their degradation is impaired in transduced cells. [Error bars ±SEM. In (B) and (C), Student‟s t test, and in (D) one-way ANOVA with Tukey‟s post-hoc analysis used to test for statistically significant differences. NS, *, ** and *** indicate p>0.05, p<0.05, p<0.01 and p<0.001, respectively].

Typical autophagy substrates accumulate in α-syn aggregate-bearing cells.

As previously noted, LC3-II and p62 are autophagy substrates whose levels are routinely used to monitor autophagic function (Klionsky et al, 2007; Mizushima et al,

2010; Rubinsztein et al, 2009). Interestingly, both of these proteins accumulated in Pff-td

HEK293 α-syn cells (Figure 4-S1C,E), but not in Pff-td naïve HEK293 cells that do not express α-syn and therefore do not form α-syn aggregates (Figure 4-3A,B). Additionally, the levels of TX-insoluble high molecular weight (HMW) ubiquitinated proteins, which are typically degraded by autophagy (Bartlett et al, 2011; Pankiv et al, 2007), were significantly increased in Pff-td HEK293 α-syn cells in the absence or presence of proteasome inhibition (Figure 4-S2A,D), but not in Pff-td naïve HEK293 cells (Figure 4-

3C,D). These data suggest an impairment of autophagy in aggregate-bearing cells.

α-Syn aggregates also interact with autophagy and proteasome components in neurons but are resistant to degradation.

To determine if α-syn aggregates in primary wild-type (wt) neurons are also resistant to degradation, we utilized a recently developed neuronal model of α-syn pathology where endogenous mouse α-syn readily aggregates to form LB-like α-syn inclusions when transduced with α-syn Pffs (Volpicelli-Daley et al, 2011). The α-syn aggregates in neurons, like those in HEK293 α-syn cells, co-localized with ubiquitin

(Volpicelli-Daley et al, 2011), p62 and the 20S proteasome, but not with Lamp1

Figure 5 Typical autophagy substrates accumulate in α-syn aggregate-bearing cells.

Figure 4-3: Typical autophagy substrates accumulate in α-syn aggregate-bearing cells. (A,B) 48 h after Pff transduction, α-syn and naïve HEK293 cells were harvested and proteins were extracted in SDS. (A) Representative IB for p-α-syn, p62, LC3 and the loading control, α-tub. (B) Quantification of IBs from 3 independent experiments, with protein levels in Pff-td cells expressed as a percentage of the corresponding PBS control level. (C,D) 48 h after transduction, α- syn and naïve HEK293 cells were extracted in TX and then SDS. (C) Representative IB for HMW ubiquitin (ub), α-syn, with α-tub as a loading control. (D) Quantification of IBs from 3 independent experiments, with Pff-td cell levels expressed as a percentage of the corresponding PBS-td cell levels. Insoluble Ub protein levels did not increase in Pff-td naïve HEK293 cells. Error bars ±SEM. Student‟s paired t test was used to test for statistically significant differences. NS and * indicate p>0.05, and p<0.05, respectively.

(Figure 4-4A). Live imaging of LC3-mRuby transfected neurons revealed that LC3- positive vesicles co-localized with α-syn aggregates and accumulated in Pff-td neuronal processes, and immuno-EM with p-α-syn antibodies demonstrated that these aggregates can be found in close association with vesicle-like structures, but not inside these vesicles

(Figure 4B). These data suggest that α-syn inclusions associate with autophagy components in neurons, but are not efficiently delivered to autophagolysosomes for degradation.

To further confirm that neuronal -syn aggregates, like those in HEK293 -syn cells, are not effectively cleared by autophagy, Pffs were introduced into neurons that had been cultured in vitro for 10 days (DIV10). At DIV13, before significant amounts of α- syn aggregates were generated (Volpicelli-Daley et al, 2011), the Pff-td neurons were treated for 48 h with 3-MA to inhibit autophagy, or with starvation media to activate autophagy (Young et al, 2009). As expected, 3-MA caused accumulation of the autophagy substrate p62 in the TX-soluble fraction, whereas starvation slightly reduced its levels (Figure 4C). However, neither treatment significantly affected p-α-syn levels

(Figure 4C,D), suggesting that α-syn inclusions in neurons are not significantly degraded by autophagy during the time–frame of these studies.

To further examine clearance of α-syn aggregates in neurons, α-syn expression was reduced in PBS or Pff-td wt neurons by transfection of an anti-α-syn siRNA. Despite a significant reduction in endogenous -syn expression after siRNA treatment (77%±5%; compare –siRNA and +siRNA at 10d in Figure 4E), insoluble α-syn progressively accumulated in the siRNΑ-treated Pff-td neurons during the course of experiment 65

Figure 4-0-6: α-Syn aggregates interact with protein degradation pathway components but cannot be effectively degraded in neurons

Figure 4-4: α-Syn aggregates interact with protein degradation pathway components but cannot be effectively degraded in neurons. At 5 days in vitro (DIV), primary hippocampal neurons were transduced with human wt α-syn Pffs, and analyzed at 19 DIV. (A) IF using indicated antibodies, demonstrating that neuronal α-syn aggregates co-localized with p62 and 20S proteasome but not with Lamp1 (B) Top; images of neurons transfected with LC3-mRuby and α-syn- GFP, showing accumulation of LC3 puncta that co-localized with p-α-syn aggregates in Pff-td neuronal processes. Bottom; Immuno-EM, showing nano-gold-labeled p-α-syn aggregates at 50,000X magnification, with proximal autophagic vesicle-like structures.(C,D) At DIV10, neurons were transducedvesicle with wt-like α- syn Pffs, and at DIV13 they were starved or treated with 10 mM 3-MA. Pff-structures.(td cells inC,D one) At well DIV10, were harvested at DIV13 to monitor pre-treatment levels of p-α-syn (denotedneurons as were “0h”). transduced The rest were harvested at DIV15. (C) IB for indicated antibodies (Tuj-1, neuronwith wt specific α-syn βIII Pffs,-tubulin). and at (D) Quantification of IBs. (E,F) At DIV5, neurons were transducedDIV13 with PBS they orwere wt αstarved-syn Pffs or and after 24 h were transfected with anti-α-syn siRNA, or vehicle. treatedCells were with harvested 10 mM 310,-MA. 14 and 17 days after transduction. (-) siRNA denotes vehicle transfectedPff-td PBS cells-td cells in oneharvested well 10 days after transduction. (E) Representative IB for mouse specificwere endogenous harvested atα- synDIV13 (m- αto- monitor pre-treatment syn). (F) Quantification of IBs from 3 independent experiments. Error bars ±SEM. One-way ANOVA with Tukey‟s post-hoc analysis used. NS indicates p>0.05.levels Scale of p bar=20-α-syn (denoted µm (A), 500 nm (B). Images used in panel (B) are provided by Laura as Volpicelli “0h”).- Daley The rest and Annawere Stieber. harvested at DIV15. (C) IB for indicated antibodies (Tuj-1, neuron specific βIII-tubulin). (D) 66 Quantification of IBs. (E,F) At DIV5, neurons were transduced with PBS or wt α-syn Pffs and after 24 h were transfected with anti- α-syn siRNA, or vehicle. Cells were harvested 10, 14 and 17 days after (Figure 4-4E,F). Thus, even a low level of α-syn expression was enough to sustain the growth of α-syn aggregates, suggesting that α-syn aggregates are not cleared from neurons at a significant rate.

Finally, Pff-td wt neurons, but not Pff-td neurons from α-syn knock-out (KO) mice, showed increased p62, LC3-II and TX-insoluble HMW ubiquitin protein levels

(Figure 4-5Α-D), demonstrating that accumulation of these proteins are not caused simply by the addition of Pffs, but rather by the m-α-syn inclusions that are seeded by

Pffs. Moreover, these data suggest that α-syn aggregates impair autophagy in neurons, as was observed in HEK293 cells.

Autophagy function is impaired in -syn aggregate-bearing cells.

To further substantiate that α-syn aggregates impair autophagy function, we examined the degradation of the established autophagy substrate, YFP-tagged huntingtin exon 1 fragment with 72 glutamine repeats (Htt Q72), which forms highly insoluble aggregates that are degraded by autophagy (Pollitt et al, 2003; Ravikumar et al, 2002).

Htt Q72 and p-α-syn did not associate with each other when examined by IF in Pff-td cells (Figure 4-6A). However, insoluble Htt Q72 levels were greatly increased in Pff-td cells compared to PBS-td cells (Figure 4-6B,C). The difference in insoluble Htt Q72 levels between the PBS-td and Pff-td cells was largely negated when autophagy was inhibited with 3-MA, and was enhanced when autophagy was further activated with Rap

(Figure 4-6B,C). This indicates that the increase in Htt Q72 levels in Pff-td cells is due to an impairment in autophagic function that cannot be overcome by enhancement of

Figure 4-0-7: Canonical autophagy substrates accumulate in neurons with α-syn accumulations.

Figure 4-5: Canonical autophagy substrates accumulate in neurons with α-syn accumulations. Primary hippocampal neurons from wt or α-syn knock-out (KO) mouse embryos were cultured in nutrient-rich neuronal media. At DIV5, neurons were transduced with human α-syn Pffs or PBS, and neurons were harvested for IB

at DIV19. (A,C) Representative IB for p-α-syn, m-α-syn, ub, p62, LC3 and α-tub. (B,D) The effect of Pff transduction on levels of immunoblotted proteins in wt and α-syn KO neurons. Proteins levels are shown as a percentage of the values of PBS-td cells. TX-insoluble HMW ub proteins, p62, LC3-II and m-α-syn levels were significantly increased in Pff-treated neurons, suggesting that removal of these proteins was impaired. No change was observed in Pff-td α-syn KO neurons. Error bars ±SEM. Student‟s paired t test was used to test for statistically significant differences. No asterisk, *, ** and *** indicate p>0.05, p<0.05, p<0.01 and p<0.001, respectively. Scale bar = 20 µm.

Figure 4-0-8: Autophagy function is reduced in α-syn aggregate bearing cells.

Figure 4-6: Autophagy function is reduced in α-syn aggregate bearing cells. (A,B,C) 48 h after PBS or Pff transduction, HEK293 α-syn cells were transfected with YFP-tagged Htt Q72. 20 h after transfection, the cells were treated with 10 mM 3-MA, 0.2 µM Rap or vehicle (-) for 24 h. (A) Diffuse, soluble (left) or aggregated (right) Htt Q72 (green) did not colocalize with p-α-syn (red) aggregates. (B) Representative IBs for p-α-syn and GFP (recognizes YFP tag of Htt), as well as Ponceau S total protein staining. The Htt Q72 within TX-insoluble cellular fractions was visualized by dot immunoblot. (C) Quantification of dot blots from 4 independent experiments. Ratio of insoluble Htt Q72 levels in Pff-td and PBS-td cells for each treatment condition is indicated below the graph. (D,E) 48 h after transduction, HEK293 α-syn or naïve HEK293 cells were transfected with YFP-tagged Htt Q72, and 48 h after transfection harvested for IB. (D) Representative IBs for GFP and Ponceau S total protein staining. The Htt Q72 within TX-insoluble cellular fractions was visualized by dot IBs. (E) Quantification of dot blots from 4 independent experiments, shown as ratio of insoluble Htt Q72 levels of Pff-td and PBS-td cells. Error bars ±SEM. Statistical significance was determined by (B,E) Student‟s paired t test and (D) one-way ANOVA with Tukey‟s post- hoc analysis. NS, *, ** and *** indicate p>0.05, p<0.05, p<0.01 and p<0.001, respectively. Scale bars = 20 µm.

autophagosome formation. That the amount of insoluble Htt Q72 was substantially increased in Pff-td cells upon 3-MA addition reveals there was not a complete cessation of autophagy-mediated degradation of Htt Q72 in the α-syn aggregate-bearing cells in the absence of 3-MA. Although there was a modest increase of Htt Q72 clearance in the Rap- treated PBS-td cells, the Rap effect was likely muted in both the PBS-td and Pff-td cells because autophagy was already activated due to maintenance under starvation conditions after transduction. As expected, p-α-syn levels remained essentially unaffected by the 3-

MA and Rap (Figure 6B). Likewise, Pff transduction did not increase Htt Q72 levels in naïve HEK293 cells that do not form α-syn aggregates (Figure 6D, E), demonstrating that

α-syn Pffs do not by themselves impair autophagy, nor do they seed Htt Q72 aggregates.

Together, these and previous data indicate that autophagy function is compromised in α-syn aggregate-bearing cells, as evidenced by the accumulation of canonical autophagy substrates and the partial inhibition of Htt Q72 degradation.

Autophagosome clearance is impaired in -syn aggregate-bearing cells.

As the autophagy pathway consists of multiple steps, including autophagosome formation, maturation, and subsequent clearance of autophagosomes, we investigated whether α-syn aggregates specifically interfere with any of these steps. Our data suggest that autophagosome formation is not prevented, as demonstrated by the large number of

LC3-II-positive autophagosomes that are associated with -syn aggregates, and the increased LC3-II levels observed in Pff-td cells (Figure 4-1, Figure 4-2C, Figure 4-

S1A,C,E and Figure 4-3A,B). However, LC3-II levels could also be elevated as the result

70 of an elevation in autophagosome synthesis, rather than or in addition to an inhibition of autophagosome clearance. To distinguish between these possibilities, we utilized a LC3-

II turnover assay (Mizushima et al, 2010). PBS-td and Pff-td HEK293 α-syn cells were treated for 4 h with a high concentration of chloroquine (100 M) to inhibit all lysosomal degradation. This treatment abolished the differences in LC3-II levels between PBS-td and Pff-td cells (Figure 4-7A,B), and such a result is typically observed when there is an impairment of autophagosome clearance (Mizushima et al, 2010; Rubinsztein et al,

2009). This was further confirmed when PBS-td and Pff-td cells were simultaneously treated with Rap and Cq for 4 h, to induce accumulation of a large pool of autophagic vesicles. After removal of both drugs, LC3-II levels rapidly decreased in the PBS-td cells, so that more than half of the LC3-II was cleared in 24 h and nearly all LC3-II was degraded by 48-72 h (Figure 4-7C,D). Remarkably, there was no net clearance of LC3-II in Pff-td cells after 24 h, and LC3-II levels remained significantly higher than the corresponding level observed in PBS-td cells at each later time-point (Figure 4-7C,D). It should be noted that the residual effects of Cq likely caused a transient slowing in the elimination of LC3-II in both PBS and Pff-td cells (Zetser et al, 2004). Nonetheless, these data suggest that α-syn aggregates do not inhibit formation of autophagosomes, but rather impair autophagosome clearance.

Lysosome function appears normal in -syn aggregate-bearing cells.

The absence of Lamp1 co-localization with α-syn aggregates (Figure 4-1) suggests that the resistance of α-syn deposits to degradation and the impairment of

Figure 4-0-9: Autophagosome clearance is impaired in α-syn aggregate-bearing cells.

Figure 4-7: Autophagosome clearance is impaired in α-syn aggregate-bearing cells. (A,B) HEK293 α-syn cells were incubated in nutrient-rich media for 4 h after Pff transduction and the cells were treated with 100 µM of the lysosome inhibitor chloroquine (Cq) 48 h after transduction. After 4 h of incubation with Cq, the cells were lysed in SDS buffer. (A) Representative IB for p-α-syn, LC3 and the loading control, α-tub. (B) Quantification from 4 independent experiments. Pff-td and PBS-td cells showed comparable LC3-II levels upon complete inhibition of autophagosome clearance by Cq, suggesting that autophagosome clearance is impaired. (C,D) HEK293 α-syn cells were incubated in nutrient-rich media 4 h after Pff transduction and cells were treated with 0.2 µM rapamycin and 100 µM Cq 48 h after transduction. This “pulse” generated a large pool of autophagosomes in both Pff-td and PBS-td cells. After 4 h, the drugs were removed, and cells were harvested 0, 24, 48 and 72 h later. As a control, a set of cells was treated only with vehicle and were harvested 72 h after treatment. (C) Representative IB for LC3 and α-tub. (D) Quantification of 3 independent experiments. Error bars ±SEM. In (B) and (D) Student‟s t test was used to test for statistically significant differences. NS, *, and ** indicate p>0.05, p<0.05, and p<0.01, respectively.

autophagosome clearance may be caused by a defect in autophagosome maturation, rather than by an impairment of lysosomal degradation. Nevertheless, lysosomal defects could contribute to decreased autophagosome clearance by inhibiting lysosome- autophagosome fusion (Kawai et al, 2007). To investigate whether α-syn aggregates affect Lamp1-positive lysosomes, we conducted IF analyses in PBS-td and Pff-td

HEK293 cells. Although lysosomes appeared primarily as abundant fine puncta in PBS- td cells, they were less abundant and enlarged in Pff-td cells. The frequency of giant lysosomes (>2.5 µm in diameter), which were rarely observed in PBS-td cells, was significantly increased in Pff-td cells (Figure 4-8A). No such change in lysosome morphology was observed in Pff-td naïve HEK293 cells (Figure 4-S3A), demonstrating that formation of giant lysosomes was caused by the formation of LB-like α-syn aggregates, rather than simply Pff treatment. Giant lysosomes could also be readily observed in PBS-td cells after treatment with lysosomal protease inhibitors (Figure 4-8A,

B). Notably, lysosomal inhibition did not result in the accumulation of p-α-syn aggregates in giant lysosomes within Pff-td cells, or increased Lamp1/p-α-syn colocalization (7.0%±2.6% with inhibitor versus 10.2%± 6.0% without inhibitor).This suggests that the lysosomal dilation observed in Pff-td cells was not due to the sequestration and subsequent degradation of α-syn aggregates in lysosomal compartments

(Figure 4-8A).

Lysosomal enlargement may indicate possible lysosomal dysfunction, which could lead to impaired autophagosome clearance. Therefore, we asked if the giant lysosomes in Pff-td cells are functional. The lysosomal processing of a fluorogenic

Figure 4-0-10: Lysosome morphology, but not catalytic function, is affected in Pff-td cells.

Figure 4-8: Lysosome morphology, but not catalytic function, is affected in Pff-td cells. (A)24 h after Pff transduction, cells were treated for 24 h with a lysosomal protease inhibitor cocktail (Lysosome Inh.) or vehicle (-). The IF images shows examples of giant (>2.5µm in diameter) Lamp1-positive (green) vesicles in Pff-td cells (arrowheads). Quantification of giant Lamp1 vesicles, shown as number of vesicles per 100 nuclei (blue), is shown on the right. Pff- td cells harbored significantly more giant lysosomes than PBS-td cells. Lysosomal inhibitor treatment eliminated this difference. Colocalization between p-α-syn (red) and Lamp1 (green) was not observed in the absence or presence of inhibitor treatment. (B) The fluorogenic lysosomal substrate, Magic Red MR-(RR)2, is cleaved by the lysosomal protease Cathepsin-B (Cath-B) to yield a fluorescent product. Both normal lysosomes and the giant lysosomes in Pff-td cells converted the fluorogenic substrate, suggesting that lysosomal function was unaffected by the α-syn aggregates. Treatment with lysosomal inhibitors (top-right) diminished lysosomal fluorescence, confirming dependence of Magic Red MR-(RR)2 cleavage on proper lysosome function. (C) A representative IB (n=3) is shown which reveals similar EGF-induced EGFR degradation by lysosomes in PBS-td and Pff-td cells 30 min and 120 min after addition of EGF. EGFR was not substantially degraded in PBS-td or Pff-td cells in the absence of EGF (EGF -). Cq treatment inhibited EGFR degradation, confirming that the reduction of EGFR after EGF treatment was dependent on lysosome function. Statistical significance was determined with Student‟s t test. NS, ** indicate p>0.05, and p<0.01, respectively Scale bars = 20 µm.

Cathepsin-B substrate, Magic Red-RR(2), as well as the lysosomal degradation of the epidermal growth factor (EGF) receptor after addition of EGF, were assessed in Pff-td cells. Notably, lysosome function was maintained in these cells, as both processing of the Magic Red substrate (Figure 8B) and degradation of the EGFR (Figure 8C) was observed. The integrity of lysosomes in Pff-td cells was also supported by experiments which demonstrated that the enlarged lysosomes had a normal pH (as monitored by

Lysosensor; Figure 4-S3C), and that the lysosomal protease, Cathepsin D, could be properly processed from its Procathepsin D precursor (Figure S3D).

As lysosomal function in Pff-td cells appeared to be normal, we asked if autophagy impairment in α-syn aggregate-bearing cells might result from a defect in autophagosome maturation. This could be due to reduced autophagosome/lysosome fusion, or to a reduced number of fully matured autophagosomes that are available for lysosomal fusion. A step in autophagosome maturation that precedes lysosomal fusion is amphisome formation, during which autophagosomes fuse with multivesicular bodies

(MVBs) (Eskelinen, 2005). Utilizing the MVB marker CD63 (Escola et al, 1998), the localizations of CD63, LC3, Lamp1 and p-α-syn were compared in PBS-td and Pff-td cells. PBS-td cells showed light and diffuse staining for LC3 and CD63, with some co- localization of these markers (Figure 4-9A). However, inhibition of lysosome function caused accumulation of LC3 vesicles which largely co-localized with both CD63 and

Lamp1, demonstrating that these vesicles matured to amphisomes and autophagolysosomes before their clearance was blocked by the lysosomal inhibitors

(Figure 9A, B). LC3-positive vesicles accumulated and co-localized with p--syn in Pff-

Figure 0-11: Effect of Pff-td on Lysosomes

Figure 4-S3: Effect of Pff-td on Lysosomes. (A) 24 h after Pff transduction, naive HEK293 cells were treated for 24 h with a lysosomal inhibitor cocktail (Lys. Inh.) or vehicle (-).The IF images show that the PBS-td and Pff-td cells show similar lysosome morphology, with few giant lysosomes. Lysosomal inhibitor treatment resulted in an increase of giant (>2.5µm in diameter) Lamp1 (green)-positive vesicles, as indicated with arrowheads. Quantification of giant Lamp1 vesicles, shown as number of vesicles per 100 nuclei (blue), is shown on the right. (B) WT neurons were examined by IF 14 days after PBS or Pff transduction. Lamp1 (green) vesicles appeared as abundant, fine puncta in PBS-td cells, whereas in Pff-td cells they were typically enlarged and clustered with p-α-syn (red) aggregates. (C) HEK293 α-syn cells were incubated with lysosensor 48 h after PBS or Pff transduction to visualize lysosomal pH. Giant lysosomes in the Pff-td cells were brightly stained with lysosensor, indicating that they retained an acidic pH. Treatment with lysosomal inhibitors (top-right) eliminated the punctate lysosomal staining, demonstrating the pH dependence of the fluorescent dye. (D) HEK293 cells were treated with Cq or vehicle 48 h after PBS or Pff transduction. 24 h after treatment, cells were lysed and underwent IB analysis to determine the extent of processing of the lysosomal protease Cathepsin-D (Cath-D), which is cleaved from Procathepsin-D (Procath-D) in functional, acidic lysosomes (Hasilik et al., 1982). [n>=2 for all experiments. NS indicates p>0.05. Scale bars = 20 µm].

Figure 4-0-12: Autophagosomes associated with p-α-syn aggregates are impaired in maturation

Figure 9A: Autophagosomes associated with p-α-syn aggregates are impaired in maturation. 24 h after PBS or Pff transduction, HEK293 α-syn cells were treated for 24 h with a lysosomal inhibitor cocktail (Lysosome Inh.) or vehicle (-) before they were fixed for IF. (A) IF images showing CD63 (red), LC3 (green), p-α-syn (blue, white dotted outline), and nuclei (denoted as “Nu”). There were few LC3 and CD63 puncta in PBS-td cells, and these did not co-localize in the absence of lysosomal inhibitor. However, LC3 and CD63 co- localization was observed in the PBS-td cells when the clearance of autophagic vesicles was prevented by lysosomal inhibitor treatment. In Pff-td cells, LC3 vesicles that accumulate with p-α-syn aggregates were not generally co-localized with CD63 vesicles. Inhibition of lysosomes resulted in some overlap of CD63 and LC3, but CD63 did not localize to LC3 vesicles associated with p-α-syn. Although they did not co-localize with the p-α-syn aggregates, CD63 vesicles in Pff-td cells appeared to be larger and more variable in size compared to those in PBS-td cells[n>=3 for all experiments. Scale bars = 20 µm].

Figure 9B: Autophagosomes associated with p-α-syn aggregates are impaired in maturation. 24 h after PBS or Pff transduction, HEK293 α-syn cells were treated for 24 h with a lysosomal inhibitor cocktail (Lysosome Inh.) or vehicle (-) before they were fixed for IF. (B) IF images showing Lamp1 (red), LC3 (green), p-α-syn (blue, white dotted outline), and nuclei (denoted as “Nu”). Lamp1 co-localized with LC3 in PBS-td cells treated with lysosomal inhibitor, and in cells without α-syn aggregates in Pff-td cells after addition of lysosome inhibitor. However, Lamp1 was largely excluded from LC3 vesicles that overlapped with α-syn aggregates in Pff-td cells in the absence or presence of lysosome inhibitor. [n>=3 for all experiments. Scale bars = 20 µm].

78 td cells in the absence of lysosomal inhibition, but there was little overlap of LC3 with

CD63 (Figure 4-9A) or Lamp1 (Figure 4-9B) at the sites of p--syn aggregates. As with

PBS-td cells, inhibition of lysosomal function in Pff-td cells resulted in an increase of

CD63 and Lamp1 co-localization with LC3, with the exception that CD63 and Lamp1 staining was generally not observed in LC3-positive p--syn aggregates (Figure 4-9A,

B). Although there was minimal localization of CD63 with p-α-syn aggregates in Pff-td cells, either in the absence or presence of lysosomal inhibition (11.8%±2.6% and

6.7%±4.1%, respectively), the observed CD63-containing MVBs were not associated with -syn aggregates. However, they appeared to be enlarged, as was seen with Lamp1- positive vesicles (Figure 4-8A and Figure 4-9A). These data support the interpretation that the LC3-positive autophagosomes that associate with α-syn aggregates do not mature to amphisomes. Moreover, the enlarged LC3/CD63-positive amphisomes that do form in

Pff-td cells and which are not associated with -syn aggregates may have altered properties that could provide an explanation for the partial impairment of autophagic function observed in the cells harboring -syn inclusions.

Autophagy activation enhances cellular toxicity in -syn aggregate-bearing cells.

Previous studies have shown that activation of autophagy may attenuate cytotoxicity of aggregate-prone proteins that are degraded by autophagy (Sarkar et al,

2009; Young et al, 2009). However, our data suggest that the resistance of α-syn

Figure 0-13: Treatments that activate autophagy increase the toxicity of α-syn aggregates.:

Figure 4-S4: Treatments that activate autophagy increase the toxicity of α-syn aggregates. (A) A lactate dehydrogenase (LDH) release assay was performed to monitor cell death in naïve HEK293 cells or HEK293 cells that express α-syn. Only Pff transduced HEK293 α-syn cells (α-syn Pff), showed significantly increased LDH release. (B) PBS- or Pff transduced HEK293 α-syn cells were maintained in nutrient- rich complete media (comp), treated with rapamycin or maintained in starvation media after transduction, until a tetrazolium-based viability assay was carried out. Both starvation and rapamycin treatment significantly reduced viability in transduced cells (C,D) Pff- or PBS-td wt neurons were starved, or treated with rapamycin and the LDH release assay (C) and the viability assay (D) were performed. Starvation significantly increased cell death and decreased viability in neurons and rapamycin, trended towards the same pattern. [Error bars ±SEM. In (A) and (B), one-way ANOVA with Tukey‟s post-hoc analysis used to test for statistically significant differences. In (C) and (D), student‟s paired t test was used to test for statistically significant differences. NS, *, ** and *** indicate p>0.05, p<0.05, p<0.01 and p<0.001 respectively]

A 80 aggregates to autophagy is not the result of a defect in autophagy initiation, but rather a problem of autophagasome maturation. Thus, a strategy of autophagy activation may not be productive in cells containing -syn inclusions, and in fact the cell death that is observed in both aggregate-bearing HEK293 cells and neurons (Figure 4-S4A and(Volpicelli-Daley et al, 2011) is exacerbated upon activation of autophagy with rapamycin or serum-starvation (Figure 4-S4B-D). Although it is possible that the increase in toxicity observed with these treatments results from cellular mechanisms other than autophagy, these data are consistent with the notion that increased autophagosome generation can be detrimental in cells when autophagosome clearance is impaired

(Boland et al, 2008; Wong & Cuervo, 2010; Zhang et al, 2011).

4.4 Discussion

Utilizing recently developed cellular models of Pff-seeded α-syn aggregation, we have demonstrated that LB-like inclusions are resistant to degradation despite their interaction with ALP and UPS components. These aggregates persist even when soluble

α-syn levels are substantially reduced, indicating that once seeding occurs, pathologic α- syn aggregates are refractory to clearance from the cells. Interestingly, overall autophagic clearance is impaired in the α-syn aggregate-bearing cells, which may be a factor contributing to the observed reduction in cellular viability. Importantly, alterations of autophagy were observed in both non-neuronal cells and in primary cultured neurons harboring α-syn aggregates. Indeed, the neuronal Pff transduction model of α-syn aggregation is unique in that α-syn aggregates can be formed with very high efficiency in the absence of α-syn overexpression or toxic insults that may perturb autophagy or 81 proteasome function. Moreover, our cellular models of α-syn seeding with recruitment of endogenous α-syn to yield LB-like aggregates may replicate key aspects of synucleinopathies, as several recent lines of evidence suggest that the spreading of amyloid protein pathology, such as seen with α-syn in PD, may result from the neuronal release and uptake of extracellular α-syn seeds (Angot et al, 2010; Brundin et al, 2010;

Luk et al, 2012; Soto, 2012).

Previous studies revealed that LBs in PD co-localize with UPS and ALP components such as ubiquitin, 20S proteasome, p62 and LC3 (Higashi et al, 2011; Ii et al,

1997; Kuzuhara et al, 1988; Zatloukal et al, 2002). However, it was unclear whether these associations resulted in degradation of LBs. Although LB-like inclusions co-localized with UPS and ALP components in our cell models, such interactions did not to lead to degradation of α-syn aggregates. A similar recruitment of ALP components has also been observed with other aggregated proteins that are resistant to autophagic degradation

(Wong et al, 2008). The failure to degrade α-syn aggregates cannot be explained by a complete inhibition of autophagy, as Htt Q72 appeared to be degraded in aggregate- bearing cells, albeit at a significantly reduced rate, suggesting that α-syn aggregates are selectively resistant to degradation. As with a number of other protein aggregates subject to autophagy, Htt Q72 aggregates occur as “aggresomes”, which are believed to be actively generated by cells to sequester misfolded proteins within the cytoplasm

(Kaganovich et al, 2008; Kopito, 2000; Markossian & Kurganov, 2004). However, Pff- seeded α-syn aggregates do not show robust co-localization with aggresomal markers

82 such as vimentin (Kawaguchi et al, 2003; Luk et al, 2009), suggesting that they are a distinct type of inclusion that is resistant to degradation.

The demonstration that cells harboring aggregated α-syn have partial impairment of autophagy complements prior studies which showed that soluble α-syn species may reduce autophagic activity (Cuervo et al, 2004; Martinez-Vicente et al, 2008; Winslow et al, 2010). The α-syn inclusions appear to impair autophagy by reducing autophagosome clearance. This disruption of autophagy may contribute to the cellular toxicity seen in the aggregate-bearing cells, and we observed further cell death upon activation of autophagy.

Autophagosome clearance defects have been previously suggested in neurodegenerative diseases. For instance, the abnormal accumulation of autophagosomes in AD brain tissue is thought to be caused by impaired autophagosome clearance (Boland et al, 2008), and

LC3-II levels are elevated in PD brain tissues with abundant LB pathology (Higashi et al,

2011). An impairment of autophagosome clearance is likely to be detrimental, as removal of both unwanted, damaged proteins and defective organelles such as mitochondria

(Elmore et al, 2001; Novak et al, 2010), is vital for cells. Therefore, impairment of autophagy may lead to cell death through several different mechanisms, including oxidative stress, which is strongly implicated in PD pathogenesis (Jenner & Olanow,

1996; Zhang et al, 2000). (Jenner & Olanow, 1996; Zhang et al, 2000). Indeed, several proteins that are implicated in PD pathogenesis, including ATP13A2, LRRK2 and Pink-

1, may also affect autophagy function (Gomez-Suaga et al, 2012; Michiorri et al, 2010;

Usenovic et al, 2012), suggesting that autophagic dysfunction may be a primary cause of cell death in PD.

The LC3-positive autophagic vesicles that accumulate with α-syn aggregates were not decorated with markers of amphisomes and autophagolysosomes, indicating that autophagosome maturation is likely impaired prior to amphisome formation. The resistance of α-syn aggregates to degradation may be caused by an inability of aggregate- associated autophagosomes to migrate within the cell, and/or an inability of MVBs to fuse with the autophagosomes. Alternatively, it is possible that α-syn aggregates interfere with the ability of autophagosomes to mature, such that the LC3-positive structures that accumulate in aggregate-bearing cells are incomplete or abnormal. In addition to the - syn inclusions being refractory to autophagic degradation, it appears that there is a more general impairment of autophagy in aggregate-containing cells. Interestingly, the Pff-td cells show an enlargement of both CD63- and Lamp1-positive vesicular bodies, although lysosome function seems to be unaltered. These changes may suggest that the α-syn aggregates have an effect on vesicle-related cellular processes, including autophagy.

Indeed, α-syn aggregates perturb endosomal trafficking in yeast by interacting with rab proteins (Gitler et al, 2008; Soper et al, 2011).

In conclusion, we have demonstrated that seeded α-syn aggregates are resistant to degradation and impair autophagosome clearance. These findings have bearing on therapeutic strategies in PD and related synucleinopathies, as prior studies have indicated that autophagy activation via rapamycin treatment may attenuate disease phenotype in certain in vivo models of PD (reviewed by (Bove et al, 2011). Although the models used in these studies may provide important insights, they typically rely on oxidative insults or proteasome inhibitors to achieve the desired phenotype, and these treatments could

84 increase the cellular dependence on autophagic removal of damaged mitochondria and unwanted proteins (Lee et al, 2012; Ogata et al, 2006). In addition, neurons in these models lack the insoluble, LB-like α-syn aggregates that are observed in our cellular models and in synucleinopathies. Our studies suggest that activation of autophagy may not be effective once α-syn aggregates form, which is likely to occur before clinical symptoms manifest. On the contrary, autophagy activation may be harmful to -syn aggregate-bearing cells because of the impaired autophagosome clearance. However, it is possible that alternative therapeutic approaches, such as restoration of normal autophagic flux (Lee & Gao, 2009; Li et al, 2011), may reduce the ALP impairment caused by the α- syn aggregates and be effective in enhancing the survival of affected cells. Further studies will determine if these approaches can reduce neurodegeneration in PD and related synucleinopathies.

4.5 Materials and Methods

Mammalian and Primary Neuronal Cell Cultures

HEK293 cells (QBiogene, CA) stably expressing wt or A53T α-syn were generated as previously described (Luk et al., 2009). Cells were maintained in complete media [DMEM (Gibco, CA), supplemented with 10% FBS, penicillin/streptomycin and

L-glutamine], and 500 µg/ml, 100 µg/ml or no G418 (Gibco, CA) was added to media for wt α-syn, A53T α-syn and naïve cells respectively. The day before the experiment,

HEK293 cells were plated at a density of 60,000-75,000 cells/well on poly-D-lysine

(PDL, 0.1 µg/ml in dH2O) and swine gelatin (0.1% w/v in dH2O) coated 12 mm 85 coverslips, and 150,000-200,000 cells/well PDL coated on 12-well plates (Thermo-Fisher

Scientific, MA).

IB. Hela T-Rex A53T α-syn cells were maintained in Tet-Free complete medium

(DMEM, supplemented with 10% Tet-Screened FBS penicillin/streptomycin, L- glutamine) including 100 µg/ul G418 and 100 µg/ul hygromycin B. The day before the experiment, Hela T-Rex cells were plated at a density of 25,000-30,000 cells/well on

PDL coated coverslips, and 50,000-60,000 cells/well on 12-well plates. 1 µg/ml dox was added at the time of plating to induce α-syn expression.

Primary mouse hippocampal neurons were cultured and maintained as previously described (Volpicelli-Daley et al, 2011). Briefly, hippocampi dissected from E16-E18

C57BL/6 or CD-1 mouse brains (Charles River, MA) and α-syn KO mice (Abeliovich et al, 2000), were treated with papain and DNAse, and dissociated neurons were plated in plating media [Neurobasal Media (Gibco, CA), supplemented with 10% FBS, penicillin/streptomycin, Glutamax, B27] on PDL (0.1 mg/mL, in 0.1 M borate buffer pH8.4) coated coverslips at a density of 60,000-100,000 cells/well or PDL coated 12-well plates at 400,000-600,000 cells/well. Plating media was changed to neuronal media

(Neurobasal Media, supplemented with penicillin/streptomycin, Glutamax, B27) 2-18 h after plating. 86

Recombinant α-syn proteins and generation of α-syn Pffs

α-Syn Pffs were generated by incubating purified α-syn (5 mg/mL in PBS) at

37°C with constant agitation for 5 days. Electron microscopy, Thioflavin S, and protein sedimentation assays confirmed generation of insoluble amyloid fibrils at the end of incubation (Giasson et al., 2001). Generated Pffs were aliquoted and stored at −80°C.

Pff Transduction

Unless indicated otherwise, HEK293 or Hela T-Rex cells expressing A53T α-syn, and α-syn Pffs generated from C-terminally myc tagged and truncated α-syn (1-120) were used for non-neuronal Pff transductions. By using this combination of cells and Pffs, a

4.7-fold increase in percentage of cells bearing aggregates (78.5% ±0.7% vs

16.7%±3.5%, 48h after transduction) was elicited compared to wt α-syn cells wt α-syn

Pffs combination. Cells were washed with Opti-MEM (Gibco, CA), before transduction.

For dox-induced Hela cells, cells were rinsed 3 times with Opti-MEM for 5 min each to completely remove dox. 0.8 µg of sonicated α-syn Pffs in 80 µl dPBS (Cellgro, MA)

87 were added to a tube of protein transduction reagent Bioporter (Sigmα-Aldrich, MO), and incubated 10 min at room temperature. After incubation, 420 µl of Opti-MEM was added to the reagent tube and one tube of diluted reagent/Pff complex was used to transduce 4 wells on a 12-well plate or 8 coverslips. After 4 h, Opti-MEM with reagent/Pff complex was removed and cells were maintained in low-serum starvation media (DMEM, 0.25%

FBS supplemented with penicillin/streptomycin, L-glutamine) unless indicated otherwise.

In neurons, wt α-syn Pffs were diluted in PBS at 100 µg/mL, sonicated 65 times, and diluted in neuronal media. For coverslips 0.5 µg and for 12-well plate wells 5 µg of α-syn

Pffs were added directly to culture media at 5 or 10 DIV. Neuronal media of transduced cells was not removed earlier than 3 days after transduction. Where indicated, neurons were starved in starvation medium (DMEM, penicillin/streptomycin) to activate autophagy (Young et al, 2009).

Drug Treatments

2.5 mg/ml Rapamycin in DMSO (Sigma-Aldrich, MO) was stored at -20oC, and was diluted 1:10 in DMSO before each experiment. Lysosome inhibitor cocktail components, Leupeptin hemisulphate, Pepstatin A, and E46d (all from Sigma-Aldrich,

MO) were prepared at: 20 mM in dH2O, 2 mM in DMSO and 10 mM in DMSO,

o respectively, and kept at -20 C. 5 M ammonium chloride (NH4Cl, Thermo-Fisher

Scientific, MA), 200 mM chloroquine (Sigma-Aldrich, MO) and 100 mM 3- methyladenine (Sigma-Aldrich, MO) solutions were prepared in dH2O immediately before each experiment. Working concentrations of each drug are described in the figure legends. 88

Plasmids and Transfections

The pYFPN1-Htt-Q72 plasmid was a generous gift of Dr. Robert Baloh. HEK293 transfections were carried out using Fugene HD transfection reagent (Promega, WI) as per manufacturer‟s instructions. Neurons were transfected with LC3-RFP and α-syn-GFP plasmids using Lipofectamine 2000 (Invitrogen, CA) according to manufacturer‟s instructions. Transfection was done at DIV5 and neurons were transduced at DIV6. Live images are acquired using Nikon Eclipse TE2000-E microscope equipped with CoolSnap

HQ camera (Roper Scientific) and NIS Elements (Nikon Inc, NY). For siRNA transfections, Lipofectamine siRNAMAX (Invitrogen, CA) was used according to manufacturer‟s instructions. The sequence of the anti-mouse-α-syn siRNA duplex was:

5‟-GCAUGAGACUAUGCACCU-3‟ and 5‟-UAUUUAUAGGUGCAUAGU-3‟.

Transfection was carried out at DIV6, 1 day after transduction.

Lysosomal function assays

Magic Red Cathepsin MR-(RR)2 (Immunochemistry Technologies, MN) was used to monitor Cathepsin-B activity in HEK293 cells, according to manufacturer‟s intstructions. Briefly, 48h after transduction, cells were incubated with the Magic Red reagent for 15 mins, washed in imaging buffer (10mM HEPES, 136mM NaCl, 2,5mM

KCl, 2mM CaCl2, 1.3mM MgCl2, 10mM glucose) and the fluorescent signal generated by proteolytic activity was observed using a fluorescent microscope. For EGFR degradation assay, 48h after transduction, HEK293 cells were incubated with 200nm/ml murine EGF

(Peprotech, NJ) or BSA for 30, and 120 minutes. As a positive control, a set of EGF

89 treated wells were pre-incubated with 100uM chloroquine for 1h. At the end of incubations, cells are harvested in 2% SDS lysis buffer for immunoblotting.

Immunofluorescence

Cells were fixed with 4% paraformaldehyde in PBS followed by permeabilization with 0.1% TX. For LC3 and Lamp1 staining, digitonin (100 µg/ml) used for permeabilization. For 20S proteasome staining, cells were permeabilized during fixation with 1% TX to reduce nuclear staining. After blocking with Bovine serum albumin

(BSA, 3% w/v in PBS), cells were incubated in primary antibodies (Table 1) followed by

Alexa fluor-conjugated secondary antibodies (Invitrogen, CA). Cells on coverslips were incubated with DAPI and mounted on glass slides in Fluoromount-G (SouthernBiotech,

AL). For neurons, 4% w/v sucrose included in the fixation solution. Images were acquired using an Olympus BX 51 microscope equipped with a digital camera DP71 and

DP manager (Olympus, PA), except for Figures 6A and 8A, where images are acquired using Nikon Eclipse TE2000-E microscope equipped with CoolSnap HQ camera (Roper

Scientific) and NIS Elements (Nikon Inc, NY).

Immunoblotting

For sequential extractions, cells were scraped into 1% TX in Tris-buffered saline

(TBS) (50 mM Tris, 150 mM NaCl, pH 7.4) and protease/phosphatase inhibitor cocktail at 4°C. Lysates were sonicated and centrifuged at 100,000xg for 30 min. The pellet was washed and suspended in 2% SDS in TBS. For total protein extractions, cells were scraped and sonicated directly in 2% SDS in TBS. Protein concentrations were estimated 90 by BCA assay (Thermo-Fisher Scientific, MA) and equal amounts of protein were loaded on gels for each sample unless indicated otherwise. For sequential extractions, protein concentrations of TX fractions were used to calculate the volume of lysate to be loaded.

(Jackson Immunoresearch, PA), WesternLightning ECL substrate (Perkin Elmer, MA) and LAS-3000 imager (Fuji; Tokyo, Japan), or by use of IRDye secondary antibodies and

ODY-2816 Imager (Li-Cor Biosciences, NE).

Immuno Electron Microscopy

Immuno-EM were conducted as described before (Volpicelli-Daley et al, 2011).

Shortly, 14 days after Pff-treatment, neurons were fixed in periodate-lysine- paraformaldehyde and permeabilized with 0.05% saponin in PBS with 2% fish gelatin and 0.05% thimerosal followed by incubation in p-α-syn antibody. Neurons were then

91 incubated with goat anti-mouse IgG coupled to nanogold (Nanoprobes, Yaphank, NY).

The nanogold labeled neurons were postfixed and gold toned with 0.05% gold chloride.

Quantifications, Image Processing, Statistics

Colocalization of p-α-syn aggregates and inclusions of Pdp related proteins was quantified by manual counting using Image J (NIH, USA). Inclusion was defined as a cytoplasmic region with clearly increased IF signal compared to surrounding cytoplasmic areas of the same cell. IF pictures from 3 or 4 independent experiments were used for quantification, a minimum of 100 p-α-syn aggregates was counted for each condition. IB signals were quantified by Multi Gauge V2.3 (Fujifilm; Tokyo, Japan) or Image Studio

V2.0 (Li-Cor Biosciences, NE). Acquired images processed with Adobe Photoshop CS2

(Adobe Systems Inc., CA) to facilitate visualization where needed. The adjustments applied equally across the entire image and to controls. All experiments were repeated a minimum of 3 times. The number following “±” sign indicates SEM. Statistical analyses used for each experiment were indicated in figure legends, all analyses were carried out using Graphpad Prism 4 (Graphpad Software Inc, CA).

4.6 Supplementary Materials and Methods

Drug Treatments, Plasmids and Transfections

C-lac (Calbiochem; Darmstadt, Germany) was dissolved in DMSO at 10 mM, and stored at -20oC. pmStrawberry-Atg4BC74A (Addgene plasmid 21076) (Fujita et al, 2008) was generated by Dr. Tamotsu Yoshimori and deposited to Addgene. Transfection was

92 carried out using Fugene HD transfection reagent (Promega, WI) as per manufacturer‟s instructions. Lamp1-RFP plasmid (Addgene plasmid 1817)(Sherer et al, 2003) was generated by Dr. Walther Mothes and deposited to addgene with a comment indicating that Lamp1 sequence in this plasmid contains two point mutations which are not important for function. Neurons were transfected with Lamp1-RFP plasmid using

Lipofectamine 2000 (Invitrogen, CA) according to manufacturer‟s instructions.

Transfection was done at DIV5 and neurons were transduced at DIV6. Live images are acquired using Nikon Eclipse TE2000-E microscope equipped with CoolSnap HQ camera (Roper Scientific) and NIS Elements (Nikon Inc, NY).

Toxicity Assays

CytoTox 96® Non-Radioactive Cytotoxicity Assay (Promega, WI) was used to monitor LDH release as an indicator of cytotoxicity and the CellTiter 96® AQueous One

Solution Cell Proliferation Assay (Promega, WI) was used to monitor MTS tetrazolium reduction to cell estimate viability, according to manufacturer‟s instructions. HEK293 cells were treated for 48 h with rapamycin 48 h after transduction. Complete media

(“Comp”) indicates that transduced cells were incubated in nutrient-rich media, starting from 4 h after transduction, for 92 h. For neurons, 7 days after transduction, cells were starved in DMEM or treated with rapamycin for 7 days. For viability assays, HEK293 cells and neurons are incubated with MTS tetrazolium substrate for 1 h, and 4 h respectively. Results are shown as ratios of Pff and PBS-td cells, that received the same treatment.

Acknowledgements

We are grateful to D.M. Riddle, V. Kelm and A. Stieber for their technical assistance. We thank

J.Q. Trojanowski, T. Cohen, K. Luk and J. Guo for their critical reading of the manuscript. We

also thank R. Baloh (Cedars-Sinai Medical Center, CA) for generously providing pYFPN1-Htt-

Q72 plasmid. This work was supported by National Institutes of Health Grants NS053488, the

JPB Foundation, the Benaroya Foundation, and the Jeff and Anne Keefer Fund.

Table 1: Antibodies used in this study

Antigen Antibody Application Dilution Source

20S Proteasome PW 8155 IF 1:2000 Biomol

alpha-tubulin T9026 IB 1:5000 Sigma-Aldrich

α-syn (human) Syn211 IB 1:2000 (Bruening et al, 2000)

α-syn (mouse) mSyn-1 IB 1:500 (Vopicelli-Daley et al., 2011)

CD-63 CD-63 IF 1:500 (Abache et al, 2007)

GFP sc-9996 IB 1:500 Santa Cruz

Lamp1 (human) H4A3 IF,IB 1:1000, 1:1000 BD biosciences

Lamp1 (mouse) 1D4B IF 1:500 Developmental Studies Hybridoma LC3 PM036 IF, IB 1:2000, 1:500 MBL

myc 9E10 IF 1:2000 DHSB

p62 (human) 610832 IF, IB 1:2000, 1:1000 BD Transduction

p62 (mouse) 2C11 IF, IB 1:1000, 1:1000 Abnova

p-α-syn 81a IF, IB 1:8000, 1:1000 (Waxman & Giasson, 2008)

p-α-syn p-α-syn 6.2 IF 1:1000 (Luk et al, 2012)

Tuj-1 MMS-435P IB 1:5000 Covance

Ubiquitin Ub1B4 IB, IF 1:1000, 1:1000 (Sampathu et al, 2006)

Figure 0-14: Antibodies used in this study:

Chapter 5: General Conclusions and Discussion

In this study, we investigated the interplay between LB-like α-syn aggregates and

Pdps, using a novel cell based model of Parkinson‟s disease. This study is unique in the sense that not only the α-syn aggregates investigated are very similar to LBs and LNs observed in brains of those who suffer from synucleinopathies, but also the method used to generate them may accurately recapitulate their formation in vivo. Furthermore, unlike common methods used to induce formation of α-syn inclusions, Pff-seeding does not alter

Pdp function, thereby rendering this model very suitable to study the interplay between α- syn aggregates and Pdps. Thus, we believe that our findings will help improve our understanding of the role of LBs in pathogenesis. In the rest of this chapter, we will further elaborate the discussion on our findings and speculate about the possible mechanism of autophagy impairment, the differences between LB-like seeded aggregates and non-fibrillar α-syn inclusions and possible therapeutic implications of our findings.

5.1 Possible mechanisms of α-syn aggregate mediated impairment of autophagy

The autophagic vesicles that accumulate with the aggregates only contained early autophagosome markers, which may indicate that autophagosome maturation is impaired.

It is possible that this impairment is directly caused by diminished autophagosome transport due to sequestration of the autophagosomes by α-syn aggregates. Another

95 possibility is that the LB-like aggregates affect development of the phagophore, so that the autophagosomes that surround the aggregates are abnormal or incomplete. It is possible that the affinity of α-syn to membranes may interfere with this membrane based process. However, abnormal lysosome/late endosome morphology we observed suggests that the aggregates may also have an indirect effect on autophagy, or vesicle trafficking in general. Indeed, it was previously shown that golgi is dispersed in Pff-td HEK293 cells

(Luk et al, 2009), and α-syn aggregates perturb endosomal trafficking in yeast by interacting with rab proteins (Gitler et al, 2008; Soper et al, 2011). It is also possible that the synaptic deficits observed in Pff-td neurons (Volpicelli-Daley et al, 2011) are contributed by defects in vesicle trafficking. Interestingly, our findings are remarkably similar to the effects of PI(3,5)P2 deficiency, which is implicated in human neurodegenerative diseases, particularly ALS (Chow et al, 2009; Chow et al, 2007).

Reduction of this signaling lipid causes impaired autophagosome clearance (Ferguson et al, 2009), formation of giant lysosomes/late endosomes but does not significantly affect lysosomal proteolytic function such as EGFR degradation (Ikonomov et al, 2003). Thus,

α-syn aggregate mediated inhibition of a single protein that regulates PI(3,5)P2 levels, or a downstream effector of this signaling lipid, could cause the phenotype we observed in

α-syn aggregate bearing cells. Furthermore, the α-syn aggregates appear to be completely resistant to autophagy, whereas other autophagy substrates can still be degraded at slower rates. Therefore, it is not clear if a single defect is responsible of the failure of autophagy in degrading α-syn aggregates and the general impairment in autophagy. One could speculate that physical sequestration of the autophagosomes, or formation of defective/incomplete autophagosomes may be responsible for the former impairment, 96 whereas defects in vesicle trafficking contributes to the latter. Further studies are needed to investigate these pathways to elucidate the molecular mechanism of autophagosome impairment caused by α-syn aggregates.

5.2 LB-like seeded aggregates and non-fibrillar α-syn inclusions

The non-fibrillar smaller inclusions that are seen in several in vivo α-syn overexpression models may also form in human brain, particularly in regions where α- syn is concentrated, such as synapses (Kramer & Schulz-Schaeffer, 2007). However, being susceptible to degradation these structures may not be readily detected. LBs and

LNs may be a completely distinct group of aggregates, initiated by cell-to-cell transmission of α-syn seeds, and being resistant to degradation, these aggregates constitute the prominent lesions in synucleinopathies. It is postulated that α-syn aggregates are neuroprotective, or inert, and can be easily degraded by autophagy.

However, these claims are mostly based on the data obtained from studies on non-fibrillar

α-syn inclusions. Unlike many others, our α-syn aggregation models recapitulate LB-like

α-syn aggregate formation, without forcing formation of other forms of α-syn accumulations. A major difference between these two types of α-syn accumulations is the former are usually generated as a result of “aggresomes response” through which misfolded proteins are actively sequestered and packaged by the cells to facilitate their degradation by autophagy (Kopito, 2000). In addition, they are likely to include many other proteins accumulated during the efforts to make α-syn accumulate, thus it is not clear if these inclusions are indeed α-syn inclusions or inclusions that merely contain α- syn. On the other hand, it is likely that seeded α-syn aggregates do not rely on 97

“aggresomes response” to form in the cells, since it is clearly shown that purified recombinant α-syn can form aggregates that are ultrastructurally and biochemically very similar to LBs (Conway et al, 1998). Moreover, seeding with α-syn confers selectivity, and it did not cause accumulation of other aggregate prone proteins such as TDP-43 (data not shown), therefore seeded aggregates should basically be aggregates of α-syn including other proteins, not aggregates of other proteins including α-syn. This could explain why they are more similar to LBs, than aggresomes-like α-syn inclusions.

Therefore, we believe our study provide a unique insight into possible impact of LBs on cellular pathways, and their susceptibility to removal by autophagy.

5.3. Insights into the therapeutic implications of our study

We believe that activation of autophagy can be a promising potential preventative therapy for synucleinopathies. It may reduce levels of monomeric α-syn, soluble oligomers and other types of α-syn accumulations, and possibly help delay LB formation by reducing available α-syn to be recruited. However, we also believe that activation of autophagy may not be a favorable approach after accumulation of LB and LN-like aggregates, which typically occurs before clinical symptoms manifest. At that point, autophagosome clearance may be impaired, and as previously discussed; activation of this pathway may exacerbate neurodegeneration under these conditions. Indeed, this result was not only observed by us in our cell based models, but also in an in vivo model of SOD-1 aggregation. SOD-1 mutations are implicated in amyotrophic lateral sclerosis

(ALS), a lethal motor neurodegenerative disease (Deng et al, 1993), and in a mouse model of SOD-1 aggregation, it was observed that autophagosome clearance is reduced 98

(as indicated by accumulation of LC3-II and p62), and treatment of these mice with autophagy activator rapamycin does not help remove aggregates, but also exacerbates mitochondrial impairment, and augment neurodegeneration (Zhang et al, 2011).

Our findings indicate that once they are formed, LB-like α-syn inclusions are refractory to removal; however, this does not indicate that their detrimental effects on the cells cannot be stopped. Elimination of the autophagy impairing effects of the aggregates may render the aggregates relatively inert, and despite their continued existence in the cells, they would have reduced cytotoxicity. For example, if recruitment of autophagosomes to the aggregates is stopped, perhaps by elimination of p62, autophagy function may not be affected by the aggregates. Alternatively, treatments that reduce autophagic activity may be beneficial if the cells have overactive autophagy that could be caused by a number of reasons such as age related impairment of other Pdps, and PD- linked LRRK2 mutations (Bravo-San Pedro et al, 2012). In addition, treatments that would improve vesicular trafficking, such as expression of rab proteins, or restoration of

PI(3,5)P2 levels could be beneficial if these factors are affected by α-syn aggregates.

Therefore, it is very important to elucidate the underlying mechanism of autophagy impairment caused by LB-like α-syn aggregates.

5.4 Insights into the possible interplay between LBs, autophagy and mitochondrial function in neurodegeneration

Recent studies suggest that role of autophagy in PD pathogenesis may be more important than initially thought. Although the evidence indicating that mitochondrial

99 dysfunction has a primary role in PD pathogenesis is unequivocal, autophagy may be another primary player. Inhibition of autophagy in mouse neurons can cause neurodegeneration (Hara et al, 2006; Komatsu et al, 2006). For example, mice lacking atg5, a gene essential for autophagosome nucleation (Mizushima et al, 2001), in brain tissue, accumulates insoluble, ubiquitinated protein aggregates, followed by neuron loss through mechanisms including apoptosis (Hara et al, 2006). Moreover, autophagy deficient mice also show accumulation of deformed mitochondria and peroxisomes in liver, supporting the in vitro evidence that autophagy is crucial in clearing damaged organelles which could otherwise have cytotoxic effects (Komatsu et al, 2005). Finally, neurodegeneration is a common symptom of lysosomal storage disorders, which are caused by lysosomal defects affecting all members of the ALP pathway, including autophagy (Settembre et al, 2008). These findings support that similar to mitochondrial dysfunction, impaired autophagy may also cause neurodegeneration.

If impaired autophagy and mitochondrial dysfunction are the major mechanism of neurodegeneration in PD and related synucleinopathies, risk factors for PD should have a direct or indirect effect on these toxic defects. Indeed, age, a major risk factor for PD, impairs mitochondria (Conley et al, 2007). Aging may also stress and activate autophagy by causing a decline in other Pdps, thereby increasing the dependence on autophagy to remove unwanted proteins (Farout & Friguet, 2006; Kiffin et al, 2007; Ma et al, 2011).

Moreover, all genes whose mutations are linked to PD have roles in at least one of these pathways (Figure 5-2). ATP13A2 is important for lysosomal function (Usenovic et al,

2012). Mutations in the lysosomal protein GBA can increase the risk of PD, and recessively inherited loss of function this protein causes a lysosomal strorage disease 100

(Gaucher‟s disease), indicating that this protein is important for ALP function (Nichols et al, 2009). Pink-1 and Parkin are not only involved in mitochondrial function but also crucial in autophagy mediated removal of defective mitochondria (mitophagy) (Narendra et al, 2008; Vives-Bauza et al, 2010). As well as the mitochondrial toxins commonly used to mimic parkinsonism (Gorell et al, 1998; Langston et al, 1999; Liou et al, 1997), DJ-1 loss of function mutations also impair mitochondria (Guzman et al, 2010; Hao et al,

2010). Finally, as we show here, the pathological hallmarks of PD, LBs, impair autophagy, which may cause cell death directly, or indirectly by causing accumulation of defective mitochondria. Interestingly, the most frequent LRRK2 mutation G2019S, which is observed in almost 40% of familial PD cases in certain populations, shown to activate autophagy (Bravo-San Pedro et al, 2012), and deletion of this gene, whose mutations are the most common cause of familial PD, reduces autophagic activity in aged mice (Tong et al, 2010) and also slows down pathogenesis in a A53T α-syn aggregation model (Lin et al, 2009). Thus, although many of these proteins, as well as α-syn, have also been implicated with processes unrelated to autophagy or mitochondrial function, with recent studies (some reviewed in(Lachenmayer & Yue, 2012), including ours, the evidence converged on these two pathways. Thus, impaired macroautophagy may be a primary cause of the neurodegeneration in PD and related synucleinopathies (Figure 5-1).

What we did not discuss thus far is why a subset of neurons in PD, including dopaminergic neurons of SNc, and noradrenergic neurons of locus ceruleus, are much more vulnerable to degeneration than the other CNS neurons. This topic is still debated; one of the earliest theories was that the presence of dopamine in dopaminergic cells renders these cells vulnerable. For one, a number of studies provided evidence suggesting 101 that α-syn plays a role in the regulation of dopamine by modulating its production (Perez et al, 2002; Wu et al, 2011) and release (Larsen et al, 2006; Senior et al, 2008; Yavich et al, 2004), such that pathological changes in α-syn function may disrupt dopamine homeostasis. Moreover, oxidized dopamine derivatives can be toxic, rendering dopaminergic cells sensitive to mitochondrial dysfunction/oxidative stress (Anderson et al, 2011). An oxidized dopamine derivative, dopaminochrome, is also shown to interact with α-syn, and stabilize potentially toxic oligomeric species while inhibiting fibrillization (Norris et al, 2005). Additionally, it was suggested that α-syn forms soluble protein complexes that are normally neuroprotective, but these complexes increase the rate of apoptosis in the presence of dopamine (Xu et al, 2002). However, dopamine is not likely the only factor causing selective degeneration in PD. Firstly, many neurons which are lost in this disease, some before SNc neurons, such as noradrenergic locus ceruleus, histaminergic tuberomammillary nucleus (Del Tredici & Braak, 2004; German et al,

1992; Surmeier et al, 2011) do not have dopamine. Second, in mice, unless α-syn expression is specifically targeted to dopaminergic neurons, SNc is often spared while α- syn aggregation and neuron loss can be observed in many brain regions (Hashimoto et al,

2003), suggesting that presence of dopamine may even be protective against α-syn pathology by inhibiting aggregation. Lastly, dopamine replacement therapies, such as administration of dopamine precursor L-dopa, do not promote disease progression (Fahn,

2005). Arguably a better supported theory attempting to explain the selective vulnerability in PD, relies on the observation that the vulnerable neurons have autonomous pacemaking activity and are decorated with L-type Ca+2 channels, enabling

Ca+2 influx coupled with this activity (Surmeier et al, 2011). Intracellular Ca+2 levels 102

Figure 0-1: A hypothetical model of neurodegeneration in PD

Figure 5-1: A hypothetical model of neurodegeneration in PD This model is based on the hypothesis that impairments in autophagy and mitochondrial function are two major contributors of cell death observed in PD. Once formed, LBs may impair autophagosome clearance and this impairment can cause cell death directly by accumulation of misfolded and damaged proteins, and indirectly by impairing mitophagy resulting in increased numbers of damaged mitochondria that generate free radicals. This defect can result in oxidative stress, and elevation of intracellular Ca+2 levels in dopaminergic cells, which both activates autophagy, further damaging the LB bearing cells that are unable to efficiently clear autophagosomes. Factors that can directly affect mitochondrial function (such as mitochondrial toxins, mutations in DJ- 1, Parkin and Pink-1, and possibly α-syn oligomers), mitophagy (mutations in Parkin and Pink-1), autophagy (mutations in ATP13A2, GBA), may also cause cell death independent of LBs. Mutations in LRRK2 may contribute to LB formation, as well as LB mediated cell death by activating autophagy. For simplicity, only select interactions are shown. It is likely that the actual interplay between these mutations and pathways is more complicated, considering that decades of relentless research on PD provided only modest progress in our understanding of the disease.

103 are to be kept exceedingly low at all times (100nM vs 10mM in extracellular space), since it is used as an signaling molecule and can form Ca3(PO4)2 precipitates at high concentrations. Restoration of low Ca+2 levels after Ca+2 influxes requires vast amounts of energy, which could put significant stress on mitochondria (Guzman et al, 2010;

Wilson & Callaway, 2000). Moreover, along with oxidative stress , elevated intracellular

Ca+2 also activates autophagy (Harr & Distelhorst, 2010), which might further increase the vulnerability of the neurons to LB-mediated autophagosome clearance impairment.

Thus, it is possible that with their stressed mitochondria, and increased abundance of oxidatively damaged α-syn, vulnerable neurons are more prone to form de novo α-syn fibrils or seeded LBs. Moreover, once formed, impairment of autophagosome clearance by LBs would be particularly destructive to these vulnerable neurons, because of their activated autophagy, and their dependence on robust mitophagy to maintain a healthy pool of mitochondria. Impaired mitophagy may cause reduction in energy reserves, and inefficient Ca+2 removal, that would further activate autophagy, resulting in a vicious cycle consequently leading to cell death.

5.5 Concluding remarks

In this study, we investigated the interplay between LB-like α-syn aggregates and autophagy, and based on our findings, we believe that these aggregates significantly contribute to neurodegeneration by impairing autophagic clearance. We hope that our work constituted a primer for future studies that will further investigate the underlying mechanisms of this effect, thereby helping provide new therapeutic targets for these devastating diseases. 104

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