tspace.library.utoronto.ca€¦ · ii Structural characterization of amyloid assemblies and amyloid...

Structural Characterization of Amyloid Assemblies and Amyloid Protein – Lipid Interactions

by

Jason Ho-Lun Yau

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Department of Biochemistry University of Toronto

© Copyright by Jason Ho-Lun Yau 2016

ii

Structural characterization of amyloid assemblies and amyloid

protein-lipid interactions

Jason Ho-Lun Yau

Doctor of Philosophy

Department of Biochemistry

University of Toronto

2016

Abstract

Amyloids are insoluble aggregates rich in β-structure that result from the self-assembly of

polypeptides in a non-native structural fold. The deposition of these aggregates onto organs has

been traditionally linked to many human diseases, however the discovery of functional amyloids

in nature suggests there may also be beneficial roles for this self-assembly process. Pathogenic

and functional amyloids both contain a cross-β backbone composed of β-sheets that run

perpendicular to the fibril axis, but there lacks clear molecular details on the structural

differences to explain their divergent biological behaviours. Solid-state NMR (SSNMR) has

become an invaluable tool for structural determination of amyloids, as poor solubility of these

assemblies make them challenging for traditional X-ray crystallography and solution NMR

methods. This thesis aimed to identify key structural features between cytotoxic and non-

cytotoxic amyloid fibrils using SSNMR, and to examine the effects of protein-lipid interactions

on membrane disruption and amyloid formation that can reflect amyloid cytotoxicity. First, three

amyloid fibrils formed from fragments of the human prion protein were examined, with the

findings that a parallel β-sheet arrangement and packing of hydrophobic residues into the fibril

core defined the basic structure in these assemblies. Next, these three peptide amyloids were

sonicated to explore fragmentation-associated cytotoxicity of these amyloids. The results here

iii

highlighted the importance of solvent-exposed hydrophobicity on amyloid-membrane

interaction, linking the increased surface hydrophobicity of fragmented fibrillar species to greater

membrane disruption and cytotoxicity. Finally, SSNMR experiments were applied to

characterize the structure of human apolipoprotein serum amyloid A when associated with high-

density lipoprotein (HDL)-like particles and when misfolded into amyloid fibrils. The results led

to a preliminary model that can explain functional differences between lipid-free and HDL-

associated SAA. Together, these findings highlight the differing effects of lipid interaction on the

pathogenicity of amyloid fibrils.

iv

Acknowledgments

First off, I would like to thank my supervisor Dr. Simon Sharpe for his strong positive impact

and constant guidance throughout these past years. From my 4th year undergraduate research

project to my graduate research, Simon has given me the freedom to explore and broaden my

scientific horizons. He has stimulated my passion for science with many interesting research

projects, for which I am most grateful to have been working under his supervision. It has been a

memorable journey growing as a young scientist.

I would also like to thank my graduate committee members Dr. JoAnne McLaurin, Dr. Régis

Pomès, and Dr. Christopher Yip for their thoughtful advice and critique of my work, allowing

me to improve the quality of my research and expand my knowledge.

A great many thanks to the lab family, especially Karen, who I joke is my lab mommy. Without

her constant support and friendship, I would never have gotten this far. Patrick, Dave, and Greg,

who have made my day-to-day research less boring, thank you for sharing all the fond memories

inside and outside of the lab with me and introducing me to the world of coffee. Sean, Lisa, and

Aditi, thanks for the amazing scientific discussions about things that stretch and recoil. My team

of undergrad students Sympascho, Sam, Simoun, Richard, thanks for all the hard work.

My close high school friends who keep me sane and remind me of the life outside of science, as

well as all the friends I’ve met along the way in judo, volleyball, soccer, ultimate Frisbee, and on

the PGCRL 20th floor – thanks.

Finally I must thank my family for their support and patience throughout my many years of

study. My parents, for teaching me to strive for excellence. My sister, for being there to listen

and share a laugh. My grandfather, for his love and support. Thank you all for your unwavering

faith in me.

v

Table of Contents

Acknowledgments ........................................................................................................................ iv

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

List of Tables ................................................................................................................................ ix

List of Figures .................................................................................................................................x

List of Equations ........................................................................................................................ xiii

List of Appendices ...................................................................................................................... xiv

Abbreviations ...............................................................................................................................xv

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

1.1 Protein misfolding and amyloids .........................................................................................2

1.1.1 Protein folding .........................................................................................................2

1.1.2 Amyloids and diseases .............................................................................................3

1.1.3 Functional amyloids .................................................................................................5

1.1.4 Amyloid structure ....................................................................................................6

1.1.5 Kinetics of amyloid assembly ..................................................................................9

1.1.6 Amyloid fibrils and oligomeric intermediates .......................................................10

1.2 Amyloid protein-lipid interaction ......................................................................................12

1.2.1 Lipid interactions with amyloidogenic proteins ....................................................12

1.2.2 Membrane-induced fibril formation ......................................................................12

1.2.3 Membrane disruption and amyloid toxicity ...........................................................13

1.2.4 Fibril fragmentation and toxicity ...........................................................................15

1.3 Solid-state nuclear magnetic resonance spectroscopy .......................................................16

1.3.1 Background on NMR .............................................................................................17

1.3.2 Anisotropy in SSNMR and magic angle spinning (MAS) .....................................18

1.3.3 Chemical shift and secondary structure .................................................................20

1.3.4 Polarization transfer and SSNMR experiments .....................................................21

1.3.5 Solid-state NMR models of amyloid fibrils ...........................................................24

1.4 Mammalian prion protein (PrP) .........................................................................................24

1.4.1 Prions and diseases ................................................................................................24

1.4.2 Prion propagation and prion strains .......................................................................27

1.4.3 Structural studies of PrP peptides ..........................................................................27

1.5 Serum Amyloid A ..............................................................................................................29

1.5.1 Acute-phase serum amyloid A and amyloid A (AA) amyloidosis ........................29

1.5.2 Structure of SAA....................................................................................................30

1.5.3 High density lipoproteins (HDLs) .........................................................................33

1.5.4 Glycosaminoglycans (GAGs) and AA amyloidosis ..............................................34

1.6 Rationale and Hypotheses ..................................................................................................35

1.6.1 Structural characterization of prion peptide fibrils ................................................36

1.6.2 Structural changes of cytotoxic fibril fragments ....................................................37

1.6.3 Structural characterizations of human SAA2 ........................................................37

vi

Chapter 2. Structural comparison of three amyloidogenic hexapeptides from the

human prion protein ...............................................................................................................39

2.1 Introduction ........................................................................................................................40

2.2 Methods..............................................................................................................................41

2.2.1 Synthesis of uniformly and selectively labeled peptides .......................................41

2.2.2 Preparation of amyloid fibrils ................................................................................42

2.2.3 Transmission electron microscopy (TEM) and X-ray fibre diffraction .................43

2.2.4 Thioflavin T (ThT) Fluorescence...........................................................................44

2.2.5 FTIR measurements ...............................................................................................44

2.2.6 Solid-state NMR measurements ............................................................................44

2.2.7 SPINEVOLUTION simulations ............................................................................46

2.2.8 Structure calculations .............................................................................................47

2.3 Results and Discussion ......................................................................................................47

2.3.1 Biophysical characterization of PrP peptide fibrils ...............................................47

2.3.2 Chemical shift assignments of 13C, 15N-labelled peptides .....................................51

2.3.3 The prion peptide amyloid fibrils all contain parallel, in register β-sheets ...........57

2.3.4 Quaternary contacts within PrP(245-250) fibrils ...................................................58

2.3.5 High-resolution structure of PrP(178-183) peptides within amyloid fibrils ..........63

2.3.6 Quaternary contacts within PrP(244-249) fibrils ...................................................67

2.3.7 Fibril structures of PrP(178-183), PrP(244-249) and PrP(245-250) ......................70

2.4 Discussion ..........................................................................................................................75

Chapter 3. Fragmentation-induced membrane interactions of amyloid fibrils ................78

3.1 Introduction ........................................................................................................................79

3.2 Methods..............................................................................................................................80

3.2.1 Sonication of PrP fibrils .........................................................................................80

3.2.2 ANS Fluorescence .................................................................................................80

3.2.3 Liposome disruption assay .....................................................................................81

3.2.4 SSNMR measurements ..........................................................................................81

3.3 Results ................................................................................................................................82

3.3.1 ThT fluorescence and TEM measurements of sonicated fibrils ............................82

3.3.2 Hydrophobic surfaces of sonicated PrP peptide amyloids .....................................83

3.3.3 Liposome disruption by sonicated fibrils formed by PrP(244-249), PrP(245-

250), and PrP(178-183) ..........................................................................................84

3.3.4 Chemical shift assignments of sonicated PrP peptide fibrils .................................86

3.3.5 β-sheet arrangement of sonicated fibrils ................................................................87

3.3.6 Different chemical environments and the loss of intermolecular contacts in the

fibril core upon sonication .....................................................................................88

3.4 Discussion ..........................................................................................................................91

vii

Chapter 4. Structural characterization of the lipid-bound and amyloid fibrillar states

of human serum amyloid A ....................................................................................................93

4.1 Introduction ........................................................................................................................94

4.2 Methods..............................................................................................................................96

4.2.1 Recombinant SAA expression ...............................................................................96

4.2.2 Purification and sample preparation of lipid-free SAA .........................................96

4.2.3 Preparation of lipid vesicles and lipid-bound SAA on reconstituted high-

density lipoprotein (rHDL-SAA) ...........................................................................97

4.2.4 SAA fibrillization kinetics measurements .............................................................97

4.2.5 Sample preparation of rHDL-SAA and SAA fibrils for SSNMR..........................97

4.2.6 Gluteraldehyde cross-linking of SAA ....................................................................98

4.2.7 Dynamic light scattering (DLS) .............................................................................98

4.2.8 Circular dichroism and thermal stability................................................................98

4.2.9 FTIR measurements ...............................................................................................99

4.2.10 Solution and solid-state NMR ................................................................................99

4.3 Results ..............................................................................................................................100

4.3.1 Characterization of soluble lipid-free SAA oligomers ........................................100

4.3.2 SAA solubilizes DMPC vesicles and forms discoidal HDL-like particles ..........101

4.3.3 Secondary structure and thermodynamic stability of apo- and lipid-associated

SAA......................................................................................................................102

4.3.4 Fibrillization of lipid-free SAA in presence of GAGs .........................................103

4.3.5 Fibrillization of lipid-associated SAA in presence of GAGs ...............................105

4.3.6 SSNMR of SAA amyloid fibrils ..........................................................................106

4.3.7 SSNMR of rHDL-SAA and chemical shift assignments .....................................111

4.3.8 Comparison between experimental and predicted chemical shift of lipid-bound

SAA......................................................................................................................115

4.4 Discussion ........................................................................................................................117

Chapter 5. Conclusions and Future Directions ..................................................................120

5.1 Structural features of amyloid fibrils ...............................................................................121

5.2 Amyloid fragmentation and membrane interaction .........................................................121

5.3 Lipid association of SAA and the effects on structure and amyloid formation ...............122

5.4 Initial structural model of amyloid SAA by SSNMR ......................................................123

5.5 Future directions ..............................................................................................................123

5.5.1 Stability and fibillization kinetics of lipid-free and lipid-bound SAA as an

effect of pH ..........................................................................................................123

5.5.2 SSNMR studies of the SAA fibril core and structural differences between

fibrils formed under different solution conditions ...............................................125

5.5.3 The effects of GAGs and lipids on SAA amyloid fibril morphology and

structure................................................................................................................126

5.5.4 Structure of lipid-associated SAA .......................................................................127

5.5.5 Functional diversity of SAA ................................................................................128

5.6 Final concluding remarks .................................................................................................131

viii

Bibliography ................................................................................................................................132

Appendices ...................................................................................................................................157

ix

List of Tables

Table 1.1. Proteins associated with human amyloidosis................................................................. 4

Table 1.2. SSNMR pulse sequences used in this thesis. ............................................................... 23

Table 2.1. 13C/15N isotope-labeling schemes of the amyloidogenic prion peptide sequences used.

....................................................................................................................................................... 43

Table 2.2. Backbone torsion angles of PrP peptide amyloids calculated using TALOS+ using 13C

and 15N chemical shift. .................................................................................................................. 57

Table 2.3. Intra-residue 13C – 15N distance constraints determined through ZF-TEDOR. ........... 66

Table 2.4. Comparing internuclear distances determined by experimental RR measurements and

PrP(244-249) NMR structural models. ......................................................................................... 69

Table 5.1. Thermal stability of lipid-free SAA2 as a function of solution pH. .......................... 124

x

List of Figures

Figure 1.1. Protein folding energy landscape and aggregation. ...................................................... 3

Figure 1.2. Cross-β assembly of amyloids and 8 classes of steric zipper arrangements of β-sheets.

......................................................................................................................................................... 7

Figure 1.3. Kinetics of amyloid formation.................................................................................... 10

Figure 1.4. Mechanisms for membrane disruption of amyloid proteins. ...................................... 14

Figure 1.5. Membrane disruption and fibril toxicity is dependent on size of amyloid assemblies.

....................................................................................................................................................... 16

Figure 1.6. Magic-angle spinning in solid-state NMR. ................................................................ 19

Figure 1.7. Human prion protein (PrP) and its misfolded PrPsc state. .......................................... 26

Figure 1.8. Human Serum amyloid A. .......................................................................................... 32

Figure 1.9. Synthesis and remodeling of high density lipoproteins in reverse cholesterol transport

(RCT). ........................................................................................................................................... 34

Figure 1.10. Glycosaminoglycans in the body. ............................................................................. 35

Figure 1.11. Sonication-enhanced amyloid cytotoxicity of short amyloidogenic peptides. ......... 37

Figure 2.1. TEM of PrP peptide fibrils. ........................................................................................ 48

Figure 2.2. FTIR spectra of PrP peptide amyloid fibrils............................................................... 49

Figure 2.3. ThT Fluorescence of PrP peptide amyloid fibrils....................................................... 50

Figure 2.4. Cross-β characterization of PrP(178-183) amyloids. ................................................. 51

Figure 2.5. Resonance assignments for PrP(244-249) SFLI-labelled peptides. ........................... 52

Figure 2.6. Resonance assignment of PrP(245-250) uniformly 13C, 15N-labelled peptides. ........ 53

Figure 2.7. Resonance assignment of PrP(178-183) DCN/VI-labelled peptides.......................... 54

Figure 2.8. 13C secondary chemical shift calculations and linewidths for PrP peptide amyloids. 56

Figure 2.9. CT-PITHIRDS measurement of backbone inter-strand distances in selectively 13CO-

labelled peptides............................................................................................................................ 58

Figure 2.10. 13C – 13C RAD/DARR correlation spectra for long-range intermolecular contacts in

PrP(245-250) fibrils. ..................................................................................................................... 60

xi

Figure 2.11. Long-range intermolecular contacts observed in 2D 13C – 13C spin diffusion spectra

of PrP(245-250) SI/F fibrils. ......................................................................................................... 61

Figure 2.12. 13C – 15N heteronuclear ZF-TEDOR measurement of intermolecular contacts for

uniformly-labelled PrP245-250 fibrils. ......................................................................................... 62

Figure 2.13. ZF-TEDOR measurements of 25% uniform-13C, 15N-labelled PrP(245-25) diluted

with 75% unlabelled material. ...................................................................................................... 63

Figure 2.14. Long-range intermolecular contacts in PrP(178-183) amyloids prepared from

DCN/VI-labelled peptides. ........................................................................................................... 64

Figure 2.15. 13C – 13C intermolecular contacts in PrP(178-183) amyloids. ................................. 65

Figure 2.16. 13C – 15N intramolecular distances of PrP(178-183) peptides in amyloid fibrils. .... 66

Figure 2.17. 13C – 13C long-range intermolecular contacts in PrP(244-249) amyloids. ............... 68

Figure 2.18. 13C RR measurements for PrP(244-249) SI/F fibrils. ............................................... 70

Figure 2.19. Structure of PrP(245-250) fibrils. ............................................................................. 71

Figure 2.20. Structure of PrP(178-183) amyloids. ........................................................................ 72

Figure 2.21. Structure of PrP(244-249) amyloids. ........................................................................ 73

Figure 2.22. Seeding of PrP(244-249) fibrils................................................................................ 75

Figure 3.1. Morphological changes to sonicated PrP peptide amyloids. ...................................... 83

Figure 3.2. Surface hydrophobicity of PrP peptide amyloids. ...................................................... 84

Figure 3.3. Membrane disruption activity of PrP peptide amyloids upon sonication................... 85

Figure 3.4. Secondary 13C chemical shift calculations observed for sonicated PrP peptide

amyloids. ....................................................................................................................................... 87

Figure 3.5. 13C inter-β-strand distance measurements of sonicated PrP peptide fibrils. .............. 88

Figure 3.6. PrP peptides experience a different chemical environment after sonication. ............. 89

Figure 3.7. Intermolecular β-sheet contacts in sonicated PrP(244-249) fibrils. ........................... 90

Figure 3.8. Intermolecular β-sheet contacts in sonicated PrP(245-250) and PrP(178-183) fibrils.

....................................................................................................................................................... 91

Figure 4.1. Oligomeric apo-SAA with a dynamic and flexible C-terminus. .............................. 101

Figure 4.2. SAA solubilizes DMPC vesicles to form discoidal HDL-like particles. .................. 102

Figure 4.3. SAA secondary structure and thermodynamic stability. .......................................... 103

xii

Figure 4.4. GAG-induced fibril formation of SAA at 37oC. ...................................................... 104

Figure 4.5. Secondary structure of apo-SAA and SAA amyloid fibrils. .................................... 105

Figure 4.6. GAG-induced fibril formation of rHDL-SAA at pH 3.6. ......................................... 106

Figure 4.7. 1D 13C spectra of uniformly 13C, 15N-labelled SAA2 fibrils examined by MAS

SSNMR. ...................................................................................................................................... 107

Figure 4.8. 13C amino acid chemical shift assignment of immobile region in SAA2 fibrils. ..... 109

Figure 4.9. 13C chemical shift assignment of residues in the mobile region of SAA2 fibrils. .... 111

Figure 4.10. 1D 13C spectra of uniformly 13C, 15N-labelled rHDL-SAA examined by MAS

SSNMR. ...................................................................................................................................... 112

Figure 4.11. Preliminary chemical shift assignments of rHDL-SAA2 from 13C – 13C correlation

spectrum. ..................................................................................................................................... 113

Figure 4.12. Chemical shift assignment from 13C – 15N heteronuclear NCACX spectrum of

rHDL-SAA. ................................................................................................................................. 114

Figure 4.13. Comparison of experimental 13C spectrum of rHDL-SAA2 with chemical shifts

predicted from the crystal structure of the SAA1.1 hexamer. .................................................... 116

Figure 5.1. Fibrllization of apo-SAA in the presence of heparin in different pH buffers. ......... 125

Figure 5.2. Cell surface receptors that mediate the functions of SAA in the immune system. .. 130

xiii

List of Equations

Equation 1-1 Chemical shift anisotropy. ...................................................................................... 18

Equation 1-2 Dipole-dipole coupling Hamiltonian....................................................................... 18

xiv

List of Appendices

Table A 1. 13C and 15N chemical shifts from PrP(244-249) fibrils. ............................................ 157



xv

Abbreviations

α-Syn – Alpha-synuclein

Aβ – Amyloid-β peptide

AA - Amyloid A

AFM – Atomic force microscopy

ANS – 1-Anilinonaphthalene-8-Sulfonic Acid

APF – Annular protofibril

ApoA – Apolipoprotein A

BSE – Bovine spongiform encephalopathy

CJD – Creutzfield-Jakob disease

CP – Cross polarization

CR – Congo Red

CT-PITHIRDS – Constant-time PITHIRD

CWD – Chronic wasting disease

DARR – Dipolar-assisted rotational resonance

DLS – Dynamic light scattering

DMPC – 1,2-dimyristoyl-sn-glycero-3-phosphocholine

DP – Direct pulse

ECM – Extracellular matrix

ER – Endoplasmic reticulum

FFI – Fatal Familial Insomnia

FID – Free induction decay

FO – Fibrillar oligomer

GAG – Glycosaminoglycan

GM1 – Monosialotetrahexosyl ganglioside

xvi

GPI – Glycosylphosphatidylinositol

GSS – Gerstmann–Sträussler–Scheinker syndrome

HDL – High-density lipoproteins

HSQC – Heteronuclear single quantum correlation

IAPP – Islet amyloid polypeptide

IDP/IDR – Intrinsically disordered protein / intrinsically disordered region

IL – Interleukin

INEPT – Insensitive nuclei enhanced by polarization transfer

MAS – Magic angle spinning

NMR – Nuclear magnetic resonance

PAR – Proton-assisted recoupling

PEG – Polyethylene glycol

PFO – Pre-fibrillar oligomer

PK – Proteinase K

PrP – Prion protein

PC – Phosphatidylcholine

PE – Phosphatidylethanolamine

PG – Phosphatidylglycerol

PS – Phosphatidylserine

POPC – 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine

POPG – 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphotidylglycerol

RAD – RF-assisted diffusion

RCT – Reverse cholesterol transport pathway

RF – Radio-frequency

rHDL – Reconstituted high density lipoprotein

RMSD – Root mean square deviation

xvii

RR – Rotational resonance

SAA – Serum amyloid A

SSNMR – Solid-state nuclear magnetic resonance

TEM – Transmission electron microscopy

ThT – Thioflavin T

TMS – Tetramethylsilane

TNF – Tumour necrosis factor

TOBSY – Total through bond correlation spectroscopy

TSE – Transmissible spongiform encephalopathy

ZF-TEDOR – Z-filtered transferred-echo double resonance

1

Chapter 1.

Introduction

2

1.1 Protein misfolding and amyloids

1.1.1 Protein folding

Protein folding can be described as a stochastic sampling of conformational ensembles across the

free energy landscape to reach a thermodynamically stable structure (Anfinsen 1973; Dill &

Chan 1997; Dobson 2003). A nascent polypeptide folds through various intermediates by small

stabilizing structural changes to lower its free energy and restrict its conformational freedom as it

samples the landscape, until it eventually achieves a unique stable structure with minimum free

energy (Figure 1.1) (Dill & Chan 1997). This reversible process is dictated by the protein

primary sequence, and is driven to hide the polypeptide backbone and hydrophobic side chains

within the folded structure (Anfinsen 1973; Dobson 2003). Often, the energy landscape is rugged

and decorated with local minima that have a higher free energy than the native fold. These

kinetic traps result in the accumulation of partially unfolded or misfolded intermediate

conformations with exposed surfaces that promote non-native interactions and aggregation (Dill

& Chan 1997). Molecular chaperones are part of the regulatory machinery that prevents

aggregation and ensures efficient folding of polypeptides into their native conformation on a

biologically relevant time scale (Hartl & Hayer-Hartl 2009; Hartl et al. 2011). Once folded,

specific energy barriers prevent the protein from converting into the aggregation-prone states

without restricting the functional dynamics of the protein (De Simone et al. 2011).

Destabilization by denaturation (pH, heat, denaturant), mutations, post-translational

modifications, or proteolysis, can lower the energy barrier and promote protein misfolding (De

Simone et al. 2011; Neudecker et al. 2012). Alternatively, changes in protein quality control due

to age and stress, allow misfolded proteins to escape protein regulatory processes and avoid

being degraded by ubiquitin-proteosome or autophagy (Hartl et al. 2011). Non-native

conformational intermediates that accumulate can form intermolecular interactions and lead to

oligomerization and aggregation of misfolded proteins into amorphous aggregates. In some

cases, this results in the formation of highly ordered fibrillar species called amyloids that are

recognized as thermodynamically stable end-stage structures.

3

Figure 1.1. Protein folding energy landscape and aggregation. The energy landscape acts as a funnel that drives

a multitude of unfolded conformations to the unique native structure stabilized by intramolecular contacts including

hydrophobic effects, charge interactions, and hydrogen bonding. Intermediates that represent local energy minima

can become populated during protein folding. Chaperones aid in the process by helping intermediates trapped in the

partially folded conformations to overcome the energy barriers of these local minima, while at the same time

preventing these partially folded states from forming intermolecular contacts that drive aggregation. Amyloid fibrils

are thought to be the thermodynamically stable end stage of misfolded proteins. Reprinted with permission from

Hartl et al. 2011. Copyright 2011 Nature Publishing Group.

1.1.2 Amyloids and diseases

Misfolded proteins self-assembling into intractable amyloid aggregates is now recognized as a

major medical challenge of the 21st century (Dobson 1999; Knowles et al. 2014). There are over

50 human diseases related to protein misfolding and amyloids, including neurodegenerative

disorders such as Alzheimer’s disease, Parkinson’s disease, and mammalian prion diseases, and

non-neuropathic diseases such as type II diabetes mellitus, Amyloid A (AA) amyloidosis, and

dialysis-associated amyloidosis (Table 1.1) (Dobson 1999). While some diseases could be due to

4

the loss of biologically functional protein, a growing collection of literature point towards the

cytotoxic effects of the amyloid fibrils and smaller misfolded intermediates that accumulate

during the misfolding process. Amyloidosis refers to the medical condition characterized by a

localized or systemic deposit of amyloid fibrils onto organs (Chiti & Dobson 2006). The

deposition of these amyloid fibrils either extra- or intra-cellularly onto organs manifests into

cellular death and organ failure as a hallmark for disease progression (Chiti & Dobson 2006;

Hardy & Selkoe 2002; Soto 2003; Bolton et al. 1982; Spillantini et al. 1997; Dobson 1999; Sipe

& Cohen 2000).

Table 1.1. Proteins associated with human amyloidosis

Amyloid Diseases Proteins Involved Native

structural fold

Organs

Associated

Localized Amyloidosis

Alzheimer’s Disease Amyloid-β (Aβ) and

tau Disordered Brain

Parkinson’s Disease α-Synuclein (α-Syn) Disordered Brain

Transmissible Spongiform

Encephalopathies (TSE) Prion protein (PrP) α-helix Brain

Chronic Traumatic

Encephalopathies Tau Disordered Brain

Huntington’s Disease Huntingtin Largely

disordered Brain

Type II Diabetes Islet amyloid

polypeptide (IAPP) Disordered Pancreas

Injection-localized

Amyloidosis Insulin α-helix Skin

Systemic Amyloidosis

Amyloid Light-chain

Amyloidosis

Immunoglobulin light

chain β-sheet

Kidney, heart,

skin

Amyloid A (AA)

Amyloidosis

Serum Amyloid A

(SAA) α-helix

Kidneys, spleen,

blood vessels

Familial Renal Amyloidosis Apolipoprotein A-I

(ApoA-I) disordered Kidney

Dialysis-associated

amyloidosis β2-microglobulin β-sheet, Ig-like

Joints, connective

tissue

Cerebral Amyloid

Angiopathy Cystatin C α+β Blood vessels

Familial Amyloid

Polyneuropathy Transthyretin α+β

Peripheral

nervous system,

heart

5

The pathogenesis of amyloid disease involves misfolding of a cellular protein into a β-rich non-

native conformation and assembly into insoluble amyloids. The protein implicated in each

disorder may begin with a different native structure. Intrinsically disordered proteins (IDPs) or

proteins with extensive intrinsically disordered regions (IDRs) do not fold into regular compact

structures under physiological conditions and only undergo transition into more ordered folded

states in the presence of a specific binding partner (Dyson & Wright 2005). The ability of these

highly dynamic polypeptides to interact with various partners and adopt the necessary structure

are important to their biological roles of signaling and regulation, but can also lead to amyloid

fibril formation in certain cases such as Aβ, α-Syn, and ApoA-I (Uversky et al. 2008).

1.1.3 Functional amyloids

In recent years, non-pathogenic amyloid-like structures have also been discovered in various

organisms. Now recognized as functional amyloids, these structures play roles in a variety of

important biological activities. For example, functional amyloids in mammals have been found

to act as a storage medium for various protein/peptide hormones in the pituitary secretory

granules, accelerate the polymerization of the melanin protein PMel17 in skin and eye pigment,

or mediate programmed necrosis signalling such as the RIP1/RIP3 necrosome complex (Li et al.

2012; Maji et al. 2009; Fowler et al. 2006; Coustou et al. 1997; Gopalswamy et al. 2014). Yeast

functional amyloids are used to passage non-mendelian genetic phenotypes such as [PSI+] and

[URE3] in Saccharomyces cerevisiae (Chiti & Dobson 2006). The yeast cell surface adhesin Als

protein and the bacteria curli protein form amyloids as part of the extracellular matrix (ECM)

biofilm when colonizing and dealing with environmental stress (Ramsook et al. 2010; Barnhart

& Chapman 2006). The filamentous fungi Podospora anserina use HET-s fibrils to activate cell

death and prevent two strains with different alleles at the het-s/S locus from fusing into

heterokaryons (Coustou et al. 1997). As well, fungal hydrophobins self-assemble into an

amphipathic monolayer of amyloid-like rodlets at the hydrophobic:hydrophilic interface to

reduce surface tension at air:water boundaries or to form a protein coat around spores (Morris et

al. 2011).

Amyloidogenic proteins also have interesting applications in biomaterials and nanostructures.

For example, fibrils functionalized with cell adhesion moieties (i.e. tripeptide RGD of

fibronectin) can be developed into nanoporous matrices suitable for cell growth and tissue

engineering (Ahn et al. 2010). Alternatively, the self-assembling properties of amyloid fibrils can

6

be used to prepare hydrogels for drug/enzyme delivery therapy, or to template synthesis of

nanowires with conducting or optical properties (Maji et al. 2008; Scheibel et al. 2003; Mankar

et al. 2011). Therefore, understanding the structural properties of amyloids and the assembly

processes that govern the size, shape, morphology, and pathogenicity of fibrils has broad

importance to scientists and engineers.

1.1.4 Amyloid structure

Amyloid fibrils, as observed using transmission electron microscopy (TEM) or atomic force

microscopy (AFM), are typically long, twisted filaments with diameters of 6-12 nm and up to

several microns long (Geddes et al. 1968; Greenwald & Riek 2010). Amyloid-like fibrils have

been reported having a wide range of morphologies, including curly twisted fibrils, linear

straight fibrils, rods, and nanotubular tape. In each case, fibrils are composed of multiple

protofilaments that are twisted or laterally associated to each other, forming bundles with

different morphologies (Figure 1.2 A) (Dobson 2003; Meinhardt et al. 2009; De Jong et al.

2006). Each protofilament contains a core structural motif of β-sheets running parallel to the

fibril elongation axis, called the cross-β structure (Geddes et al. 1968; Tycko 2000). The β-

strands are spaced 4.7 Å apart to form β-sheets that elongate along the length of the fibril, while

the sheets are stacked 9 - 12 Å apart (Geddes et al. 1968). This cross-β arrangement gives rise to

a characteristic fibre diffraction pattern with reflections at the meridian and equator that arise

from the inter-strand and inter-sheet distances, respectively (Figure 1.2 A) (Geddes et al. 1968;

Sunde et al. 1997; Greenwald & Riek 2010). The cross-β structure is also recognized by

characteristic dyes that bind selectively to amyloids. For example, thioflavin T (ThT) exhibits a

shift in its fluorescence emission maximum to 482 nm upon binding, and Congo red (CR) stains

fibrils and exhibits a red-green birefringence under polarized light (Klunk et al. 1989; Biancalana

& Koide 2010). However, there have been reports of β-sheet fibrillar structures that induce no or

low fluorescence quantum yield of bound-ThT (LeVine 1995).

7

Figure 1.2. Cross-β assembly of amyloids and 8 classes of steric zipper arrangements of β-sheets. A)

Schematic zoom of cross-β amyloid backbone. β-strands are assembled at 4.7 Å to give rise to the meridian

reflection (black arrow) and the β-sheet stacking at 9-12 Å give rise to the equatorial reflection (white arrow) in X-

ray fibre diffraction. Protofilament association further govern the morphology of fibrils to give it the twist.

Reprinted with permission from Wille et al. 2009 and Fitzpatrick et al. 2013. Copyright 2009 and 2013 Proceedings

of the National Academy of Sciences of USA. B) 8 possible classes of steric zipper, four containing parallel β-sheets

and four containing anti-parallel β-sheets. These β-sheets can be further oriented to have the parallel or anti-parallel

stacking of β-sheets, which can be aligned in a face-to-face or face-to-back manner. Reprinted with permission from

Sawaya et al. 2007. Copyright 2007 Nature Publishing Group.

Extensive intermolecular hydrogen bonds of the backbone, aligned parallel to the fibril axis,

contribute to the stabilization of the amyloid cross-β fold (Sunde et al. 1997; Dobson 2004;

Knowles et al. 2014). The free energy associated with fibril formation is thus related to the

protein concentration, and the amyloid state can be more thermodynamically favourable than the

native protein fold beyond a critical protein concentration (Baldwin et al. 2011; Knowles et al.

2014). It has been proposed that the amyloid fold is an intrinsic structural fold that all

polypeptide sequences can adopt, and not only a disease-related phenomenon. Numerous

research groups have shown that polypeptides and proteins of various sequences, lengths, and

amino acid compositions can adopt the amyloid cross-β architecture both in vitro and in vivo

(Blancas-Mejía & Ramirez-Alvarado 2013; Sipe & Cohen 2000) and much research has been

carried out to obtain atomic-level structures of these diverse amyloids.

8

X-ray diffraction studies of microcrystals formed by peptide fragments of Aβ, tau, PrP, insulin,

IAPP, and yeast prion Sup35, have provided insight into the molecular details of amyloidogenic

protein assemblies. Sawaya et. al. proposed that tightly interdigitated hydrophobic side chains

exclude water and form “steric zippers” that make up the core of each protofilament (Sawaya et

al. 2007). This burial of hydrophobic residues into the dehydrated fibril core acts much like the

hydrophobic effect in protein folding, and is an important factor in the stability of the fibril

assembly (Sunde et al. 1997; Dobson 2004; Knowles et al. 2014). These steric zippers can be

classified by the strand arrangement - parallel or anti-parallel alignment of β-strands into sheets,

parallel or anti-parallel stacking of β-sheets, and the stacking of the same (face-to-face) or

different faces (face-to-back) of β-sheets – to give eight possible classes of steric zippers (Figure

1.2 B).

The class 1 steric zipper (parallel strands stacked in an anti-parallel face-to-face arrangement)

has been most frequency observed in non-crystalline fibrillar assemblies by SSNMR structural

studies, including PrP(106-126) (Walsh et al. 2009) and yeast GNNQQNY (Lewandowski et al.

2011). At the same time, class 4 zippers (anti-parallel strands stacked in an anti-parallel face-to-

back arrangement) and class 5 zippers (anti-parallel strands stacked in an anti-parallel face-to-

face arrangement) have been observed for IAPP peptide amyloids (Nielsen et al. 2009; Wiltzius

et al. 2009). In other cases, anti-parallel β-strand arrangement has been reported but with poorly

defined inter-sheet information to classify the steric zipper (Balbach et al. 2000; Madine et al.

2008; van der Wel et al. 2010; Petkova et al. 2004). Adding to the complexity, the same

sequence can also adopt multiple steric zipper arrangements as observed for Aβ, Sup35, and de

novo designed peptides (Verel et al. 2008; Colletier et al. 2011; Lewandowski et al. 2011). To

date, no examples in class 3 zippers have been observed.

Side chain interactions outside the steric zippers further govern the association of β-sheets into

higher-ordered assemblies, such as protofilaments assembling into mature fibrils (Sawaya et al.

2007). In yeast prions that are rich in Asn/Gln (N/Q) sequences, studies suggest that parallel β-

sheet alignment of the peptides allows N/Q carboxyamide side chain to form a network of

hydrogen bonds, providing additional stabilization to amyloid fibrils (Perutz et al. 1993; van der

Wel et al. 2007). Nevertheless, much research is still being carried out to examine how

polypeptide sequence affects fibril formation, and the molecular determinants that govern the

different biological behaviours of amyloids.

9

1.1.5 Kinetics of amyloid assembly

Fibril formation can be described as a nucleation-dependent event that typically follows a

sigmoidal curve with a lag phase, growth phase, and plateau phase (Figure 1.3). During the

initial lag phase, soluble protein undergoes partial or complete unfolding that may occur via

native-like intermediate structures or non-native intermediates (Neudecker et al. 2012; Lorenzen

et al. 2014; Das et al. 2014; De Carufel et al. 2015). An altered charge distribution and

hydrophobic group exposure at the surface of the misfolded protein promotes intermolecular

interactions and assembly, forming oligomers that range in size from 2 to 100-mers. However,

this value can differ between different amyloid systems and the conditions these oligomers are

isolated. The assembled oligomers aggregate and become a nucleus that can template the

misfolding of more precursor proteins and nucleate the assembly of monomers into β-rich

structures, leading to an exponential rate of protofibril formation. Additional events during the

growth phase – such as secondary nucleation of misfolded oligomers by preformed fibrils, or

fragmentation that increases the surface area to template misfolding – further speeding up the

polymerization and aid in the rapid accumulation of misfolded assemblies (Cohen et al. 2013).

Finally, free monomers are depleted as fibrillization process reaches the plateau phase, and the

protofibrils assemble into higher-ordered structures. The rate of fibrillization can be accelerated

using temperature, salt, pH, and agitation. Temperature and pH destabilize protein to promote

misfolding/unfolding events; salt increases the propensity of aggregation by screening charge

repulsion; and agitation increases air-water or air-surface exposure. Adding small amounts of

fragmented fibrils to the system, or agitating to increase fragmentation events, seeds amyloid

growth and decreases the length of the initial lag phase for rapid fibril formation.

10

Figure 1.3. Kinetics of amyloid formation. Amyloid formation modelled as a sigmoidal curve when monitored

using ThT fluorescence. It goes through three phases – lag phase, growth phase, and plateau phase – where

misfolded monomer, small oligomers, protofibrils are accumulated. Addition of fibril seeds in green shortens the

rate-limiting lag phase to promote elongation and fibril formation.

1.1.6 Amyloid fibrils and oligomeric intermediates

While diagnosis of amyloid disease is typically carried out using histological stains to detect

amyloids in tissue biopsy, the clinical symptoms are often observed before the onset of amyloid

plaque accumulation (Dickson et al. 1995). Small oligomeric intermediates, either on- or off-

pathway of fibrillization, have been shown to decrease viability of cell cultures and are believed

to be the pathogenic culprits of diseases (Hardy & Selkoe 2002; Walsh et al. 2002). These pre-

fibrillar aggregates are often transient and heterogeneous in nature due to the presence of

multiple possible misfolding pathways leading to different structures and morphologies. Hence,

it has been difficult to obtain highly pure samples for high-resolution structural characterization

of these oligomeric intermediates.

Soluble oligomers are rich in β-sheet secondary structure with broadly overlapping size

distribution (Kayed et al. 2007; Diociaiuti et al. 2014; Chimon et al. 2007; Yu et al. 2009;

Ahmed et al. 2010). They can be classified as pre-fibrillar oligomers (PFO), fibrillar oligomers

11

(FO), or annular protofibrils (APF) (Kayed et al. 2007; Diociaiuti et al. 2014; Chimon et al.

2007; Yu et al. 2009; Ahmed et al. 2010) by their mutually exclusive reactivity to conformation-

specific antibodies (Glabe 2008). In fact, each of these antibodies can recognize oligomers from

various amyloid-forming sequences, suggesting that the fundamental epitope arises from a

distinct structural organization of the polypeptide backbone common to each group, rather than

specific side chain conformations. (Kayed et al. 2004; Kayed et al. 2007; Mamikonyan et al.

2007; Glabe 2008). These common structures may play a significant role in governing the

toxicity of these intermediates, and hints at the potential of soluble oligomers as

immunotherapeutic targets to treat Alzheimer’s and other amyloid diseases (Goure et al. 2014).

For example, peptide fragments from PrP, Aβ, α-Syn, and IAPP have all been reported to form

spherical oligomers that permeabilize lipid bilayer membranes and induce toxicity in cell

cultures, which both activities can be attenuated by anti-PFO antibodies (Kayed et al. 2004;

Kayed et al. 2003). Meanwhile, soluble FOs can have similar morphology and size as PFOs but

share immunoreactivity to mature fibrils (Kayed et al. 2007). Aβ42 FOs were reported to contain

cross-β structure with anti-parallel β-strands similar to fibrils, but nucleates Aβ monomers into

FOs instead of seeding fibril formation (Kayed et al. 2007). APFs can adopt ring-shaped or pore-

like structures as reported for Aβ42, α-synuclein, and TTR (Lowe et al. 2004; Kayed et al. 2009;

Pires et al. 2012). They can also be precursors to larger closed rings, ribbons, or protofilaments

(Huang et al. 2000; Sokolowski et al. 2003). These annular oligomers were not reactive to

antibodies against monomers, fibrils, or spherical oligomers, but instead recognized antibody that

binds to β-barrels (Kayed et al. 2009; Laganowsky et al. 2012). Yet surprisingly, APFs have little

membrane-permeabilizing activity compared to PFOs and are less toxicity than PFOs (Kayed et

al. 2009).

Size, structural flexibility, and compactness all alter oligomer instability and toxicity (Krishnan

et al. 2012; Mannini et al. 2014; Ladiwala et al. 2012; Campioni et al. 2010). Yet despite

misfolded proteins adopting many conformations, cytotoxicity of amyloid oligomers is reported

to correlate most to hydrophobic exposure probed by 1-anilinonaphthalene-8-sulfonic acid

(ANS) (De Simone et al. 2011; Mannini et al. 2014; Bolognesi et al. 2010). By examining the

molecular organization of pathogenic and functional amyloids, it can provide us with insights

into their different biological properties and begin to identify key interactions with biological

components that are important to amyloid cytotoxicity.

12

1.2 Amyloid protein-lipid interaction

Surfaces accelerate amyloid formation compared to incubating in solution alone (Zhu et al.

2002). This can be due to the increased local concentration of amyloid proteins caused by surface

interactions that make them behave as diffusing on a 2D plane rather than a 3D space (Shen et al.

2012). It is, therefore, important to consider the interactions between amyloidogenic proteins and

lipid surfaces, especially the cell membrane. In particular, how does lipid affect the structural

fold of the native protein and alter the assembly of amyloid structures, yet at the same time, how

these interactions with amyloid oligomers or fibrils negatively impact the bilayer.

1.2.1 Lipid interactions with amyloidogenic proteins

Several amyloidogenic proteins have been reported to interact with lipids as a monomeric

precursor. For example, Aβ peptide retains part of the C-terminal transmembrane region after it

is processed from the Aβ precursor protein by β- and γ-secretases, and can be inserted into

phosphatidylcholine (PC) membranes (Matsuzaki & Horikiri 1999; Bokvist et al. 2004; Lemkul

& Bevan 2013). α-Syn is an intrinsically disordered protein in solution that forms amphipathic

helices at its N-terminus upon binding to anionic phosphatidylserine (PS) lipids (Bodner et al.

2009; Comellas et al. 2012). This may be an important functional property, since α-Syn binds to

regions of high membrane curvature or exposed hydrophobic patches during synaptic vesicle

regulation (Bellani et al. 2010). Amphipathic lipid–surface binding proteins known to form

amyloids - apolipoprotein A (ApoA) and serum amyloid A (SAA) - adopt more stable α-helical

secondary structures in the presence of zwitterionic PC lipids, which is essential for their roles in

guiding high-density lipoprotein (HDL) structure and function (Guha et al. 2008; Gao et al.

2012; Mizuguchi et al. 2015).

1.2.2 Membrane-induced fibril formation

Interactions with membrane-surface functional groups may stabilize non-native structural folds

that are more aggregation-prone (Cecchi & Stefani 2013). These might serve as nucleation sites

that may otherwise not exist in solution, leading to fibrils with different morphologies than those

formed in solution (Comellas et al. 2012; Niu et al. 2014). For example, Aβ has been shown to

form more amorphous aggregates on gold surfaces modified with hydrophobic CH3-groups, but

formed small oligomers and small protofibrils when modified with COOH (Moores et al. 2011;

Zhu et al. 2002). Interactions with electrostatics of the lipid headgroups may also alter amyloid

13

formation by increasing peptide adsorption onto the monolayer and possibly drive the peptide

deep into the monolayer (Maltseva et al. 2005; Ege et al. 2005; Chi et al. 2008; Galvagnion et al.

2015). Alternatively, it could recruit oligomers of different surface charge and size to enhance

amyloid formation (Moores et al. 2011; Niu et al. 2014; Yip & Mclaurin 2001). Anionic

phospholipids such as phosphatidylglycerol (PG) and phosphatidylinositol (PI) can shift the

conformation of Aβ peptides from soluble, largely unstructured monomer towards β-sheet-rich

aggregates, while PC and sphingomyelin have very weak or no interaction with Aβ (McLaurin et

al. 1998). Other lipid components such as the sphingolipid monosialotetrahexosyl ganglioside

(GM1) has been shown to interact with Aβ peptides, induce structural rearrangement into β-

sheet, and possibly seed formation of cytotoxic Aβ oligomers (Choo-Smith & Surewicz 1997;

Matsubara et al. 2013; Yanagisawa et al. 1995). Fragments from different regions of the Aβ

peptide – extracellular domain, hydrophobic region, or transmembrane domain – was shown to

aggregate on supported total brain lipid extract with different fibril morphology, suggesting a

diverse effect of sequences on the aggregation of amyloid proteins onto membranes (Yates et al.

2013).

1.2.3 Membrane disruption and amyloid toxicity

Various mechanisms for amyloid toxicity have been proposed with a large amount of data

obtained using the Aβ peptide as a model system. These include binding to glutamate receptors

and increasing intracellular Ca+2 levels (Cecchi & Stefani 2013), inducing reactive oxygen

species (ROS) production, causing oxidative stress from oligomer-induced mitrochondrial

deregulation (Umeda et al. 2011; Mutisya et al. 1994), triggering cell apoptosis via caspase-

dependent signalling pathway (Loo et al. 1993), causing lysosomal membrane leakage (Yang et

al. 1998; Umeda et al. 2011; Ditaranto et al. 2001), and causing endoplasmic reticulum (ER)

stress to elicit the unfolded protein response (UPR) and disrupt protein homeostasis (Imaizumi et

al. 2001; Umeda et al. 2011; Fonseca et al. 2014).

In addition to the above hypothesis, amyloids have also been proposed to induce cytotoxicity via

a common mechanism mediated by the physical interactions of peptides with the cell membranes

(Rushworth & Hooper 2011; Terzi et al. 1997; Pastor et al. 2008; Yip & Mclaurin 2001; Engel et

al. 2008; Qiang et al. 2015). Shown in Figure 1.4, amyloid proteins binding to membranes can

induce several adverse effects that lead to toxicity. First, peptides or oligomer deposition onto

membranes as a carpet on one leaflet of the bilayer can cause asymmetric pressure between two

14

leaflets and possibly increase membrane stress by inducing curvature (McLaurin et al. 1998;

Matsuzaki & Horikiri 1999; Relini et al. 2009). This may change the fluidity of the membrane

and cause deformations that disrupt its structural integrity (Yip & Mclaurin 2001; Engel et al.

2008; Chaudhary et al. 2014; Milanesi et al. 2012). Second, amyloids may be able to form small

pores that allow ions to diffuse across membranes and disrupt cellular ion homeostasis as shown

in single-channel conductance experiments (Jeffrey 2013; Lin et al. 1997; Kagan 2012; Kourie &

Culverson 2000). Toroidal pore formation by antimicrobial peptides or the cylindrin β-barrel

structures reported for some Aβ oligomers may offer structural models of how amyloidogenic

peptides can form membrane pores (Sengupta et al. 2008; Bertelsen et al. 2012). Third,

amphiphilic peptides can bind to the surface of the lipid bilayer and penetrate into the membrane

to reduce surface tension in a manner similar to detergents. This leads to a loss of bilayer

integrity and causes either membrane thinning or membrane holes, which has been documented

for α-Syn and PrP peptides in anionic PC/PG lipid membranes. (Reynolds et al. 2011; Walsh et

al. 2014; Hellstrand et al. 2013). Finally, the fibrillization of PrP(106-126) oligomers on

membrane surface has been shown to disrupt the phase separation of ordered and disordered

domains in fluid lipid bilayers (Walsh et al. 2014; Engel et al. 2008).

Figure 1.4. Mechanisms for membrane disruption of amyloid proteins. Amyloid proteins binding to membrane

can lead to several mechanisms that induce cytotoxic effects. These include (i) membrane carpeting effect, (ii)

formation of a stable pore in the membrane, (iii) detergent-like effects, and (iv) membrane surface raft-like

aggregation. Reprinted with permission from Berthelot et al. 2013. Copyright 2013 Elsevier.

15

1.2.4 Fibril fragmentation and toxicity

Fragmentation events during the polymerization phase of fibril formation are known to generate

new nucleation sites that promote protein misfolding and accumulate more amyloid fibrils (Xue

et al. 2009; Marshall et al. 2014). As such, small fibril seeds are often used to shorten the

nucleation lag phase and accelerate in vitro fibrillization, and can be considered critical on-

pathway intermediates that populate during protein misfolding. Amyloid fibrils formed by β2-

microglobulin, α-Syn, hen egg white lysozyme, or peptide fragments of Aβ, tau protein, PrP,

ApoA-I, have all demonstrated increased cellular toxicity upon fragmentation, suggesting these

fibril seeds may share a common mechanism for cytotoxicity (Xue et al. 2009; Pastor et al. 2008;

Milanesi et al. 2012). One hypothesis was that fragmentation generates fibrils of smaller size that

more readily interact with lipid bilayer and cause membrane disruption (Figure 1.5).

Alternatively, fragmentation can expose more fibril ends to allow the depolymerization and

release of smaller species to generate cytotoxic effects (Xue et al. 2009). At the same time,

fragmented β2-microglobulin fibrils have been shown to be internalized by cells more readily

than mature fibrils, and that they can disrupt the lysosomal protein degradation (Jakhria et al.

2014). While intermediate species can contain β-structure similar to fibrils, there are no studies

that characterize the structure of these small fibril fragments, or that address the molecular basis

for their cytotoxicity.

16

Figure 1.5. Membrane disruption and fibril toxicity is dependent on size of amyloid assemblies. Mature

fibrillar assemblies have limited membrane interaction and are less cytotoxic as they become larger and more

ordered. However, fragmented fibrils have more exposed hydrophobic surfaces capable to interact with membranes,

and are more likely to be toxic entities in amyloid diseases. Reprinted with permission from Xue et al. 2009.

Copyright 2009 Journal of Biological Chemistry.

1.3 Solid-state nuclear magnetic resonance spectroscopy

Solution NMR has been used to examine the early stages of protein misfolding and small

oligomeric misfolding intermediates (Karamanos et al. 2015; Neudecker et al. 2012), and

crystallography has been successful in examining the structures of small amyloid peptides in

fibril-like crystals (Sawaya et al. 2007; van der Wel et al. 2007). However, the non-crystalline

and insoluble nature of most amyloid fibrils makes these high molecular weight polymers

challenging targets for structural determination by these methods. Instead, solid-state NMR

(SSNMR) has become a valuable tool for studying these biological solids.

17

1.3.1 Background on NMR

NMR spectroscopy is based on the interaction of a nucleus with the external magnetic field that

stems from the quantum mechanical description of all atomic nuclei having an intrinsic spin

angular momentum (Rule & Hitchens 2006). For nuclei with odd mass numbers (i.e. 1H, 13C,

15N, 19F, 31P), their angular momentum generates a magnetic dipole moment that naturally aligns

to an external magnetic field and establishes a low energy (ground) and a high energy (excited)

state. The transition energy (ΔE) between these states in NMR is directly proportional to the

external magnetic field by the equation: ΔE/ħ = - ω = γ·Bo where ħ is the reduced Planck’s

constant, ω is resonant frequency of the transition (Larmor frequency), Bo is the magnetic field

strength, and γ is the gyromagnetic ratio that relates the magnetic dipole moment of a nucleus to

its spin angular momentum.

NMR experiments use radio-frequency (RF) electromagnetic radiations to excite the nuclear

spins at the corresponding resonant frequency and perturb the magnetic moment in the magnetic

field. This creates a coherent magnetization that can be thought of as the sum of all aligned

magnetic moments in the excited state. As the excited states lose coherence and return to

equilibrium, we detect the magnetization decreasing over time in the form of a free induction

decay (FID) that is a convoluted measurement of the transition frequencies for all nuclei in the

experimental sample. Fourier transformation is used to deconvolute the FID signal by converting

the time evolution of magnetization into the individual frequencies encoded in the FID.

NMR spectroscopy is a powerful tool for studying macromolecules because nuclei are highly

sensitive to their local environment. First, electrons surrounding a nucleus create a secondary

magnetic field that affect the total magnetic field experienced by the nuclei. This shielding

interaction is much weaker than the external magnetic field, but perturbs the resonant frequency

of a nucleus enough to give rise to a characteristic chemical shift. Second, nuclear spins can

influence each other. 1H, 13C, 15N in biological samples can be “coupled” to each other either

through direct bond connections (scalar coupling) or due to their proximity through space

(dipolar coupling). These couplings allow us to measure the distance between multiple nuclei.

Third, nuclear spin interactions with the environment (spin-lattice, T1 relaxation) and nearby

nuclei (spin-spin, T2 relaxation) affect the lifetime of the excited state to return back to thermal

equilibrium, and are sensitive to local motion and dynamics. NMR pulse sequences developed

18

over the past several decades have focused on exploiting and measuring these spin interactions to

infer molecular details about the structure and dynamics of biological macromolecules.

1.3.2 Anisotropy in SSNMR and magic angle spinning (MAS)

The asymmetric distribution of electrons surrounding a nucleus influences the degree of

chemical shielding of the external magnetic field. This effect is sensitive to the orientation of the

nucleus and its surrounding electrons relative to the magnetic field.

(Equation 1.1)

Equation 1-1 Chemical shift anisotropy. Chemical shift chemical shift frequency (ωcs ) for an axially symmetric

shielding tensor. ω0 is the resonant frequency of the nucleus, the angle θ is the angle between the external magnetic

field and the z-axis of the principle axis frame (PAF), and σzz is the chemical shielding tensor aligned to the z-axis of

the PAF. σzz in this case contains both the isotropic and anisotropic components of the chemical shift tensor. (Duer

2004).

Likewise, the dipolar coupling between two nuclei is dependent on both the internuclear distance

and the orientation of the vector defining the coupling within the external magnetic field (Duer

2004).

(Equation 1-2)

Equation 1-2 Dipole-dipole coupling Hamiltonian. The truncated heteronuclear dipolar Hamiltonian interaction

in angular frequency units (rad s-1). μ0 is the magnetic dipole moment, γI and γS are the gyromagnetic ratios for the 2

nuclear spins I and S, the angle θ is the angle between the external magnetic field and the dipole vector between the

two nuclear spins, and r is the distance that separates the two spins (Duer 2004).

Therefore, both chemical shift and dipolar coupling interactions contain anisotropic components

that depend on their orientation relative to the external magnetic field (θ) defined by the term

(3cos2 θ – 1). In solution NMR, chemical shift anisotropy (CSA) and dipolar couplings are

averaged out due to the rapid rotational diffusion of molecules in solution, resulting in sharp

peaks positioned at the isotropic chemical shift, albeit large molecular weight proteins and

complexes in solution can suffer from broad lines due to slower tumbling and correspondingly

faster T2 relaxation times.

19

In SSNMR, however, the samples have a restricted or non-existent tumbling motion compared to

molecules in solution, and are inherently less mobile on the NMR timescale. This manifests into

broadening of the NMR peaks due to observing chemical shifts as well as homonuclear and

heteronuclear dipolar interactions associated to each orientation of every spin, yielding a “powder

pattern” spectra for a static solid. Magic angle spinning (MAS) overcomes this limitation by

mechanically spinning the solid sample tilted at an angle (θ = 54.74o) with respect to the external

magnetic field (Figure 1.6 A). This causes the orientation-dependent anisotropic term (3cos2 θ –

1) to be time-averaged to zero at MAS spinning frequencies sufficiently greater (at least 3-4

times) than the anisotropies that are on the order of 102 to 105 Hz (Figure 1.6 B). Mechanical

spinning also adds a periodicity to the resonant frequency of nuclear spins that manifests as

spinning sidebands spaced at the MAS frequency, and approximately traces out the CSA pattern

when spinning rates are smaller than the anisotropy.

Figure 1.6. Magic-angle spinning in solid-state NMR. A) Schematic diagram showing the alignment of the NMR

sample at the magic angle (54.74o) with respect to the external magnetic field. (B) 1D 13C spectra of uniform 13C-

labelled valine powder collected with no spinning (static), and increasing MAS spinning speed, showing the

contribution of each nuclei in all orientations causing line broadening. MAS removes the anisotropic component of

chemical shift and dipole-dipole couplings to give a single peak at the isotropic chemical shift for each 13C nuclei

highlighted in different colors. Spinning sidebands (*) appear at intervals of the spinning frequency away from the

isotropic chemical shift peak when the MAS speed is not fast enough to completely average away the CSA.

20

1H is a popular nucleus for studying biological systems by NMR due to its high natural

abundance (99.9% in nature) and high γ for greater energy transition and signal sensitivity (γ1H is

4x greater than γ13C and 10x greater than γ15

N). However, typically used MAS frequencies of <

20 kHz are too slow to sufficiently eliminate 1H – 1H homonuclear dipolar couplings that are on

the order of 105 Hz, rendering 1H peaks broad and spectral resolution too poor for analysis.

Recent technical developments also allow NMR MAS probes to reach fast (> 60 kHz) and ultra-

fast MAS (> 100 kHz), along with pulse sequences that suppress 1H – 1H homonuclear

couplings, making 1H detection in SSNMR achievable in some instances (Zhou et al. 2009;

Linser et al. 2011; Zhou et al. 2012; Mroue et al. 2015). 13C and 15N are more suited for studying

biomolecules by SSNMR since their weaker dipolar interactions (< 103 Hz) are easily removed at

slower MAS frequencies. As such, we will be employing SSNMR experiments that use 13C and

15N detection for this thesis. Enriching peptides or recombinant proteins with 13C and 15N

isotopes (natural abundance of 1.1% and 0.37%, respectively) is achieved by either chemical

synthesis using isotope-labeled amino acids or expression and purification from E. coli. grown

on media supplemented with 13C-glucose and 15NH4Cl as the sole carbon and nitrogen sources,

respectively.

SSNMR pulse sequences that selectively reintroduce the dipolar couplings removed by MAS has

represented a major advancement for protein structure determination. As shown in Equation 1-2,

dipolar interactions are through-space interactions between two nuclei with the strength directly

related to the inverse cube of their distances (1/r3), hence data gathered from measuring dipolar

couplings can provide accurate internuclear distances (Duer 2004). Backbone-backbone,

backbone-side chain, and side chain-side chain distance constraints can therefore be obtained

using SSNMR for accurate protein structure determination.

1.3.3 Chemical shift and secondary structure

As mentioned above, the NMR chemical shift of a given nucleus results from shielding of the

external magnetic field due to surrounding electrons (Rule & Hitchens 2006). Since the chemical

environment of the protein backbone is strongly dependent on the torsion angles and ,

chemical shifts of backbone nuclei (i.e. CO, Cα, Cβ, Hα, Hβ, NH) can be a strong indicator of

secondary structure. Analysis of 1H, 13C, and 15N isotropic chemical shifts are routinely carried

out on solution and solid-state NMR data by comparing experimental values to standard random

coil values expected for amino acids in a peptide (Wishart et al. 1992; Wishart & Sykes 1994;

21

Wishart et al. 1995). The computer program TALOS (Torsion Angle Likelihood Obtained from

Shifts and sequence similarity) can predict the backbone torsion angles from chemical shifts of

an assigned protein by match strings of 3 residues to a database of high resolution structures with

chemical shift and sequence homology to the triplet of interest (Cornilescu et al. 1999).

Together, these data can give highly accurate secondary structure information for protein

structure determination. Inversely, it is also possible to accurately calculate 1H, 13C, and 15N

chemical shifts of backbone and side chain atoms based on existing protein structures (Neal et al.

2003; Han et al. 2011; Schneider et al. 2013).

1.3.4 Polarization transfer and SSNMR experiments

Spin pairs that are in spatial proximity can influence each other either via scalar-coupled

(through-bond) or dipolar-coupled (through-space) mechanisms. This property can be exploited

to induce the exchange of magnetization or polarization. There are two key benefits to

transferring 1H polarization to 13C and 15N for subsequent detection – first it increases the

magnitude of the observed signal by a factor of 4 and 10, respectively, due to the relative γ

between nuclei. Secondly, 1H has the added benefit of faster T1 relaxation rates allowing use of

shorter inter-scan delays and minimizing data acquisition times. Insensitive nuclei enhanced by

polarization transfer (INEPT), a scalar-coupling based transfer, and cross polarization (CP), a

dipolar-coupling based transfer, are both utilized in SSNMR experiments to transfer 1H

polarization to 13C or 15N (Morris & Freeman 1979; Pines et al. 1972). Polarization transfer is

also vital in acquiring multidimensional NMR because it correlates two spins based on their

proximity, generating a cross peak at a chemical shift position corresponding to the resonant

frequency of the two coupled spins. Many SSNMR experiments have been developed to exploit

this property to collect distance constraints (Table 1.2).

13C – 13C homonuclear correlation experiments provide information about 13C located in both

the backbone (ie. CO, Cα,) and side chains (Cβ, Cγ, etc) of amino acids. For example, RF-

assisted diffusion/dipolar-assisted rotational resonance (RAD/DARR), proton-driven spin

diffusion (PDSD), and proton-assisted recoupling (PAR) are experiments that quantitatively

measure dipolar interactions between 13C pairs within 7 Å of each other. In RAD/DARR, an RF

field applied to 1H at a frequency of the MAS spinning (ωr) or 2ωr reintroduces 1H – 13C and 1H

– 1H dipolar couplings to achieve 13C recoupling for polarization transfer (Morcombe et al. 2004;

Takegoshi et al. 2001). In contrast, the PAR experiment applies RF fields to both 1H and 13C

22

nuclei such that 13C polarization transfer relies not on 13C – 13C couplings but on 1H – 13C

couplings via a 13C1–[1H]–13C2 pathway (De Paepe et al. 2008). The advantage of PAR being

more efficient at mid- and long-range 13C – 13C distance measurements in a densely labelled

sample and applicable even under faster MAS conditions. By selecting a mixing period to allow

polarization exchange to occur between nuclear spins, the resulting spectrum shows cross peaks

between nearby 13C nuclei to facilitate intramolecular amino acid chemical shift assignment, or

between more distant 13C nuclei for intermolecular 13C – 13C contacts between residues.

There are also pulse sequences that can selectively reintroduce couplings between a specific pair

of 13C nuclei, but require more careful experimental setups. For example, constant-time

PITHIRDS (CT-PITHIRDS) utilizes RF-pulses synchronized to MAS spinning to achieve 13C –

13C dipolar recoupling (Tycko 2007). As the recoupling time increases, 13C polarization is

transferred away and the signal will decrease as a function of 13C – 13C distances. By

incorporating individual 13C-isotopes at known positions on the protein, one can compare the

experimental dephasing curve with simulated curves to measure the distance between the two

nuclei. Alternatively, rotational resonance (RR) experiments measure 13C – 13C distances by

changing the MAS frequency to match the recoupling condition for a unique pair of 13C nuclei

(Raleigh et al. 1988). These experiments therefore provide highly accurate internuclear distances

constraints between 2 specific nuclei to build structural models.

15N – 13C heteronuclear correlation experiments establish connectivities between 15N and 13C

nuclei located on the peptide backbone and/or side chains, and are commonly used to identify

sequential residues. Selective polarization transfer from an amide nitrogen to Cα of the same

residue (i) or CO of the previous residue (i-1) (called NCACX and NCOCX, respectively, in

conjunction with a transfer to CX through a 13C – 13C recoupling sequence following the 15N –

13C recoupling) help determine residues that are connected through the peptide backbone for

sequential chemical shift assignments (Petkova et al. 2003). In addition, we can also measure

backbone-side chain or side chain-side chain distances in an experiment called transferred-

echoed double resonance (TEDOR), where the 15N – 13C cross peak intensity builds up as a

function of recoupling time to reflect the distance between the two nuclei (Jaroniec et al. 2002;

Hing et al. 1993). Using this experiment, we can determine accurate distances between backbone

amide and side chain methyl groups, or between side chain amino and carboxylate functional

groups.

23

Table 1.2. SSNMR pulse sequences used in this thesis.

Pulse Sequence Correlated

nuclei Dimensions Application

Scalar coupling (through-bond) polarization transfers

INEPT 1H – 13C or 15N 1D Selective transfer for mobile regions of

protein (Pines et al. 1972).

HSQC 1H – 15N or 13C 2D Correlates 2 directly bonded nuclei (i.e.

backbone amide, side chain)

(Bodenhausen & Ruben 1980).

TOBSY 13C – 13C 2D Intra-residue 13C correlation for amino

acid assignment (Leppert et al. 2004).

Dipolar coupling (through-space) polarization transfers

Cross

Polarization (CP)

1H – 13C 1D Selective transfer for immobile regions

of protein (Pines et al. 1972).

RAD/DARR,

PDSD, PAR

13C – 13C 2D Intra-residue 13C correlation for amino

acid assignment (short mixing time) or

inter-residue 13C – 13C distance

constraints within 7 Å (long mixing

time) (Morcombe et al. 2004; Takegoshi

et al. 2001; De Paëpe et al. 2008).

Rotational

Resonance (RR)

13C – 13C 1D Quantitative 13C – 13C measurements

for side chain or backbone constraints;

require specific MAS speed for each

pair of 13C nuclei being examined

(Raleigh et al. 1988).

CT-PITHIRDS 13C – 13C 1D Quantitative 13C – 13C measurements

using dephasing curves to determine

inter-strand distances within β-sheets;

require incorporation of individual 13C-

isotope at known positions (Tycko

2007).

ZF-TEDOR 13C – 15N 2D Quantitative distance measurements

between 13C and 15N for backbone-

backbone or backbone-side chain

constraints (Jaroniec et al. 2002).

NCACX/NCOCX 13C – 15N 2D or 3D Intra- and inter-residue 13C – 15N

correlation for backbone sequential

assignment (Petkova et al. 2003).

24

1.3.5 Solid-state NMR models of amyloid fibrils

Using the above SSNMR methods, and others, researchers have been able to collect constraints

on backbone conformation, β-sheet organization, and ultimately the arrangement of cross-β

structural units in biological solids, including amyloid fibrils. SSNMR data have been used to

build high-resolution structural models of various amyloid systems, including the human

Alzheimer’s Aβ40 and Aβ42 peptides, transthyretin, mammalian PrP peptides, yeast Sup35, and

the yeast Het-S protein (Petkova et al. 2006; Schmidt et al. 2009; van der Wel et al. 2010; Van

Melckebeke et al. 2010; Jaroniec et al. 2004; Caporini et al. 2010; Wasmer et al. 2008; Cheng et

al. 2011; Walsh et al. 2009).

Alzheimer’s Aβ 40/42 peptides have been studied extensively by SSNMR, and are the most well

understood model for amyloid fibrils. Petkova et al have shown that the Aβ40 peptide can

generate polymorphic fibrils that are reflected in SSNMR measurements, as indicated by having

more than one set of chemical shifts to suggest inequivalent chemical environments (Petkova et

al. 2006; Paravastu et al. 2008). Recent structural studies of Aβ40 fibrils seeded with brain

extract also highlighted structural polymorphism in disease-related amyloid fibrils from different

Alzheimer’s patients (Lu et al. 2013). A cytotoxic fibril polymorph of Aβ42 was also solved by

SSNMR to contain a double-horseshoe-like cross-β sheet conformation (Wälti et al. 2016).

Meanwhile, structural studies of Het-S(218-289) fibrils demonstrate a highly ordered β-solenoid

core structure, with flexible N- and C-termini that do not play role in the structural core of

amyloid fibrils (Wasmer et al. 2008; Siemer et al. 2006).

Despite the wealth of knowledge we have gained about amyloids from SSNMR, understanding

the structural polymorphism and differences in cytotoxicity seen in pathogenic and functional

amyloids is still critical to address the biological behaviours of these protein aggregates.

Examining the structural properties of amyloid systems and their interactions with lipid

membrane can provide us insight into how amyloids interact with biological systems.

1.4 Mammalian prion protein (PrP)

1.4.1 Prions and diseases

“Prion” is used to describe the “proteinacious and infectious” agents in transmissible spongiform

encephalopathies (TSE), or prion diseases, of humans and mammals (Prusiner 1982). TSEs are

25

neurodegenerative disorders characterized by the accumulation of misfolded, protease-resistant

PrP as amyloids in the central nervous system. They are transmitted without nucleic acid and

require only the host-encoded precursor PrP. Most TSEs are limited to a specific host and in rare

cases to a few related species; some examples include Creutzfeldt-Jacob disease (CJD), fatal

familial insomnia (FFI), Gerstmann-Sträussler-Scheinker syndrome (GSS), and kuru in humans,

chronic wasting disease (CWD) in elk and deer, bovine spongiform encephalopathy (BSE) in

cattle, and scrapie in sheep (Béringue et al. 2008). In humans, approximately 85% of all

recognized prion disease are sporadic, with an annual worldwide incidence of 1 – 2 per million

(Béringue et al. 2008). The disease can also be acquired via contaminated surgical instrument,

blood transfusion, corneal grafts, consumption of contaminated foods (e.g. variant CJD from

consumption of BSE infected beef), or injection of cadaveric growth hormone (Kraus et al. 2013;

Aguzzi & Heikenwalder 2006). Meanwhile, hereditary forms of human prion diseases are caused

by mutations linked to specific disease phenotypes, and include FFI and GSS.

Mammalian PrP is a glycophosphatidylinositol (GPI)-anchored glycoprotein found mostly

associated in lipid raft of the plasma membrane. It is expressed predominantly in the central

nervous system, but is also found in a wide range of tissues including thymus, liver, intestine,

spleen, and heart (Peralta & Eyestone 2009). The 253-residue primary sequence contains an N-

terminal signalling sequence and C-terminus membrane anchoring sequence that are both post-

translationally removed during trafficking through the ER and golgi apparatus, leaving residues

23 to 231 in the mature protein. The resulting cellular form (PrPc) has a disordered N-terminal

region (residues 23 to 90) containing 5-8 repeats of a copper-binding octapeptide sequence, and a

globular 3-helix fold connected by 2 anti-parallel β-strands (Figure 1.7 A, B) (Zahn et al. 2000).

There are two N-glycosylation sites (Asn181 and N197), one disulfide bridge (Cys179 -

Cys214), and a GPI anchor at residue 231. PrP function is still highly elusive, with evidence

suggesting its importance in transmembrane signal transduction due to its localization to

membrane rafts, copper binding and trafficking, apoptotic and anti-apoptotic activity, and some

neuroprotective activity (Westergard et al. 2007; Aguzzi & Heikenwalder 2006; Freir et al.

2011).

26

Figure 1.7. Human prion protein (PrP) and its misfolded PrPsc state. (A) Schematic of PrP primary structure

and its post-translational modifications. Shown in green are peptide segments reported to form amyloid fibrils.

Adapted with permission from Aguzzi & Heikenwalder 2006. Copyright 2006 Nature Publishing group. (B)

Solution NMR structure of human PrP 90-231 (1QM0). Residues 90-120 were highly disordered and therefore not

deposited with the model. (C) Trimer of left-handed β-helix model for PrPSc determined from electron

crystallography and X-ray fibre diffraction. Reprinted with permission from Wan et al. 2015. Copyright 2015 Prion.

(D) Parallel β-sheet model of PrPamyloid determined using SSNMR. Reprinted with permission from Groveman et al.

2014. Copyright 2014 Journal of Biological Chemistry.

Soluble PrPc converts into the infectious scrapie form (PrPSc) in prion diseases through template-

assisted (PrPSc) misfolding and structural reorganization into β-sheet rich amyloid fibrils. In

particular, the globular region of the protein adopts a proteinase-K (PK) resistant rigid amyloid

core that resists H/D exchange, and the disordered N-terminus thought to play a role in

modulating the oligmerization and aggregation of PrP (Cordeiro et al. 2005; Lu et al. 2007).

Recent solution NMR studies on non-native stable oligomers of syrian hamster ShaPrP(90-231)

have shed light into the misfolding mechanism of PrP. Native PrPC misfolded into β-monomers

are entropically driven to form stable β-octomers through burial of hydrophobic groups exposed

27

after the loss of native structure, and further driven by enthalpy to assemble into larger oligomers

that become the nuclei for amyloid formation (Larda et al. 2013). The structure of PrPSc is not

yet known, although multiple structures of the amyloid core have been proposed from different

studies. These include an extended parallel β-sheets model from solid-state NMR measurements

(Groveman et al. 2014), a left-handed β-helix model from X-ray fibre diffraction and electron

microscopy (Wille et al. 2004; Wille et al. 2009), β-spiral model from molecular dynamics (MD)

simulation (DeMarco & Daggett 2004), and a domain-swapped model (Yang et al. 2005) (Figure

1.7 D).

1.4.2 Prion propagation and prion strains

The protein-only hypothesis in prion disease describes that propagation of the misfolded PrP

requires only the infectious prion state to template the misfolding of host native protein, and that

the disease spreads by finding naïve hosts as a new pool for replication (Kraus et al. 2013). Yet

despite having identical protein sequence, it was recognized that the same species can develop

different pathologies and clinical symptoms from the same infectious agent (Pattison & Millson

1961). The idea of prion strains was introduced to rationalize these differences in vacuolar

lesions in the brain area, disease incubation times after inoculation, and clinical symptoms

(Collinge & Clarke 2007; Morales et al. 2007). At the atomic level, prion strains describe the

polymorphism in misfolded PrP conformations due to the stochastic nature of the initial

nucleation process that lead to different nucleation pathways (Lu et al. 2013), and result in

assemblies with different biochemical and physical properties such as thermodynamic stability,

proteolytic sensitivity, glycosylation pattern, conformational selection, transmission barrier, and

epitope exposure (Safar et al. 1998; Bessen & Marsh 1994; Collinge & Clarke 2007). These

strain-specific properties are faithfully propagated in a template-mediated mechanism similar to

amyloid seeding, and ultimately leads to distinct fibril structure with different exposed epitopes

and fibril stability. The complexity of prion fibrils warrants further structural studies in

describing amyloid fibril polymorphism in order to gain a deeper understanding about prions

strains, prion propagation, and infectivity.

1.4.3 Structural studies of PrP peptides

The structure of the PrPSc state has puzzled investigators for many years because of the various

conformations it can adopt. As a result, many have preferred to investigate short PrP peptides as

28

a reductionist approach to understand the structure of PrPSc. Peptide fibrils are easier to prepare,

have reduced sequence complexity for unambiguous NMR chemical shift assignment and

structural determination, and have similar biophysical properties to amyloid fibrils of full-length

protein. Many segments from the globular fold of PrP exhibit amyloidogenic propensities and

have been used to study amyloid toxicity (Figure 1.7 A) (Vilches et al. 2013; Cheng et al. 2011;

Sonkina et al. 2010; Tagliavini et al. 1993; Pastor et al. 2008; Thompson et al. 2000; Laws et al.

2001).

One of the most well-studied peptides consists of residues 106-126 (PrP(106-126)) located in the

hydrophobic domain. This peptide forms β-rich amyloid fibrils that are detergent-insoluble and

PK-resistant, as well as non-fibrillar spherical oligomers that are cytotoxic in neuronal cell

cultures ((Ettaiche et al. 2000; Forloni et al. 1993; Thellung et al. 2000). Structural studies

indicate the palindromic sequence AGAAAAGA (residues 116 to 123) forms the key steric

zipper core in both the amyloid fibrils as well as the oligomers (Walsh et al. 2009; Walsh et al.

2010), suggesting the soluble oligomer may represent an on-pathway intermediate to fibril

formation. In addition to inducing apoptosis, the amyloid oligomers of PrP(106-126) were also

demonstrated to form ion channels or permeabilize membranes formed by mixture of

PC/PE/PS/cholesterol (Lin et al. 1997; Kayed et al. 2004). A recent study by Walsh et. al. showed

that PrP(106-126) has different modes of membrane disruption when exposed to membranes

composed of different lipid mixtures. PrP(106-126) oligomers in the presence of anionic lipid 3:1

PC/PG acted in a detergent-like manner and solubilized membrane to form small vesicles; while in

cholesterol-containing mixture 1:1:1 DSPC (1,2-distearoyl-sn-glycero-3-phosphocholine) / DOPC

(1,2-dioleoyl-sn-glycero-3-phosphocholine) / cholesterol, the oligomers fibrillized on the surface of

the membrane and caused a loss of ordered-domain structure (Walsh et al. 2014). This reorganization

of cholesterol-containing ordered domains and disruption of the membrane may reflect possible

mechanisms in which amyloid oligomers can cause cell death in prion diseases.

Unfortunately, other PrP peptides have not been extensively studied to draw parallels between

their structures, membrane interactions, and toxicities. SSNMR reports on fragments 113-127

and 89-143 have characterized their amyloid β-sheet secondary structure but with poorly defined

models on the peptide assembly (Laws et al. 2001; Cheng et al. 2011). Additional fragments in

the hydrophobic domain of the prion protein – 105-132, 118-135, and 120-133 – formed

amyloids that destabilized membranes and caused cell death, yet very little structural data has

29

been collected to correlate amyloid assembly to membrane interaction and toxicity (Vilches et al.

2013). Meanwhile, fragment 185-206 was shown to destabilize anionic PC/PS model membranes

but the active structure is unknown (Sonkina et al. 2010); fragment 178-193 caused cell death in

the presence of copper, lending support to the involvement of copper in prion neurotoxicity

(Thompson et al. 2000); and fragments 245-250, 244-249, and 178-183 formed amyloids that

exhibit varying levels of cytotoxicity, but their structure and membrane disruption activity have

not been addressed (Pastor et al. 2008). In order to fully understand the structural complexity of

PrP conformations in different strains and address the mechanism of neurotoxicity, a more

cohesive study on amyloid structure, toxicity, as well as amyloid-membrane interaction is

necessary.

1.5 Serum Amyloid A

1.5.1 Acute-phase serum amyloid A and amyloid A (AA) amyloidosis

Serum amyloid A (SAA) is an acute-phase apolipoprotein up-regulated as part of the

inflammatory cascade response (Uhlar et al. 1994). Challenges such as tissue injury and

bacterial/viral infection trigger the production of cytokines interleukin-1 (IL-1), IL-6, and tumour

necrosis factor (TNF) in acute inflammation, leading to the induction of SAA and its rise in

serum from μg/mL to nearly mg/mL within hours (Uhlar & Whitehead 1999). SAA is

predominantly synthesized by hepatocytes and is primarily associated with high-density

lipoproteins (HDLs), although expression has been detected in extrahepatic cell-lines (Upragarin

et al. 2005; Sjoholm et al. 2005; Eckhardt et al. 2010). It is highly conserved across vertebrates,

suggesting a biological advantage to having SAA participate in part of the short-term anti-

inflammatory response (Uhlar et al. 1994). Multiple SAA genes and proteins have been

described for several species; human and mouse have four and five members, respectively, and

are the most extensively studied. Additionally, there are at least three genes in dog, mink, rabbit,

and horse, two in hamster, and one in cow and sheep (Uhlar & Whitehead 1999).

The human SAA family contains members SAA1, 2, 3, and 4. SAA1 and SAA2 are up-regulated

during the inflammatory response (A-SAA); SAA3 is considered a pseudogene due to single

base frameshift causing a premature stop codon and preventing expression in humans; and SAA4

is constitutively expressed in humans (C-SAA) (Uhlar & Whitehead 1999; Westermark &

Westermark 2009; Uhlar et al. 1994; De Beer et al. 1995). A-SAA has been shown to displace

30

ApoA-I and becomes the major apolipoprotein on HDLs during its up-regulation, and circulates

in the serum associated to the HDL3 subclass (Malle & de Beer 1996; Clifton et al. 1985;

Coetzee et al. 1986), while C-SAA is found associated with all HDL classes and very low

density lipoproteins (VLDL), and normally accounts for 90% of all SAAs on HDL particles

(Whitehead et al. 1992; De Beer et al. 1995). Although the primary physiological role of SAA is

still largely unknown, some functions have been proposed based on a range of studies in mice

models and in cell culture. These include altering cholesterol transport and metabolism,

upregulating of pro- and anti-inflammatory cytokines, removing endotoxins during infection,

inducing ECM-degrading enzymes after tissue damage, and facilitating the migration and

adhesion of neutrophils, lymphocytes, and mast cells at sites of inflammation (Urieli-Shoval et

al. 2000; Derebe et al. 2014; Preciado-Patt et al. 1996; Xu et al. 1995; Eckhardt et al. 2010).

Persistent high levels of A-SAA are observed in patients suffering from chronic inflammatory

diseases such as rheumatoid arthritis, familial Mediterranean Fever (FMF), and Crohn’s Disease,

or from chronic infections such as leprosy and tuberculosis. This long-term up-regulation of A-

SAA from both hepatocytes and non-hepatic cells leads to a systemic amyloid A (AA)

amyloidosis in approximately 15% of patients suffering from a chronic inflammatory disease

(Röcken & Shakespeare 2002). The misfolding and aggregation of SAA results in the deposition

of amyloid aggregates throughout the body, with affected organs including the kidney, liver,

spleen, gastrointestinal tract, and the vascular system (Perfetto et al. 2010). Kidneys are the most

frequently affected organ in AA amyloidosis, and renal amyloid deposits inevitably manifest as

renal dysfunction, proteinuria, and ultimately organ failure if left untreated. Currently, the

primary intervention against AA amyloidosis is to suppress the inflammatory condition by

treating the underlying disease/infection, or to use anti-TNF and anti-IL-1 immunosuppressive

therapies to decrease the levels of circulating SAA (Bilginer et al. 2011; Rumjon et al. 2012).

Patients with AA amyloidosis have a median survival time of 2 - 10 years, with the intervening

strategies prolonging renal survival. However, dialysis or renal transplantation are the only

treatment options once end-stage renal disease develops.

1.5.2 Structure of SAA

Human SAA1 and SAA2 contains 104-residues that share 96% sequence homology, with allelic

variants exhibiting sequence changes at 1-6 amino acids, with the exception of SAA2.2 (Figure

1.8 A). Both SAA1 and SAA2 were initially predicted to adopt a predominantly α-helical

31

structure with a flexible C-terminus (Nordling & Abraham-Nordling 2012). Lipid-free (apo)

SAA oligomerizes in solution, forming both hexamers and octamers, with the hexamer being

slightly more stable (Wang et al. 2005; Wang et al. 2012). The structure of human apo-SAA1.1

was recently solved as a hexameric assembly formed from 4-helix-bundled monomers containing

an unstructured C-terminus stabilized by bifurcated salt bridges between Tyr104, Tyr35, Arg39

and Arg96 (Figure 1.8 B) (Lu et al. 2014). The N-terminal helix 1 (residues 1-27) is amphipathic

and has been shown to interact with 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC)

(Segrest et al. 1976). Residues 29-42 contain sequences similar to the cell binding domains of

extracellular matrix cell adhesive glycoproteins laminin (YIGSR) and fibronectin (RGD), and

can inhibit adhesion of human T-lymphocytes to these glycoproteins (Uhlar & Whitehead 1999).

Meanwhile, C-terminal residues 76-104 have been shown to interact with cystatin-C, neutrophils,

and glycosaminoglycans (GAG) such as heparan and heparin sulfate, and may play a role in

folding, oligomerization, and fibrillization of the protein (Ohta et al. 2009; Patke et al. 2012;

Maszota et al. 2015).

32

Figure 1.8. Human Serum amyloid A. (A) Sequence alignment of human SAA1 (isoforms 1-5), SAA2 (isoforms

1-2), and SAA4. Yellow boxes showcase sequence variations between SAA1 and SAA2 isoforms. Regions of

functional importance are underlined in colour and mapped to the sequence, while the arrow points to the possible

N-glycosylation site (NSS) in SAA4. (B) 4-helix bundle crystal structure of human SAA1.1 (4IP9) assembled as a

hexameric ring composed of a dimer of trimers. Two clusters of positive charge identified in the hexamer – cluster 1

(Arg1, Arg-62, and His-71) in the outer apex and cluster 2 (Arg-15, Arg-19, Arg-47) in the centre-ring – were

shown to affect SAA binding to both HDL and GAGs. Reprinted with permission from Lu et al. 2014. Copyright

2014 Proceedings of the National Academy of Sciences of USA.

It is important to note that there may be differences in the functionality of SAA between the apo-

and HDL-bound states. Apo-SAA can activate pro-inflammatory pathways, rather than acting in

an anti-inflammatory manner observed for HDL-bound SAA (Christenson et al. 2013). This may

be applicable during chronic inflammation, during which extrahepatic SAA expression in

adipose tissue, intestinal epithelia, and macrophages are increased with a corresponding increase

in apo-SAA (Sjoholm et al. 2005; Eckhardt et al. 2010). At the time of this study, the structure of

33

the HDL-bound human SAA had not yet been reported. Likewise, the structural basis for SAA

function based on HDL-binding and sequence variations are not yet known. In particular, what is

the mechanism of SAA association to HDL particles, the structure that SAA adopts when bound

to HDL, and how HDL-association affects the function of SAA. These are all important

questions that we hope to address in order to fully understand the diverse roles that SAA plays

during inflammation.

1.5.3 High density lipoproteins (HDLs)

HDLs are protein-lipid assemblies that transport peripheral tissue cholesterol back to the liver for

excretion or hormone metabolism as part of the reverse cholesterol transport pathway (RCT)

(Figure 1.9) (Barter et al. 2003). They are heterogeneous complexes composed of

apoliproproteins (ApoA-I, II, IV, ApoC – M, SAA), lipids (triacylglyerides (TG), phospholipids,

sphingolipids), sterols (cholesterol, cholesterol ester (CE)) and other enzymes (Kontush et al.

2015). HDLs are constantly remodelled and turned over as part of cholesterol transport such that

they vary in size, shape, and composition that range from 7-13 nm in diameter (Guha et al. 2008;

Gursky 2015). Nascent HDLs are the smallest in size, low in cholesterol, and mainly composed

of 2 ApoA-I wrapped around phospholipids forming a discoid shape; meanwhile HDLs are

matured into spherical particles with an exterior envelop of lipids and apolipoproteins and an

apolar core containing CE and TG (Shih et al. 2009). Mature HDLs can be separated by

ultracentrifugation into 2 major subclasses - the smaller and denser HDL3 (1.125 < density <

1.21 g/mL) and larger and less dense HDL2 (1.061 < density < 1.125 g/mL) - to reflect the

protein-to-lipid ratio of the particle (Barter et al. 2003). Apolipoproteins can freely associate and

dissociate during HDL remodeling and fusion to compensate for the imbalance between particle

core volume and surface area. As such, apolipoproteins have properties that facilitate this

behaviour including a compact globular shape with loosely folded tertiary structure, low-

cooperative unfolding and low heat capacity increment, and amphipathic helices with high

propensity to bind to apolar surfaces (Gursky 2015).

34

Figure 1.9. Synthesis and remodeling of high density lipoproteins in reverse cholesterol transport (RCT). The

lipid transporter ABCA1 promotes transfer of phospholipids and cholesterol to the cytoplasmic leaflet at peripheral

cell membrane. This creates a protrusion of the membrane that lipid-free apoA-I can interact with to form nascent

discoidal HDLs that can subsequently take up additional cholesterol. Remodeling and maturation of HDLs is carried

out by various enzymes: lecithin-cholesterol acyltransferase (LCAT) esterifies cholesterol into cholesterol esters,

cholesterylester transfer protein (CETP) exchanges CE for triacylglycerol (TG), and phospholipid transfer protein

(PLTP) inserts additional phospholipids into HDLs. CE and TG are sequestered into the apolar core due to their

hydrophobicity to drive the formation of mature spherical HDL particles. At the hepatocyte, apolar core lipids are

taken up with the help of scavenger receptor B1 SR-B1 and the HDLs disintegrates. Adapted with permission from

Guha et al. 2008. Copyright 2008 American Chemical Society.

1.5.4 Glycosaminoglycans (GAGs) and AA amyloidosis

Clinical amyloid isolates from patients with AA amyloidosis contain the N-terminal 76 residues

of SAA, in addition to glycosaminoglycans (GAGs) and serum amyloid P (Husebekk et al. 1985;

Snow et al. 1987; Snow et al. 1991). GAGs are known to be amyloid-enhancing factors that

accelerate fibrillization in many in vitro amyloid systems such as Aβ, PrP, and α-Syn (Ancsin &

Kisilevsky 1999; Sipe & Cohen 2000; Madine et al. 2012; Madine et al. 2013). They are highly

sulfated, unbranched heteropolysaccharides found in proteoglycans, on the cell surface, in the

extracellular matrix, and in circulation that play an important biological role in wound healing by

promoting collagen polymerization and helping cells anchor to surrounding matrix (Prydz &

Dalen 2000). The highly negatively charged functional groups are proposed to recruit protein via

electrostatic interactions with basic residues Arg, His, and Lys (Rabenstein 2002). This can

affect the conformation of the interacting proteins by modifying the native interactions to

35

promote aggregation, or behave by scaffolding monomers to increase local concentrations for

misfolding. SAA-GAG studies carried out using heparan sulfate (HS) show that it can induce an

SAA transition to β-sheet structure in vitro, and it plays a role in inducing AA amyloidosis is cell

culture and mouse models (Ancsin & Kisilevsky 1999; De Beer et al. 1993). Inhibition of the HS

– SAA interaction renders mouse models resistant to AA amyloidosis (Li et al. 2005). In

addition, treatment of mice with the small sulfonated molecule Eprosidate slowed the decline in

renal function, suggesting a reduction in amyloidosis (Merlini et al. 2007). SAA interaction with

GAGs such as HS, and their co-deposition in AA amyloidosis, is therefore likely to be important

for disease onset and for disease progression. Studying their interaction and structural properties

may offer potential new avenues for clinical treatments of AA amyloidosis.

Figure 1.10. Glycosaminoglycans in the body. Various GAGs are synthesized from the golgi apparatus and

exported to cell surfaces and ECM either as proteoglycans or free GAGs. They include heparin/heparan sulfate

(Hep/HS), chondroitin sulfate, dermatan sulfate, and keratan sulfate that differ in the position of the functional

groups (sulfate or carboxylate) and charge distribution. (X = H or SO3-; Y = SO3

- or Ac)

1.6 Rationale and Hypotheses

A shared core structure in amyloids has generally been considered to be implicated as a

determinant of their cytotoxicity. However, the existence of functional amyloid in mammals and

yeast suggests there are more factors that play role in the gain-of-toxic effects of the amyloid

36

state. Instead, differences in amyloid assemblies at the atomic level, and/or their interaction with

lipid membranes, likely reflects the difference in biological behaviour between functional and

pathological amyloid fibrils. This highlights the importance of studying well-defined amyloid

systems to gain further insight into sequence determinants of amyloid assembly and structure. At

the same time, it is important to study the lipid interactions of amyloid proteins to understand the

underlying mechanisms that lead to cellular toxicity. As such, there are three goals to address in

my thesis:

1. Examine the structural differences between amyloids of three prion protein fragments

PrP(178-183), PrP(244-249), and PrP(245-250) cited to exhibit differing levels of

cytotoxicity (Pastor et al. 2008).

2. Examine the effects of fibril sonication on the prion peptide fibrils and determine the

ability of the fragmented fibrils to interact with lipid membranes.

3. Characterize the structure of human SAA2 in its lipid-bound and amyloid states to begin

understanding its transitions between the native state and misfolded disease-causing

amyloid state.

1.6.1 Structural characterization of prion peptide fibrils

Three segments from the human prion protein, PrP(244-249), PrP(245-250), and PrP(178-183),

were previously identified from an amyloid prediction algorithm, along with several other

hexapeptide stretches from naturally occurring amyloid proteins, to spontaneously fibrillize in

vitro (Lopez de la Paz et al. 2004). These segments were later demonstrated to be cytotoxic in

cell cultures to varying degrees (Pastor et al. 2008). In chapter 2, we used biophysical techniques

including ThT fluorescence, transmission electron microscopy (TEM), fourier-transformed

infrared spectroscopy (FTIR), and SSNMR, to characterize the high-resolution structures of

these three peptides and thus to compare the molecular organization of cytotoxic and non-

cytotoxic amyloids. We hypothesized that two of these newly identified peptides, PrP(244-249)

and PrP(245-250) that demonstrated levels of toxicity in rat PC12 cell cultures comparable to the

Alzheimer’s Aβ derived peptide Aβ(16-21), would exhibit a different molecular arrangement of

β-sheets than the third, non-cytotoxic peptide, PrP(178-183).

37

Figure 1.11. Sonication-enhanced amyloid cytotoxicity of short amyloidogenic peptides. PC12 cell viability

upon incubation with various sonicated hexapeptide amyloids, as measured by a MTT reduction assay after 24 hrs.

The varying degree of cell death between these sonicated fibrils suggest differences in toxicity between various

amyloidogenic sequences. Black bars refer to 50 μM, grey bars to 10 μM, and white bars to 5 μM. Reprinted with

permission from Pastor et al. 2008. Copyright 2008 Elsevier.

1.6.2 Structural changes of cytotoxic fibril fragments

Sonication of mature fibrils has previously been demonstrated to enhance cytotoxicity of

amyloid fibrils formed by a number of proteins and peptides (Pastor et al. 2008; Xue et al. 2009;

Jakhria et al. 2014). We hypothesized that the sonication of long mature fibrils produces smaller

intermediate species that will much more readily disrupt lipid membrane, either by increasing the

amount of fibril ends that can release soluble toxic species such as small oligomers, or by

exposing hydrophobic patches that can interact with cell membrane, nucleating fibrils directly on

membrane or disrupting the lipid bilayer (Ahmed et al. 2010; Stefani 2010; Milanesi et al. 2012).

In chapter 3, we examined the structural changes of the three PrP peptide fibrils by ThT

fluorescence, electron microscopy, and SSNMR upon sonication of mature fibrils. We also

compared the membrane disruption caused by these sonicated fibrils in a fluorescence-based

liposome leakage assay, and the degree of hydrophobic residue exposure upon sonication, using

ANS fluorescence. This allowed us to correlate structural differences and membrane disruption

activities of these fibrils with cytotoxicity.

1.6.3 Structural characterizations of human SAA2

A-SAA is involved in both the acute-phase inflammatory response, but gets deposited as amyloid

fibrils in AA amyloidosis. It can exist in the body in a lipid-free apo state, an HDL-associated

state, and an amyloid fibrillar state, each with specific biological activities. In order to examine

the functional differences between these three SAA states and to elucidate the mechanism of

38

conversion into amyloid fibrils, we first need to address the structures of the HDL-bound SAA as

well as amyloid-state SAA. In chapter 4, we used solid-state NMR along with complementary

biophysical techniques to examine the structure of SAA when it is incorporated into HDL-like

particles, and when it is fibrillized in the presence of heparin. We hypothesize that HDL-bound

SAA adopts a different structural organization than the apo-SAA to carry out different functions,

and that in the HDL-bound form, SAA is at least partly protected from fibril formation relative to

the apo-form of the protein. At the same time, it must undergo large structural rearrangement

from α-helical native state into β-sheet amyloid state.

39

Chapter 2.

Structural comparison of three

amyloidogenic hexapeptides

from the human prion protein

References:

Sections of this chapter were originally published from Yau, J., and Sharpe, S. Structures of

amyloid fibrils formed by the prion protein derived peptides PrP(244-249) and PrP(245-250).

Journal of Structural Biology. 2012. 18(2). 290-302 and reprinted with permission. Copyright

2012 Elsevier.

Contributions:

The Advanced Protein Technology Center at Sickkids (now SPARCS centre) provided initial

unlabeled PrP peptides. Dr. Christopher Yip (University of Toronto) provided assistance in

FTIR measurements. Dr. Marie-Claude Jobin (University of Toronto) and Tatiana Skarina

(University of Toronto) assisted in collection of X-ray fibre diffraction data.

40

2.1 Introduction

Amyloid fibrils share a common cross-β structural backbone comprised of self-repeating β-

strands aligned perpendicular to the fibril elongation axis (Geddes et al., 1968; Tycko, 2000).

The strands are spaced 4.7 Å apart to form an extended β-sheet, which are stacked ~10 Å apart.

(Geddes et al., 1968; Greenwald and Riek, 2010). This “steric zipper” model for amyloid

structure was proposed based on X-ray diffraction of peptide microcrystals, and highlights how

interdigitating side chains allow β-sheets to tightly assemble to exclude water molecules at the

interface. This burial of hydrophobic residues into the fibril core, much like the hydrophobic

effect in protein folding, provides an important stabilizing force to the amyloid structure. It is

also important to consider that these side chain interactions further govern the association of β-

sheets into higher-ordered assemblies, such as protofilaments assembling into mature fibrils

(Sawaya et al. 2007). Thus, there is an important need to understand how amino acid sequence

affects fibril formation, and how it determines the molecular features of amyloids that govern

their different biological behaviours.

In a recent study by Lopez de la Paz et al., an amyloid prediction pattern was used to identify

small stretches of putative amyloid-forming hexapeptides from sequences of naturally occurring

amyloid proteins (Lopez de la Paz and Serrano, 2004). Several of these segments were associated

to previous reports of amyloid-forming regions of proteins, while others had not been identified

before – including fragments of the human prion protein (PrP) and ApoA-I. These

amyloidogenic sequences were subsequently shown to spontaneously fibrillize in vitro and the

majority exhibited cytotoxicity to neuronal cell culture when fragmented by ultrasonication

(Pastor et al., 2008). Two of these newly identified peptides, corresponding to residues 244-249

and 245-250 of human PrP, demonstrated levels of toxicity in rat PC12 cell cultures comparable

to the Alzheimer’s Aβ derived peptide Aβ(16-21), while a third PrP peptide from residues 178-

183 did not.

PrP(244-249) ISFLIF and PrP(245-250) SFLIFL are located in the C-terminus of the PrP pre-

polypeptide and are removed from the mature GPI-anchored PrP (Figure 1.7). However, the

amino acid sequences of both peptides share a highly hydrophobic and aromatic chemistry with

Aβ(16-21) KLVFFA, a segment of the Aβ fibril core. On the other hand, PrP(178-183) DCVNIT

is a short segment from helix 2 of PrP protein. This region of the protein forms part of the

41

extended beta sheet core region of amyloid fibrils in full-length PrP, and has been proposed to

play an important role in the oligomerization and aggregation of PrP (Groveman et al. 2014;

Singh et al. 2014). Currently, the links between sequence, structure, and cytotoxicity of amyloid

fibrils are poorly understood beyond the context of individual amyloid peptides. Studying the

structure of these three peptides can provide us insight into the assembly process of fibrils and

how the sequence affects both the amyloid structure and variable cytotoxicity of amyloid fibrils.

In this chapter, we determined the structures for the amyloid fibrils formed by PrP(244-249),

PrP(245-250), and PrP(178-183) using high resolution SSNMR data. All three peptides

exhibited an in-register parallel arrangement of strands within the individual β-sheets, but

differed in the packing of adjacent β-sheets within the fibril core (Figure 1.2). PrP(245-250)

adopted a class 1 zipper motif, with an anti-parallel face-to-face packing of opposing sheets, and

is stabilized by salt bridges formed between the N- and C-termini. Despite having similar

sequence and the same net hydrophobicity, PrP(244-249) formed fibrils with two distinct inter-

sheet interfaces. In these fibrils, both class 2 and class 3 steric zippers were observed, giving

parallel face-to-back and face-to-face packing of sheets, respectively. The presence of two

inequivalent sets of chemical shifts at equal proportion for all sites in PrP(244-249) strongly

suggests an equal population of the two packing types within fibrils formed by this peptide –

both largely determined by optimal packing of hydrophobic side chains. PrP(178-183) adopted a

class 1 steric zipper with hydrophobic residues Val and Ile packed within the core. Meanwhile,

Cys, Asn, and Thr are aligned on the opposite face of the β-sheet and can provide additional

stability by forming polar zippers. These structures provide new insight into the forces driving

amyloid assembly, and are a starting point for probing the molecular basis of cytotoxicity in

these peptides.

2.2 Methods

2.2.1 Synthesis of uniformly and selectively labeled peptides

Prion peptides PrP178-183 D178CVNIT183, PrP244-249 I244SFLIF249, and PrP245-250

S245FLIFL250 were prepared by solid phase peptide synthesis on Wang resin (Novabiochem),

using standard 9-fluorenylmethoxycarbonyl (FMOC) chemistry with O-(7-Azabenzotriazol-1-

yl)-N,N,N',N'-tetramethyluronium hexafluorophosphate (HATU) and N,N-

diisopropylethylamine (DIEA) activation. FMOC-protected, uniformly 13C and 15N-labelled

42

amino acids, [1-13C]-labelled phenylalanine, [1-13C]-labelled isoleucine, and [1-13C]-labelled

valine were purchased from ISOTEC stable isotopes. Peptides were cleaved from the resin using

a cocktail of thioanisole, ethanedithiol, anisole, and 90% TFA (trifluoroacetic acid)

(5:2.5:2.5:90) for 2.5 hours and precipitated from the cleavage solution using cold diethyl ether.

Precipitated peptides were washed twice with cold diethyl ether before dissolving in 25%

acetonitrile: 75% water: 0.1% TFA and lyophilizing. Peptides were purified by reverse-phase

HPLC using a water/acetonitrile gradient with 0.1%TFA on an 11 x 300mm C18 peptide column

(Vydac). The identities of purified peptides were confirmed by MALDI mass spectrometry

before lyophilizing and storing at -20oC.

2.2.2 Preparation of amyloid fibrils

To induce fibril formation, lyophilized peptide was first dissolved in hexafluoroisopropanol

(HFIP) (Fluka) at 10mg/mL and bath sonicated for 5 minutes to disaggregate the peptide. The

monomeric peptide was then incubated at room temperature for an additional 10 minutes before

evaporating HFIP under a gentle stream of N2(gas). The dried peptide film was re-dissolved in

filtered 20mM glycine-HCl buffer (pH 2.6) at a concentration of 1mg/mL for PrP(244-249) and

PrP(245-250), and 2mg/mL for PrP(178-183). Fibril formation was allowed to proceed at room

temperature for 1 week, after which the presence of amyloid fibrils confirmed by transmission

electron microscopy (TEM), and samples were stored at room temperature for further

experiments.

For seeding experiments a solution of mature fibrils (parent sample) was diluted 1:1 in 20 mM

glycine-HCl buffer (pH 2.6) and probe sonicated (10% amplitude output) on ice in 5-sec on/off

pulses for a total of 2 min. This was then added to a solution of monomeric peptide (1mg/mL) at

a concentration of 5% (v/v) seeds. This solution was then incubated for 24 h until fibril

formation was complete. Four rounds of seeding were performed, each time using seeds

produced from the previous generation of fibrils. Samples were taken for TEM analysis of fibril

morphology, and the first and last generations were analyzed by SSNMR spectroscopy.

SSNMR samples with mixed labelling schemes were prepared by dissolving the two 13C, 15N-

labelled peptides (Table 2.1) separately in HFIP at 10mg/mL and sonicating for 5 minutes, then

mixed together and incubated at room temperature for additional 10 minutes before evaporating

HFIP and fibrilizing in 20 mM glycine-HCl (pH 2.6).

43

Table 2.1. 13C/15N isotope-labeling schemes of the amyloidogenic prion peptide sequences used.

Peptide Name Position of Isotope label

PrP(244-249) SFLI IS245F246L247I248F (Uniform-13C, 15N)

ISIF I244S245FLI248F249

SI IS245FLI248F

IL I244SFL247IF

F ISF246LIF

F246CO IS(1-13C)F246LIF

I248CO ISFL(1-13C)I248F

PrP(245-250) Uniform All residues labeled

SI S245FLI248FL

F SF246LIFL

F246CO S(1-13C)F246LIFL

F249CO SFLI(1-13C)F249L

PrP(178-183) DCN D178C179VN181IT

VI DCV180NI182T

T DCVNIT183

V180CO DC(1-13C)V180NIT

I182CO DCVN(1-13C)I182T

2.2.3 Transmission electron microscopy (TEM) and X-ray fibre

diffraction

A 4 µL sample was absorbed onto freshly glow-discharged 200-mesh carbon films prepared

from copper rhodium grids (Electron Microscopy Sciences) for 2 minutes. The grids were

washed with 10µL MilliQ water, stained with 10µL freshly filtered 2% uranyl acetate followed

by subsequent blotting, and air dried at ambient temperature. TEM images were obtained using a

JEOL 1011 microscope operating at 80keV.

Dehydrated fibril stalks for X-ray fibre diffraction were prepared by suspending a 10 µL droplet

of fibril solution (5 mg/mL) between two wax-tipped glass capillary to allow to evaporate at

room temperature (Morris & Serpell 2012). Diffractions were collected on an R-Axis HTC

image plate X-ray detector with a 2 min exposure. Diffractograms were processed by iMosflm

and CLEARER software (Makin et al. 2007; Battye et al. 2011).

44

2.2.4 Thioflavin T (ThT) Fluorescence

Stock solutions were prepared by dissolving ThT at 2.5 mg/mL in 20mM sodium phosphate

buffer (pH 7.5), giving a concentration of 8 mM. ThT stocks were freshly prepared and stored in

the dark prior to use. Final samples for fluorescence spectroscopy contained 20 µM ThT and 0 –

200 µM peptide, with the remaining volume adjusted using to 1.0 mL with 20 mM sodium

phosphate buffer (pH 7.5) unless otherwise stated. ThT fluorescence emission spectra were

recorded from 450 to 600 nm using a Photon Technology International C60 spectrofluorometer

with excitation at 442 nm. A 2 nm slit width was used for excitation and 5 nm for emission. ThT

fluorescence spectra from PrP peptide containing samples were compared with a control solution

of 20 µM ThT in phosphate buffer in the absence of peptide. An emission maximum observed at

482nm was used as the signature of dye binding to cross-β containing peptide assemblies in each

case.

2.2.5 FTIR measurements

FTIR spectra were acquired on a Nicolet Nexus 670 spectrometer system equipped with a mid-

range IR source, a KBr beamsplitter and an MCT-A detector on the Continuum microscope

attachment. Thin films of fibril samples were air-dried on 32 mm cesium chloride (CsCl) discs

before collecting spectra on the microscope attachment in transmission mode. Spectra were

averaged from 256 scans over a range of 4000–650 cm-1 wavenumbers at 4 cm-1 resolution, and

were baseline-corrected using the OMNIC software package (Nicolet/Thermo Electron). Second

derivatives of the amide I and II region (1800–1500 cm-1) was calculated using OriginPro 8

software for spectral deconvolution and secondary structure determination.

2.2.6 Solid-state NMR measurements

Fibril samples were all dialyzed against water to remove buffer salts, then lyophilized and

packed into 3.2 mm MAS rotors. SSNMR experiments were performed on an 11.6T Varian

VNMR spectrometer using a narrow bore triple-resonance Varian T3 MAS probe. Samples were

maintained at ambient temperature, with high flow-rate of dry air to alleviate sample heating.

Spectra were externally referenced to the downfield 13C resonance of adamantane at 38.56 ppm

relative to tetramethylsilane (TMS) (Morcombe & Zilm 2003). All SSNMR were processed

using NMRPipe software (Delaglio et al. 1995).

45

1H – 13C and 1H – 15N cross-polarization (CP) were implemented using a linear ramped radio

frequency (rf) of 40 - 60 kHz on the low channel and a 50 - 80 kHz field on the 1H channel, with

contact times of 1 to 1.5 ms. The π/2 pulse widths on all channels were typically 2.0 - 5.0 μs. 1H

decoupling fields of 100-120 kHz were applied during all t1 and t2 periods, using a two pulse

phase modulation (TPPM) decoupling scheme (Bennett et al., 1995). Two dimensional (2D) 13C

– 13C correlation spectra were obtained using an RAD/DARR recoupling sequence (Morcombe

et al., 2004; Takegoshi et al., 2001) at 11 kHz MAS and mixing periods of 25 - 500 ms. On

average, spectra were collected in 48 hours using 512 scans per FID and 160 data points in the

indirect dimension.

2D 13C – 15N correlation spectra were collected at 11kHz MAS, using a double cross polarization

pulse sequence with selective transfer from 15N to 13C implemented using a frequency offset and

weak CP fields to record NCO or NCA spectra (Petkova et al., 2003). 15N to 13C CP after the t1

period was achieved using a linear ramp on the 15N channel with RF fields of 10 to 25 kHz on

each channel, and a contact time of 2 ms. A RAD mixing period of 50ms was implemented after

the 15N to 13C CP transfer to achieve 13C – 13C mixing. On average, spectra were collected in 48

hours using 512 scans per FID and 96 data points in the indirect dimension.

Constant time 13C recoupling experiments were performed using the CT-PITHIRDS pulse

sequence (Tycko, 2007), with k1 equal to 4, k2 decreasing from 20 to 0, and k3 increasing from 0

to 20, giving a 45.6 ms total dipolar recoupling time. CT-PITHIRDS spectra were obtained at

MAS rate of 20 kHz, such that 16.67 μs π pulses were used during the recoupling period. Each

spectrum was taken with a sweep width of 20161 Hz and 512 scans per FID. The root mean

square (rms) noise for each data point was calculated from the baseline regions of each spectrum,

using integral widths identical to the peak of interest, and was normalized against the area of the

first peak.

13C – 15N coherence transfer experiments were performed using the 3D z-filtered transferred-

echo double resonance (ZF-TEDOR) recoupling pulse sequence (Jaroniec et al., 2002). ZF-

TEDOR spectra were obtained at an MAS rate of 10 kHz. Rotor-synchronized blocks of 5 μs

15N π-pulses were incremented during the recoupling period to achieve a total dipolar recoupling

time of 0 - 14.4 ms. Each 2D ZF-TEDOR spectrum was collected in 4 - 6 hours using 64 scans

per FID and 32 points in the indirect dimension. Peak volumes were measured using the peak

46

detection function of NMRPipe software, and all values were normalized to the maximum

volume of the one-bond transfer N-Cα for the corresponding amino acid. 13C transverse

relaxation (T2) times were measured using a spin-echo pulse sequence under similar conditions

of 1H decoupling and MAS rate (Carr and Purcell, 1954; Meiboom and Gill, 1958). As per the

PITHIRDS data, the rms noise in the ZF-TEDOR spectra was calculated from baseline area of

each spectrum, using an integrated area identical to the peak of interest.

Rotational resonance (RR) experiments were performed as previously reported (Raleigh et al.,

1988; Walsh et al., 2009). Briefly, the MAS rate for each experiment was selected to match the

frequency difference between C1 and C2 spins, giving an n=1 RR condition for recoupling. A 2 -

4 ms Gaussian π pulse was used for selective inversion of the C1 peak prior to a mixing period of

0 - 40 ms. A reference signal (S0) was recorded without inversion of the C1 signal along with

each experimental S1 curve, and the difference spectrum (S0 - S1) for each time point was

calculated. Polarization transfer was calculated as the difference in peak areas (C1 - C2),

normalized to the C1 peak area at an RR mixing time of 0 ms. 13C zero-quantum coherence

relaxation time (T2ZQ) values for each spin pair C1 and C2 were estimated from single-quantum

T2 values (measured using the spin-echo pulse sequence under identical 1H decoupling and with

a similar MAS rate to the RR experiments) (Carr and Purcell, 1954; Meiboom and Gill, 1958).

2.2.7 SPINEVOLUTION simulations

The SPINEVOLUTION software package was used for all simulations of NMR data (Veshtort

and Griffin, 2006). CT-PITHIRDS dephasing curves were calculated using a linear arrangement

of five 13C atoms at equal distance from each other and observing only the polarization on the

central spin as a function of recoupling time. A slight zigzag offset of the five atoms, by 0.25 Å

from the centre line, gave us the best relative intensity near the end timepoints of the dephasing

curve, and was used for fitting the NMR data. ZF-TEDOR polarization transfer curves were

simulated using a spin system containing only a single 13C and 15N nucleus positioned at

increasing distances. 13C transverse relaxation (T2) times were included in all

SPINEVOLUTION simulations to account for signal loss over the time of the NMR experiment.

Addition of further coupled 13C and 15N nuclei, to emulate the surrounding chemical

environment of an amino acid, resulted in minimal changes to the simulated curves, and were

therefore omitted to decrease calculation times. The RR polarization transfer was simulated

using six spins chosen to represent the C1 and C2 nuclei and two directly bonded 13C spins for

47

each nucleus. The C1 – C2 distance (r12) and T2ZQ values measured for each nucleus were

included in the calculations. In all simulations, the best fit to the experimental data was obtained

by minimizing the weighted sum of squared residuals between experimental and simulated

recoupling curves, as previously reported (Walsh et al., 2009).

2.2.8 Structure calculations

An initial model of each peptide fibril was built in UCSF Chimera as a pair of ideal parallel in-

register β-sheets composed 8 strands (Pettersen et al., 2004). Two sheets were stacked using

close inter-sheet contacts consistent with our 13C RAD/DARR data. Structural constraints on

internuclear distances and backbone torsion angles made use of harmonic potential energy

functions to restrain models. Multiple rounds of simulated annealing were carried out with

Xplor-NIH, using the CHARMM19/20 energy forcefield for calculations (Schwieters et al. 2003;

Schwieters et al. 2006; Brooks et al. 1983). An initial round of annealing was performed with

torsion angles of the middle four residues set at φ = -119 and ψ = 113, representing an ideal

parallel β- sheet conformation. Then, subsequent annealing was repeated using the backbone

torsion angles predicted from TALOS (Cornilescu et al., 1999). Inter-sheet contacts obtained

from 13C spin diffusion experiments were defined as < 7 Å distances between pairs of atoms.

Calculated distance from CT-PITHIRDS and ZF-TEDOR measurements were also included as

restraints.

2.3 Results and Discussion

2.3.1 Biophysical characterization of PrP peptide fibrils

Initial characterization of the amyloid fibrils formed by PrP(178-183), PrP(244-249), and

PrP(245-250) were performed using TEM, ThT fluorescence, and FTIR. TEM micrographs of

negative-stained samples showed straight, unbranched fibrils composed of either a single

filament, or paired filaments twisted together with irregular periodicity (Figure 2.1 A, B, C). In

all cases, single filaments were typically 6 - 10 nm in diameter and up to several μm in length,

with little variation in the morphology of the individual filaments within fibrils. This resembles

the previous description of these peptides, where the fibrils formed were unbranched and had

irregular intertwining of filaments (Pastor et al., 2008).

48

Figure 2.1. TEM of PrP peptide fibrils. Negative-stained TEM images of amyloid fibrils formed by PrP(244-249)

(A), PrP(245-250) (B), and PrP(178-183) (C). TEM images show sample at 20,000 – 40,000x magnification, and

insets on top right show the same sample at 100,000x magnification.

FTIR spectra of these fibrils are shown in Figure 2.2 A-C, and were marked by a strong band in

the Amide I region near 1630 cm-1. Secondary derivative analysis of these spectra identified a

strong peak at 1631 cm-1 for PrP(244-249), 1635 cm-1 for Prp(245-250) and 1625/1633 cm-1 for

PrP(178-183) (Figure 2.2 D-F). This is indicative of a predominantly β-sheet secondary structure

being adopted by all three peptides within the fibril assemblies (Dong et al., 1990). The small

side peak in PrP(245-250) and PrP(178-183) near 1650 cm-1 corresponds to the disordering of

the terminal residue, while the peak at 1674 cm-1 in PrP(178-183) arises from the side chain

amide of Asn181. We did not observe a peak at 1690 cm-1 in the secondary derivative analysis of

neither three fibrils that correspond to anti-parallel β-strands (Figure 2.2 D-F), suggesting that a

parallel arrangement of β-strands is more likely adopted in each case (Halverson et al. 1991;

Kubelka & Keiderling 2001).

49

Figure 2.2. FTIR spectra of PrP peptide amyloid fibrils. The amide I and II regions of FTIR spectra and second

derivative are shown for PrP(244-249) (A, D), PrP(245-250) (B, E), and PrP(178-183) (C, F), with the wavenumber

of identified peaks labelled.

To confirm the amyloid nature of these fibrils, ThT binding was monitored using fluorescence

emission. ThT is an amyloid-specific dye with high affinity binding to cross-β architecture, and

exhibits a shift in its fluorescence emission spectrum to give a maximum near 480 nm upon

binding (Naiki et al., 1989). Intense ThT fluorescence observed for PrP(244-249) and PrP(245-

250) fibrils suggests the presence of cross-β structure in both peptide assemblies (Figure 2.3).

50

Figure 2.3. ThT Fluorescence of PrP peptide amyloid fibrils. ThT fluorescence emission spectra are shown in

the presence of increasing concentrations of PrP244-249 fibrils (A) and PrP245-250 fibrils (B).

Surprisingly, the fibrils formed by PrP(178-183) fibrils did not bind to the cross-β-specific dyes

Thioflavin T (ThT) and Congo red (CR) (Figure 2.4 A, B). Amyloid-binding of these dyes are

typically reported as an increased fluorescence emission at 482nm (ThT) or absorbance intensity

(CR) at 540 nm. However, the mechanism of binding is not well understood, and may be

significantly modulated by the chemical properties of exposed side chains on the surface of a

fibril (Wu et al. 2009; Biancalana & Koide 2010). X-ray fibre diffraction of dehydrated PrP(178-

183) fibril stalks showed a strong repeating element spaced at 4.66 Å and 11.0 Å, corresponding

to the inter-strand and inter-sheet spacing that are typically found in the cross-β motif of amyloid

fibrils (Figure 2.4 C). While these data did not show the typical meridonal and equatorial

alignment of these reflections, this was likely due to poor alignment of the samples prepared for

fibre diffraction. To further investigate the structure of these amyloid-like fibrils in more detail,

MAS SSNMR was used.

51

Figure 2.4. Cross-β characterization of PrP(178-183) amyloids. PrP(178-183) amyloids show a lack of binding

to the amyloid-specific dyes ThT (A) and Congo Red(B), as indicated by lack of fluorescence increase at 482nm and

absorbance at 540nm, respectively. (C) X-ray fibre diffraction data with arrows showing the meridian (4.66 Å) and

equator (11.0 Å) reflections typical of the cross-β backbone of amyloids.

2.3.2 Chemical shift assignments of 13C, 15N-labelled peptides

Lyophilized fibrils were prepared from selectively or uniformly 13C, 15N- labelled amino acids

(Table 2.1). The spin systems for individual amino acids were identified using 2D 13C – 13C

correlation spectra obtained with a short 25-50 ms RAD/DARR mixing period (Figure 2.5 A,

Figure 2.6 A; Figure 2.7 A). Sequential assignments were performed using selective 13C – 15N

transfers from amide nitrogens to Cαi or COi-1 nuclei, followed by 50 ms 13C – 13C RAD/DARR

mixing, giving rise to the NCACX and NCOCX spectra shown below each RAD/DARR

spectrum (Figure 2.5 B, C; Figure 2.6 B,C; Figure 2.7 B, C). Chemical shift assignments in

PrP245-250 fibrils were carried out using a single uniformly 13C, 15N labelled sample, while

complementary labelling schemes (U-13C, 15N-ISIF and U-13C, 15N-SFLI) were used to assign

the amino acids in PrP(244-249) fibrils. Selective incorporation of 13C, 15N nuclei for PrP(178-

183) in 50% U-13C, 15N-DCN and 50% U-13C, 15N-VI mixed fibrils (denoted as DCN/VI) lead to

a single backbone N-COi-1 cross-peak appearing in the NCOCX experiment but five N-Cαi cross-

52

peaks appearing in the NCACX spectrum (Figure 2.7 B, C). Chemical shifts for Thr183 were

determined using DCN/T mixed fibrils (spectra not shown).

Figure 2.5. Resonance assignments for PrP(244-249) SFLI-labelled peptides. (A) 2D 13C – 13C RAD/DARR

spectrum with 50 ms mixing time was recorded to identify each labelled residue in the peptide. Sequential

assignments were performed using NCACX (B) and NCOCX (C) spectra. In each case only the 13C aliphatic region

is shown for simplicity. Assignments for chain A are indicated by dashed lines, while chain B assignments are

indicated by solid lines. All measurements were performed at 11 kHz MAS.

53

Figure 2.6. Resonance assignment of PrP(245-250) uniformly 13C, 15N-labelled peptides. (A) 2D 13C – 13C

RAD/DARR spectrum recorded with a 50 ms mixing time was recorded to identify all residues in the peptide.

Sequential assignments were performed using NCACX (B) and NCOCX (C) spectra. Only the 13C aliphatic region

is shown for simplicity. All measurements were performed at 11 kHz MAS.

54

Figure 2.7. Resonance assignment of PrP(178-183) DCN/VI-labelled peptides. (A) Fibrils prepared from a 50:50

mixture of U-13C, 15N DCN-labelled and U-13C, 15N VI-labelled peptides were used to obtain chemical shift

assignments of the residues. 13C – 15N correlation experiments NCACX (B) and NCOCX (C) were used to assign

backbone and side-chain 15N nuclei, with only 1 sequential backbone transfer identified in NCOCX spectrum due to

site-specific labelling of peptides.

Isotropic 13C and 15N chemical shifts are sensitive to local structure and conformational

environment. The fibrils formed by PrP(245-250) and PrP(178-183) exhibited only one set of

frequencies for all 13C and 15N nuclei, suggesting that the peptides exist in a chemically

homogenous environment within fibrils (Table A 2, Table A 3). All 13C line widths at half peak-

height were measured to be 1 - 2 ppm, indicating a high degree of structural order and

55

homogeneity within the fibrils (Figure 2.8 B). 13C secondary chemical shift analysis for

backbone CO, Cα, and Cβ resonances was performed by calculating the difference between the

experimental data and theoretical values reported for a random coil (Saitô, 1986; Wishart and

Sykes, 1994). A positive Cβ difference (downfield shifts in experimental chemical shift) and

negative CO and Cα difference (upfield shifts) are characteristic of an extended β-sheet

conformation. This secondary shift pattern can be seen for Phe246 through Phe249 of PrP(245-

250) and Cys179 through Ile182 of PrP(178-183) (Figure 2.8 A). The C-terminal residue of all

peptides exhibited a more α-helical 13C secondary shifts, suggesting a potentially irregular

structure for those sites due to a lack of H-bond constraints.

In contrast, the PrP(244-249) fibrils gave rise to two distinct sets of 13C and 15N chemical shifts,

with both having approximately equal intensities in all samples, suggesting that two non-

equivalent peptide chains are found within the fibrils formed (Table A 1). Spectral overlap

caused by the combination of non-equivalent chains and repeating amino acids necessitated the

use of complementary labelling schemes with the SFLI and ISIF peptides (Table 2.1) to reduce

spectral complexity and allow sequential assignments. As shown in Figure 2.8, both inequivalent

chains of PrP(244-249) have an extended β-sheet for residues Ser245 through Ile248.

56

Figure 2.8. 13C secondary chemical shift calculations and linewidths for PrP peptide amyloids. (A) 13C

chemical shift deviation from published random-coil chemical shift values are shown for PrP(244-249) chain A and

B, PrP(245-250), and PrP(178-183). (B) 13C NMR cross peak linewidths measured at half-height.

Backbone chemical shift values (N, CO, Cα, Cβ) were used to predict the backbone torsion angles

Φ and Ψ for each PrP peptide using the TALOS algorithm (Cornilescu et al., 1999). The results

are listed in Table 2.2 and comparable to typical values for β-sheets at Φ = -120 o and Ψ = 135 o,

further supporting an extended β-strand structure within each of the fibrils.

57

Table 2.2. Backbone torsion angles of PrP peptide amyloids calculated using TALOS+ using 13C and 15N

chemical shift.

Residue Phi Φ (o) Psi Ψ (o)

PrP(245-250) Phe246 -141 ± 11 163 ± 10

Leu247 -154 ± 12 136 ± 13

Ile248 -128 ± 27 130 ± 29

Phe249 -119 ± 26 115 ± 19

PrP(244-249) Chain A Ser245 -129 ± 11 138 ± 21

Phe246 -114 ± 13 123 ± 10

Leu247 -123 ± 26 145 ± 16

Ile248 -119 ± 29 147 ± 18

PrP(244-249) Chain B Ser245 -128 ± 28 150 ± 16

Phe246 -121 ± 15 130 ± 12

Leu247 -111 ± 13 122 ± 12

Ile248 -118 ± 25 132 ± 18

PrP(178-183) Cys179 -132 ± 24 144 ± 14

Val180 -130 ± 12 141 ± 11

Asn181 -125 ± 8 132 ± 10

Ile182 -125 ± 12 143 ± 18

2.3.3 The prion peptide amyloid fibrils all contain parallel, in register β-

sheets

To determine the type of strand assembly present within the β-sheet rich core of each PrP peptide

fibril, we examined the average distance between the backbones of adjacent β-strands using the

CT-PITHIRDS homonuclear recoupling experiment (Tycko 2007). This experiment measures

decay of the NMR signal as a function of a recoupling period (here from 0 - 45.6 ms), during

which signal decays due to the presence of 13C – 13C dipolar couplings between adjacent nuclei.

The strength of the observed dipolar coupling, and therefore the rate of signal decay, is inversely

proportional to the internuclear distance between two nuclei (r) by a factor of 1/r3. Thus when

applied to fibrils prepared from peptides containing a 13C incorporated at only one backbone

position, we can accurately measure the inter-strand distance within a β-sheet to distinguish

between parallel in-register β-sheet architecture and other arrangements.

Fibrils were prepared from peptides with 13C incorporated at a single carbonyl position and all

CT-PITHIRDS dephasing curves are shown in Figure 2.9. Through comparing the experimental

data with numerical simulations from SPINEVOLUTION, we find our data for PrP(245-250)

best fit to an intermolecular distance of 5.0 Å ± 0.2 Å between both the carbonyl atoms of

58

Phe246 on adjacent strands, and between Phe249 carbonyl atoms. Similarly, CT-PITHIRDS

experiments of PrP(244-249) fibrils resulted in intermolecular distances of 4.9 Å ± 0.2 Å and 5.0

Å ± 0.2 Å at the carbonyl sites of Phe246 and Ile248, respectively, while PrP(178-183) fibril data

best fit to an intermolecular distance of 5.0 Å ± 0.2 Å and 4.9 Å ± 0.2 Å between the carbonyls

of Val180 and Ile182, respectively. With the interstrand distance of an ideal β-sheet being 4.7 Å,

this data strongly suggests that all three peptides are arranged in-register parallel β-sheets within

their respective amyloid fibrils. The 13CO chemical shifts of chain A and B in PrP(244-249)

fibrils overlapped, so that the dephasing curves for these fibrils contain contributions from both

inequivalent chains, supporting the same architecture for chain A and B.

Figure 2.9. CT-PITHIRDS measurement of backbone inter-strand distances in selectively 13CO-labelled

peptides. Dipolar dephasing curves obtained from CT-PITHIRDS homonuclear recoupling experiment are shown

for single-site labelled (A) PrP(245-250) F246CO (red square) and F249CO (green triangle) fibrils, (B) PrP(244-

249) F246CO (blue circle) and I248CO (orange square) and (C) PrP(178-183) Val180CO (yellow triangle) and

Ile182CO (purple square). Error bars were determined as the RMS noise in each spectrum and are within the area of

each data point. Simulated PITHIRDS curves shown in each case were generated by SPINEVOLUTION using a

chain of five 13C nuclei, with each fixed at an equal distance from the next. Distances of 4.5 – 7.0 Å are presented

here, with a 0.5 Å increment between simulated dephasing curves.

2.3.4 Quaternary contacts within PrP(245-250) fibrils

In order to determine the quaternary structure of PrP(245-250) fibrils, as defined by the assembly

of β-sheets within the hydrophobic core of the fibril, additional structural constraints were

collected from solid-state NMR experiments. 2D 13C – 13C RAD/DARR measurements of longer

mixing periods (250 - 500 ms) can identify 13C nuclei that are spaced up to 7.0 Å apart. New

cross peaks that arise from spatially proximate but sequentially distant amino acids are important

for identifying amino acid that come into contact in the β-sheets core.

59

A set of 13C – 13C RAD/DARR spectra obtained using 25, 250, and 500 ms mixing times are

shown for PrP(245-250) fibrils in Figure 2.10. At longer mixing times, a new cross peak between

the Leu250 and Phe246 side chains is observed (Figure 2.10 D-F, upper spectra). The loss of its

intensity in a sample prepared from diluting labelled material with 75% unlabelled peptide

(Figure 2.10 D-F, lower spectra), confirmed that this interaction arises from intermolecular

packing of residues Leu250 and Phe246 rather than from intramolecular contacts. This indicates

tight packing of these residues at the interface of the two β-sheets. We also prepared mixed

sample with site-specific labelling PrP(245-250) SI/F to validate this cross peak. For fibrils

formed by an equimolar mixture of these two peptides, a cross peak between Ile248 and Phe246

side chains was observed but not Phe246 and Ser245 (Figure 2.11). Taken together, these data

support an anti-parallel face-to-face packing of the β-sheets within the PrP(245-250) fibrils,

forming a class 1 steric zipper.

60

Figure 2.10. 13C – 13C RAD/DARR correlation spectra for long-range intermolecular contacts in PrP(245-250)

fibrils. 2D 13C – 13C correlation spectra for 13C, 15N-uniformly labelled PrP(245-250) fibrils are shown. Spectra

were recorded at 11kHz MAS, with a mixing time of (A) 25 ms, (B) 250 ms, and (C) 500 ms. Horizontal slices at

the Leu250 Cγ frequency are shown for (D) 25 ms, (E) 250 ms, and (F) 500 ms mixing times. Interresidue cross peaks

are indicated on the slices, and are discussed in the text. Corresponding slices obtained from identical experimental

conditions for diluted fibril samples (prepared as a mixture of 25% 13C, 15N-uniformly labelled and 75% unlabelled

PrP(245-250) peptide) are shown at the bottom of each panel for comparison.

61

Figure 2.11. Long-range intermolecular contacts observed in 2D 13C – 13C spin diffusion spectra of PrP(245-

250) SI/F fibrils. (A) 13C – 13C correlation spectrum obtained for PrP(245-250) fibrils prepared from a mixture of SI

and F labelled peptides. This spectrum was recorded at an MAS rate of 11 kHz with a mixing time of 500 ms. (B)

Horizontal slices of Ser, Ile, and Phe side chains (indicated with different 13C frequencies) are shown to with Ile-Phe

cross peaks but not Ser-Phe cross peaks.

Additional evidence supporting this quaternary structure for the PrP(245-250) fibrils came from

the ZF-TEDOR experiment, which measures 13C – 15N heteronuclear dipolar couplings.

Internuclear distance is determined by fitting the changes in experimentally-observed cross peak

volume over recoupling time to SPINEVOLUTION simulations. Our ZF-TEDOR transfer curve

arise from a distance of 3.0 Å ± 0.3 Å between the Ser245 amino 15N and the Leu250 carboxyl

13C (Figure 2.12). As an internal reference, the fixed one-bond and two-bond distances for

Ser245 N – Cα and Ser245 N – CO were measured and fit well to one- and two-bond distance of

2.0 ± 0.4 Å and 2.8 Å ± 0.4 Å, respectively. ZF-TEDOR measurements on samples containing

labeled peptide diluted 1:4 with unlabelled PrP(245-250) did not contain a Ser245 N to Leu250

CO cross peak, confirming that this is an intermolecular contact (Figure 2.13). This constraint

places the termini of opposing PrP(245-250) sheets in close contact and suggests a possible salt

bridge interaction stabilizing the core structure (Petkova et al., 2006; Walsh et al., 2009).

62

Figure 2.12. 13C – 15N heteronuclear ZF-TEDOR measurement of intermolecular contacts for uniformly-

labelled PrP245-250 fibrils. (A) Representative regions of 2D heteronuclear 13C – 15N ZF-TEDOR spectra are

shown, highlighting cross peaks at the S245 15N resonance. The dipolar recoupling time used for each spectrum is

indicated. (B) Peak intensities from 2D spectra were integrated and normalized to the maximum intensity observed

for the one-bond Ser245 N – Ser245 Cα cross peak. The best-fit simulated build-up curves shown for each spectrum

were obtained using the 13C – 15N distances indicated on the graph. 13C transverse relaxation (T2) times were

included in the simulations to account for loss of NMR signal over time.

63

Figure 2.13. ZF-TEDOR measurements of 25% uniform-13C, 15N-labelled PrP(245-25) diluted with 75%

unlabelled material. Normalized cross peak intensities are shown for 13C –15N ZF-TEDOR spectra recorded on

fibrils formed by a 1:3 dilution of uniformly 13C, 15N labeled PrP(245-250) with unlabelled peptide. When compared

to the undiluted sample, similar buildup curves are observed for the one-bond S245 N – S245 Cα and two-bond

S245 N – S245 CO cross peaks. However, contact between N-terminal S245 N to C-terminal L250 CO was lost

(position indicated in panel A), strongly suggesting that this cross peak results from intermolecular contact between

the two peptides.

2.3.5 High-resolution structure of PrP(178-183) peptides within amyloid

fibrils

Intermolecular structural constraints were obtained for PrP(178-183) fibrils using 13C – 13C

RAD/DARR spectra recorded with 500 ms mixing times and PAR spectra obtained with 10ms

mixing. PAR is a similar dipolar transfer technique to RAD/DARR, but relies on nearby 1H to

assist in polarization transfer. As shown in the PAR spectrum in Figure 2.14, unambiguous

intermolecular cross peaks were observed between Ile182 side chain and the Asp178 side chain,

Cys179 backbone, and Val180 side chain in a DCN/VI mixed label sample. To confirm the

Val180 – Ile182 cross peak, we prepared samples by pre-mixing 20% 13C, 15N-VI-labelled

64

peptides with 80% unlabelled peptide in HFIP, followed by fibrillization. The Val-Ile cross

peaks decreased significantly as shown in the 1D slices taken from RAD/DARR experiments of

diluted peptides (Figure 2.15). This confirmed the cross peaks arise from inter-sheet contact

between Val180 and Ile182 rather than intra-strand contact between these residues.

Figure 2.14. Long-range intermolecular contacts in PrP(178-183) amyloids prepared from DCN/VI-labelled

peptides. 1D 13C slices from homonuclear 13C – 13C PAR experiment show cross peaks between Ile182 and

Asp178, Cys179, and Val180 indicated by the arrows. The schematic shows the orientation of opposing peptides

that would give rise to these intermolecular contacts. The experiment was performed using a 10 ms PAR mixing

time, recorded with an MAS frequency of 21 kHz.

65

Figure 2.15. 13C – 13C intermolecular contacts in PrP(178-183) amyloids. 1D slices of RAD/DARR spectra for

the Ile182 δCH3 showing cross peak intensities between Ile182 and Val180 (indicated by arrow) is reduced by the

addition of unlabelled peptide, suggesting these are inter-sheet contacts between Ile and Val.

We also measured distances between proximate 13C and 15N nuclei using ZF-TEDOR. Cross

peak between backbone 15N and side chain 13C atoms builds up as a function of transfer time to

provide intra-residue distance constraints of the peptide. For each pair of nuclei, we determined

the distance by fitting the experimentally-observed cross peak intensity build-up to

SPINEVOLUTION simulations. As shown for Asp178 in Figure 2.16 A-B, the different rates of

cross peak buildup between the amide NH group and backbone Cα or side chain COγ atoms

provided unambiguous high-resolution structural constraints for each residue within the PrP(178-

183) peptides (Figure 2.16, Table 2.3). In this case, although PrP(178-183) fibrils also formed

class 1 steric zipper as PrP(245-250), we did not observe N- to C-terminus salt-bridge

interaction.

66

Figure 2.16. 13C – 15N intramolecular distances of PrP(178-183) peptides in amyloid fibrils. (A) Representative

2D 13C – 15N spectra showing the buildup of cross peaks for Asp178 at various ZF-TEDOR transfer times. Peak

intensities from 2D spectra were integrated and normalized to the maximum intensity observed for one-bond NH-Cα

cross peak of the corresponding amino acid. (B) The best-fit simulated build-up curves shown for each spectrum

were obtained using the 13C – 15N distances indicated on the graph. (C) Distances for ≥ 3-bond nuclei measured in

ZF-TEDOR experiments that were used as experimental constraints for building the structural model.

Table 2.3. Intra-residue 13C – 15N distance constraints determined through ZF-TEDOR.

Residue Nuclei pair ZF-TEDOR Distance (Å)

Asp178 N - γCO 3.0 ± 0.4

Val180 N - γ1CH3 3.2 ± 0.4

N - γ2CH3 3.2 ± 0.4

Ile182 N - γCH3 3.1 ± 0.4

N - γCH2 2.8 ± 0.4

N - δCH3 3.2 ± 0.4

Thr183 N - γCH3 3.4 ± 0.4

67

2.3.6 Quaternary contacts within PrP(244-249) fibrils

To reduce the amount of cross peak overlap in the 2D 13C – 13C spectra from PrP(244-249)

fibrils due to two inequivalent peptide chains and repeating amino acids, we collected SSNMR

data from fibrils prepared using an equimolar mixture of peptides with the site specific labelling

schemes PrP(244-249) SI/F, IL/F, and SI/IL (Figure 2.17). Long-range intermolecular contacts

observed in RAD/DARR spectra of these samples include Ile248 – Phe246, Phe246 – Ile244,

Phe246 – Leu247, Leu247 – Ile248, and Ile244 – Ser245. Specifically, we observed cross peaks

between Ile248 γCH2 – Phe246 aromatic ring, and reciprocally Phe246 aromatic ring – Ile248

side chain (γCH2, γ, δCH3) in the SI/F mixture (Figure 2.17 A); between Ile244 γCH3 – Phe246

aromatic ring and Leu247 δCH3 – Phe246 aromatic ring in the IL/F mixture (Figure 2.17 B); and

finally Ile244 γCH3 – Ser245 Cβ and Ile248 γCH3 –Leu247 δCH3 in SI/IL mixture (Figure 2.17

C). As Ile244 and Ile248 γCH3 had overlapping chemical shifts, the 1D slice in SI/IL sample

shows cross peaks for both nuclei.

68

Figure 2.17. 13C – 13C long-range intermolecular contacts in PrP(244-249) amyloids. 13C – 13C RAD/DARR

experiment obtained with mixing time of 500 ms are shown for PrP(244-249) fibrils containing an equimolar

mixture of peptides with the SI/F (A), IL/F (B), or SI/IL (C) labeling schemes. Horizontal slices at different 13C

frequencies are shown to the right of each 2D spectrum, and inter-chain cross peaks are indicated on the slices. All

spectra were obtained at an MAS frequency of 11 kHz.

69

To further validate these intermolecular contacts, 13C RR experiments carried out on the

PrP(244-249) fibrils. RR measurements selectively reintroduce dipolar couplings between nuclei

whose resonance frequencies differ by a multiple of the MAS frequency (here using an n=1 RR

condition where ωr = ω1 - ω2) (Madine et al., 2008). The experimental recoupling curves shown

in Figure 2.18 were collected on the PrP(244-249) SI/F fibrils to show we were able to determine

an internuclear distance of 5.9 Å, and 5.2 Å from the Phe246 aromatic Cγ to Ile248 γCH2, and

γCH3, respectively, when compared to SPINEVOLUTION simulations. Although the methyl

resonances of Ile248 on the two inequivalent peptide chains cannot be independently resolved,

the fitting of the RR data clearly shows close proximity between the Phe246 Cγ and an

isoleucine side chain. Additional RR measurements made on SI/F, IL/F, and SI/IL fibrils

provided accurate internuclear distance restraints between Phe246 – Ile248, Phe246 – Leu247,

Phe246 – Ile244, and Ser245 – Ile244, respectively (Table 2.4).

Table 2.4. Comparing internuclear distances determined by experimental RR measurements and PrP(244-

249) NMR structural models.

Nuclei RR Fits

Average Distance

Measured on

Minimized Model

(Class 3)

Average Distance

Measured on

Minimized Model

(Class 2)

F246 Cγ – I248 γCH2 5.9 Å 5.0 ± 0.8 Å >10 Å

F246 Cγ – I248 δCH3 4.8 Å 4.5 ± 0.5 Å >10 Å

F246 Cγ – I248 γCH3 5.2 Å 4.2 ± 0.4 Å >10 Å

F246 Cγ – I244 δCH3 4.9 Å 4.5 ± 0.6 Å >10 Å

F246 Cγ – I244 γCH3 5.1 Å 4.5 ± 1.3 Å >10 Å

S245 Cβ – I244 δCH2 6.7 Å 8.2 ± 0.7 Å 6.1 ± 0.8 Å

F246 Cγ – L247 δCH3 5.4 Å >10 Å 7.4 ±1.6 Å

70

Figure 2.18. 13C RR measurements for PrP(244-249) SI/F fibrils. Peak intensity difference curves showing

polarization exchange between specific C1 and C2 spins under n=1 RR conditions. (A) Experimental data and

simulated RR curves for F246 Cγ - I248 γCH2 at an MAS rate of 14,050 Hz. Simulations are shown for r12

distances of 4.5 - 6.5 Å, using T2ZQ values of 2.0 ms (solid) or 1.8 and 2.2 ms (upper and lower dashed lines). The

experimentally determined T2ZQ was 2.07 ± 0.13 ms for this spin pair. (B) Experimental data and simulated RR

curves for F246 Cγ - I248 γCH3 at an MAS rate of 15,158 Hz. Simulations are shown for r12 distances of 4.5 - 6.5 Å,

using values of 2.8 ms (solid) or 2.6 and 3.0 ms (upper and lower dash lines). The experimentally determined T2ZQ

for this spin pair was 2.74 ± 0.22ms.

ZF-TEDOR measurements on PrP(244-249)ISIF fibrils did not result in cross peak buildup

between the N- and C-termini of adjacent peptides, suggesting a separation for these groups of

more than 6.0 Å, and ruling out the possibility of an inter-sheet salt-bridge interaction at the

termini (data not shown). Unfortunately, due to the reduced resolution in the PrP(244-249)

spectra caused by the presence of two sets of resonances, no unambiguous 13C – 15N distances

could be obtained from ZF-TEDOR measurements of these samples.

2.3.7 Fibril structures of PrP(178-183), PrP(244-249) and PrP(245-250)

High resolution structures for each fibril were determined by simulated annealing using torsion

angles from TALOS calculations and 13C – 13C and 13C – 15N distance constraints obtained from

CT-PITHIRDS, ZF-TEDOR and RAD/DARR experiments. RR data was withheld from structure

71

calculations of PrP(244-249), and was instead used to validate the structures determined for each

packing interface in those fibrils, as described below.

The structure of PrP(245-250) fibrils is shown in Figure 2.19, and has the classical class 1 steric

zipper structure seen in the majority of fibril structures solved to date – parallel in-register β-

sheets stacked in a face-to-face, anti-parallel arrangement. The backbone root mean square

deviation (RMSD) of the model is 0.48 Å ± 0.15 Å and total RMSD is 1.34 Å ± 0.56 Å. In

Figure 2.19 B, a slight twist is evident, even over only 4 strands, as expected for an extended β-

structure. Unfortunately, no distinct and periodic twisting was seen in TEM images of these

fibrils (Figure 2.1), preventing the use of this parameter as an additional restraint. The packing

of hydrophobic side chains within the fibril core is evident, with Phe246 and Ile248 forming the

dehydrated interface, and the Ser245 side chain on the outer face, potentially allowing solvent

exposure or hydration.

Figure 2.19. Structure of PrP(245-250) fibrils. (A) Schematic representation of the packing model for PrP(245-

250) fibrils. (B) A ribbon diagram of a representative fibril structure, averaged from 8 low energy structures,

showing 4 strands per sheet, (C) Stick representation of two representative peptide structures showing the relative

orientation of the side chains on opposing β-sheets. Fibrils assembled into face-to-face, anti-parallel stacking of

parallel β-sheets, referred to as the class I steric zipper packing model.

PrP(178-183) fibrils also exhibit class 1 steric zipper structure of parallel in-register β-sheets

stacked in a face-to-face, anti-parallel arrangement (Figure 2.20). The calculated structures gave

a backbone RMSD of 0.65 ± 0.12 Å and a total RMSD of 3.10 ± 0.16 Å. The fibril core is

stabilized by the packing of the hydrophobic side chains of Ile182 and Val180, while the polar

72

residues Cys179, Asn181, and Thr183 face the exterior and are exposed to solvent. In-register

alignment of the β-strands also positioned the Asn side chains together to allow the carboxamide

groups to form intra-sheet hydrogen bond, possibly contributing to the 1671 cm-1 peak seen in

FTIR spectra. No salt bridge interaction between the N- and C-termini was suggested by ZF-

TEDOR experiments.

Figure 2.20. Structure of PrP(178-183) amyloids. (A) Schematic representation of the packing model for

PrP(178-183) fibrils. (B) A ribbon diagram of a representative fibril structure, averaged from 8 low energy

structures, showing 4 strands per sheet, (C) Representation of the β-sheet structures showing the relative orientation

of peptides assembled into the class I steric zipper packing model with face-to-face, anti-parallel stacking of parallel

β-sheets.

The structure determination of PrP(244-249) within amyloid fibrils was complicated by the

presence of two sets of distinct chemical shifts. Based on RAD/DARR 13C – 13C correlation

experiments, a single inter-sheet packing model did not satisfy all observed intermolecular

contacts. Instead, two different packing arrangements of the peptide must be present within the

fibril samples to account for the experimental data. A class 3, parallel stacking of β-sheets in

face-to-face arrangement (Figure 2.21 A, C), satisfies the Phe246 – Ile244 and Phe246 – Ile248

contacts observed in the RAD/DARR data. In this inter-sheet interface, hydrophobic residues

Ile244, Phe246, and Ile248 are packed facing into the fibril core, with the serine hydroxyl group

potentially facing out of the fibril core and exposed to solvent (Figure 2.21 E). On the other

hand, a class 2 parallel stacking of β-sheets in face-to-back arrangement satisfies the observed

73

Ile244 – Ser245, Phe246 – Leu247, and Leu247 – Ile248 contacts, and allowing the hydrophobic

residues Phe246, Leu247, Ile248 to be buried in the fibril core (Figure 2.21 B, D, F). In this case,

the serine hydroxyl side chains are positioned such that they likely form either stabilizing

hydrogen bonds across the interface (i.e. to backbone across the zipper), or polar-aromatic

interactions to phenylalanine ring for additional stability (Imai et al., 2007).

Figure 2.21. Structure of PrP(244-249) amyloids. The structures determined for both inter-sheet arrangements

observed in PrP(244-249) fibrils are shown, with the class 3 steric zipper shown in A, C, E and the class 2 steric

zipper structure in B, D, F. (A) and (B) show schematic representations of each structure. (C) and (D) depict the

structures for two 4-stranded sheets in each case. Representative peptide structures, averaged from 8 low energy

structures, show opposing sheets from class 3 (E) and class 2 (F) structures.

RR experiments were used to confirm the two packing interfaces in the PrP(244-249) fibrils.

Internuclear distances measured using RR were compared with the distances measured from our

energy-minimized structures of both types of packing. Having two different packing

arrangements can also explain the presence of two resonances for each site in our PrP(244-249)

74

NMR experiments. The calculated structures have a backbone RMSD of 0.70 Å ± 0.27 Å (class

2) and 0.66 Å ± 0.17 Å (Class 3), and a total RMSD (all heavy atoms) of 3.03 Å ± 0.56 Å (class

2) and 2.70 Å ± 0.31 Å (class 3). The parallel-stacking of β-sheets in both models also fits with

our ZF-TEDOR data, which support structures in which the N- and C-termini of opposite chains

are not positioned in close proximity, preventing salt-bridge interactions.

The combination of a homogeneous fibril morphology observed in our TEM images, and the 1:1

ratio of peak intensities for the two inequivalent sets of chemical shifts in all spectra, leads us to

believe that both internal structures (class 2 and class 3 steric zippers) are likely to be present in

each filament. This was supported by the seeding experiments shown in Figure 2.22, in which

the chemical shifts and relative populations of the A and B chain NMR signals remain

unchanged after 4 generations of seeding, as opposed to weighting towards a kinetically favoured

species. However, we cannot completely exclude the possibility that two different types of fibril

are present, each consisting of peptides in only one packing class.

75

Figure 2.22. Seeding of PrP(244-249) fibrils. Negative-stained TEM images of PrP(244-249) fibrils and the

corresponding 13C – 13C correlation spectra are shown for the parent, SFLI-labelled fibrils (A) and the seeded, 4th

generation daughter, SI-labelled fibrils (B). 2D 13C – 13C spectra were recorded at an MAS rate of 11 kHz with a

mixing time of 25 ms. Horizontal slices at the Ser245 Cβ frequencies of the A and B chains are shown in (C). After

4 generations of seeding, the relative intensity of the signal arising from each of the two chains remains equivalent,

suggesting that both species are still present in sample in equal quantity. Likewise, the chemical shifts for each chain

are unchanged, within experimental error.

2.4 Discussion

Our solid state NMR investigation of fibrils from three prion protein derived hexapeptides has

revealed important information on the inter-sheet packing of these small peptide amyloid fibrils.

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PrP(245-250) peptides formed a class 1 steric zipper, with parallel in-register β-sheets stacked in

the anti-parallel, face-to-face manner that has been observed for many other amyloid fibrils. The

hydrophobic residues isoleucine and phenylalanine are confined in the core to form the

dehydrated surface, with the N- and C-termini forming stabilizing salt bridge interactions

between the opposing sheets. PrP(244-249) peptides also assembled as parallel in-register β-

sheets. Yet despite having the same hydrophobicity and similar sequence to PrP(245-250), it

formed both class 2 and class 3 steric zippers. In the class 2 zipper, sheets are stacked in a

parallel face-to-back manner, and leucine, isoleucine, and phenylalanine facing the fibril core

with a possibility of π-π interactions between aromatic residues on opposing sheets; in the class 3

zipper, sheets are stacked a parallel face-to-face manner such that isoleucine and phenylalanine

are facing the fibril core. The effect of these variations in fibril architecture on the toxicity of

different amyloid peptides is explored in Chapter 3.

A similar role for phenylalanine in amyloid formation has been seen for Aβ(16-22) in which

Phe19 and Phe20 are positioned on opposite faces in an extended beta-sheet arrangement based

on solid-state NMR studies (Balbach et al. 2000; Senguen et al. 2011). This arrangement with

Phe19 involved in the π- π interactions of the aromatic side chain for stabilizing the fibril

packing, and Phe20 involved in further stacking of protofilaments into higher ordered assemblies

(Inouye et al., 2010). It is likely in 244-249 and 245-250 sequences that a similar set of π- π

interactions may drive the formation of parallel versus anti-parallel sheets, or may drive the

association of protofibrils into a macrofilament, as has been proposed in extensive studies of

fibrils and nanotubes formed by phenylalanine-rich peptides (Reches & Gazit 2003; Meital &

Ehud 2006).

The packing of hydrophobic side chains within the fibril core leads to more polar residue Ser245

pointing outwards of the protofilaments in PrP(245-250) class 1 and PrP(244-249) class 3 fibrils,

potentially allowing solvent exposure or hydration (Figure 2.19 A, Figure 2.21 A). However,

minimizing hydrophobic exposure is a stronger driving force for amyloid formation than solvent

exposure, as Ser245 in PrP(244-249) can be packed into the core in class 2 fibrils (Figure 2.21

B). Fibril polymorphism similar to PrP(244-249) has been previously observed for other small

amyloid peptides yeast Sup35 fragment GNNQQNY (Lewandowski et al. 2011; van der Wel et

al. 2007; van der Wel et al. 2010) and human IAPP fragment SNNFGAILSS (Madine et al.,

2008).

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Finally, PrP(178-183) peptides formed a class 1 steric zipper with isoleucine and valine packed

to form the dehydrated hydrophobic core. The Asp178 13Cγ chemical shift (175.9 ppm) was

slightly upfield shifted from the reported value of a protonated side chain (177.1 ppm) compared

to the value expected for a deprotonated side chain (180.3 ppm) (Platzer et al. 2014). Protonation

of Asp178 under acidic conditions (pH 2.6) suggest the carboxylate side chain would not impose

large charge repulsion to disrupt the packing of in-register β-strands in the protofilament during

fibrillization. The polar residues cysteine, asparagine, threonine are located on the surface of the

protofilament, and the in-register parallel alignment of the peptides allows the carboxamide of

Asn to form additional hydrogen bond network for stabilization.

High resolution structural models of the three peptide fibrils point out that sequence plays a key

role in governing how β-strands can be assembled into the cross-β backbone. However, no

obvious differences in fibril packing can explain the reported difference in cytotoxicity of the

three amyloids. Instead, the amount of hydrophobic versus polar residues exposed on the outside

of protofilaments may be a large factor in cytotoxicity. While residues exposed on the surface of

the protofilament are known to govern the assembly behaviour of amyloid fibrils into larger

bundles, they may also be important to their interaction with other biological systems.

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Chapter 3.

Fragmentation-induced

membrane interactions of

amyloid fibrils

Contributions:

Undergraduate student Anton Dobrin (University of Toronto) assisted with the liposome

disruption assay.

79

3.1 Introduction

Amyloid formation is a nucleation-dependent process during which small monomeric,

oligomeric, and protofibrillar species are populated during the early stages of misfolding and

aggregation (Hardy & Selkoe 2002; Selkoe 2003; Kayed et al. 2003). These on- or off-pathway

intermediate species are now widely considered to be closely connected to be the pathogenicity

of amyloid diseases, rather than the mature fibrillar deposits. Not only do smaller intermediate

species template the misfolding of precursor protein, they have been shown to disrupt membrane

bilayers, induce apoptosis, and decrease cell viability. The cytotoxic activity of soluble

misfolded intermediates is strongly related to both their size and hydrophobicity, where

oligomers of smaller size and greater hydrophobicity can more readily interact membrane

surfaces and induce cell death (Mannini et al. 2014; Xue et al. 2009).

Fibril seeds are critical on-pathway intermediates that decrease the lag phase for nucleation and

accelerate the rate of fibril formation. However, they exhibit different biological behaviours from

mature fibrils. Fibril seeds formed from various amyloidogenic proteins and peptides, such as

lysozyme, β2-microglobulin, α-Syn, Aβ, and PrP have been shown to cause a greater reduction

in cell viability than mature fibrils in a 3-(4, 5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium

bromide (MTT) reduction assay (Xue et al. 2009; Mannini et al. 2014; Pastor et al. 2008). At the

same time, small fibril fragments can be more readily internalized by cells than the mature

forms, where uptake might be required for cytotoxicity (Pastor et al. 2008; Jakhria et al. 2014).

Some mechanisms that have been proposed for amyloid cytotoxicity include increasing the

permeability of lysosomal and cell membrane, causing mitochondrial dysfunction, or inducing

ER stress that elicits the UPR and halts protein synthesis and trafficking (Gharibyan et al. 2007;

Umeda et al. 2011).

Structural studies of amyloidogenic proteins and peptides have primarily been performed on

mature end-stage fibrils, given that fibril seeds propagate their structural characteristics into

mature fibrils (Lu et al. 2013; Groveman et al. 2014). As such, there has been a lack of structural

information on small fibril fragments to address why they behave differently than their mature

counterparts. To understand the underlying differences between the two, we have examined the

high resolution structure of sonicated fibrils using solid-state NMR spectroscopy (SSNMR).

Three amyloid fibrils, PrP(178-183), PrP(244-249), and PrP(245-250), derived from the

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mammalian prion protein, exhibit varying degrees of cytotoxicity in cell culture upon

sonication– with PrP(244-249), and PrP(245-250) being toxic and PrP(178-183) not toxic (Pastor

et al. 2008). As I have shown in Chapter 2, these fibrils exhibit a classic amyloid cross-β

backbone with stacking of parallel β-sheets. Whether fragmenting the fibrils by ultrasonication

induces structural changes, and how these fibril fragments exhibit enhanced cytotoxicity, is the

topic of the current chapter.

Here, we report that sonication of mature fibrils did not cause substantial disruption to the

fibrillar morphology nor to the secondary structure of all three peptide fibrils. All fibrils retained

the parallel β-sheet arrangements present in the mature fibrils. Instead, we observed that

fragmentation disrupted the internal packing within some fibril packing and altered the amount

of exposed hydrophobic surface area. In PrP(244-249) fibrils, sonication disrupted the class 3

steric zipper and increased the surface-exposed hydrophobic area as shown by the binding to

ANS. At the same time, these sonicated fibrils increased membrane disruption of liposomes

composed of both anionic (3:1 PC/PG) and zwitterionic (PC) lipid mixtures. PrP(245-250) fibrils

were not affected by sonication, but exposed hydrophobic surfaces disrupted liposome that was

increased in anionic membrane composition. Finally, the molecular packing and surface

hydrophobicity of non-toxic PrP(178-183) fibrils were unchanged as a result of sonication, and

fragmentation did not increase membrane disruption. These results support the concept that

surface properties of fibrils play a critical role in governing fibril-membrane interaction and

amyloid toxicity.

3.2 Methods

3.2.1 Sonication of PrP fibrils

PrP peptide fibrils were prepared following the same method described in Chapter 2. Sonicated

fibrils were freshly prepared by diluting fibrils to 0.5 mg/mL with 20 mM Glycine-HCl buffer,

and probe sonicated (10% amplitude output) at 4oC using 5-sec on/off cycles for a total of 2 min.

3.2.2 ANS Fluorescence

A 10 mM stock of ANS (FLUKA) was freshly prepared in 20 mM sodium phosphate buffer (pH

7.5). All fluorescence spectra were recorded on a Photon Technology International C60

spectrofluorometer, with excitation/emission slit widths of 2/5 nm. ANS were excited at 375 nm

and recorded emission spectra from 400 to 600 nm. A fluorescence emission maximum observed

81

at 470 nm was used as the signature of dye binding. All triplicate measurements contained 15µM

ANS with volume adjusted to 1 mL.

3.2.3 Liposome disruption assay

1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) and 1-palmitoyl-2-oleoyl-sn-glycero-

3-phosphotidylglycerol (POPG) lipids stocks were purchased from Avanti Polar Lipids pre-

dissolved in chloroform at 25 mg/mL stocks. Large unilamellar vesicles (LUV) were prepared

from co-dissolved 3:1 POPC/POPG lipids or pure POPC lipids in chloroform and drying into a

thin lipid film under a constant stream of N2(g). The film was lyophilized for 2 hours,

resuspended in MilliQ H2O at 20 mg/mL, and re-lyophilized overnight to remove residual

solvents. The lipids were then brought up at 100 mg/mL in buffer containing 10 mM HEPES pH

7.4, 50 mM TbCl3 and 85 mM sodium citrate, followed by 7 rounds of freeze-thaw between

room temperature and liquid N2, with vortex mixing during room temperature cycles, before

extruding through a 400 nm filter. LUVs were separated from free Tb+3 ions by gel filtration

using a SuperdexTM Peptide 10/300 GL column (GE Healthcare) in buffer containing 10 mM

HEPES pH 7.4 and 100 mM KCl. Liposome peaks were collected and used within 3 days.

The membrane disruption assay was performed in buffer containing 10 mM HEPES pH 7.4, 100

mM KCl, 50 mM dipicolinic acid (DPA), and 100 mM NaCl, in a final 1 mL volume.

Fluorescence emission scans were measured from 320-600 nm using an excitation of 314 nm on

PTI C60 spectrofluorometer, with the 1:1 DPA/Tb+3 complex having three emission maxima at

492 nm, 544, and 574 nm. Fibril samples were diluted in 20mM glycine-HCl buffer prior to

adding to liposome solution. Measurements were performed in triplicates, with the buffer

background subtracted and normalized to 100% LUV disruption with 0.1% Triton X-100.

Kinetic traces of dye release show fluorescence signal at 544 nm reached a plateau after 5 min,

and all measurements were recorded after 5 min incubation.

3.2.4 SSNMR measurements

SSNMR experiments were performed as described in Section 2.2 of Chapter 2.

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3.3 Results

3.3.1 ThT fluorescence and TEM measurements of sonicated fibrils

Cell cultures treated with sonicated PrP peptide fibrils of PrP(244-249) and PrP(245-250) had

decreased MTT reduction activity compared to mature fibrils, suggesting fibril fragments were

more cytotoxic than mature fibrils (Pastor et al. 2008). To generate fragmented fibrils, we probe-

sonicated samples that had been fibrillizing for more than 1 week (Figure 3.1 A-C) and studied

them using negative-stained TEM. The average fibril length after sonication were reduced in

length to ~ 130-150 nm (Figure 3.1 D-F). ThT fluorescence emission was measured for all three

peptides and reflected the same ThT-binding results as mature fibrils in Chapter 2. Sonicated

PrP(244-249) and PrP(245-250) fibrils bound to ThT with a concentration-dependent increase in

fluorescence at 482 nm, but PrP(178-183) fibrils did not (Figure 3.1 G-I). This suggested that the

sonicated fibrils retained their cross-β structure and that there were no major morphological

changes upon sonication.

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Figure 3.1. Morphological changes to sonicated PrP peptide amyloids. Electron micrographs of sonicated fibrils

prepared after 2 minutes of ultrasonication of mature fibrils formed by (A) PrP(244-249), (B) PrP(245-250), and (C)

PrP(178-183). Scale bars represent 500nm in each case. (D-F) Fibril length distributions after sonication. The mean

length of sonicated fibrils were measured from > 100 particles to be 154 ± 4 nm, 130 ± 4 nm, and 128 ± 2 nm for

PrP(244-249), PrP(245-250), and PrP(178-183), respectively. (G-I) ThT fluorescence emission spectra of sonicated

fibrils.

3.3.2 Hydrophobic surfaces of sonicated PrP peptide amyloids

The cytotoxicity of intermediate fibril species has been shown to increase with exposed

hydrophobic surface area (Mannini et al. 2014). To investigate the surface hydrophobicity of

sonicated PrP peptide fibrils, we measured the binding to the fluorescent dye ANS. Shown in

Figure 3.2, a concentration-dependent binding of ANS to mature PrP(244-249) fibrils was

observed, as expected since there are surface-exposed hydrophobic residues in both class 2 and

class 3 fibril structures. ANS fluorescence was increased upon sonicating the fibrils, suggesting

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that fragmentation led to an increase in exposed hydrophobic surface. PrP(245-250) also

exhibited a concentration-dependent binding in the mature form, but sonication did not increase

the binding within error of the measurement. Finally, we did not observe any ANS binding to

PrP(178-183) fibrils in its mature and sonicated form, suggesting minimal hydrophobic exposure

– consistent with surface-exposed polar residues indicated by our NMR structures determined in

Chapter 2.

Figure 3.2. Surface hydrophobicity of PrP peptide amyloids. ANS binding to mature and sonicated fibrils of (A)

PrP(244-249), (B) PrP(245-250) and (C) PrP(178-183). Fluorescence emission maxima at 470 nm are plotted in the

presence of increasing concentration of mature fibrils (colour) and sonicated fibrils (black). Measurements were

performed in triplicate (* corresponding to p < 0.05).

3.3.3 Liposome disruption by sonicated fibrils formed by PrP(244-249),

PrP(245-250), and PrP(178-183)

We monitored the membrane disruption activity of these fibrils using a dye-release assay.

Disruption of large unilamellar vesicles (LUVs) containing Tb+3 ion releases the ion, allowing it

85

to bind to the chelator DPA in bulk solution, forming a fluorescent complex. The fluorescence

emission of the DPA/Tb+3 complex at 544 nm can be interpreted as disruption of the lipid

bilayer. PrP(244-249) fibrils were shown to disrupt LUVs composed of zwitterionic (POPC) as

well as an anionic (PC/PG) lipid mixture, and this activity was increased after sonication (Figure

3.3). PrP(245-250) fibrils also exhibited sonication enhanced disruption of anionic LUVs.

However, this enhancement of disruption was not seen for zwitterionic LUVs. Meanwhile, non-

cytotoxic PrP(178-183) did not disrupt LUVs of either anionic or zwitterionic lipid

compositions, regardless of sonication.

Figure 3.3. Membrane disruption activity of PrP peptide amyloids upon sonication. Liposome dye release

assay of anionic PC/PG (A) or zwitterionic PC-only (B) lipid compositions treated with mature and sonicated PrP

fibrils. Fraction of dye released was normalized to complete LUV disruption with 0.1% Triton X-100 in each

measurement and carried out in triplicate (**, corresponding to p < 0.01; *, p < 0.05). Error bars represent standard

deviation of measurements.

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3.3.4 Chemical shift assignments of sonicated PrP peptide fibrils

The molecular-level effects of sonication on PrP peptide fibrils were examined using SSNMR.

We prepared sonicated fibrils using 13C, 15N-incorporated peptides, and flash-froze the samples

immediately after sonication to encapsulate all fragmented fibril species generated. The

lyophilized samples were then packed into an MAS rotor to carry out 13C chemical shift

assignments. Due to the spectral complexity of the PrP(244-249) peptide (as described in

Chapter 2 above), we prepared two peptide mixtures for PrP(244-249) – [U-13C,15N]-SI/F and

[U-13C,15N]-IL/F – to help with assignment by reducing the number of resonances in each

spectrum. In the case of PrP(178-183) we used a mixture of [U-13C,15N]-DCN/VI. Chemical shift

assignments of all three fibrils after sonication suggested retention of β-secondary structure as

indicated by the secondary chemical shift differences: Cβ exhibited a downfield shift (positive

difference) relative to random coil values, while CO and Cα exhibited upfield shifts (negative

difference) (Figure 3.4).

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Figure 3.4. Secondary 13C chemical shift calculations observed for sonicated PrP peptide amyloids. 13C

secondary chemical shift values are shown for PrP(244-249) chain A and B, PrP(245-250), and PrP(178-183).

3.3.5 β-sheet arrangement of sonicated fibrils

Next, we measured the distance between neighbouring strands in the β-sheet using the 13C – 13C

dipolar recoupling experiment CT-PITHIRDS for PrP(244-249) and PrP(245-250). As for the

mature fibrils, each measured dephasing curve corresponded to the spacing between a single

specifically 13C-labelled backbone carbonyl on adjacent β-strands. We determined that both

sonicated fibrils had an inter-strand distance of 5.0 Å (Figure 3.5) when compared to simulated

distance calculations (solid line) by SPINEVOLUTION. These measurements suggest that the

fibrils retained their in-register parallel β-sheet arrangement after sonication.

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Figure 3.5. 13C inter-β-strand distance measurements of sonicated PrP peptide fibrils. Dipolar dephasing

curves obtained from 13C – 13C CT-PITHIRDs homonuclear recoupling experiment for fibrils PrP(244-249) and

PrP(245-250). (A) Single-site labeled PrP(244-249) F246CO fibrils before (open circle) and after (blue diamond)

sonication. (B) Single-site labeled PrP(245-250) F246CO fibrils before (open square) and after (green triangle)

sonication. Simulated dephasing curves best fit to 13C atoms spaced 5.0 Å apart and was plotted here as a solid line.

Error bars were determined as the RMS noise in each spectrum.

3.3.6 Different chemical environments and the loss of intermolecular

contacts in the fibril core upon sonication

We noticed that the chemical shifts of sonicated fibrils were slightly different from those of

mature fibrils. Mature PrP(244-249) fibrils contained two sets of 13C and 15N chemical shifts

corresponding to two inequivalent steric zipper arrangements – parallel β-sheets stacked face-to-

face (class 3) and parallel β-sheets stacked face-to-back (class 2). After sonication, one set of

chemical shifts exhibit less than 1ppm deviation from mature fibrils (species 1), and the other

showed greater than 1ppm difference from mature fibril chemical shifts (species 2) (Figure 3.6).

We were unable to assign which of these species corresponded to the two steric zipper

arrangements based on chemical shift alone due to their overlap, requiring further experiments as

discussed in the next section. Interestingly, we observed a new set of chemical shifts appearing

in PrP(245-250) fibrils (species 2) in addition to the original chemical shifts (species 1).

Meanwhile, PrP(178-183) fibrils did not exhibit large chemical shift changes upon sonication,

and no new chemical shifts were observed. These data suggest that fragmentation of the amyloid

fibrils could be inducing slight structural changes to PrP(244-249) and PrP(245-250) fibrils such

that a subset of the peptides were exposed to a new chemical environment (species 2) different

from the mature fibril core (species 1), while PrP(178-183) remained the same. Combined with

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ANS data, these results suggest that the changes to the chemical environment of the peptides are

likely due to altered solvent exposure.

Figure 3.6. PrP peptides experience a different chemical environment after sonication. 13C chemical shift

differences between mature and sonicated fibrils were calculated to indicate a subset of peptides in PrP(244-249)

and PrP(245-250) sonicated fibrils had different chemical shifts than the mature fibrils (species 2). PrP(178-183)

sonicated fibrils did not exhibit this new set of chemical shifts.

13C – 13C correlation experiments were collected using 500ms RAD/DARR mixing time to

examine any structural changes in the fibril core upon sonication. As shown in Figure 3.7, the

long-range quaternary contacts between F246 and I248 in PrP(244-249) fibrils have been

reduced after sonication. The cross peak intensity between the F246 aromatic ring and I248

γCH3 was decreased in the SI/F sample (Figure 3.7 A). Similarly, the F246 aromatic – I244 γCH3

cross peak intensity in the IL/F sample was weaker after sonication. These observations suggest

that the I244, F246, and I248 contacts in the class 3 packing model were disrupted after

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sonication. On the other hand, the F246 aromatic – L247 δCH3 cross peak in the IL/F sample was

still present after sonication, suggesting that the class 2 packing interface was maintained (Figure

3.7 B). We still observed the Phe246 – Ile248 cross peak in PrP(245-250) SI/F fibrils (Figure 3.8

A), as well as the Val180 – Ile182 cross peak in PrP(178-183) DCN/VI fibrils (Figure 3.8 B).

Figure 3.7. Intermolecular β-sheet contacts in sonicated PrP(244-249) fibrils. 13C – 13C RAD/DARR spectra of

PrP(244-249) SI/F (A) and IL/F (B) fibrils obtained with 500 ms mixing time. 2D spectra at the aliphatic-aromatic

region highlight changed cross peak intensity between Phe, Ile, and Leu. 1D slices at Ile and Leu 13CH3 chemical

shifts are indicated with a dash line and shown on the right, with arrows pointing to methyl-phenyl ring cross peaks.

All spectra were obtained at an MAS frequency of 11 kHz.

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Figure 3.8. Intermolecular β-sheet contacts in sonicated PrP(245-250) and PrP(178-183) fibrils. 13C – 13C

RAD/DARR spectra of PrP(245-250) SI/F (A) and PrP(178-183) DCN/VI (B) fibrils obtained with 500 ms mixing

time. 1D slices at Ile182 are shown with arrows pointing to intermolecular side chain-side chain contacts. All spectra

were obtained at an MAS frequency of 11 kHz.

3.4 Discussion

Fibril fragmentation is a critical step in amyloid formation since it creates additional nucleation

sites to promote fibril elongation and reduce the lag time. At the same time, fragmented fibrils

are recognized to exhibit greater cytotoxicity than mature fibrils, akin to other small oligomeric

intermediates. The difference in toxicity between mature and fragmented fibrils is hypothesized

to relate to increased membrane interaction of the fragmented fibrillar species. However, current

literature has largely overlooked the structural properties of fragmented fibrils that may explain

the observed difference between the two species. In this chapter, we decided to examine the

effects of sonication on the structure of fibrils formed by three PrP derived hexapeptides

PrP(178-183), PrP(244-249) and PrP(245-250) by SSNMR and their interaction with membranes

to address their varying degree of cell culture toxicity (Pastor et al. 2008).

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Our investigations suggested that fibril fragmentation had minimal effects on the cross-β

backbone structure within the three fibrils. However at the molecular level, fragmentation could

disrupt the steric zipper packing within the fibril core, leading to an increased surface area of

solvent-exposed hydrophobic residues as reported by ANS binding. This increase in surface

hydrophobicity correlated with an increased ability to disrupt lipid bilayers, as demonstrated by

PrP(244-249) fibrils with both PC and PC/PG compositions. In the case of PrP(178-183) fibrils

that did not show any ANS binding before and after fragmentation, no membrane disruption was

observed. Solvent-exposed hydrophobicity was present in mature PrP(245-250) fibrils, but it was

not increased upon fragmentation. As a result, mature and fragmented PrP(245-250) fibrils

showed similar liposome disruption activity in zwitterionic lipids. Electrostatic interactions with

lipid headgroup possibly helped drive membrane interaction of fragmented PrP(245-250) fibrils,

leading to increase lipid disruption of anionic liposomes (Maltseva et al. 2005). The difference in

lipid disruption activity between PrP(244-249) and PrP(245-250) fibrils suggest different

sequences may have different driving forces for lipid interaction that is modulated by lipid

composition. Nonetheless, our results suggest that mature fibrils are less membrane-active and

less cytotoxic than the intermediate species possibly due to hydrophobic residues packed into the

fibril core and buried away from solvents.

A slight alteration in sequence for PrP(244-249) compared to PrP(245-250) (ISFLIF vs. SFLIFL)

affected both the peptide assembly within the fibrils and also the fragmentation-associated lipid

interaction. Thus, subtle sequence changes not only impacted fibril structure and stability, it

could also influence the fibril’s susceptibility to fragmentation and change the amount of

exposed hydrophobic groups that modulate biological behaviours. Although our studies only

examined amyloid toxicity induced through membrane interaction, it is likely that other

mechanisms associated with specific amyloidogenic proteins may be involved in the pathology

of different amyloid diseases. Surface-exposed hydrophobicity of various misfolded intermediate

conformations (pre-fibrillar oligomer, spherical oligomers, fibrillar oligomers, or fibril

fragments) can widely influence the interactions with biological components such as cell surface

receptors as well as membranes, and could manifest in different observable cytotoxic effects.

Therefore, it is critical to further our fundamental understanding of amyloid assemblies and their

interactions with other macromolecular systems to develop better therapeutic approaches for

disease-associated amyloids and to design new biomaterials from amyloids.

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Chapter 4.

Structural characterization of

the lipid-bound and amyloid

fibrillar states of human serum

amyloid A

Contributions:

Karen Simonetti (Lab manager of Sharpe lab, Hospital for Sick Children) constructed the

pET302 vector and carried out initial SAA2 expression trials. She also helped collect TEM of

rHDL-SAA and DLS of apoSAA. Undergraduate student Sympascho Young (University of

Toronto) carried out the ThT fibrillization kinetic measurements and TEM analysis of SAA

fibrils. Undergraduate student Quang Huynh (University of Toronto) carried out the liposome

clearance assay.

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4.1 Introduction

SAA is an acute-phase reactant up-regulated in the serum as part of the inflammatory cascade

(Uhlar & Whitehead 1999). It is an apolipoprotein synthesized predominantly in the liver and

normally bound to high-density lipoproteins (HDLs). In particular, native SAA has been shown

to displace apolipoprotein A-I (ApoA-I) to become the major apolipoprotein on the circulating

HDL3 particles during the inflammatory response. There are several roles proposed for SAA to

deal with inflammation and infection, which include recycling cholesterol and lipids at sites of

inflammation and tissue injury, activating the release of pro- and anti-inflammatory cytokines,

acting as a chemoattractant for leukocytes, and aiding in migration and binding of neutrophils

and macrophages at sites of inflammation (Urieli-Shoval et al. 2000; Derebe et al. 2014;

Preciado-Patt et al. 1996; Xu et al. 1995; Eckhardt et al. 2010). In addition, expression of lipid-

free SAA has been detected in extrahepatic tissues, although the function for the apo-form of

SAA are not yet known (Malle & de Beer 1996; Clifton et al. 1985; Coetzee et al. 1986).

SAA is highly conserved across vertebrates and invertebrates, with multiple genes and proteins

being documented in each species (Malle & de Beer 1996; Uhlar et al. 1994). Human SAA

family consists of four members that are 104-112 residues long, but only two members SAA1

and SAA2 are up-regulated during acute inflammation and are referred to as acute-phase SAA

(A-SAA) in the literature. They each have their own allelic variants (SAA1.1 - 1.5, SAA2.1 -

2.2) that share 96% sequence homology, with the exception of SAA2.2 that is C-terminally

truncated (Figure 1.8 A). Recent reports on the structure and assembly of human SAA1 and

mouse SAA3 revealed that the protein adopts a 4-helix bundle structure, containing an N-

terminal amphipathic helix that can interact with lipids and a disordered C-terminal tail that has

been proposed to interact with other cellular components such as GAG (Figure 1.8 B) (Segrest et

al. 1976; Lu et al. 2014; Derebe et al. 2014). Soluble apo-SAA also oligomerizes into marginally

stable hexamers or octamers in solution, with the C-terminus thought to play a role in regulating

this oligomerization and assembly (Wang & Colon 2005; Wang et al. 2012; Wang et al. 2002;

Takase, Tanaka, et al. 2014; Patke et al. 2012; Maszota et al. 2015).

Persistent high levels of A-SAA in patients with chronic infections or inflammatory diseases

such as rheumatoid arthritis, Crohn’s Disease, and tuberculosis, has been shown to result in

systemic amyloid A (AA) amyloidosis in 15% of patients (Röcken & Shakespeare 2002).

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Amyloids isolated from patients show that the N-terminal 76 residues of SAA are deposited into

amyloids along with other serum components such as GAGs and serum amyloid P. Apo-SAA

oligomers were also reported to form amyloid fibrils under physiological temperatures, and

promoted by acidic conditions and the presence of GAG (Wang et al. 2005; Elimova et al. 2009;

Ye et al. 2011; Aguilera et al. 2014). Decreasing the length of the GAG chains in matrix

proteoglycans made animals resistant to AA deposition (Merlini et al. 2007; Li et al. 2005).

These point to a strong biological relationship between GAG and SAA and the presence of

specific interactions promoting amyloid formation.

Recently, Takase et. al. demonstrated that SAA1 can solubilize lipid vesicles to form discoidal

particles (rHDL) with similar morphologies to nascent HDLs (Takase, Furuchi, et al. 2014).

SAA associated with these rHDL particles remained α-helical with higher thermal stability than

soluble apo-SAA, and were less prone to protease digestion under physiological temperatures.

SAA displacing ApoA-I also did not change the stability and fusion of HDLs isolated from

serum, suggesting that SAA and ApoA-I provided similar stability to the HDL particles

(Jayaraman et al. 2015). Nevertheless, the crystal structure of the apo-SAA1.1 does not fully

explain the ability for SAA to replace ApoA1 and scaffold HDL or HDL-like particles. We

hypothesize the SAA structure in the HDL-bound state may be similar to ApoA-I and different

from the 4-helix bundle structure of the apo-form. The structure of amyloid and HDL-bound

forms of SAA has not been reported, making it hard to determine the mechanism of SAA

misfolding and whether apo- or HDL-bound SAA is the origin for AA amyloidosis.

Here, we used SSNMR to characterize the structure of the amyloid and lipid-bound states of

SAA2.1 (referred herein as simply SAA unless otherwise indicated). Under acidic conditions and

in the presence of GAGs (heparin and heparan sulfate), we observed that SAA readily formed

morphologically homogeneous fibrils. Fibrillization kinetics were probed using ThT

fluorescence which showed a rapid build-up of ThT-binding species within 2 days of

fibrillization at pH 3.6 and physiological temperatures. Heparin-induced fibrils were examined

by FTIR and SSNMR, identifying a major structural rearrangement of helix-rich SAA into β-

sheet rich fibrils. NMR chemical shift analysis further suggested that these fibrils are composed

of a rigid N-terminal fibril core and a mobile C-terminal tail (residues 76-104). Meanwhile, lipid-

bound SAA solubilized lipid vesicles to form discoidal rHDLs and adopted α-helical secondary

structure with higher thermal stability that apo-SAA. Using ThT fluorescence kinetics

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measurements, we showed that GAG-induced fibrillization was hindered in rHDL-SAA. Initial

NMR chemical shift analysis supports a highly α-helical nature of rHDL-SAA. The C-terminus

of the protein was disordered in the lipid-free and fibrillar forms, yet appeared ordered and

relatively rigid in the rHDL-SAA sample. Such structural changes may be implicated in

functional differences between soluble and HDL-bound SAA.

4.2 Methods

4.2.1 Recombinant SAA expression

N-terminal 6xHis-tagged human SAA isoform 2.1 (6xHis-thombin-hSAA2.1) was cloned into a

pET302 vector with T7 promoter and transformed into E. coli strain BL21(DE3) (Invitrogen) for

expression. All growth media was supplemented with 50μg/mL carbenicillin. 50mL Luria broth

(LB) starter culture was inoculated from a glycerol stock and grown at 37oC overnight. Sufficient

starter culture to produce an OD600 of 0.15 in 1 L media was centrifuged (5000g at 4oC for 20

min) and resuspended in fresh LB media. The 1 L LB expression culture was grown at 37oC to

an OD600 of 0.7 and induced with 1mM isopropyl-β-D-thiogalactopyranoside (IPTG) for 3 h at

37oC. Cells were harvested and pellets stored at -20oC. 13C, 15N-labelled protein was prepared

using the same 50mL LB starter culture, but subsequent expression culture was carried out in M9

minimal medium supplemented with 1 g/L 15NH4Cl and 2g/L uniform-13C-glucose (Cai et al.

1998).

4.2.2 Purification and sample preparation of lipid-free SAA

The cell pellet from 1 L growth was resuspended in 20mL lysis buffer (20 mM Tris pH 8, 250

mM NaCl, 2.5 mM MgCl2) containing 12.5 μg/mL DNase I and 1 tablet of complete EDTA-free

protease inhibitor (Roche). Cells were lysed by homogenization and centrifuged (17000 g at 4oC

for 45 min) to isolate the inclusion bodies containing the 6xHis-hSAA2.1 protein. Ni+2–NTA

(Qiagen) affinity chromatography was carried out on the inclusion body in 6 M guanidinium

chloride (GuHCl), 50 mM sodium phosphate pH 8.0, and 250 mM NaCl. The eluted protein was

refolded at room temperature via rapid dilution into refolding buffer (55 mM Tris pH 8.2, 21 mM

NaCl, 0.88 mM KCl, 1.1 M GuHCl, 400 mM L-arginine, 1 mM EDTA, 1 mM reduced

glutathione, and 1 mM oxidized glutathione) and extensively dialyzed against low salt buffer (20

mM sodium phosphate pH 7.5, 21 mM NaCl, 1 M Urea) at 4oC to remove residual GuHCl before

removing the 6xHis-tag with thrombin protease as per the manufacturer’s instructions (GE

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Healthcare). Thrombin was removed using a benzamidine-sepharose (GE healthcare) column

and residual protease activity was stopped with 1 mM PMSF (Sigma-Aldrich). The sample was

concentrated using Centricon 3 kDa MWCO filter (Amicon) at 4oC, and checked for

contaminants using SDS-PAGE. Samples were stored at 4oC in low salt buffer containing 1M

urea, dialyzed to remove urea, and centrifuged prior to use to remove aggregated material.

Protein concentration was calculated from OD280 using the extinction coefficient ε280 = 23,950

M2 mol-1 (www.expasy.org) and a final molecular weight of 11,772 Da after thrombin cleavage.

4.2.3 Preparation of lipid vesicles and lipid-bound SAA on reconstituted

high-density lipoprotein (rHDL-SAA)

Preparation of rHDL-SAA was carried out following the protocol reported by Takase et. al.

(2014). Briefly, DMPC vesicles were prepared from a chloroform stock (20 mg/mL, Avanti) by

drying lipids into a thin film under a gentle stream of N2(g), vacuum dried for 1 hour, then

respuspended in 1 mL MilliQ H2O followed by lyophilization overnight to remove residual

organic solvents. Lipids were then rehydrated at 5 mg/mL in 20 mM sodium phosphate pH 7.5,

21 mM NaCl buffer, with 7 freeze-thawed cycles and vigorous vortexing between cycles. DMPC

vesicles were incubated with SAA2.1 (0.5 mg/mL) at 24 oC for 2 hours to form rHDL particles.

Light scattering was measured at 406 nm on an Ultrospec™ 2011 pro UV/Visible

Spectrophotometer (GE Healthcare).

4.2.4 SAA fibrillization kinetics measurements

SAA samples were dialyzed into pH 3.6 buffer (10 mM sodium acetate pH 3.6, 21 mM NaCl) or

pH 7.5 buffer (20 mM sodium phosphate pH 7.5, 21 mM NaCl) for fibrillization. ThT binding

kinetics were carried out in a 96-well plate (Greiner black, clear-bottom) using the SpectraMax

i3 plate reader at 37oC. 200 μL sample in each well contained a final concentration of 0.35

mg/mL protein, 0.1 mg/mL GAG (heparin or heparan sulfate), and 8 μM ThT, and then topped

with 50 μL paraffin oil to prevent sample evaporation. Fluorescence measurements in triplicates

were recorded every 30 min with rocking for a total of 96 hours. Excitation and emission

wavelengths were set to 442 and 482 nm, respectively.

4.2.5 Sample preparation of rHDL-SAA and SAA fibrils for SSNMR

Polyethylene glycol (PEG) precipitation of rHDL-SAA particles was adopted from similar

methods used to prepare NMR samples of lipid-protein nanodiscs, as described by Sligar et. al.

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(2006). Briefly, rHDL-SAA particles were prepared with uniformly 13C, 15N-labelled SAA and

precipitated with the addition of 3-volume equivalents of 40% PEG 3350 (in 20 mM sodium

phosphate pH 7.5, 21 mM NaCl). Samples were left to precipitate overnight at 4oC and collected

by centrifugation at 100,000 g for 4 hrs at 4oC. SAA amyloid fibrils were prepared using 13C,

15N-labelled lipid-free SAA; the protein was dialyzed into pH 3.6 buffer (10 mM sodium acetate

pH 3.6, 21 mM NaCl) and fibrillized at 37oC with agitation in the presence of heparin (0.35

mg/mL protein, 0.1 mg/mL heparin) for 14 days. SAA fibrils were collected by centrifugation at

100,000 g for 4 hours at 4oC. The centrifuged pellets were packed into a 36 μL Kel-F insert for a

4 mm MAS zirconia rotor (Bruker).

4.2.6 Gluteraldehyde cross-linking of SAA

Gluteraldehyde cross-linking experiments were carried out in 20 mM sodium phosphate buffer

pH 7.5, 21 mM NaCl with and without 1 M Urea. 50 μg of protein in a total volume of 200 μL

(0.25 mg/mL or 20 μM) was treated with 10 μL of a freshly prepared solution of 2.5 %

gluteraldehyde at 37oC. 30 μL aliquots of cross-linked sample was removed at indicated time

intervals and quenched by the addition of 3 μL 1 M Tris-HCl pH 8.0. Cross-linked proteins were

then solubilized by adding an equal volume of Laemmli sample buffer and run in 12% SDS-

polyacrylamide gels.

4.2.7 Dynamic light scattering (DLS)

DLS experiments were performed using a DynaPro-99 instrument (Wyatt/ProteinSolutions) at

20oC. 50 μL sample contained 20 μM SAA in 20 mM sodium phosphate buffer pH 7.5, 21 mM

NaCl, 1 M Urea. Data were recorded and processed using Dynamics 5.26.38 software, using a 10

μs sampling time averaged over 20 scans.

4.2.8 Circular dichroism and thermal stability

Circular dichroism (CD) measurements were collected on Jasco J-810 spectropolarimeter using a

1 mm path length quartz cuvette (Hellma Analytics). Spectra were measured from 190-250 nm at

a scan rate of 50 nm/min and averaged over three measurements. Thermal denaturation at [θ]220

was carried out from 10 - 85oC with heating at 3 oC/min, with every 5 oC performing a full

spectral 190-250 nm scan. CD samples contained approximately 20 μM protein.

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4.2.9 FTIR measurements

SAA was fibrillized in pH 3.6 buffer at 37oC, in the presence of heparin, for 15 days (0.35

mg/mL SAA to 0.1 mg/mL heparin). Samples were absorbed onto a hydrophilic

polytetrafluoroethylene (PTFE) membrane on the Direct Detect assay-free card and FTIR spectra

recorded with Direct Detect infrared spectrometer (EMD Millipore) as per the manufacturer’s

instructions. Buffer components were recorded separately and subtracted from the final FTIR

spectra. Second derivatives of the FTIR absorbance spectra were calculated using OriginPro for

data analysis.

4.2.10 Solution and solid-state NMR

Solution NMR experiments were recorded on Bruker Avance III 600 MHz (14.3 T) spectrometer

equipped with a 5 mm TXI triple resonance probe with Z-axis gradients. 2D 1H – 15N HSQC and

3D backbone experiments HNCA, HN(CO)CA, and CBCA(CO)NH were collected at 14oC on

13C, 15N-labeled lipid-free SAA in pH 7.5 buffer containing 20 mM sodium phosphate, 21 mM

NaCl, 2% DPC, and 10% D2O. All spectra were externally referenced to 4,4-dimethyl-4-

silapentane-1-sulfonic acid (DSS). Data processing were carried out on NMRPipe and CCPNMR

software (Delaglio et al. 1995; Vranken et al. 2005).

Solid-state NMR experiments were carried out on Bruker Avance III-HD 700MHz spectrometer

(16.3 T) using a 4 mm DVT MAS triple resonance probe. Sample cooling gas was maintained at

5oC and a sample temperature of approximately 15oC due to heating from spinning friction.

MAS spinning frequency was maintained at 10,000 ± 5 Hz, and spectra were externally

referenced to the downfield 13C resonance of adamantane at 38.56 ppm relative to

tetramethylsilane (TMS) (Morcombe & Zilm 2003). Typical π/2 pulse widths were 2.5 μs on 1H,

4.0 μs on 13C, and 5.0 μs on 15N. 1H – 13C and 1H – 15N cross-polarization (CP) transfers were

implemented using a linear ramped RF field centered around 40 - 60 kHz on the low channel and

a 50 - 80 kHz field on the 1H channel, with contact times of 1 to 1.5 ms. 1H – 13C INEPT transfer

was achieved using delays correspond to JHC = 145 Hz. 1H decoupling was achieved using a two

pulse phase modulation (TPPM) decoupling scheme with 100 kHz field strength during all t1 and

t2 periods, (Bennett et al., 1995).

2D 13C – 13C correlation spectra were collected via dipolar recoupling experiments (RAD/DARR

transfer scheme) with 25 - 500 ms mixing (Morcombe et al. 2004), or via scalar coupling

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experiments (TOBSY, P19 recoupling) with a 9.6ms mixing time (Leppert et al. 2004). On

average, spectra were collected in 24 hours using 128 scans per FID and 160 complex points in

the indirect dimension.

2D and 3D 13C – 15N correlation spectra were collected using a selective double CP pulse

sequence under 10 kHz MAS, with selective transfer from 15N to 13C using SPECIFIC CP for

NCO or NCA spectra (Petkova et al., 2003). 15N – 13C SPECIFIC CP after the t1 period was

achieved using a linear ramp on the 15N channel with RF fields of 25-35 kHz on each channel,

and a contact time of 2 ms. A RAD/DARR mixing period of 25 - 35 ms was implemented after

15N – 13C SPECIFIC CP to achieve 13C – 13C transfers. 2D spectra were collected using 128

scans and 80 complex points in indirect dimension. 3D spectra were collected in 8 days with 48

scans per FID and 24 complex points in both the first and second indirect dimensions. Non-

uniform sampling (NUS) was implemented for 3D spectra using 25% data sampling to reduce

the experimental time with identical experimental data points. NUS data was processed with

MddNMR software to reconstruct a complete 3D dataset (Orekhov & Jaravine 2011), and

analyzed with CCPNMR.

4.3 Results

4.3.1 Characterization of soluble lipid-free SAA oligomers

To examine whether soluble SAA2.1 existed as monomeric or oligomeric in solution, lipid-free

SAA (apo-SAA) was cross-linked using gluteraldehyde at 37 oC in buffer containing 1 M urea,

which was the buffer for storage and thrombin cleavage, and in buffer containing no urea that

reflected buffer conditions in subsequent experiments. Gluteraldehyde was selected as an

indiscriminant crosslinking agent to couple any Lys residues within the oligomer without any

distance constraints. We observed distinct laddering of cross-linked SAA with identifiable bands

at molecular weights corresponding to monomer, dimer, trimer, and tetramer (Figure 4.1 A).

Larger species were also observed at longer cross-linking times. Size measurement of apo-SAA

by DLS showed a dominant species of approximately 75 kDa, corresponding to a hexamer

(monomer = 11.7 kDa). High molecular weight species were also present in DLS experiments,

suggesting a tendency of apo-SAA to higher-ordered oligomers or aggregates (Figure 4.1 B).

Use of size-exclusion gel filtration to determine molecular weight was unsuccessful as the

protein tended to interact with the column, preventing analysis (not shown). As expected, 1H –

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15N HSQC spectrum collected for apo-SAA showed broadened peaks indicative of a large

macromolecule with slow tumbling and fast T2 relaxation (Figure 4.1 C). Meanwhile, the peaks

that were observable had poor chemical shift dispersion suggesting a disordered structure for

those sites. Using standard 3D NMR experiments for backbone assignment, we were able to

identify the narrow resonances as arising from the C-terminal region of apo-SAA. Complete

resonance assignment will require preparation of deuterated SAA, coupled with TROSY NMR

experiments, and is beyond the scope of this thesis.

Figure 4.1. Oligomeric apo-SAA with a dynamic and flexible C-terminus. (A) Gluteraldehyde cross-linking

(XL) of apo-SAA in buffer with and without urea shows a laddering of SAA as a monomer, dimer, trimer, and

possible tetramer. Larger oligomers and aggregates are seen at later crosslinking times, consistent with the reported

aggregative nature of apo-SAA. (B) DLS of apo-SAA showing a potentially heterogeneous oligomeric SAA

composed of hexamers and larger species in solution. (C) Solution NMR 1H – 15N HSQC of 380 μM apo-SAA (4.5

mg/mL) at 14oC in pH 7.5 buffer containing 20 mM sodium phosphate, 21 mM NaCl, and 2% DPC.

4.3.2 SAA solubilizes DMPC vesicles and forms discoidal HDL-like

particles

Apolipoproteins such as ApoA-I and SAA1 have been shown to solubilize DMPC vesicles and

spontaneously form discoidal HDL-like particles (Pownall et al. 1978; Takase, Furuchi, et al.

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2014). To investigate whether SAA2 exhibited similar vesicle-clearance activity, we incubated

apo-SAA with DMPC vesicles at the gel to liquid-crystalline phase transition temperature (24

oC) and monitored sample turbidity with absorbance at 406 nm. The observed decrease in light

scattering upon addition of protein indicated that SAA2 was also able to solubilize DMPC lipid

vesicles (Figure 4.2 A). At 1:1 (w/w) lipid-to-protein ratio (18:1 molar ratio), all vesicles were

solubilized and formed discoidal rHDL particles that were 10-15 nm in diameter (Figure 4.2 B).

Figure 4.2. SAA solubilizes DMPC vesicles to form discoidal HDL-like particles. (A) Vesicle solubilization

activity of SAA was tested using 80 μg (blue, triangle) and 120 μg (orange, square) DMPC incubated with apo-SAA

for 2 hrs at 24.6oC, in triplicates. Vesicle clearance was monitored by relative light scattering at 406nm. (B)

Negative-stained TEM of rHDL particles formed at 1:1 lipid-to-protein (w/w) ratio after 2 hrs of incubation.

Micrographs were stained with 2% (w/v) uranyl acetate.

4.3.3 Secondary structure and thermodynamic stability of apo- and lipid-

associated SAA

CD spectroscopy showed that SAA was predominantly α-helical in both apo- and lipid-

associated states at 10oC (Figure 4.3 A). However, the stability of SAA was significantly

different between the two states when tracking temperature-dependent unfolding using the

ellipticity at 220 nm. Apo-SAA had a sigmoidal thermal denaturation curve with a midpoint

melting temperature (TM) of 36.3oC, whereas lipid-bound rHDL-SAA had a markedly higher TM

of 49.8oC (Figure 4.3 B). Apo-SAA also exhibited a broader unfolding transition, with

significantly less α-helical structure at lower temperatures than rHDL-SAA. This is in agreement

with lipid-free apolipoproteins being partially unfolded under physiological temperatures of 37oC

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to allow exposure of amphipathic helices for lipid association. Both states of SAA were

completely unfolded by 60oC and could be refolded back to its native α-helical structure, despite

a slight reduction in protein concentration due to small amounts of aggregation (not shown).

Figure 4.3. SAA secondary structure and thermodynamic stability. (A) Molar ellipticity of apo-SAA and

rHDL-SAA collected in pH 7.5 sodium phosphate buffer, 21 mM NaCl, 10oC, with 0.25 mg/mL protein

concentration. (B) Thermal denaturation curves of apo-SAA (blue diamond) and rHDL-SAA (yellow triangle) were

tracked at 220 nm under the same buffer condition to show.

4.3.4 Fibrillization of lipid-free SAA in presence of GAGs

SAA fibrillization kinetics were measured at both pH 7.5 and pH 3.6 to examine the effects of

GAGs and pH on SAA2 fibril formation (Figure 4.4 A, B). We tracked ThT fluorescence at 482

nm over 4 days and observed that SAA did not fibrillize at pH 7.5 even in the presence of the

amyloid-inducing GAGs heparin and heparan sulfate. Meanwhile, SAA in pH 3.6 buffer

exhibited a long lag phase, with subsequent fibrillization after agitating for 40 hrs. Presence of

heparin and heparan sulfate drastically reduced the nucleation lag phase, and ThT fluorescence

increased within the first timepoint of measurement (30 min). ThT fluorescence plateaued after

25 hrs in both heparin and heparan sulfate-induced fibrillization, although the fluorescence

intensity in heparan sulfate-induced fibrils was lower than heparin-induced fibrils. Negative-

stained TEM images of samples prepared at the end of this time course lacked the typical fibrillar

morphology of mature amyloids (not shown), and only after 10 days of further incubation at

37oC did we observe morphologically homogeneous fibrils (Figure 4.4 C). Surprisingly, fibril

morphologies resulting from the two GAGs were slightly different under TEM, with heparin-

induced fibrils being thinner and smoother (13 ± 2 nm) than heparan sulfate-induced fibrils (17 ±

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1 nm). The different levels of ThT fluorescence may be a result of different fibril morphology, or

different fibril organization at the molecular level.

Figure 4.4. GAG-induced fibril formation of SAA at 37oC. ThT binding kinetics measurement of SAA

fibrillization in the presence of heparin (red), heparan sulfate (blue), or absence of any GAG (green), were

performed at pH 3.6 (A) and pH 7.5 (B). Each fibrillization condition contained 0.35 mg/mL SAA, 0.1 mg/mL

GAG, 21 mM NaCl, 8 μM ThT, and were incubated at 37oC with agitation. All measurements were performed in

triplicates, with error bars representing the standard deviation. (C) Negative-stained TEM images were taken after

10 days of incubation at 37oC, showing morphological differences of SAA fibrils induced from heparan sulfate (left)

and heparin (right). Heparan sulfate-induced fibrils were thick and had rough edges, while heparin-induced fibrils

were thin and smooth. The mean widths of SAA fibrils were measured, giving 17 ± 1 nm in samples with heparan

sulfate and 13 ± 2 nm in samples with heparin. Micrographs were stained with 2% (w/v) uranyl acetate

We compared the secondary structure of SAA fibrils to that of soluble apo-SAA by FTIR to

identify structural changes that occurred as a result of fibrillization (Figure 4.5 A). As expected,

soluble apo-SAA showed a strong peak at 1655 cm-1 in the amide I region (1600 – 1700 cm-1)

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arising from its α-helical structure, while heparin-induced fibrils were marked by a strong peak at

1625 cm-1 suggesting a predominantly β-sheet secondary structure. In the second derivative

analysis, we also identified peaks around 1660 – 1685 cm-1 and attributed them to β-turn

structures (Figure 4.5B). There was no 1695 cm-1 peak recognized in the FTIR spectrum of SAA

fibrils, suggesting a possible parallel β-sheet arrangement (Kubelka & Keiderling 2001; Dong et

al. 1990).

Figure 4.5. Secondary structure of apo-SAA and SAA amyloid fibrils. (A) The amide I region of FTIR spectra

of apo-SAA (blue) and heparin-induced SAA fibrils (black) showing changes in secondary structure composition.

(B) Second derivative analysis of FTIR spectra, allowing deconvolution of the underlying peaks and analysis of the

secondary structure of SAA. Range of values for FTIR bands corresponding to specific secondary structure

elements: 1624-1642 cm-1 – β-sheet; 1648 cm-1 – random coil; 1656 cm-1 – α-helix; 1667-1685 cm-1 – β-turns (Kong

& Yu 2007).

4.3.5 Fibrillization of lipid-associated SAA in presence of GAGs

Fibril formation by SAA incorporated in rHDL particles was also monitored by ThT

fluorescence. As shown in Figure 4.6, rHDL-SAA alone did not fibrillize after 40 hrs incubation

at 37oC in the absence of any GAGs, but the addition of heparin and heparan sulfate promoted

fibril formation within the first timepoint of measurement as similar to apo-SAA. There was a

markedly slower increase in ThT fluorescence of fibrillization compared to lipid-free SAA. Most

notably, ThT fluorescence intensity in heparan sulfate-containing rHDL samples was much

lower than observed for apo-SAA after 48 hrs, while the heparin-containing samples both

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reached the same fluorescence intensity after 30 hrs. This suggested that fibrils can still be

formed from HDL-bound SAA when exposed to GAGs.

Figure 4.6. GAG-induced fibril formation of rHDL-SAA at pH 3.6. ThT fibrillization kinetics of rHDL-SAA

and apo-SAA performed in the presence of heparin (red, yellow), heparan sulfate (blue, light blue), or absence of

any GAGs (green, black) at 37oC. Sample contained 0.35 mg/mL SAA/rHDL-SAA, 0.1 mg/mL GAG, 21 mM NaCl,

and 8 μM ThT. Error bars represent the standard deviation for the triplicate measurements.

4.3.6 SSNMR of SAA amyloid fibrils

To examine the structure of SAA amyloid fibrils in greater detail, we fibrillized uniformly-13C,

15N-labelled, lipid-free protein in the presence of heparin (under the same pH 3.6 buffer

conditions as the SAA kinetics studies above). We initially collected 1-dimensional (1D) 13C

spectra to study the local dynamics within SAA fibrils. Direct pulse (DP) 13C excitation

identified all 13C nuclei in the sample and served as a reference spectrum, while experiments

starting with 1H – 13C polarization transfer through CP and INEPT were selective for immobile

or mobile regions of the protein, respectively. As the fibril core is hypothesized to be more rigid,

we expected residues located in the β-sheet core to have greater efficiency in the dipolar-based

CP transfer.

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As shown in Figure 4.7, CP and INEPT transfers yielded drastically different 1D 13C spectra for

SAA amyloid fibrils. The 1H – 13C CP spectrum exhibited broad protein 13C peaks in the

aliphatic (0-70ppm), aromatic (110-140 ppm), and carbonyl regions (165-180 ppm), while the 1H

– 13C INEPT spectrum had sharper peaks in the same regions. As carbonyl 13C atoms lack

directly-bonded 1H, no signals were observed in that region of the INEPT spectrum. As

expected, the DP spectrum of SAA fibrils was a convolution of the broad peaks observed in CP

and the sharp peaks seen in INEPT spectra, with the exception of the carbonyl region.

Interestingly, we did not observe strong peaks from the saccharides in GAGs, which would be

expected to resonate in the 60-110 ppm region (Holme & Perlin 1989). We speculate that either

heparin did not co-aggregate with the fibrils, or that there was too low abundance in the fibrils to

observe signal of the unlabeled GAG relative to the isotope-labeled protein.

Figure 4.7. 1D 13C spectra of uniformly 13C, 15N-labelled SAA2 fibrils examined by MAS SSNMR. Direct

pulsed (top, black), 1H – 13C cross-polarization (middle, red) and 1H – 13C INEPT spectra (bottom, green) were

recorded under 10kHz MAS with VT gas set to 5oC. All spectra collected with 1024 scans. CP contact time was 1.5

ms while INEPT transfer time was set to τ1/2 = 1.742 ms (J = 145 Hz).

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We further examined the mobile and immobile regions in SAA fibrils using 2D 13C – 13C

homonuclear correlation experiments. To achieve this, we used a CP-RAD/DARR dipolar-based

transfer scheme to identify residues in the immobile region, and INEPT-TOBSY scalar-based

transfer for residues in the mobile region. CP-RAD/DARR experiment recorded with a 50 ms

mixing time exhibited spectrum with broad peaks, as expected based on the broad resonances in

the 1D CP spectrum (Figure 4.8 A). Due to the poor spectral resolution, we were only able to

perform amino acid-type chemical shift assignments for residues with unique resonant

frequencies that did not overlap with other residues (Ala and Ser, indicated by circles in the 2D

spectrum, Figure 4.8 B). To this end, secondary chemical shift analysis suggested a mixed

population in the core, with one subpopulation of Ser (Ser-A) and Ala (Ala-A) in the fibril core

that had a strong positive chemical shift difference for Cβ and negative chemical shift differences

for CO and Cα, suggesting a β-sheet secondary structure. Chemical shifts for the second

subpopulation (Ser-B, Ala-B and Ala-C) were more ambiguous secondary structure. The lack of

well-resolved peaks prevented site-specific information about the immobile region from being

obtained.

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Figure 4.8. 13C amino acid chemical shift assignment of immobile region in SAA2 fibrils. (A) Aliphatic region

of 2D 13C – 13C CP-RAD/DARR homonclear correlation spectrum, with a closer inspection of the alanine and serine

Cα – Cβ cross peaks. Spectrum was acquired with 50ms RAD/DARR mixing time at 5oC, 10 kHz MAS. (B)

Secondary chemical shift analysis of the alanine and serine populations identified, reported as the difference in

experimental chemical shift value from theoretical random coil values.

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We also acquired 2D 13C – 13C INEPT-TOBSY spectrum to examine the mobile region of SAA

fibrils. A 9.6 ms TOBSY mixing time was used such that 13C cross peaks arise only from ≤ 3-

bond transfers (Figure 4.9 A). With reduced spectral overlap and sharp linewidths relative to the

CP-RAD/DARR spectrum, we were able to carry out 13C chemical shifts assignments for all

amino acid types present in this spectrum. Secondary chemical shift analysis showed that most

amino acids exhibited values similar to random coil, suggesting a disordered structure in the

mobile portion of the SAA fibrils (Figure 4.9 B). 2D 1H – 13C INEPT spectrum also showed that

the mobile region contained residues with aromatic side chains (Figure 4.9 C), however their

backbone chemical shifts could not be identified without acquiring 3D correlation spectra (time-

prohibitive due to poor signal to noise ratios in SAA fibril samples).

Complete sequential assignment of SAA amyloid fibrils has yet to be carried out, and would be

inhibited by broad resonances in the core of the fibril, but unique residues clustered towards the

C-terminus (3 of 4 Pro, 1 of 3 Ile, 1 of 1 Thr) were identified in the 2D 13C – 13C INEPT-TOBSY

spectrum. This suggests that a highly mobile and flexible C-terminus is present in the SAA

amyloid fibril. Meanwhile, unique residue Val57 was not identified in the INEPT-TOBSY

spectrum and likely excluded from the flexible C-terminus. Hence the amino-acid type

assignments of the SSNMR experiments allowed us to hypothesize an initial structural model for

SAA amyloid fibrils containing a β-sheet rich core formed by the N-terminal portion of SAA,

and a flexible C-terminal tail that includes residue Ile65 (Figure 4.9 D).

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Figure 4.9. 13C chemical shift assignment of residues in the mobile region of SAA2 fibrils. (A) Aliphatic region

of the 2D 13C – 13C INEPT-TOBSY homonuclear correlation spectrum with 9.6ms TOBSY mixing time.

Experiments were collected at 5oC, 10 kHz MAS. (B) Secondary chemical shift analysis of the amino acid type

assigned in the 2D spectra. (C) Aromatic region of the 1H – 13C INEPT spectrum, showing correlations arising from

aromatic side chains. (D) Sequence and cartoon representation of proposed fibril architecture, with the mobile

flexible region indicated and residues unique to the C-terminus in bold.

4.3.7 SSNMR of rHDL-SAA and chemical shift assignments

Next, we studied the structure of lipid-associated SAA using MAS SSNMR. We prepared

uniformly-13C, 15N-labelled rHDL-SAA with non-deuterated DMPC lipids, and examined the

local protein dynamics using 1D NMR experiments as described above for SAA fibrils (Figure

4.10). 13C DP spectrum (top) of rHDL-SAA exhibited strong protein peaks across the protein

aliphatic, aromatic, and carbonyl regions, as well as a sharp peak near 70 ppm from PEG3500.

1H – 13C CP spectrum were similar to the 13C DP spectrum, while 1H – 13C INEPT spectrum did

not contain any peaks except those arising from PEG3500 in the buffer. These results suggest

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that SAA is relatively immobile in its lipid-associated state and has a close association with the

rHDL particles.

Figure 4.10. 1D 13C spectra of uniformly 13C, 15N-labelled rHDL-SAA examined by MAS SSNMR. Direct

pulsed (top, black), 1H – 13C cross-polarization (middle, red) and 1H – 13C INEPT spectra (bottom, green) were

recorded under 10 kHz MAS with VT gas set to 5oC. Asterisk denotes the natural abundance 13C signal from

PEG3500.

2D 13C – 13C correlation spectra were recorded using a CP-RAD/DARR transfer scheme with a

short mixing time, such that only intra-residue cross peaks were observed. Initial 13C chemical

shift assignments identified several unique residues (4 of 4 Pro, 3 of 3 Ile, 1 of 1 Thr, and 1 of 1

Val) in the spectrum (Figure 4.11 A). We further collected 2D and 3D 13C – 15N heteronuclear

correlation spectra to perform chemical shift assignments (Figure 4.12 A, B), and resolved 44

distinct residues (~42% of SAA residues), allowing us to determine secondary structure

propensity across many regions in the rHDL-bound form of SAA. The strong propensity for

most identified residues to exhibit α-helical chemical shifts is in agreement with our CD

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measurements, further demonstrating that SAA retained α-helical structure upon lipid

association. Unfortunately, sequential assignment of the full protein was not feasible with our

current datasets due to degenerative chemical shifts and low signal-to-noise in 3D experiments.

Ongoing work seeks to address this with higher sample concentration and selective labeling of

the protein.

Figure 4.11. Preliminary chemical shift assignments of rHDL-SAA2 from 13C – 13C correlation spectrum. (A)

The aliphatic region of 2D 13C – 13C CP-RAD/DARR homonuclear correlation spectrum recorded with a 10 ms

RAD/DARR mixing time at 5oC and 10 kHz MAS. (B) Secondary chemical shift analysis of amino acid types

assigned from 2D CP-RAD/DARR.

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Figure 4.12. Chemical shift assignment from 13C – 15N heteronuclear NCACX spectrum of rHDL-SAA. (A)

2D 13C – 15N NCACX heteronuclear correlation spectrum recorded with a 35 ms RAD/DARR mixing time. (B) 3D

13C – 15N NCACX spectra showing 13C – 13C 2D-slices resolved in the 15N dimension (with corresponding 15N

chemical shift labelled) to show assignment of Gly and Ala with overlapping peaks in either 13C or 15N dimension.

2D and 3D experiments were collected at 5oC and 10 kHz MAS.

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4.3.8 Comparison between experimental and predicted chemical shift of

lipid-bound SAA

In order to address whether lipid-associated and apo-SAA contained similar tertiary structure, we

compared the predicted 13C – 13C correlation spectrum based on the crystal structure of

hexameric apo-SAA to our experimental spectra. To do so, we modelled the SAA2.1 sequence

on the crystal structure of SAA1.1 (PDB ID: 4IB8; 96% homology) using the Phyre2 program

(Kelley et al. 2015) to generate a PDB coordinate file, which was then submitted to ShiftX2

program to predict the backbone chemical shifts (Han et al. 2011). In the overlay of experimental

(13C – 13C CP-RAD/DARR) and simulated spectra shown in Figure 4.13 A, a majority of

residues matched up to the experimental spectrum, as expected from the highly helical structure

of the protein.

In the crystal structure of apo-SAA, Pro49 is located in the hairpin loop between helix 2 and 3,

while Pro92, Pro97, and Pro101 are located in the disordered C-terminal region (Figure 4.13 B).

However, experimental proline chemical shifts clearly deviated from those predicted for the

lipid-free SAA structure, in particular Pro97 and Pro101 (Figure 4.13 C). As a comparison,

proline residues in the ApoA-I helical belt model (2MSE, Figure 4.13 B, C) gave better

alignment to our experimental chemical shifts. These prolines in the ApoA-I structure were

primarily found in kinks or turn regions connecting amphipathic helices, rather than random coil

structure that Pro97 and Pro101 were observed in the lipid-free state.

At the same time, the greater dispersion of the Ala cross peaks observed in the experimental

spectra of rHDL-SAA suggest the presence of distinct local chemical environments for

subgroups of Ala residues that were not present in the lipid-free structure (Figure 4.13 C). These

differences in the Cα – Cβ cross peak regions suggest that while rHDL-SAA2 remained α-

helical, the 4-helix bundle tertiary conformation of apo-SAA may not reflect the lipid-associated

SAA structure. Instead, we hypothesize that HDL-bound SAA adopts an extended helical belt

model much like that proposed for ApoA-I. The ability for SAA to scaffold rHDL particles, and

to replace ApoA-I during acute inflammation give additional rationale to support this hypothesis.

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Figure 4.13. Comparison of experimental 13C spectrum of rHDL-SAA with chemical shifts predicted from the

crystal structure of the SAA1.1 hexamer. (A) Simulated 13C – 13C spectrum of soluble SAA2.1 (threaded to the

structure of SAA1.1 (PDB ID: 4IB8) using Phyre2 overlaid against experimental spectrum of rHDL-SAA collected

in CP-RAD/DARR experiment. 13C chemical shifts of soluble SAA were calculated using ShiftX2 and only the Cα

and Cβ correlations were plotted (green circles). (B) Helix-bundle model of SAA and helical belt model of ApoA-I

(PDB ID: 2MSE) with proline (red) and alanine (cyan) highlighted. (C) Zoomed view the experimental rHDL-SAA

13C – 13C spectrum overlaid with the calculated chemical shifts of proline (red) and alanine (cyan) Cα – Cβ cross

peaks in helix-bundle (left) and helical belt models (right).

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4.4 Discussion

In this chapter, we presented preliminary characterization of the structural properties of SAA

when associated to HDL-like particles (rHDL) and when assembled into amyloid fibrils. Our

cross-linking and DLS data agreed with previous reports of SAA oligomerizing (Wang et al.

2002; Wang et al. 2005; Lu et al. 2014), and suggested that apo-SAA2.1 is predominantly

hexameric with some tendency to form large oligomers or aggregates in solution. Other studies

have shown that apo-SAA can populate multiple oligomeric states, dependent on solution

conditions, consistent with the heterogeneity observed in our data (Wang et al. 2012; Patke et al.

2012). 1H – 15N HSQC spectrum of apo-SAA exhibited significant line-broadening for most

resonances, as expected of a large (>75 kDa) oligomeric protein in solution. Surprisingly, some

resonances remained clearly identifiable in these spectra, and we assigned them as arising from

the flexible C-terminal segment of SAA, in contrast to the previous crystal structure of the apo-

SAA1.1 hexamer with the C-terminus bound to the core 4-helix bundle fold by a bifurcate salt-

bridge (Lu et al. 2014). This may reflect differences in the structure and dynamics for SAA2.1,

as sequence variations between different SAA isoforms are predominantly located in the C-

terminal portion of the protein. This may also reflect functional differences between SAA

isoforms, allowing sequence-dependent differences in protein-protein interactions. Alternatively,

it may simply reflect the different sample conditions in these studies.

SAA2.1 exhibited relatively low thermal stability in the lipid-free (apo) state and was partially

unfolded under conditions of physiological temperature and pH. However, this degree of

destabilization was not sufficient to induce SAA fibrillization under the time-scale of our ThT

kinetics measurements. Further destabilization by acid (pH 3.6), as well as the presence of

amyloid-inducing GAGs, was needed to promote SAA amyloid formation. FTIR spectroscopy of

the fibrils showed a major structural rearrangement from α-helix to β-sheet, further supporting

that acid destabilization and loss of native secondary structure was necessary to induce rapid

conversion of SAA into amyloid fibrils. GAGs heparin and heparan sulfate both significantly

reduced the lag time of SAA2.1 aggregation. It was interesting to note that addition of each GAG

gave rise to significant differences in morphology, where typical thin and smooth fibrils were

formed in the presence of heparin and thicker fibrils with rough edges in the presence of heparan

sulfate. This difference in morphology may result from the different sulfation patterns of heparin

and heparan sulfate, different lengths of sulfated domains in GAGs, different modes of SAA

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binding, or other as yet determined factors (Patke et al. 2012; Aguilera et al. 2014; Takase,

Tanaka, et al. 2014; Takase et al. 2016). Whether different intensities of ThT fluorescence for the

two fibrils reflect differences in morphologies or molecular structure also remain to be

determined.

Heparin-induced SAA fibrils studied by SSNMR were found to contain both immobile (core)

and mobile (C-terminus) segments. Residues 1-11, 1-12, 1-27 in helix 1, and helix 3 were

previously reported to have high propensity to form amyloid fibrils in vitro (Rubin et al. 2010;

Lu et al. 2014; Egashira et al. 2011), and were deduced to form the immobile fibril core. Clinical

AA deposits are also known to contain a protease-resistant fibril core comprised of N-terminal

residues 1-76 (Husebekk et al. 1985). Based on these observations and our amino acid-type

chemical shift assignments, we rationalized an initial model where C-terminal residues 65-104

were flexible and disordered in the amyloid fibril, leaving it potentially exposed for proteolytic

degradation, while the amyloid cross-β core consists of the N-terminal residues 1-64. Structural

heterogeneity seen in our SAA fibrils may relate to the pathogenic nature of SAA fibrils, or may

reflect the denaturing conditions under which the fibrils were formed.

SAA2.1 formed discoidal HDL-like particles (rHDL-SAA) upon associating with DMPC

liposomes, in a manner similar to previous reports on SAA1, and contained predominantly α-

helical secondary structure with higher thermal stability than apo-SAA (Takase, Furuchi, et al.

2014). rHDL-SAA retained its α-helical structure at 37oC and had a much sharper unfolding

transition. Lipid association also provided partial protection from amyloid formation, as the

fibrillization was hindered relative to apo-SAA in our measurements, and in the case with

heparan sulfate, never reached the same fluorescence intensity as apo-SAA. SSNMR data of

rHDL-SAA revealed the whole protein was ordered and relatively immobilized upon lipid

association, including the previously flexible and disordered C-terminus, with chemical shifts

indicative of a highly helical structure throughout. Although this could be an artifact arising from

the PEG precipitation in the SSNMR sample preparation, our spectra showed well-resolved

resonances for all residues, including those in the C-terminus, suggesting a high degree of

structural homogeneity for lipid-associated SAA. A heterogeneous precipitation of the

disordered C-terminus would be expected to generate SSNMR spectra with broadened peaks.

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We compared our experimental 13C – 13C spectrum of rHDL-SAA to the simulated spectra of

two structural models proposed for apolipoproteins, the 4-helix bundle apo-SAA structure and

the helical belt ApoA-I structure. Backbone proline Cα – Cβ chemical shifts of rHDL-SAA

overlaid more closely with the helical belt model. In addition, alanine chemical shifts showed

more dispersion than the 4-helix bundle structure. Together, this strongly suggests the structure

of lipid-associated SAA deviates from that reported in the crystal structure. We proposed that a

conformational change in the tertiary structure could occur during lipid association such that C-

terminus becomes immobilized and interacts with lipids. This ordering of the C-terminus upon

lipid association also gives a potential reason why rHDL-SAA was partially protected from

fibrillization, as the protein became more thermodynamically stable and C-terminus became less

accessible to GAGs.

HDL-association is considered to be crucial to the physiological function of SAA during

inflammation. At the same time, HDL-association may be a stabilizing mechanism to prevent its

amyloid formation during up-regulation. A flexible C-terminus in both the lipid-free and amyloid

state SAA emphasized a link between GAG interaction and SAA fibrillization – supporting

previous work that the C-terminus is important for GAG to recruit misfolded SAA for

fibrillization, but it does not participate in forming the protease-resistant structural core of the

amyloid fibril (Patke et al. 2012; Egashira et al. 2011). Meanwhile, increased thermodynamic

stability and a less accessible C-terminus may prevent HDL-bound SAA from unfolding and

fibrillizing under physiological temperatures. However, the C-terminus must still be able to

exchange on and off from HDLs in order to interact with other cellular receptors involved in the

signalling of inflammation such as cytokine production and release. The potential mechanism for

such conformation exchange has not yet been determined. Our findings emphasize the

importance of the C-terminus of SAA, particularly due to sequence variations between isoforms

being clustered towards the C-terminal portion of the protein, and provide some initial structural

observations that are the basis for ongoing studies of SAA structure and function.

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Chapter 5.

Conclusions and Future

Directions

Contributions:

Dr. Simon Sharpe (Hospital for Sick Children) carried out the Trp fluorescence thermal

denaturation measurements. Undergraduate student Simoun Icho (University of Toronto) carried

out the ThT fibrillization kinetic measurements in different pH buffer conditions.

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Amyloid fibril formation is a protein folding phenomenon in which misfolded proteins self-

assemble into a β-sheet structure more thermodynamically stable than their metastable native

fold. Yet there lacks detailed structural information on amyloids to distinct between features of

functional and pathogenic amyloids. The goal of this thesis was to further study the structures of

amyloid assemblies using SSNMR, and to study the interaction of amyloidogenic proteins with

lipids to examine the effects of membrane on amyloid function or toxicity.

5.1 Structural features of amyloid fibrils

In Chapter 2, we solved the structure of three amyloidogenic peptides PrP(178-183), PrP(244-

249),and PrP(245-250) by SSNMR, allowing me to identify structural features in the three

peptide fibrils that may confer different toxicities. An in-register parallel alignment of β-strands

was observed as the repeating cross-β building block in all three amyloid fibrils. This

arrangement supported Q/N-rich sequences (seen in PrP178-183) forming additional intra-sheet

hydrogen bonds as polar zippers. The stacking of β-sheets in a face-to-face or face-to-back

manner served to maximize the burying hydrophobic side chains (i.e. Ile, Phe, Leu, Val) into a

dehydrated core. Residues that did not face the protofilament core were more likely to be

solvent-exposed, and could dictate fibril interaction with other biological entities that play role in

amyloid toxicity. This work highlights the effects of amino acid composition on the inter-sheet

packing of amyloids, and sheds light on the diverse possibilities of protein assembly in

pathogenic and functional amyloids. Understanding these structural properties of amyloids

would advance the development of engineered nanomaterials and help design amyloids with

biological applications.

5.2 Amyloid fragmentation and membrane interaction

In Chapter 3, we examined the effects of fragmentation on amyloid structure and its interaction

with phospholipid bilayer. We showed through various SSNMR measurements that

fragmentation did not affect the cross-β backbone or the parallel β-sheet arrangement of the three

PrP peptide fibrils. Rather, in specific cases, fragmentation can disrupt the steric zipper packing

in the fibril core. SSNMR and ANS fluorescence data suggested that fragmentation led to a

disrupted fibril packing, greater solvent-exposed surface hydrophobicity, and greater membrane

disruption activity. Non-cytotoxic fibrils, on the other hand, had minimal hydrophobic exposure

and membrane disruption as either mature or fragmented fibrils. Previous studies hypothesized

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that fragmentation-associated amyloid toxicity can be due to an increase in the concentration of

exposed fibril ends, permitting depolymerization and release of small cytotoxic species into

solution. Although our data could not rule out this possibility, our study offers an alternate

possibility to explain fragmentation-associated cytotoxicity. The small fragmented fibril species,

which are accumulated during the early stages of amyloid assembly or generated via fibril

breakage, can exhibit a greater degree of surface-exposed hydrophobicity than mature fibrils.

This exposure increases membrane interaction, lipid disruption, and ultimately leading to

cytotoxicity. The shorter length and greater membrane association of fragmented fibrils may also

allow them to be more readily internalized by cells than mature fibrils. These results point

towards potential differences between functional amyloids and pathogenic amyloids, where

sequence context, surface hydrophobicity, and fibril structure plays a critical role to confer

amyloid cytotoxicity.

5.3 Lipid association of SAA and the effects on structure and

amyloid formation

In chapter 4, we explored the effects of lipid interaction on the biophysical and structural

properties of human SAA2.1. HDL association is crucial to SAA’s physiological function in

dealing with inflammation, yet current data have only reported the structure of lipid-free SAA

and left the structure of the lipid-associated state largely unaddressed. SAA forming discoidal

HDL-like particles with DMPC provided a unique basis to begin examining the properties of

lipid-associated SAA. CD spectra collected here and by others indicate SAA retained its α-

helical secondary structure after lipid association, with higher thermal stability comparable to

other apolipoproteins SAA1 and ApoA-I. Using solution and solid-state NMR, we also showed

that lipid association corresponded to an ordering and decreased mobility of SAA C-terminus.

NMR chemical shift analysis supported rHDL-SAA retaining an α-helical secondary structure,

but backbone proline chemical shifts suggested a tertiary structure similar to an extended helical-

belt structure as ApoA-I rather than maintaining the 4-helix bundle structure. A possible

structural rearrangement of the helices in SAA would accommodate interactions with lipids and

form discoidal particles akin other apolipoproteins.

It has been reported that rHDL-SAA was more resistant to PK digest than apo-SAA, providing

evidence for decreased accessibility of the protein associated to the particle (Takase, Furuchi, et

al. 2014). A reduced mobility of the C-terminus in the helical belt model would decrease

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accessibility to GAGs and hence slow down fibrillization. Our observation may explain why AA

amyloidosis is not prominently observed in patients suffering from chronic inflammatory

disorders, as HDL-association can increase the thermodynamic stability to prevent SAA from

unfolding under physiological temperatures and prevent fibrillization. Nonetheless, we suspect

that the C-terminus must still be able to exchange on and off from HDLs in order to interact with

other cellular proteins for its function.

5.4 Initial structural model of amyloid SAA by SSNMR

To understand the mechanism of SAA fibrillization, we also investigated SAA amyloids by

biophysical and NMR techniques. Although apo-SAA was marginally stable under physiological

temperature and pH, it was not sufficient to trigger amyloid formation. A decrease in α-helical

secondary structure by acid destabilization and the presence of GAGs was necessary to trigger

SAA fibrillization into β-sheet amyloids. A closer examination of apo-SAA by solution NMR

identified a flexible and dynamic C-terminus for the oligomeric SAA in solution, a major

difference from the crystal structure reported previously. Our initial structural model of SAA

amyloids identified N-terminal residues 1-64 as the β-sheet core, while residues 65-104 remains

flexible and disordered. The C-terminus remaining disordered in SAA amyloids hinted towards

its role in dictating the kinetics of amyloid formation rather than a structural role in the amyloid

assembly. GAG-induced fibrillization likely required C-terminus to scaffold and recruit soluble

SAA close together, while only the N-terminus of SAA is assembled into the β-sheet fibril core

and becomes resistant to proteolysis.

5.5 Future directions

5.5.1 Stability and fibillization kinetics of lipid-free and lipid-bound SAA

as an effect of pH

Structural rearrangement of the native protein into a β-sheet rich nucleus is often the rate limiting

step in amyloid formation, and thus the structure and thermal stability of the protein influences

the time for nucleation. Our ThT kinetics indicated that low pH was necessary for fibril

formation of the apo- and lipid-associated forms of SAA, as was the presence of GAGs (heparin

or heparin sulfate). We also observed that lipid association of SAA increased the lag phase and

slowed its fibrillization. Understanding if and how lipid association can affect the thermal

stability of SAA under different pH conditions, and the effect of lipid composition on SAA

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stability, will help us further understand the protective role of HDL-association on hindering

SAA amyloid formation.

To test this, we can perform CD spectroscopy to monitor SAA secondary structure and thermal

denaturation across a wide range of pH conditions (i.e. pH 3.5 – 7.5 with incrementing pH units).

Preliminary thermal denaturation measurements using Trp fluorescence of apo-SAA suggested a

pH-dependent unfolding of apo-SAA (Table 5.1). We also performed ThT fibrillization kinetics

of apo-SAA under varying pH in the presence of heparin at 37 oC, and observed a pH-dependent

fibrillization in buffers below pH 5.5 (Figure 5.1). This dependence could be due to the

protonation of histidine imidazole side chain, which affects the unfolding of SAA as well as

binding to the anionic GAGs. These results matched previous reports that heparan sulfate

promoted SAA aggregation from HDL particles under mildly acidic conditions (pH 5.5)

(Elimova et al. 2009) Similar thermal denaturation and ThT fibrillization kinetic measurements

with varying pH can be performed on lipid-bound SAA prepared with DMPC vesicles. Follow

up experiments with varying lipid compositions (PC, PS, and/or PE), acyl tail lengths (16:0,

16:1, 18:0, 18:1, 20:4), and possibly using lipid mixtures with cholesterol to more closely mimic

the composition of HDL particles, may further examine the effects of lipid –SAA interaction.

Table 5.1. Thermal stability of lipid-free SAA2 as a function of solution pH.

pH TM of apo-SAA

7.5 39.9 ± 0.6

6.5 34.6 ± 0.2

5.5 31.4 ± 0.9

4.5 24.4 ± 3.2

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Figure 5.1. Fibrllization of apo-SAA in the presence of heparin in different pH buffers. pH-dependent

fibrillization was observed for apo-SAA in the presence of heparin as detected by ThT-binding, but only in buffer

conditions below pH 5.5. ThT measurements were performed in triplicates at 37oC with 0.3 mg/mL SAA, 0.1

mg/mL heparin, and 8 μM ThT.

5.5.2 SSNMR studies of the SAA fibril core and structural differences

between fibrils formed under different solution conditions

SAA1 and SAA2 are both deposited as amyloid fibrils in AA amyloidosis and have 100%

sequence identity from residues 1-50. Correspondingly, the N-terminus of the protein has been

alluded to be protease-resistant in AA patients, and matched our initial NMR data that residues

1-64 formed the β-sheet core. Solving the high resolution structure of the fibril core will provide

invaluable insight into the misfolding of SAA that leads to AA amyloidosis. Although TEM

micrographs of heparin-induced fibrils formed at pH 3.6 appeared morphologically homogenous,

our SSNMR spectra contained broadened peaks to indicate structural heterogeneity at an

atomistic level. As a result, sequential assignment of NMR spectra arising from the fibril core

was not possible. We hypothesized this was likely due to SAA being almost completely unfolded

at pH 3.6. While this allowed for rapid fibrillization, it could also lead to structurally

heterogeneous intermediates being assembled into fibrils. A closer inspection of SAA stability

under varying pH (as described in Section 5.5.1), and subsequently preparing fibrils at higher pH

with less destabilization, may allow us to produce structurally homogeneous fibrils suitable for

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SSNMR studies. However, we cannot rule out the possibility that pathogenic fibrils of this type

may always exhibit far more heterogeneity than observed for functional amyloid fibrils like the

Het-S yeast prion, which gives rise to extremely well resolved SSNMR spectra (Wasmer et al.

2008; Van Melckebeke et al. 2010)

Following the optimization of fibrillization, we can perform 2D experiments (i.e. CP-

RAD/DARR) and 3D backbone transfer experiments (i.e. NCACX and NCOCX) to obtain

complete backbone chemical shift assignments. These assignments can be used to calculate

secondary structure propensity and backbone torsion angles (Cornilescu et al. 1999). Further

distance measurements may require sparse isotope labelling schemes to reduce spectral

complexity. Non-uniform 13C-enrichment can be achieved by growing bacteria using [1,3-13C]-

glycerol and [2-13C]-glycerol as the carbon source instead of uniformly 13C-labelled glucose

(Castellani et al. 2002), and results in a complementary partial labelling of amino acids that can

be useful for NMR experiments. In particular, well-resolved long-range 13C – 13C distance

measurements can be collected on partially labeled samples using RAD/DARR or PAR

experiments with long polarization mixing times. Additional constraints can be obtained using

the ZF-TEDOR experiment for 13C – 15N distance measurements. We can also determine the

inter-strand distance of the cross-β backbone using the dipolar recoupling experiment CT-

PITHIRDS as we did in chapter 2 and 3. Here, we will specifically label the 13C-carbonyl of

amino acids with low occurrences in the sequence (i.e. 1 valine, 3 isoleucine, 2 glutamine) to get

site-specific distance constraints, similar to previous reports for mouse PrP(90-231) and human

Aβ40 fibrils (Groveman et al. 2014; Lu et al. 2013). Using these techniques, it will be possible to

obtain a high-resolution structure of SAA amyloid fibrils, assuming well-ordered samples can be

prepared.

5.5.3 The effects of GAGs and lipids on SAA amyloid fibril morphology

and structure

We observed in Chapter 4 that GAGs promoted SAA fibril formation with different

morphologies and ThT-binding properties. As such, sulfation pattern of heparin and heparan

sulfate could play a role in recruiting different folding intermediates or promote conformations

with different exposed surfaces for fibril formation, resulting in different fibril morphologies and

biological properties. To address this, we can examine the morphology by AFM and TEM, or

possibly collect micrographs using electron cryomicroscopy (cryo-EM) for single fibril

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averaging. We can also perform mass-per-length calculations on these fibrils based on dark-field

TEM, a technique broadly used to study fibril polymorphism of Aβ40 amyloid fibrils and other

amyloid systems at the molecular level (Chen et al. 2009; Petkova et al. 2005). Subsequently,

SSNMR can provide site-specific information of the local structure in these fibrils to report on

structural differences between heparin-induced and heparin sulfate-induced fibrils. Similar NMR

techniques employed in Chapter 4 of this thesis can be used to explore these properties, such as

2D CP-RAD/DARR or INEPT-TOBSY experiments to look for mobile and immobile regions in

the fibrils.

Equally interesting is to examine differences in fibril structure generated with apo-SAA and

lipid-associated SAA. Our ThT kinetics support that rHDL-SAA is capable of fibrillizing, and it

has been recently shown that GAGs can promote SAA dissociation from HDL particles at low

pH and induce fibrillization (Noborn et al. 2012). It is unclear whether the acutely upregulated

HDL-bound SAA circulating in the body during inflammation or apo-SAA from extrahepatic

tissues is the source of misfolded SAA. It was recently shown that mouse cells cultures incubated

with recombinant full-length apo-SAA1.1 and HDL formed cytotoxic extracellular deposits that

exhibited multiple fibril bundle morphologies, and contained GAG, serum amyloid P, and

vesicular lipid inclusions (Kollmer et al. 2016). Comparing the structure of SAA fibrils

generated from lipid-free and lipid-associated states by FTIR, AFM, cryo-EM, and SSNMR,

could reveal possible differences in SAA amyloid structure to reflect the effects of membrane

interaction on SAA assembly.

5.5.4 Structure of lipid-associated SAA

We have collected 3D backbone 13C – 15N experiments NCACX and NCOCX for chemical shift

assignment of rHDL-SAA, yet degenerative sequences from repeating residues (Ala, Arg, Gly,

Ser) has made sequential assignments more difficult to complete with the current dataset. To

complete the chemical shift assignments of rHDL-SAA, we need to perform additional 3D

backbone transfer experiments CONCA and CANCOCX that transfers more selectively to

generate cross peaks that correlate COi-1 – N – Cαi and Cαi – N – Ci-1, respectively (Shi et al.

2009). These two experiments will provide unambiguous spectra that will allow us to complete

the sequential chemical shift assignments and begin to elucidate the structure of lipid-associated

SAA. Alternatively, selective labelling as described above for the fibrils may be required where

needed to reduce spectral complexity.

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Extensive studies on ApoA-I can lend insight into the structural arrangements of lipoproteins on

HDLs, and allow us to propose testable hypotheses regarding the structure of HDL-bound SAA.

The extended double helical belt model proposed for ApoA-I allows the protein to wrap around

the HDL particle, and SAA may adopt a similar tertiary structure when associated to HDLs (Li et

al. 2006). To further discern between the 4-helix bundled structure of apo-SAA from the helical

belt model inferred from our proline chemical shift comparison, we propose to measure the

solvent-exposure of the protein by SSNMR experiments. We hypothesize that the folded 4-helix

bundle will have residues buried into the hydrophobic core and more shielded from water

exposure, while the extended helix model will have more residues and backbone exposed. We

can measure this difference in water exposure by performing polarization transfer from H2O to

protein and examine the distance between them. Alternatively, 1H polarization transfer from lipid

acyl chains to protein could identify residues on SAA that are interacting with the lipid tails.

These approaches have been used to examine depth of insertion of membrane peptides as well as

solvent exposure of pore-forming membrane proteins (Williams & Hong 2014). Modification of

the pulse sequence such that the polarization transfer time is optimized for short-range transfer

may provide us with further distance constraints to build a structural model for the HDL-

associated SAA.

Ultimately, our goal would be to examine SAA structure when bound to HDL particles in order

to determine its biological function. Advances in microfluidics to generate spherical micrometer-

sized particles has promising biological applications such as for drug delivery, particle

diagnostics, and cell encapsulation (Dendukuri & Doyle 2009). The advantage of microfluidic

technology is its ability to control size, shape, internal structure, and functionality of the

synthesized particle, and has been used synthesize lipid droplets such as liposomes and lipid

nanoparticles. It has been shown recently that the technology exists to synthesize HDL-

mimicking nanoparticles (μHDL) with same size, morphology, and biological activity as native

HDLs (Kim et al. 2013; Luthi et al. 2010). This one-step synthesis method of manufacturing

homogeneous preparations of μHDL can be used to study structure and function of SAA when

associated to more biologically relevant HDL particles.

5.5.5 Functional diversity of SAA

The pressing question about SAA is the role of this highly conserved acute-phase apolipoprotein

in modulating the immune system to deal with inflammation, and how it exerts its

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immunological activity as it circulates the body associated to HDLs. SAA has been reported to

interact with various cell-surface receptors, and induce the production of pro-inflammatory

cytokines such as IL-1, IL-6, IL-8, TNF-α or anti-inflammatory cytokines such as IL-10 (Figure

5.2). For example, class B scavenger receptors such as CD36, CD36-LIMPII analogous 1 (CLA-

1), and scavenger receptor B1 (SR-B1), are implicated in lipid transport and can bind to lipid-

free SAA and HDL-associated SAA to cause cellular uptake of SAA (Marsche et al. 2007). SAA

binding to CD36 and CLA-1 also results in the secretion of cytokines IL-6 and IL-8 in HEK293

cells (Marsche et al. 2007; Baranova et al. 2010). Toll-like receptor 2 (TLR2) and TLR4 are

functional receptors for acute-phase SAA that stimulate expression of anti-inflammatory

cytokines IL-10 and IL-1R antagonist (Cheng et al. 2008). In addition, SAA can participate in

the innate immune response by promoting the production of pro-IL-1β and inducing its

maturation to IL-1β by activating the NLRP3 inflammasome via the ATP-receptor P2X7 (Eklund

et al. 2012). Finally, SAA binds to the formyl peptide receptor like-1 (FPRL1) receptor found on

cell surface to mediate migration of neutrophils, or induce the secretion of cytokines IL-10 and

TNF-α in monocytes (Shao et al. 1999).

The multiple interaction partners of SAA, and therefore its diverse biological function, make it

an interesting target for structural studies. The C-terminus of SAA being disordered may be an

important property for its function similar to intrinsically disordered proteins that have different

interacting partners. The heavily conserved prolines in the C-terminus of all SAA isoforms are

important for maintaining its disordered structure. At the same time, sequence variation across

the acute-phase isoforms (SAA1.1 – 1.5, SAA2.1 – 2.2), located in the C-terminal portion of the

protein (residues 52-104), further point to the possible functional diversity of the C-terminus

(Figure 1.8 A). In fact, SAA1 isoforms have been shown to have different selectivity for these

receptors to reflect sequence specificity in the functional and structural diversity of SAA proteins

(Chen et al. 2014).

130

Figure 5.2. Cell surface receptors that mediate the functions of SAA in the immune system. Effector pathways

that are activated as a result of SAA upregulation during inflammation, and sorted by the receptor binding event that

initiates the pathway. IL - interleukin; NF - nuclear factor; NO - nitric oxide; TNF - tumor necrosis factor; NLRP3 –

Nucleotide-binding, domain leucine-rich repeat-containing family, pyrin domain containing 3. Modified from

Eklund et al. 2012.

Our results indicate that the C-terminus of SAA is mostly disordered in the lipid-free state, and

that it has reduced mobility and increased secondary structure content in the lipid-bound state.

Whether this confers structural and functional differences in a way that is sequence-dependent

across different SAA isoforms is yet to be examined. Building on our current structural data, it

may be important to examine the interactions of SAA C-terminus with various receptors. We can

perform a systematic docking search for peptide-receptor binding sites using programs such as

CABS-dock (Blaszczyk et al. 2016) or PepFlexDock (Raveh et al. 2010). Follow this, we can use

recombinant SAA in its apo- and lipid-associated state to examine their interaction with receptor

ectodomains or peptides by biophysical techniques such as isothermal calorimitry (ITC), surface

plasmon resonance (SPR), or solution NMR experiments.

131

5.6 Final concluding remarks

The work presented in this thesis showed that membrane interaction plays a significant role in

the biology of amyloids. The knowledge gained from examining the structural features of short

PrP peptide fibrils strongly hinted towards a correlation between surface exposure of

hydrophobicity, membrane disruption, and amyloid toxicity. Equally important, fibril

fragmentation may play a role in in vivo cytotoxicity of amyloid diseases. Studies on lipid

interaction of human SAA provided important structural differences between lipid-associated

and lipid-free SAA in the context of its biological function. This work represents a small fraction

of our understanding of the complex nature of amyloid formation and toxicity, but it is

informative in furthering our knowledge about the biology of amyloids.

132

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Appendices

Table A 1. 13C and 15N chemical shifts from PrP(244-249) fibrils.

Chain A

Residue NH C=O Cα Cβ

Ile244 38.47 170.800 57.970 37.590

γCH2 25.620 δCH3 -

γCH3 15.920

Ser245 121.4 169.730 56.230 66.360

Phe246 122.57 171.360 50.610 39.850

1C 136.350 2, 6CH -

3,5CH - 4CH -

Leu247 126.77 172.450 53.460 42.870

γCH2 28.430 δCH3 25.100

δCH3 22.500

Ile248 118.36 172.990 57.680 38.380

γCH2 25.850 δCH3 13.290

γCH3 15.920

Phe249 123.31 178.710 54.320 34.680

1C - 2, 6CH -

3,5CH - 4CH -

Chain B


Ile244 35.59 170.180 57.850 36.780

γCH2 25.620 δCH3 -

γCH3 16.210

Ser245 119.1 170.120 54.860 63.100

Phe246 119.53 171.910 54.680 40.830

1C 137.400 2, 6CH -

3,5CH - 4CH -

Leu247 125.37 172.450 53.038 41.020

γCH2 26.830 δCH3 25.440

δCH3 22.800

Ile248 122.59 173.460 57.890 38.980

γCH2 25.430 δCH3 11.990

γCH3 15.000

Phe249 126.38 179.900 58.370 37.630

1C - 2, 6CH -

3,5CH - 4CH -

158



Ser245 33.82 168.956 55.650 62.866

Phe246 113.83 172.095 53.930 37.600

1C 137.850 2, 6CH 132.040

3,5CH 130.160 4CH 128.12

Leu247 115.86 172.090 53.070 37.420

γCH2 25.510 δCH3 24.680

δCH3 23.280

Ile248 117.73 171.390 55.500 35.560

γCH2 25.160 δCH3 10.150

γCH3 15.796

Phe249 123.19 171.74 53.9 38.15

1C 137.200 2, 6CH 129.060

3,5CH 127.180 4CH 125.62

Leu250 127.82 180.66 54.84 40.7

γCH2 28.290 δCH3 24.930

δCH3 23.850



Asp178 36.21 168.956 55.650 62.866

γCO 175.910

Cys179 118.74 170.920 55.280 28.590

Val180 120.74 172.160 58.200 35.350

γCH3 19.100 γCH3 17.790

Asn181 125.78 171.950 50.860 39.700

γCO 171.320 γNH2 115.420

Ile182 121.5 172.320 57.490 40.530

γCH2 25.970 δCH3 13.31

γCH3 14.25

Thr183 118.66 175.760 58.220 68.410

γCH3 19.750

tspace.library.utoronto.ca€¦ · ii Structural characterization of amyloid assemblies and amyloid...

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