Extracellular Chaperones and Their Cell Surface Receptors

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2020

Extracellular Chaperones and their Cell Surface Receptors

Jennifer Renee West

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Extracellular Chaperones and their Cell Surface Receptors

By Jennifer Renee West Bachelor of Arts (Biological Sciences) – Hanover College Bachelor of Science (Honours) – University of Wollongong

Supervised By Senior Professor Mark Wilson

Co-supervisor Associate Professor Heath Ecroyd

This thesis is presented as part of the requirements for the degree of

Doctor of Philosophy

School of Chemistry & Molecular Bioscience And The Illawarra Health & Medical Research Institute

University of Wollongong Wollongong, Australia June 2020

Declaration of Authority

This thesis is submitted in accordance with the regulations of the University of Wollongong in partial fulfilment of the Degree of Doctor of Philosophy. It does not include any material published by another person except when due reference is made in the text. The experimental work described in this thesis is original and has not been submitted for a degree to any other university. Some experimental work conducted for this project was completed with the assistance of a facilitator, provided by the University of Wollongong.

Jennifer Renee West

Acknowledgments

First and foremost, thank you to Mark for being not only an excellent supervisor, but also a patient teacher/mentor and someone who genuinely cares about your success as a student. I am so grateful to have had the opportunity to learn from and work along-side such an intelligent and inspiring person. Mark – also, thank you for pulling me out of my Debbie-downer thinking and providing many delicious baked goods! I will miss working in your lab very much!

Facilitators: Daniel Whitten, Luke McAlary, Rebecca Brown, Kate Roberts, Rafaa Zeineddine, Jay Perry, & Sandeep Satapathy – thank you all so very much for dedicating your valuable time in assisting me with my experimental work – I literally could not have done this without you guys!

Megan Kelly – thank you for all of your help with protein aggregation and cytotoxicity assays, and also your company, encouragement, and conversations in the lab.

Lab 210 colleagues – thank you all for making my journey at UOW both memorable and fun. Thank you all for your advice, recommendations, and overall friendship. I will miss all of you very much!

IHMRI Staff – thank you for making IHMRI such a safe, well-organised, and fantastic environment to work in.

A huge thank you to Tom, for putting up with me when I was stressed and at times discouraged throughout this monstrous project. Thank you to my parents, Tom’s parents, and all of my extended family for your support and words of encouragement – it really meant a lot. I love you all!

This research has been conducted with the combined support of the Australian Government Research Training Program Scholarship and a scholarship received from Samsung, to which I am very thankful to have received.

Abstract

Nearly all biological systems require proteins to achieve and maintain their native, three- dimensional conformation, however, various stresses and genetic mutations can cause proteins to misfold and aggregate. Both intracellular and extracellular protein aggregates are associated with protein deposition diseases (PDDs), such as Alzheimer’s disease (AD), Parkinson’s disease (PD), and amyotrophic lateral sclerosis (ALS). Although the processes involved in maintaining intracellular protein quality control are well established, much less is known regarding the maintenance of extracellular proteostasis.

Extracellular chaperones (ECs) are a unique class of proteins that have been shown to inhibit the misfolding and aggregation of various client proteins in vitro by forming stable, soluble high molecular weight (HMW) complexes with them. Furthermore, ECs direct aggregates to (yet unknown) cell surface receptors, which internalise the complex by receptor-mediated endocytosis for degradation within lysosomes. Using in vitro protein aggregation assays, this study identified neuroserpin (NS), transthyretin (TTR), and thyroglobulin (Tg) as new ECs, with NS and TTR being the first ECs classified as amyloid-specific. Though less efficient compared to clusterin (CLU), NS did partially inhibit the aggregation of citrate synthase (CS), an amorphously- aggregating client protein, but had no inhibitory effect on the amorphous aggregation of chloride intracellular channel 1 (Clic1). Neither native tetrameric TTR (hTTR) nor a monomeric TTR mutant (mTTR) specifically inhibited the amorphous aggregation of bovine serum albumin (BSA), creatine phosphokinase (CPK), or CS. In contrast, NS and mTTR were efficient inhibitors of amyloid formation by both coiled-coil beta peptide (ccbw) and alpha-synuclein (a-syn); hTTR specifically inhibited a-syn aggregation only. Sequence comparisons between NS and TTR identified a contiguous 14-residue region in both proteins that is 64% identical and 71% similar. The majority (57% in NS and 71% in TTR) of the residues within this region corresponded to an a-helix in both NS and TTR and, in the case of TTR, residues within this helix are known to be involved in the binding of beta-amyloid (Ab) to TTR. Synthetic peptides were generated corresponding to the conserved sequences incorporating the NS and TTR a-helices for use in in vitro protein aggregation assays. Rather than having an inhibitory effect, the peptide corresponding to the NS sequence increased amyloid formation by ccbw in a concentration-dependent manner, suggesting that the peptide may be interacting with ccbw amyloid species.

Tg appears to have a more broad-spectrum chaperone action similar to that of CLU and alpha-2- macroglobulin (a2M). Tg concentration-dependently inhibited amorphous protein aggregation by

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CS and ovotransferrin (OT) and amyloid formation by Ab and calcitonin (Cal). Like CLU, Tg forms stable, soluble HMW complexes with CS, which have an average particle size of 82.5 nm according to DLS analyses. The Tg-CS complexes were significantly larger than both Tg (average particle size of 45.6 nm) and CS (average particle size of 8.5 nm). It was also shown that the Tg dimer dissociates into monomers when heated to 60°C for 5 min, and that heat-treated Tg has a significantly higher surface hydrophobicity compared to untreated Tg.

Attempts were made to identify the cell surface receptors involved in the endocytosis of EC-client complexes, however, these initial experiments used biotinylated EC-client complexes. Subsequent results showed that biotinylation of HMW CLU-client complexes induced structural changes leading to differences in (i) size (increased), (ii) their ability to bind to cells, (iii) dissociation of the complex into its constituent individual proteins under non-reducing and reducing conditions and (iv) antibody recognition of the complex. The work presented here demonstrates that the cell lines HepG2 and EOC13.31 and two receptors (apolipoprotein E receptor 2 (ApoER2) and very low-density lipoprotein receptor (VLDLR)) may bind CLU-GST complexes specifically, however, further testing would be required to definitively establish this.

Collectively, the work presented here suggests that NS, TTR, and Tg likely have important protective roles as ECs in vivo. Determining whether the conserved a-helix shared between NS and TTR plays a role in the proteins’ chaperone action may lead to a better understanding of the structure or biophysical properties required for amyloid-interaction and lead to the future development of therapies for diseases associated with amyloid deposition. Like CLU, Tg appears to have a promiscuous chaperone activity and can form soluble HMW complexes with amorphously-aggregating client proteins. However, whether Tg plays a role in the cellular uptake of Tg-client complexes for intracellular degradation remains unknown. Future work is required to (i) identify the particular biophysical properties and/or binding site on EC-client complexes that mediate their uptake and (ii) the specific receptors involved in the internalisation of EC-client complexes.

Table of Contents

Declaration of Authority…………………………………………………………………………...i Acknowledgments………………………………………………………………………………...ii Abstract…………………………………………………………………………………………...iii Table of Contents………………………………………………………………………………….v List of Figures…………………………………………………………………………………...viii List of Tables……………………………………………………………………………………...x Abbreviation……………………………………………………………………………………...xi Chapter 1: Introduction……………………………………………………………………………1 1.1 Protein structure, function, and folding…………………………………………….....1 1.2 Protein unfolding, misfolding and aggregation……………………………………….4 1.3 Protein aggregates, cytotoxicity, and disease………………………………………....8 1.4 Intracellular protein quality control………………………………………………….10 1.4.1 Molecular chaperones……………………………………………………...13 1.4.2 Degradation of proteins via the UPS………………………………………14 1.4.3 Lysosomal degradation pathways………………………………………….15 1.5 Extracellular protein quality control…………………………………………………17 1.5.1 Extracellular proteolytic degradation………………………………………18 1.5.2 Known ECs………………………………………………………………...18 1.6 New candidate ECs under investigation……………………………………………..22 1.6.1 Thyroglobulin……………………………………………………………...22 1.6.2 Neuroserpin………………………………………………………………...26 1.6.3 Transthyretin……………………………………………………………….30 1.7 A model for the clearance of extracellular protein aggregates………………………33 1.8 Experimental objectives……………………………………………………………...36 Chapter 2: General Materials & Methods………………………………………………..………37 2.1 Materials……………………………………………………………………………..37 2.2 General methods……………………………………………………………………..38 2.2.1 Estimation of protein concentration……………………………………….38 2.2.2 SDS-PAGE analysis……………………………………………………….38 2.2.3 Western blot analysis………………………………………………………39 2.2.4 Dot blot analysis…………………………………………………………...39 2.2.5 Protein aggregation assays…………………………………………………39 2.2.6 Mammalian cell culture……………………………………………………40 2.2.7 CLU purification…………………………………………………………...42 2.2.8 Purification and restriction digestion of the pGEX-2T(GST-RAP) plasmid for the purification of glutathione-S-transferase (GST) and receptor-associated protein (RAP)……..…………………………………..42 2.2.9 Transformation, expression, and purification of recombinant Schistosoma japonicum GST-tagged human RAP…………………....…..43 2.2.10 Purification of recombinant Clic1……………………………………..….44 2.2.11 Expression and purification of recombinant NS………………………….45 Chapter 3: NS and TTR: Potential New ECs…………………………………………………….46 3.1 Introduction…………………………………………………………………………..46 3.1.1 NS………………………………………………………………………….46 3.1.2 TTR………………………………………………………………………...47 3.1.3 NS and TTR are putative ECs……………………………………………..49 3.2 Materials……………………………………………………………………………..49 3.3 Methods……………………………………………………………………………...49 3.3.1 Optimised expression and purification of recombinant NS………………..49 3.3.2 Immunoblotting of recombinant NS……………………………………….51 v

3.3.3 Characterisation of recombinant NS structure and activity………………..52 3.3.4 Protein aggregation assays…………………………………………………52 3.3.5 Bioinformatic analysis of NS and TTR sequences………………………...55 3.3.6 Synthetic peptides corresponding to NS residues 300 – 313 in aggregation assays…………………………………………………………56 3.4 Results……………………………………………………………………………….56 3.4.1 A more efficient method for the expression and purification of recombinant NS……………………………………………………………56 3.4.2 Structural and functional characterisation of recombinant NS…………….58 3.4.3 The effects of NS in protein aggregation assays…………………………...61 3.4.4 The effects of hTTR and mTTR in protein aggregation assays……………65 3.4.5 NS and TTR share sequence homology……………………………………75 3.4.6 PNS does not inhibit the formation of ccbw amyloid……………………..83 3.5 Discussion……………………………………………………………………………85 Chapter 4: Tg is a Newly Identified EC………………………………………………..………..97 4.1 Introduction…………………………………………………………………………..97 4.2 Materials……………………………………………………………………………..98 4.3 Methods……………………………………………………………………………...98 4.3.1 Tg in protein aggregation assays…………………………………………..98 4.3.2 Investigating Tg structure using native PAGE…………………………...100 4.3.3 Measuring regions of solvent-exposed hydrophobicity on Tg with bisANS……………………………………………………………………100 4.3.4 Tg-client protein complexes……………………………………………...101 4.3.5 Cytotoxicity assays……………………………………………………….105 4.3.6 Bioinformatic analysis comparing Tg and other known ECs…………….106 4.4 Results………………………………………………………………………………107 4.4.1 Tg inhibits the aggregation of a wide range of proteins………………….107 4.4.2 Thermal stress induced structural changes in the Tg molecule…………..112 4.4.3 Thermal stress increased surface hydrophobicity of both bTg and hTg…………………………………………………………………..113 4.4.4 bTg forms stable, soluble HMW complexes with CS……………………114 4.4.5 Assessing cytotoxicity of Cal and Ab…………………………………….119 4.4.6 Tg does not share sequence homology with other ECs…………………..124 4.5 Discussion…………………………………………………………………………..124 Chapter 5: Investigating the Binding and Uptake of HMW Chaperone-client Complexes by Cell Surface Receptors……………..…………………………………………...132 5.1 Introduction…………………………………………………………………..……..132 5.2 Materials………………………………………………………………………..…..134 5.3 Methods…………………………………………………………………………….134 5.3.1 Mammalian cell culture…………………………………………………..134 5.3.2 Expression and purification of GST-tagged RAP………………………...135 5.3.3 Formation of HMW chaperone-client complexes………………………..135 5.3.4 Biotinylation of HMW complexes and control proteins………………….135 5.3.5 Transient transfection of HEK293 cells…………………………………..136 5.3.6 Cell binding assays using biotinylated HMW chaperone-client complexes……………………………………………………….………..136 5.3.7 Inhibition of HMW chaperone-client complexes binding to cell surface receptors………………………………………………………….137 5.3.8 Affinity adsorption to identify cell surface receptors for HMW CLU-CS complexes………………………………………………………137 5.3.9 Analysis of non-biotinylated and biotinylated HMW chaperone- client complexes…………………………………………………………141 vi

5.3.9.1 SDS-PAGE analyses…………………………………………...141 5.3.9.2 Dot blot analyses………………………………………………..141 5.3.9.3 NTA analyses…………………………………………….……..142 5.3.9.4 Binding to RAW 264.7 cells……………………………………142 5.3.10 Cell binding assays using non-biotinylated HMW chaperone-client complexes………………………………………………………..……..142 5.3.11 Flow cytometry analyses………………………………………………..143 5.4 Results………………………………………………………………………………143 5.4.1 Neither biotinylated CLU-CS nor biotinylated Tg-CS bind to SRA-1 or SREC-1 transfected HEK293 cells………………………………….…143 5.4.2 Biotinylated chaperone-client complexes bind to the surface of RAW 264.7 macrophages………………………………………………..145 5.4.3 Biotinylated CLU-GST and biotinylated Tg-CS bind to EOC13.31 microglial cells……………………………………………………………147 5.4.4 Biotinylated CLU-GST, but not biotinylated Tg-CS, binds to HepG2 liver cells…………………………………………………………………148 5.4.5 Anti-CS recognised and captured CLU-CS complexes…………………..149 5.4.6 Anti-CS affinity adsorption assays did not capture receptor-bound CLU-CS………………………………………………………………….150 5.4.7 The biotinylation of HMW chaperone-client complexes modifies their structure…………………………………………………………….154 5.4.8 Non-biotinylated Tg-CS and CLU-GST complexes do not bind specifically to RAW 264.7 macrophages………………………………..163 5.4.9 Non-biotinylated CLU-GST complexes bind to the surface of EOC13.31 cells…………………………………………………………..166 5.4.10 Non-biotinylated CLU-GST complexes do not bind to the surface of HepG2 liver cells……………………………………………………..167 5.4.11 Non-biotinylated CLU-GST complexes bind to transfected HEK293 cells expressing ApoER2 and VLDLR………………………………….169 5.5 Discussion…………………………………………………………………………..172 Chapter 6: Conclusions…………………………………………………………………………180 Chapter 7: References…………………………………………………………………………..190 Appendix I……………………………………………………………………………………...224 Appendix II……………………………………………………………………………………..228 Appendix III…………………………………………………………………………………….229

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List of Figures

Figure 1.1 The protein folding funnel 4 Figure 1.2 Formation of protein aggregates 6 Figure 1.3 Model of amyloid formation 7 Figure 1.4 Scanning electron microscopy image of amorphous aggregates 7 Figure 1.5 An integrated network of intracellular protein quality control mechanisms 11 Figure 1.6 Degradation of proteins through the UPS 15 Figure 1.7 Schematic representation of CMA 17 Figure 1.8 Thyroid hormone synthesis 24 Figure 1.9 Ribbon representation of NS structure 27 Figure 1.10 Ribbon representation of cleaved NS 28 Figure 1.11 Ribbon representation of homotetrameric TTR structure 31 Figure 3.1 Plasmid map of pQE-81L(NS) 50 Figure 3.2 Imidazole elution of NS from a HisTrap column 57 Figure 3.3 Immunoblotting of purified NS 58 Figure 3.4 Native PAGE analysis of heated and unheated NS 59 Figure 3.5 Detection of covalently linked NS-tPA complexes 60 Figure 3.6 NS is an inefficient inhibitor of amorphous CS and Clic1 aggregation 62 Figure 3.7 NS inhibits amyloid formation by ccbw and a-syn 64 Figure 3.8 Effects of hTTR on the DTT-induced aggregation of BSA 65 Figure 3.9 Effects of hTTR and mTTR on the heat-induced aggregation of CPK 67 Figure 3.10 Effects of hTTR and mTTR on the heat-induced aggregation of CS 69 Figure 3.11 Effects of hTTR on amyloid formation by ccbw 70 Figure 3.12 Effects of non-chaperone control proteins on amyloid formation by ccbw 71 Figure 3.13 Effects of hTTR on amyloid formation by a-syn 72 Figure 3.14 Effects of mTTR on amyloid formation by ccbw and a-syn 74 Figure 3.15 Short conserved region shared between NS and TTR 75 Figure 3.16 Amphipathicity of predicted NS and TTR a-helices 82 Figure 3.17 Effects of PNS and PCTRL on amyloid formation by ccbw 84 Figure 3.18 Predicted effects of ECs on amyloid formation 94 Schematic diagram showing the different configurations of reagents used in a sandwich Figure 4.1 104 ELISA to specifically detect Tg-client protein complexes Figure 4.2 bTg is an efficient inhibitor of Ab aggregation 108 Figure 4.3 bTg does not specifically inhibit Cal aggregation 109 Figure 4.4 bTg and hTg are efficient inhibitors of CS aggregation 110 Figure 4.5 bTg and hTg are efficient inhibitors of OT aggregation 112 Images of InstantBlueä-stained native PAGE gels showing heat induced structural Figure 4.6 113 changes in Tg BisANS assay shows a significant increase in solvent-exposed hydrophobicity on Tg Figure 4.7 114 following heating at 60°C Figure 4.8 SEC profiles of native and heat-stressed bTg and CS 115 Figure 4.9 Images of Coomassie-stained SDS-PAGE gels analysing SEC fractions 116 Figure 4.10 Sandwich ELISA confirms that bTg and CS are physically associated in Tg-CS complexes 117 DLS estimates of the mean diameters of HMW Tg-CS complexes and uncomplexed Figure 4.11 118 native bTg and CS NTA distribution curves for estimation of the mean diameters of HMW Tg-CS complexes Figure 4.12 119 and uncomplexed native bTg and CS Figure 4.13 ThT assay measuring amyloid formation by Cal 120 Figure 4.14 MTS assay to test Nthy-ori 3-1 viability after incubation with 20 µM Cal 121 MTS assay to test the effects of 50 μM Cal on the viability of PC12, HEK293, and SH- Figure 4.15 122 SY5Y cells Figure 4.16 ThT assay measuring Ab amyloid formation 123 Figure 4.17 Calcein-AM assay to measure SH-SY5Y viability after incubation with 30 µM Ab 124 viii

Schematic representation of thyroidal endocytosis of Tg and associated trafficking Figure 4.18 131 pathways Schematic drawings showing the formation, internalisation, and degradation of HMW Figure 5.1 133 chaperone-client complexes Figure 5.2 Schematic depiction of receptor affinity adsorption assay methodology 138 Figure 5.3 Flow cytometry analysis of transfected HEK293 cells expressing SRA-1 and SREC-1 144 Flow cytometry analysis of biotinylated CLU-CS and control proteins binding to RAW Figure 5.4 145 264.7 cells Figure 5.5 Flow cytometry analysis of biotinylated Tg-CS binding to RAW 264.7 cells 146 Figure 5.6 Flow cytometry analysis of biotinylated CLU-GST binding to RAW 264.7 cells 147 Flow cytometry analysis of biotinylated CLU-GST and Tg-CS binding to EOC13.31 Figure 5.7 148 cells Figure 5.8 Flow cytometry analysis of biotinylated CLU-GST and Tg-CS binding to HepG2 cells 149 Reducing 10% SDS-PAGE analysis confirms that anti-CS-CNBr Sepharose captures the Figure 5.9 150 CLU-CS complex Reducing 15% SDS-PAGE reveals no difference in the protein composition of affinity Figure 5.10 adsorption eluates from lysates prepared from cells incubated with or without the CLU- 151 CS complex Figure 5.11 Flow cytometry analysis to detect CLU-CS binding to RAW 264.7 cells 152 Flow cytometry analysis showing that non-biotinylated CLU-CS is not detected on the Figure 5.12 153 surface of RAW 264.7 cells with anti-CLU or anti-CS antibodies Figure 5.13 SDS-PAGE analysis of CLU-CS and biotinylated CLU-CS complexes 155 Figure 5.14 SDS-PAGE analysis of CLU-GST and biotinylated CLU-GST complexes 156 Figure 5.15 SDS-PAGE analysis of Tg-CS and biotinylated Tg-CS complexes 158 Figure 5.16 Dot blot detection of CLU-CS complexes with anti-CS antibodies 159 Figure 5.17 Dot blot detection of CLU-CS complexes with G7 anti-CLU and streptavidin-HRP 160 Figure 5.18 Dot blot detection of CLU-GST complexes with two different anti-GST antibodies 161 NTA distribution curves estimating the mean diameters of CLU-CS and biotinylated Figure 5.19 162 CLU-CS complexes Flow cytometry analysis comparing the binding of CLU-CS and biotinylated CLU-CS to Figure 5.20 163 RAW 264.7 cells Figure 5.21 Flow cytometry analysis of non-biotinylated Tg-CS binding to RAW 264.7 cells 164 Figure 5.22 Flow cytometry analysis of non-biotinylated CLU-GST binding to RAW 264.7 cells 165 Flow cytometry analysis of the binding of biotinylated and non-biotinylated CLU-GST Figure 5.23 167 complexes to EOC13.31 cells Flow cytometry analysis of biotinylated and non-biotinylated CLU-GST binding to Figure 5.24 168 HepG2 cells Flow cytometry analysis showing that CLU-GST binds to transfected cells expressing Figure 5.25 170 ApoER2 Flow cytometry analysis showing that CLU-GST binds to transfected cells expressing Figure 5.26 171 VLDLR Figure A1 SEC profiles of CLU-CS and CLU-GST 228 Figure A2 Flow cytometry analysis of protein binding to RAW 264.7 macrophages 229 Figure A3 Confocal microscopy images of RAW 264.7 macrophages 230

List of Tables

Table 1.1 Examples of PDDs and the proteins implicated in their pathology 8 Table 1.2 Examples of cell surface receptors that bind known ECs 34 Table 3.1 Summary of the client proteins and conditions used to induce aggregation 53 Algorithm parameter changes used in NCBI’s Standard Protein BLAST Table 3.2 55 tool Pairwise alignment results for the 14-residue sequences in NS and TTR Table 3.3 77 compared with corresponding sequences of other proteins Expression levels of recombinant NS in various bacterial strains at 28°C Table 3.4 85 and 20°C Table 4.1 Summary of the client proteins and conditions used to induce aggregation 99 Table 4.2 Algorithm parameter settings used in NCBI’s standard protein BLAST tool 107 Table 5.1 List of antibodies and conjugates used in this study 134 Table 5.2 List of plasmids used in this study 136 Table 5.3 Brief details of scavenger receptors 173 A brief summary of results from cell binding experiments using Table 5.4 174 biotinylated HMW chaperone-client complexes

Abbreviations a2M alpha-2-macroglobulin a-Lac alpha-lactalbumin a-syn alpha-synuclein b2M beta-2-microglobulin b-Dog 1-O-n-Octyl-b-D-Glucopyranoside Ab beta-amyloid AD Alzheimer’s disease ADH alcohol dehydrogenase ADP adenosine diphosphate AFU arbitrary fluorescent units ALS amyotrophic lateral sclerosis ApoER2 apolipoprotein E receptor 2 APS ammonium persulfate ATCC American Type Culture Collection aTF apo-transferrin ATP adenosine triphosphate BCA bicinchoninic acid BiP binding immunoglobulin heavy-chain binding protein bisANS 4,4’-dianilino-1,1’-binaphthyl-5,5’-disulfonic acid, dipotassium salt BSA bovine serum albumin bTg bovine thyroglobulin Cal calcitonin ccb coiled-coil beta peptide ChEL cholinesterase-like Clic1 cys-less chloride intracellular channel 1 CLU clusterin CMA chaperone-mediated autophagy CNS central nervous system CNSA central nervous system selective amyloidosis CPK creatine phosphokinase CS citrate synthase CSF cerebrospinal fluid DLS dynamic light scattering DMEM Dulbecco’s modified eagle medium DMSO dimethyl sulfoxide DNA deoxyribose nucleic acid DTT dithiothreitol EC extracellular chaperone ECL enhanced chemiluminescence EDTA ethylenediaminetetraacetic acid ER endoplasmic reticulum ERAD ER-associated degradation

EtOH ethanol FAC familial amyloidotic cardiomyopathy FAP familial amyloidotic polyneuropathy FBS fetal bovine serum FENIB familial encephalopathy with NS inclusion bodies GST glutathione-S-transferase HB haemoglobin HDC PBS containing 1% (w/v) heat-denatured casein and 0.01% (w/v) thimerosal HFIP 1,1,1,3,3,3-hexafluoro-2-propanol HMW high molecular weight Hp haptoglobin HRP horseradish peroxidase hsc70 heat-shock cognate of 70 kDa HSP heat-shock protein hTg human thyroglobulin hTTR human transthyretin (tetrameric) HypF-N N-terminal domain of E. coli hydrogenase maturation protein IPTG isopropyl b-D-1-thiogalactopyranoside LAMP lysosomal-associated membrane protein LB Luria broth LDL low-density lipoprotein LRP low-density lipoprotein receptor-related protein mRNA messenger RNA 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)- MTS 2H-tetrazolium mTTR mutant monomeric TTR containing the mutations F87M and L110M NCBI National Center for Biotechnology Information NP-40 Tergitol-type nonyl phenoxypolyethoxylethanol NS neuroserpin NTA nanoparticle tracking analysis OD optical density OT ovotransferrin Oval ovalbumin phosphate buffered saline; 2.7 mM KCl, 1.75 mM KH PO , 135 mM NaCl, 10 PBS 2 4 mM Na2HPO4, pH 7.4 PBS/az PBS containing 0.05% (w/v) sodium azide PBS/skim milk PBS containing 5% (w/v) skim milk powder PCTRL synthetic peptide with a scrambled sequence of NS residues 300 – 313 PD Parkinson’s disease PDB Protein PDD protein deposition disease PDI protein disulfide isomerise PI propidium iodide PNS synthetic peptide corresponding to NS residues 300 – 313 PPU Protein Production Unit xii

RAP receptor-associated protein RCL reactive centre loop RNA ribose nucleic acid ROS reactive oxygen species RPMI Roswell Park Memorial Institute SA Streptavidin-Alexa SAP serum amyloid P component SDS-PAGE sodium dodecyl sulphate polyacrylamide gel electrophoresis SEC size exclusion chromatography SEM standard error of the mean SERPIN serine protease inhibitor shRNA short-hairpin RNA sHSP small heat-shock protein siRNA short interfering RNA SOC super optimal broth with catabolite repression SOD1 superoxide dismutase 1 SSA senile systemic amyloidosis T3 triiodothyronine T4 thyroxin TAE 40 mM Tris, 20 mM acetic acid, 1 mM EDTA, pH 8 TB Terrific broth TBS tris buffered saline; 0.5 M Tris, 1.5 M NaCl, pH 8 TBST TBS containing 0.1 % Triton-X 100 TEMED N,N,N’,N’-Tetramethylethylenediamine Tg thyroglobulin TH buffer 50 mM Tris-HCl, 5 mM HEPES, pH 8 ThT thioflavin T TMC thyroid medullary carcinoma tPA tissue-type plasminogen activator TREM2 triggering receptor on myeloid cells TTR transthyretin Ub ubiquitin UPR unfolded protein response UPS ubiquitin-proteasome system VLDLR very low-density lipoprotein receptor

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Chapter 1

Chapter One Introduction

Correct protein folding, and the regulation of protein unfolding, are integral biological processes that are required for the maintenance of cellular function and protein homeostasis, or “proteostasis”. Proteostasis refers to the collective processes required to maintain the concentration, structure and function of proteins in biological systems (Balch et al. 2008; Wyatt et al. 2012). Much is known regarding the complex network of intracellular systems for protein quality control, however, much less is known about corresponding systems in the extracellular environment. This chapter aims to summarise our current understanding of intracellular protein quality control systems and discusses the importance of extracellular chaperones (ECs) and their likely role in the prevention and clearance of pathological protein aggregates in the extracellular environment.

1.1 Protein structure, function, & folding Proteins are involved in all biochemical processes that take place within living organisms. The synthesis of proteins according to the genes encoded in deoxyribose nucleic acid (DNA) leads to many thousands of different molecules in biological systems (Shenoy and Jayaram 2010). Each individual protein has a characteristic shape, size and function. In order for a protein to become biologically functional, it is necessary that it first fold efficiently and accurately into its normal three-dimensional structure, even in the case where a functional protein contains disordered regions (there are currently over 200 known proteins that contain at least one intrinsically disordered domain) (Uversky 2019). Proteins are linear polymers of amino acids. The unique sequence of the amino acid residues that form the polypeptide backbone is the key determinant of its three dimensional secondary, tertiary and quaternary structures, which result from the folding process (Anfinsen 1973; Wang et al. 2008). Interactions that lead to the formation of the stable, native structure include hydrogen bonds, ionic interactions, disulfide bonds, and hydrophobic interactions. Hydrogen bonds formed between amino acid residues lead to the formation of secondary structures (i.e. alpha helices and beta sheets), whilst ionic interactions and disulfide bonds between distant amino acid residues on the same polypeptide chain lead to the formation of the tertiary structure of the protein. An additional level of protein folding (quaternary structure) can be observed when interactions between multiple polypeptide chains occur, leading to the formation of a single protein with multiple subunits. The shape of the active protein and the distinct placement of amino acid residues determine the function of the protein and the interactions other

Chapter 1 molecules can have with it. All of this leads to the highly specific functions performed by each protein produced.

Proteins are involved in initiating and regulating nearly all biological processes (Dobson 1999), including the maintenance of proteostasis. When proteins do not fold properly two problems can result: (i) normal protein function is impaired, disrupting physiological functions, and (ii) partially folded and misfolded proteins have the potential to form insoluble aggregates, which are associated with many serious (and potentially fatal) health problems and diseases (Cheung and Truskett 2005).

Exactly how a protein reaches its native structure is not entirely understood, however, it is generally accepted that proteins follow a folding pathway. It has been shown that in vitro, the folding of small proteins can occur quickly and is directed by nothing more than the amino acid sequence of the protein (Karplus and Weaver 1994). The “Levinthal paradox” points to the apparent contradiction between the observations that (i) there are an infinite number of potential conformations a protein could adopt during the folding process, and (ii) if the search for the native state was “random” the process would take a very long time, but instead proteins generally fold very quickly, often on a timescale of micro- or milliseconds (Lu and Liu 2010; Carver et al. 2003; Mayor et al. 2003; Yang and Gruebele 2004; Levinthal 1968). Scientists have invested years of research to explain this paradox. In the past decade it has been shown that the majority of small, single-domain proteins, about 120 amino acid residues or less, follow a two-state folding pathway (Jackson 1998). Two state proteins, such as chymotrypsin inhibitor 2, fold in a matter of milliseconds without experimentally detectable intermediates, making it very difficult to study this process (Jackson and Fersht 1991; Weikl 2010). In contrast, large, multi-domain proteins have a more complex folding pathway that involves a transition between several metastable intermediates as domains fold independently to reach the native structure (Jackson and Fersht 1991). Despite the increased complexity of the multi-domain folding pathway, some multi-domain proteins fold with two-state kinetics, involving the formation of localised, native-like folded domains (intermediate structures containing localised hydrophobic regions).

All models of multi-domain protein folding incorporate protein-folding intermediates and can be represented schematically by a “protein folding funnel” (Figure 1.1). The protein folding funnel suggests that proteins can adopt many random intermediates in a non-sequential manner before reaching their native folded state, however, native interactions are relatively more stable than non- native interactions, and therefore, the protein is “funnelled” towards the most thermodynamically 2

Chapter 1 stable structure. The fully folded native state is not achieved until all native-like interactions have been formed both within and between folded domains (Dobson 2003). This process is driven by free energy, the stable, native state being of lower energy than the primary and intermediate structures under physiological conditions (Anfinsen 1973). A protein is said to be “on” the folding pathway during this process, whether it is unfolded, partially folded (folding intermediates) or completely folded into its native structure. Since unfolded and partially folded proteins are thermodynamically unstable, it is natural for proteins to transition between these states (Cheung and Truskett 2005). Due to their instability, there is insufficient structural data to describe these transition intermediates (Daggett and Levitt 1992; Religa et al. 2005). Understanding this process is further complicated by the fact that in vitro studies do not mimic the crowded molecular environment within a cell and that many proteins require assistance from molecular chaperones to reach their native conformations (Hartl 1996; Zimmerman and Minton 1993). Molecular chaperones bind and release exposed hydrophobic regions of the intermediate protein structures to allow for appropriate folding and prevent protein aggregation.

Chapter 1

Figure 1.1 The protein folding funnel. The axis at the right of the figure represents the energy of the protein and the axis on the left represents the percentage of amino acid residues in their correct positions as the protein moves towards its single native state (green zone) via intramolecular interactions. The further the protein moves down the “funnel” the fewer intermediate forms it has available to transition between. Partially folded proteins can interact with other proteins (intermolecular interactions) via solvent-exposed hydrophobic regions, resulting in the formation of amorphous aggregates, toxic oligomers, or ordered amyloid fibrils (red zone). The figure also indicates the processes facilitated by molecular chaperones (i.e. de novo folding and inhibition of protein aggregation either by refolding or assisting in the degradation of partially folded proteins) (modified from Hartl et al. 2011).

1.2 Protein unfolding, misfolding, & aggregation Proteins are dynamic molecules that can both fold and unfold. Cells are equipped with a sophisticated network of machinery to control and regulate protein unfolding, for example, during processes such as transport across membranes and protein degradation (Ecroyd and Carver 2008). Despite these mechanisms, uncontrolled protein unfolding and aggregation can occur in response to various stresses, such as elevated temperature, changes in pH and ion concentrations and oxidation (Ramirez-Alvarado et al. 2010; Kelly and Balch 2003; Finkel and Holbrook 2000; Levine et al. 1994; Davies et al. 1987). Although it is believed that molecular crowding actually stabilises the native state of proteins, it has been shown that molecular crowding can interfere with the initial folding process (Kuznetsova et al. 2014; Gorensek-Benitez et al. 2017). Mutations in

Chapter 1 genetic material and/or errors in transcription or translation may also lead to the expression of proteins that are trapped on the “off-folding pathway” (e.g. incapable of folding into their native structure) (Stefani and Dobson 2003). The pattern and extent of unfolding (and propensity of a protein to aggregate) is influenced by structural modularity (consisting of different, independent folding domains, which have distinct stability and susceptibility to degradation), secondary structure composition (α-helices and β-sheets exhibit vastly different propensities to unfold), and post-translational modifications (Wolff et al. 2014). Amino acid composition can also play a role in the susceptibility and propensity of a protein to unfold and aggregate (i.e. arginine, cysteine, hsitidine, and lysine are particularly susceptible to oxidation) (Levine et al. 1994; Stadtman and Levine 2003).

When proteins become irreversibly misfolded or unfolded, their behaviour deviates from the normal and they have the potential to form pathological protein aggregates (Yerbury et al. 2005a). Insoluble, non-native protein aggregates form when solvent-exposed hydrophobic regions of proteins (normally hidden within the interior of the folded protein) interact with other protein molecules (Dobson 2003; Rajan et al. 2001). There are two main types of insoluble protein aggregates: (i) ordered/fibrillar aggregates (amyloid fibrils or plaques), and (ii) disordered/non- fibrillar (amorphous) aggregates (Figure 1.2) (Cheung and Truskett 2005; Stefani 2004). Protein aggregates can form both intracellularly and extracellularly. Intracellularly, aggregates can accumulate to form deposits, known as inclusion bodies (Markossian and Kurganov 2004; Stefani 2004) or neurofibrillary tangles (Binder et al. 2005). In mammalian cells inclusion bodies can be amorphous or fibrillar in structure and contain more than one type of protein, including chaperones, components of the ubiquitin-proteasome system, centrosomal material and cytoskeletal proteins (Rajan et al. 2001). Examples of inclusion bodies include Lewy bodies, such as those associated with Parkinson’s disease (PD) (Kim et al. 2014), Pick bodies associated with Pick’s disease (Kertesz and Munoz 1998) or superoxide dismutase 1 (SOD1) inclusions associated with familial amyotrophic lateral sclerosis (ALS) (Liu et al. 2004).

Chapter 1

Figure 1.2 Formation of protein aggregates. Due to the dynamic instability of proteins there are several possible structures they can assume: native (N) and non-native (NN), the latter including intermediates (I1 and I2) and unfolded (U) structures. Non-native structures are prone to associate with each other via hydrophobic interactions to form aggregates. Amorphous aggregates result from non-specific interactions between many different protein types and conformations, whereas highly structured amyloid fibrils can be formed by β-sheet-rich intermediates. Soluble structures are shown in the internal blue rectangle, insoluble structures are shown in the outer white region (Yerbury et al. 2005a).

Amyloid fibrils are considered to be highly structured, non-native, quaternary structures that are rich in β-sheet structures (Figure 1.3) (Greenwald and Riek 2010; Kelly and Balch 2003; Huang et al. 2000). The accepted model suggests that the formation of amyloid fibrils is based on a nucleation-dependent polymerisation mechanism. This involves two phases: (i) nucleation and (ii) elongation (Jarrett and Lansbury 1993; Murphy 2007; Wilson et al. 2008; Khurana et al. 2003). Nucleation begins when hydrophobic regions of partially folded or misfolded proteins interact with one another to form an oligomeric amyloid “nucleus” (Jarrett and Lansbury 1993; Jayaraman et al. 2012; Chiti and Dobson 2006). Once established, the oligomers begin to polymerise, forming intermediate structures known as protofibrils (Khurana et al. 2003). Elongation occurs when monomers are added to the protofibril (Langkilde and Vestergaard 2009). The amyloid fibril forms when several protofibrils (usually between two and six) interact with one another (Steffani and Dobson 2003; Khurana et al. 2003).

Chapter 1

Amyloid Fibrils Oligomers

Elongation (Growth) Phase

Nucleation (Lag) Phase Native State n Folding Nucleus Protofibrils Intermediates/ Misfolded Proteins

Figure 1.3 Model of amyloid formation. Scanning electron microscopy image of amyloid fibrils (upper left) and a schematic representation of a model for amyloid formation. The microscopy image shows amyloid-β fibrils found in brain tissue from a patient with Alzheimer’s disease (AD) (from Sachse 2010). These structures are self-assembled and insoluble (Kominos 1995). The formation of amyloid involves a two-step mechanism: (i) nucleation and (ii) elongation. Nucleation occurs when hydrophobic regions of partially folded or misfolded proteins interact to form a nucleus. Elongation commences when nuclei sequester other aggregate-prone intermediates, leading to rapid fibril growth, and subsequently the formation of insoluble mature amyloid fibrils (modified from Wilson et al. 2008).

In contrast, amorphous aggregates have no specific structure or organisation and can form deposits responsible for the development of, for example, cataracts and wine haze (Figure 1.4). In the past, very little research has been done on the mechanism of amorphous aggregation, therefore, there are currently no established models available (Stranks et al. 2009). It has been suggested that the aggregation pathway a protein follows is influenced by protein concentration, the type of stress (such as pH, oxidation or temperature) and the structure and characteristics of the protein forming the aggregate (Huang et al. 2000). Interestingly, it was also been shown that the binding of some chaperones to amyloidogenic precursor proteins leads to the formation of non-cytotoxic amorphous aggregates rather than cytotoxic amyloid (Cascella et al. 2013; Cappelli et al. 2016; Mannini and Chiti 2017).

Figure 1.4 Scanning electron microscopy image of amorphous aggregates. Aggregates of human serum albumin were imaged. A 10 μm scale bar is shown at the bottom right of the image (Ishtikhar et al. 2015). 7

Chapter 1

1.3 Protein aggregates, cytotoxicity, and disease The accumulation of misfolded proteins and protein aggregates is implicated in causing cell dysfunction and is associated with many serious diseases (Table 1.1) (Ecroyd and Carver 2008). Changes such as genetic mutations (causing incorrect protein folding) and age-related deficiencies in proteostasis may predispose individuals to protein deposition diseases (PDDs) such as AD, familial and sporadic ALS, PD and type II diabetes. These diseases are associated with insoluble deposits of aggregated proteins in tissues (Gupta et al. 2011; Hartl et al 2011; Kim et al. 2013a). Many PDDs are late-onset, suggesting that the underlying cause of the disease may be due to cumulative disruption or overwhelming of protein quality control mechanisms that were once able to maintain proteostasis. Although the link between aging and progressive deficiencies in protein quality control is not fully understood, it is likely that the cumulative exposure of proteins to stresses (heat, changes in pH or ion concentrations, molecular crowding, oxidation, or physical shear stress) may be responsible for late-onset PDDs (Wyatt et al. 2009a).

Table 1.1 Examples of PDDs and the proteins implicated in their pathology. Below is a list of well- known diseases defined by the presence of deposits of aggregated proteins. The right column shows the predominant protein component of the aggregates associated with each disease. * Denotes extracellular deposition (Ruberg and Berk 2012; Wyatt et al. 2009a; Ecroyd and Carver 2008; Kiss et al. 2005; Benson et al. 1993).

Disease Protein AD beta-amyloid (Aβ) * PD alpha-synuclein (α-syn) Dementia with Lewy bodies α-syn Transmissible spongiform encephalopathies prion protein * (Creutzfeldt-Jakob disease and Mad Cow disease) Huntington’s disease huntingtin Type II diabetes amylin * Senile systemic amyloidosis and transthyretin (TTR) * Cardiac amyloidosis ALS SOD1 Haemodialysis-related amyloidosis beta-2-microglobulin (β2M) * Hereditary renal amyloidosis fibrinogen * Medullary carcinoma of the thyroid calcitonin (Cal) * Hereditary non-neuropathic systemic amyloidosis lysozyme *

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Although the factors responsible for the formation of disease-related protein aggregates is still unknown, it is clear that the deposition of these aggregates often precede the onset of disease symptoms. It has been well established that protein aggregates lead to cellular dysfunction and have detrimental effects on the tissues and organs in which they appear, leading to disease pathogenesis. Although amorphous aggregates are associated with many debilitating diseases, such as age-related macular degeneration and retinitis pigmentosa (Surgucheva et al. 2005), cataracts (Moreau and King 2012), and renal disease (Salahuddin 2015; Yao et al. 2004), previous research studies have predominantly focussed on the cytotoxicity of amyloidogenic proteins. Although Aβ (the predominate protein aggregate associated with AD) is the most frequently studied amyloidogenic peptide to date, many other amyloid-forming proteins (both those found associated with other PDDs and synthetic peptides) have been shown to exhibit very similar cytotoxic effects, despite lacking any sequence homology with Aβ (Schubert et al. 1995). This leads to the conclusion that cytotoxicity is likely due to either the characteristic structure of amyloid (the cross-β conformation) or the assembly process of the protein aggregates and has less to do with the identity of the individual protein associated with the disease (Zamotin et al. 2006). Over the past two decades, many studies have investigated the mechanism by which amyloid aggregation occurs and which aggregate species are responsible for the resulting toxicity. From these studies, there is overwhelming evidence that amyloid cytotoxicity is primarily associated with either misfolded monomers or early-stage, prefibrillar oligomers, rather than mature amyloid fibrils (Verma et al. 2015; Walsh et al. 2014; Kayed and Lasagna-Reeves 2013; Yerbury et al. 2007; Bucciantini et al. 2002; Schubert et al. 1995; Reixach et al. 2004; Manral and Reixach 2015). For example, it has been shown that the accumulation of oligomeric Aβ in brain tissue leads to structural changes in macromolecules (i.e. proteins and lipids) resulting in progressive neuronal and synaptic dysfunction, which are symptoms associated with the disease.

One theory suggests that cytotoxicity associated with some amyloidogenic proteins (such as Aβ) is due to the production of a more oxidative environment. Oxidative stress is caused by an imbalance between the overproduction of reactive oxygen species (ROS), such as H2O2, and their inactivation/removal by antioxidants, as well as an inability to repair the resulting damage caused by such species (Garcia-Garcia et al. 2012). As mentioned previously, oxidative stress can lead to conformational changes in proteins and a loss of their function, leading to cell dysfunction and death. Studies have found that oxidative stress plays a large role in the pathological progression of many PDDs, such as AD (Huang et al. 2016; Perry et al. 2002), thyroid medullary carcinoma (TMC) (Hosseini-Zijoud et al. 2016; Wang et al. 2011), PD (Blesa et al. 2015; Hwang 2013) and type II diabetes (Folli et al. 2011; Wright et al. 2006). In the case of AD, oxidative stress correlates

Chapter 1 with the accumulation of Aβ oligomers and is thought to occur due to several reasons. Firstly, Aβ oligomers have been shown to bind metal ions (i.e. Cu2+, Fe3+, and Zn2+), which are potent catalysts of oxidation (due to the production of H2O2) and sequester them within amyloid plaques. The accumulation of metal ions within tissues could lead to a highly oxidative environment (Prasansuklab and Tencomnao 2013; Cristovao et al. 2016). Secondly, Aβ oligomers have been shown to anomalously bind to cell surface receptors, to mimic and/or blockade the binding of normal ligands. This not only interferes with a multitude of signal transduction pathways to result in abnormal cell function, but also allows for the internalisation of Aβ oligomers, introducing oxidative stress to the intracellular environment (Falsone and Falsone 2015; Kayed and Lasagna- Reeves 2013; Stefani and Rigacci 2013). Lastly, it has been shown that oligomeric Aβ, as well as other amyloidogenic proteins, can be incorporated into the plasma membrane (due to structural similarities to integral membrane protein channels) causing disruptions to membrane fluidity and permeability, as well as leading to disruption of ion gradients across membranes, uncontrolled ion influx (particularly Ca2+), and cell death (Stefani and Dobson 2003; Kourie and Henry 2002; Kawahara et al. 2000; Arispe et al. 1993).

Although there is strong evidence to suggest that oxidative stress plays a role in amyloid toxicity and PDD progression, the underlying cause for the initiation of amyloid formation is still unclear. In most cases, it is likely that the cumulative effects of stresses (e.g. changes in pH or temperature, oxidation, or shear stress) over time lead to functional decline or the overwhelming of the protein quality control system and is key to the onset of PDDs (Yerbury et al. 2016). Increasing our understanding of the mechanisms that regulate and maintain protein quality control will be critical in efforts to develop therapies for PDDs.

1.4 Intracellular protein quality control Intracellular protein quality control is essential to maintain cellular homeostasis (Hatakeyama and Nakayama 2003). Due to the structural instability of proteins, protein folding and the maintenance of proteins in their functional conformations require that both intracellular and extracellular mechanisms be in place to prevent, repair or degrade misfolded proteins. However, protein quality control is made difficult by the large degree of heterogeneity across the vast number of proteins, including differences in size, shape, stability and lifespan (Wolff et al. 2014). To cope with this heterogeneity, a complex network of quality control checkpoints, stress responses, molecular chaperones, degradation machinery and other regulatory factors are required (Gupta and Tuteja 2011; Kim et al. 2013a; Wolff et al. 2014). Figure 1.5 outlines the integrated network of intracellular protein quality control processes. 10

Chapter 1

Figure 1.5 An integrated network of intracellular protein quality control mechanisms. The proteostasis network integrates chaperone pathways for (i) the folding of newly synthesised proteins (biogenesis; indicated by the green arrows), which utilises chaperones during translation, trafficking and initial folding, (ii) the remodelling of misfolded proteins (conformational maintenance; indicated by the blue arrows), which utilises chaperones to unfold misfolded or aggregated proteins for refolding, and (iii) protein degradation (indicated by the red arrows), which utilises chaperones to facilitate the degradation of proteins that have become trapped in a non-native form via the ubiquitin-proteasome system (UPS) or lysosomal autophagy (Kim et al. 2013a).

The first checkpoints in protein quality control occur during protein synthesis. Many regulators and proofreading systems are in place to ensure DNA is transcribed accurately into messenger RNA (mRNA) (Zenkin et al. 2006) and that mRNA is translated into the correct amino acid sequence. Recent studies have suggested that ribosomes utilise multiple sophisticated mechanisms to reduce translational errors, one mechanism being the regulation of translation rate. For example, slowing the rate of translation for proteins containing large regions of (aggregation-prone) exposed hydrophobicity allows for the increased binding of chaperones prior to the release of the protein by the ribosome, preventing aggregate formation (Wolff et al. 2014). However, transcriptional and translational proofreading is not fail-proof and occasionally mistakes occur, leading to the synthesis of proteins that may be incapable of folding into their native structures.

Chapter 1

For this reason, multiple quality control mechanisms are in place, several of which can be found in the endoplasmic reticulum (ER). Asparagine-linked carbohydrate moieties are added to many proteins entering the secretory pathway and serve as a signal for protein folding progress (Bernales et al. 2006). Only when they have reached their functional native state are proteins secreted, providing an exquisite mechanism of quality control (Kaganovich et al. 2008). If proteins are incapable of folding correctly, they become trapped and accumulate within the ER, inducing ER stress and triggering the unfolded protein response (UPR). The UPR is activated by the coordinated action of three ER transmembrane stress sensors: inositol-requiring enzyme 1α, PKR-like ER kinase, and activating transcription factor 6α. Under normal conditions, these ER stress sensors are associated with the chaperone BiP (binding immunoglobulin heavy-chain binding protein), which keeps them in an inactive state. However, under conditions of ER stress, BiP dissociates from the ER stress sensors and binds to misfolded proteins accumulating in the ER lumen, thereby activating the stress sensors and the UPR. Through complex signalling transduction pathways, the UPR, in an attempt to restore ER proteostasis, increases the surface area of the ER plasma membrane, attenuates the influx of proteins into the ER, increases the production of key components of protein folding and quality control machinery (chaperones and protein-modifying enzymes), and increases the production of proteins related to ERAD (ER-associated degradation), a pathway that mediates the retrotranslocation of permanently misfolded proteins from the ER to the cytosol for degradation via the UPS, and autophagy (Kadowaki et al. 2018; Bernales et al. 2006; Kim et al. 2013a; Ma and Hendershot 2004; Grootjans et al. 2016). If ER proteostasis cannot be restored, the UPR triggers apoptosis and cell death (Hetz 2012).

Proper folding of secretory and ER-resident proteins is one of the primary functions of the ER, however, conditions within the ER are significantly different from those in other cellular compartments (such as the cytosol) and this affects the energetics and kinetics of protein folding. For example, molecular crowding in the ER is 3–6 times higher than in the cytosol (which can reach protein concentrations up to 400 g/L), redox potential is 1000 times more oxidising, and the Ca2+ concentration can fluctuate (ranging from 100 – 800 µM in the ER vs ~ 100 nM in the cytosol) (Samtleben et al. 2013). The ER also employs unique enzymes and other components necessary for glycosylation (Gidalevitz et al. 2013; Hartl et al. 2011). For these reasons, it is imperative that the ER utilises molecular chaperones (and their cofactors) to assist with protein folding. These chaperones include several heat-shock protein (HSP) family members, such as BiP (HSP70 member), GRP94 (HSP90 family member), protein disulfide isomerase (PDI), and calreticulin (Plate and Wiseman 2017; Gidalevitz et al. 2013; Ciplys et al. 2014; Conte et al. 2007).

Chapter 1

1.4.1 Molecular chaperones Associated with all protein quality control mechanisms is a complex network of molecular chaperones. These chaperones are found in various compartments within the cell including the ER and the cytosol (Hartl et al. 2011; Szolajska and Chroboczek 2011). Chaperones accompany proteins from their initial synthesis to folding and during membrane translocation and proteolytic degradation (Rehling and Rospert 2010; Gidalevitz et al. 2013; Kim et al. 2013a; Benyair et al. 2011). There are several classes of chaperones, and despite the differences in their structure and function, all chaperones have two features in common: (i) they selectively bind to non-native protein conformations (via exposed regions of hydrophobicity), and (ii) they inhibit the irreversible aggregation of non-native proteins (Wilson et al. 2008). The most abundant molecular chaperones are the HSPs (heat shock proteins; HSP40, HSP60, HSP70, and HSP90 – the numbers refer to their molecular weights) and the small heat-shock proteins (sHSPs), named due to their increased expression during times of elevated temperature or other stresses (Fink 1999; Sathish et al. 2003; Kim et al. 2013a). The chaperone functions of the HSPs and PDI are explained below. Calreticulin exerts its chaperone activity in two ways: (i) it prevents non-native protein species from leaving the Golgi apparatus and (ii) increases protein folding efficiency (Conte et al. 2007).

The rapid synthesis of HSPs in cells due to stress is regulated by the heat shock transcription factor- 1 (HSF1) (Zhang et al. 2002; Mesika and Reichmann 2019; Gomez-Pastor et al. 2018). The chaperone activity of HSP70, HSP60 and HSP90 is modulated by the binding and hydrolysis of ATP (Hartl et al. 2011) along with other co-factors. HSP70 members bind to nascent protein chains on ribosomes, preventing their premature folding, misfolding and/or aggregation (acting as a “holding” chaperone), however it does not participate in the folding process (Mayer 2010). HSP70 members also accompany newly synthesised proteins during translocation from the cytosol to the mitochondria or ER (Hartl et al. 2011; Fink 1999). HSP70 function is regulated by ATP; when ATP binds to the chaperone’s N-terminal nucleotide-binding domain, its α-helical “lid” opens, allowing the hydrophobic regions of partially folded proteins to loosely bind to the C-terminal substrate-binding domain (Penke et al. 2018; Zhuravleva and Gierasch et al. 2015). When ATP is hydrolysed, HSP70 undergoes a conformational change that closes the α-helical “lid”, binding the substrate tightly (shielding the hydrophobic regions of the protein). After ADP is cleaved from HSP70, the α-helical “lid” reopens to release the protein for folding by HSP60 family members (also termed chaperonins) (Hartl et al. 2011; Kim et al. 2013a). Chaperonins are found in all compartments within the cell except for the ER and function similarly to HSP70, with the exception that they can facilitate the folding and assembly of their protein substrate (Mayer 2010;

Chapter 1

Fink 1999). In conjunction with ATP, HSP60s are regulated by the co-chaperone chaperonin 10. Like HSP60, HSP90 binds to misfolded proteins to prevent their aggregation but can also facilitate the folding of its substrates. However, it is believed that HSP90 interacts only with a specific group of client proteins, including steroid hormone receptors, transcription factors, and protein kinases involved in the control of cellular homeostasis, proliferation, differentiation, and apoptosis (Mayer 2010; Kovacs et al. 2005; Fink 1999). Both HSP60 and HSP90 have been shown to facilitate the unfolding and refolding of non-native proteins (Mayer 2010; Fink 1999; Obermann 1998). Unlike the HSPs, the sHSPs function independently of ATP and act as holdases, merely shielding exposed hydrophobic regions from the aqueous cellular environment (Hartl et al. 2011). Various other proteins are often classified as molecular chaperones as they are known to be involved in the folding of newly synthesised proteins. Two of these are PDI and peptidyl prolyl isomerase, which catalyse the rearrangement of disulfide bonds and the isomerisation of peptide bonds adjacent to proline residues, respectively (Fink 1999).

1.4.2 The degradation of proteins via the UPS When protein quality control mechanisms fail, protein aggregates can form. In these cases, mechanisms exist to degrade protein aggregates. Ubiquitin (Ub) is a heat-stable polypeptide that is covalently and selectively attached to misfolded and aggregated proteins via a highly ordered multi-step enzymatic cascade to trigger their proteolytic degradation (Hershko 1988; Varshavsky 2012; Hochstrasser 1996; Yerbury et al. 2016). The degradation is performed by the 26S proteasome. Proteasomes are large, barrel-shaped proteases comprised of multiple subunits (the 20S core particle bound at the ends by 19S cap particles). The 19S cap particles contain isopeptidase to remove Ub from the target protein for recycling and ATPase activities that unfold the target protein and feed it into the 20S core for degradation (Figure 1.6; Kornitzer and Ciechanover 2000; Hilt and Wolf 2004; Eldridge and O’Brien 2010). Although very important in protein quality control, ubiquitination is one of the most common protein modifications and is used in most cellular processes to maintain homeostasis, such as transcriptional regulation (degradation of transcription factors when gene expression should be turned off), translation, cell metabolism, the cell cycle, and apoptosis (Hilt and Wolf 2004; Yerbury et al. 2016).

Chapter 1

Figure 1.6 Degradation of proteins through the UPS. Ub is activated by Ub-activating enzyme (E1), conjugated by Ub-conjugating enzyme (E2) and linked to the target protein (substrate) by Ub ligase (E3). Ubiquitinated proteins are then recruited (via Ub receptors) to the 26S proteasome for degradation. Deubiquitinating enzyme (DUB) opposes the activity of E3 to release Ub for reuse (Eldridge and O’Brien 2010).

1.4.3 Lysosomal degradation pathways Lysosomal degradation (or autophagy) is another proteolytic pathway involved in the maintenance of proteostasis and contributes to the degradation of about 30% of cytosolic proteins (Dice 2007). Proteins (and other cellular debris) can be trafficked to the lysosome via several pathways. Macroautophagy involves the selective cytosolic sequestering of substrates, such as damaged organelles, cytosolic proteins, protein aggregates, or invasive microbes, into vesicles (autophagosomes), which then fuse with lysosomes to transfer their contents for degradation (Feng et al. 2014; Filimonenko et al. 2010; Kroemer et al. 2010). Microautophagy occurs when cytosolic regions (which may include proteins and other cytosolic components) become non-selectively engulfed by the lysosome via vesicles that form as a direct invagination of the lysosomal membrane. The vesicles can then pinch off into the lysosomal lumen for degradation (Park and Cuervo 2013). It is important to note that microautophagy has not yet been identified in mammals, however, it has recently been shown that a microautophagy-like process does occur in mammalian late endosomes (Mijaljica et al. 2011; Sahu et al. 2011). Unlike the two systems mentioned above, chaperone-mediated autophagy (CMA) does not involve the formation of vesicles around substrates. In CMA, cytosolic proteins that contain a specific pentapeptide sequence termed the KFERQ motif (which can be exposed in folding intermediates or misfolded proteins) are selectively targeted to lysosomes (Cuervo and Wong 2014). CMA is responsible for regulating many cellular processes by modulating the levels of intracellular enzymes, transcription factors, 15

Chapter 1 and other regulatory proteins. Due to the selectivity of this process, CMA is also responsible for the removal of damaged or misfolded proteins from the cytosol without disruptions to other functioning proteins, making it an integral system for the maintenance of proteostasis and protein quality control (Cuervo 2010). There are four main steps involved in CMA: (i) substrate recognition and binding, (ii) substrate unfolding and lysosomal targeting, (iii) substrate translocation, and (iv) substrate degradation (Kaushik and Cuervo 2012). First, hsc70 (heat-shock cognate of 70 kDa), the primary cytosolic chaperone involved in CMA, binds to the KFERQ lysosomal-degradation motif in a protein and targets it to the lysosomal membrane. At the lysosomal membrane, the target protein then interacts with lysosomal-associated membrane protein type 2A (LAMP-2A), which is embedded in the lysosomal membrane and associated with other proteins required for translocation (Dice 2007; Arias and Cuervo 2011). The target protein must then be unfolded for translocation across the lysosomal membrane into the lumen. As unfolding of the target protein is not required for hsc70 binding and trafficking, or interactions with LAMP2-2A, it is currently unknown whether hsc70 is involved in the unfolding process, or if the target protein is unfolded by other chaperones present at the lysosomal membrane after binding to LAMP-2A (Figure 1.7) (Bejarano and Cuervo 2010). Once the target protein is unfolded, it can then be translocated across the lysosomal membrane and degraded by lysosomal proteases. It has been proposed that for complete translocation of the target protein across the membrane, a form of hsc70 must also be present in the lysosomal lumen (most likely to prevent the target protein from returning to the cytosol) (Arias and Cuervo 2011).

Chapter 1

Figure 1.7 Schematic representation of CMA. Partially folded or misfolded proteins with exposed KFERQ motifs (top left) are recognised by the intracellular chaperone hsc70 (top). Once bound to its substrate, hsc70 transports the protein to the lysosomal membrane where it binds to LAMP-2A. LAMP-2A mediates translocation of the protein across the membrane into the lysosomal lumen. The protein is then degraded by proteases within the lysosome (from Massey et al. 2004).

1.5 Extracellular protein quality control In mammals, the extracellular environment is comprised of plasma and other specialised body fluids. Proteins are abundant constituents of extracellular fluids, including hormones, enzymes, transporter proteins, and receptor ligands. Much like proteins in the intracellular environment, extracellular proteins are exposed to various stresses, including increases in temperature (i.e. due to fever, exercise, or environmental exposure), physical shear stress (due to fluids being pumped throughout the body and pressure differences in body compartments), and oxidative stress (body fluids are significantly more oxidising than the cytosol) (Yerbury et al. 2016; Kushimoto et al. 2013; Bekard 2010; Urbich et al. 2006). Due to their ongoing exposure to stresses, extracellular proteins require continuous chaperone surveillance to prevent protein aggregation and maintain proteostasis. As previously mentioned, partially folded and misfolded proteins have an increased propensity to aggregate due to exposed hydrophobic regions that can interact with corresponding regions on other proteins, leading to disease (Poon et al. 2000). However, under normal, non- 17

Chapter 1 disease conditions, extracellular protein aggregates do not accumulate. This suggests the existence of an extracellular protein quality control system and led to the discovery of ECs, such as clusterin (CLU) (Humphreys et al. 1999; Poon et al. 2000), haptoglobin (Hp) (Yerbury et al. 2005b), and alpha-2-macroglobulin (α2M) (French et al. 2008; Wyatt et al. 2014). Unlike the well- characterised network of intracellular protein quality control mechanisms, much less is known about corresponding mechanisms that maintain extracellular proteostasis. In addition to ECs, recent studies suggest that an extracellular proteolytic system exists to assist in the maintenance of proteostasis.

1.5.1 Extracellular proteolytic degradation Although there are several classes of specialised extracellular proteases, targeted extracellular proteolysis of non-native proteins was earlier thought unlikely, as lysosomes are only found intracellularly and proteasomes are 300 times less abundant in plasma than inside cells. Another consideration is that that proteasomal activity requires ubiquitinated proteins and ATP (which is 1000 times less abundant in plasma than the intracellular environment) (Lavabre-Bertrand et al. 2001; Wyatt et al. 2009a). However, recent studies undertaken by Patrick Constantinescu at the University of Wollongong suggest that the tissue plasminogen activation system may also play a role in extracellular proteostasis. Amorphous and fibrillar protein aggregates stimulate the conversion of plasminogen to plasmin, which can recognise, bind, and subsequently degrade the aggregates (Constantinescu 2015; Radcliffe and Heinze 1981; Radcliffe 1983; Machovich et al. 1997; Tucker et al. 2000).

1.5.2 Known ECs There are currently six proteins known to exhibit promiscuous EC activity, and these include four secreted glycoproteins: CLU, Hp, α2M and serum amyloid P component (SAP) (Wilson et al.

2008), as well as both αs1-casein (Matsudomi et al. 2004) and β-casein (Zhang et al. 2005). Several studies suggest that ECs function similarly to the well-known intracellular sHSPs (Carver et al. 2003; Poon et al. 2000; Cappelli et al. 2016). ECs and the sHSPs all function independently of ATP hydrolysis and can inhibit the aggregation and precipitation of a wide range of misfolded proteins (Carver et al. 2003; Wyatt et al. 2009b; Shashidharamurthy et al. 2005) by selectively binding to hydrophobic regions exposed at the surface of these species (Kumita et al. 2007). The exact mechanisms by which ECs selectively recognise and bind to non-native proteins to prevent aggregation are, however, incompletely understood. There are also few reports regarding the relative binding affinities of chaperones for different substrates (Xu et al. 2010). Despite gaps in knowledge, the current accepted model suggests that once ECs have bound to aggregating non-

Chapter 1 native, misfolded proteins, they form stable, soluble, high molecular weight (HMW) complexes (Carver et al. 2003). Subsequently the chaperone-client complexes bind to cell surface receptors, are internalised by receptor-mediated endocytosis, and are transported by vesicles to the lysosome for degradation (Wyatt et al. 2009b; Wyatt et al. 2011). It should be noted that the formation of HMW complexes has only been observed between chaperones and amorphously aggregating clients and it is currently unknown if amyloidogenic proteins form such complexes as well. Although there is evidence to support the above model, the fate of chaperone-client complexes is not fully understood and may be different depending on the chaperone involved. For example, CLU and α2M have been shown to bind to amorphously aggregating proteins and facilitate their cellular uptake by receptor-mediated endocytosis via various (unidentified) scavenger receptors and low-density lipoprotein receptor-related proteins (LRPs), respectively (Wyatt et al. 2011; French et al. 2008).

CLU CLU (also called Apolipoprotein J) earned its name due to its ability to “cluster” a variety of cell types in vitro (Blaschuk et al. 1983; Fritz et al. 1983; Rosenior et al. 1987). It is a highly conserved heterodimeric glycoprotein (~ 62 kDa after glycosylation) (Kapron et al. 1997) comprised of α and β chains that are held together via five disulfide bonds. CLU is an abundant extracellular protein that is secreted into many mammalian fluids such as plasma (between 0.06 – 0.1 mg/ml), semen, breast milk, ocular fluid, urine and cerebrospinal fluid (CSF) (Polzin and Cowgill 2016; Fini et al. 2016; Wilson et al. 2000; Murphy et al. 1988; Choi et al. 1990; Aronow et al. 1993). CLU expression is upregulated in response to various cellular stresses (oxidative stress in particular) (Shin et al. 2009; Lee et al. 2012; Strocchi et al. 2006) and diseases, such as AD (Nuutinen et al. 2009; Nuutinen et al. 2007; Miners et al. 2017), asthma (Kwon et al. 2014; Hong et al. 2016), and familial amyloidotic polyneuropathy (Magalhaes and Saraiva 2011). Like with the HSPs, upregulation of CLU is regulated by HSF1-HSF2 complexes (Loison et al. 2006). In addition to its broad physiological distribution, CLU exhibits high sequence homology across a wide range of mammalian species (Jenne and Tschopp 1992), suggesting that it has an important biological function in vivo. Studies also suggest that CLU facilitates the clearance of late-stage apoptotic cells and cellular debris by macrophages, exhibiting a protective role during inflammation and autoimmune diseases (Cunin et al. 2016; Bartl et al. 2001)

CLU was the first reported EC and was shown to inhibit the aggregation of a broad range of stressed proteins. As previously mentioned, the chaperone activity of CLU is similar to that of the sHSPs (Humphreys et al. 1999) in that both selectively bind to exposed hydrophobic regions on

Chapter 1 aggregation-bound proteins trapped on the off-folding pathway to form HMW complexes (preventing in vitro aggregation of proteins independently of ATP); neither have the ability to refold proteins after the removal of stress (Yerbury et al. 2007; Carver et al. 2003; Wyatt et al. 2009b; Wyatt et al. 2009c; Poon et al. 2002a; Poon et al. 2000). Interestingly, CLU is found associated with many insoluble protein deposits and amyloid plaques. At low molar ratios of CLU:client protein, CLU inhibits the formation of amyloid fibrils formed from a variety of proteins by binding to pre-fibrillar intermediates. However, it was also shown that at very low CLU:client protein ratios, for some proteins (such as Aβ) (Narayan et al. 2012; Yerbury et al. 2007), CLU enhanced the formation of amyloid and promoted aggregate toxicity. Prefibrillar amyloid species are toxic, thus, when present at very low concentrations, CLU may be unable to shield all areas of exposed hydrophobicity and may instead stabilize aggregates that continue to assemble into larger amyloid structures (Yerbury et al. 2007; Wilson et al. 2008; Cappelli et al. 2016). These data suggest that CLU-client protein interactions are complex and dependent upon the ratios of CLU to client protein, as well as the aggregation stage of the proteins. The chaperone activity of CLU is enhanced at mildly acidic pH, and this correlates with increased exposed hydrophobicity on CLU, suggesting that these regions may be involved in the interactions between CLU and its clients (Poon et al. 2002b). Interestingly, it has been shown that during ER stress CLU is released to the cytosol, however, its intracellular functions are not yet fully understood (Nizard et al. 2007). It has been suggested that intracellularly, CLU promotes autophagy (Zhang et al. 2014) and can rescue TDP43/ALS phenotype in transgenic flies (Gregory et al. 2017).

Hp Hp is an acute phase glycoprotein expressed in the liver and humans express one of three major phenotypes, Hp1-1, Hp2-1, or Hp2-2 (molecular weights vary depending on the phenotype) (Alayash 2011). Hp is present in most human body fluids (concentrations vary depending on the phenotype being expressed) with levels increasing during inflammation, infection, trauma, and tissue damage (Dobryszycka 1997; Yerbury et al. 2009). Hp has a heterogeneous and polymeric structure that differs for each phenotype due to the number of disulfide bonds that can form between subunits. Each Hp molecule contains a variable number of (“light”) α-chains (either α1 or α2) and (“heavy”) β-chains. The structure of the three phenotypes can be described as follows:

Hp1-1 [(α1β)2], Hp2-1 [(α1β)2(α2β)n], and Hp2-2 [(α2β)n] (Polticelli et al. 2008). One study reported up to 10 repeated units for Hp2-1 and as many as 20 repeats for Hp2-2 (Cheng et al. 2009). Hp has been associated with a variety of functions including binding to free, circulating haemoglobin (HB) to facilitate its clearance via the scavenger receptor CD163 (expressed by tissue macrophages). Macrophages internalise the Hp-HB complex for degradation, preventing free HB-

Chapter 1 mediated pathologies (Alayash 2011; Polticelli et al. 2008; Louderback and Shanbrom 1968). Hp has also been implicated in immune regulation and anti-inflammatory reactions (Kurosky et al. 1980; Dobryszycka 1997; Bowman and Kurosky 1982; Louagie et al. 1993; Lim 2001).

Hp was first investigated as an EC due to the similarities it shares with CLU. Both Hp and CLU are dimeric glycoproteins comprised of α and β chains, have disulfide linkages, are abundant in human plasma and exhibit increased expression during a variety of stresses and disease states (Dobryszycka 1997; Kapron et al. 1997; Blaschuk et al. 1983; Choi-Miura et al. 1992; Rosenburg and Silkensen 1995). It was shown that Hp has a chaperone mechanism similar to that of CLU and the sHSPs, protecting a wide range of proteins from amorphous aggregation in vitro by forming soluble HMW complexes with aggregating clients during heat and oxidative stress (Ettrich et al. 2002; Yerbury et al. 2005b). Like CLU, Hp does not refold proteins, is ATP-independent, and is found associated with amyloid deposits (Yerbury et al. 2005b; Powers et al. 1981).

α2M α2M is a 720 kDa homo-tetrameric glycoprotein linked via disulfide bonds and is best known as a broad-spectrum protease inhibitor that functions in haemostatic and inflammatory responses (Harpel 1973; Imber and Pizzo 1981; Borth 1992). α2M is an abundant plasma protein (1.6 – 3.4 mg/ml) and inhibits a variety of extracellular proteases via a unique trapping mechanism (Du et al. 1997; French et al. 2008). The binding of proteases to α2M results in a conformational change in the α2M molecule, “activating” the protein by exposing an LRP1 binding site on each subunit. This provides a mechanism for the clearance of α2M-protease complexes from body fluids via receptor mediated endocytosis (Imber and Pizzo 1981; Sottrup-Jensen and Birkedol-Hansen 1989; French et al. 2008).

Like CLU, Hp and the sHSPs, α2M has a broad-spectrum chaperone action and ATP- independently interacts with many different misfolded proteins by binding to exposed hydrophobic regions, to prevent aggregation and form soluble complexes (French et al. 2008; Hope et al. 2003). The chaperone action of α2M is significantly increased when the protein is exposed to physiologically relevant amounts of hypochlorite (a naturally occurring oxidant) (Wyatt et al. 2014). This increase in chaperone efficiency is due to structural changes within the α2M molecule that cause the protein to dissociate into dimers (Wyatt et al. 2015).

Chapter 1

The Caseins

αs1-casein and β-casein are two subunits of the 100 kDa casein protein component of milk (Morgan et al. 2005). It has been shown that casein proteins have similar chaperone activity to that of the sHsps and CLU, functioning independently of ATP (Treweek 2012; Sakono et al. 2011; Morgan et al. 2005). One study showed that the chaperone activity of αs1-casein increased as temperature o o increased (more chaperone effective at 37 C than at 25 C), however, under reducing conditions its chaperone activity was more efficient at lower temperatures (more chaperone effective at 25oC o than at 37 C) (Morgan et al. 2005; He et al. 2011). Sakono et al. 2011 showed that αs1-casein not only prevents protein aggregation, but also had the ability to refold some denatured proteins in the presence of ATP. β-casein was also found to inhibit the aggregation of several misfolded proteins as well as solubilise several aggregates that had already formed (Zhang 2005). These findings suggest that casein proteins might be responsible for maintaining the structural integrity of milk proteins by acting as molecular chaperones.

SAP SAP is an oligomeric (five 25 kDa subunits) plasma glycoprotein that exhibits specific calcium- dependent binding to amyloid fibrils. SAP is unique in that it has the ability to refold proteins independently of ATP. This result however, was obtained using a ten-fold molar excess of SAP relative to the client protein (lactate dehydrogenase) and the physiological relevancy of this is yet to be determined (Coker et al. 2000). More research needs to be conducted before SAP can be definitively classified as an EC (Yerbury et al. 2005a).

1.6 New candidate ECs under investigation The list of ECs continues to grow. Discovering and characterising new ECs that reside within specific tissues and bodily fluids is particularly important as PDDs can affect many areas of the body. Utilising resident ECs could be useful in developing therapies to treat or prevent the progression of PDDs.

1.6.1 Thyroglobulin Thyroglobulin (Tg) is a member of the type-B carboxylesterase/lipase family. It is a secreted homodimeric glycoprotein with a mass of 660 kDa (330 kDa per monomeric unit) (Pitt-Rivers 1976), a carbohydrate content of ~ 10%, and a sedimentation coefficient of 19S (Feldt-Rasmussen 1983; Schneider and Edelhoch 1970). Each human Tg (hTg) monomer consists of 2749 amino acid residues (after cleavage of the 19-residue signal peptide), 67 of them being tyrosyl residues that can be iodinated during thyroid hormone synthesis (Dunn and Dunn 1999). Tg is specifically 22

Chapter 1 expressed by thyroid follicular cells (Moravej et al. 2010) and is the largest and most abundant protein found in the thyroid (approximately 75% of total colloid protein content and it can reach to concentrations higher than 100 mg/mL) (Vali et al. 2000; Gentile et al. 1998). Tg is also the only mammalian protein to carry significant amounts of iodine (~ 70 – 80% of the body’s iodine is stored in the thyroid) (Ahad and Ganie 2010), which is required for its role in the synthesis of the thyroidal hormones thyroxin (T4) and triiodothyronine (T3) (Rivolta and Targovnik 2006). No crystallographic structure of Tg is available, however analysis of the amino acid sequence has predicted several structural features. Each hTg monomer is comprised of two distinct regions: (i) region I-II-III (made up by the first ~ 2170 residues) and (ii) a C-terminal acetylcholinesterase- like (ChEL) domain. Region I-II-III makes up ~ 80% of the subunit and is a contiguous disulfide- rich region comprised of multiple repeat domains with 122 cysteines at conserved positions (stabilising the subunit with more than 60 intradomain disulfide bonds (de Crombrugghe and Edelhoch 1966; Pit-Rivers 1976; Lee et al. 2008). The ChEL domain consists of approximately 520 amino acid residues and exhibits 31% identity and 47% similarity to acetylcholinesterase (Di Jeso and Arvan 2016; Park and Arvan 2004; Lee et al. 2008; Swillens et al. 1986; Ludgate et al. 1986).

Tg is the only protein precursor to the thyroidal hormones T3 (carries 3 iodine molecules) and T4 (carries 4 iodine molecules). After expression in the thyroid follicular cells, it is secreted into the lumen of the follicle, known as the colloid, in highly cross-linked form with many disulfide bridges (Dunn and Dunn 1999; Dedieu et al. 2011). Once secreted, Tg acts as a substrate to thyroid peroxidase, an apical membrane-bound glycoprotein which catalyses the iodination and subsequent conjugation of adjacent iodinated tyrosyl residues within the Tg molecule (Rawitch et al. 1983). The most heavily iodinated and important site for hormone synthesis occurs within the first 130 amino acids of the Tg molecule (Park and Arvan 2004; Vono-Toniolo et al. 2005). Although not all hormonogenic tyrosines within the Tg molecule are known, five have been identified and correspond to resides 5 (the major T4-forming site), 1290, 2553, 2567, and 2746 (the main T3-forming site) (Gentile and Salvatore 1993; Dunn et al. 1991). After conjugation, the iodinated protein is stored in the colloid (Berndorfer et al. 1996). Of the entire Tg molecule, only 20 – 40 residues become iodinated, and of these, only 4 – 16 are coupled to form thyroid hormones. As iodine incorporation into Tg and the coupling of tyrosine residues are the only means by which mammals store iodine and thyroid hormones, storing large quantities of Tg in the follicular lumen is necessary to maintain constant levels of iodine, T3, and T4 (Saber-Lichtenberg et al. 2000). When thyroid hormones are needed, Tg undergoes endocytosis and is transported to the lysosome for proteolytic cleavage. Although it has been difficult to isolate the proteolytic fragments of Tg

Chapter 1 under native conditions (e.g. without the use of reducing agents) for structural study due to the complex network of disulfide linkages within the Tg molecule (Veneziani et al. 1999), it has been hypothesised that lysosomal proteolysis of Tg occurs in two sequential steps: (i) early and selective cleavage to release thyroid hormones and (ii) delayed and complete degradation of the entire Tg molecule (Rousset et al. 2015). Once cleaved from Tg, the hormones are secreted into the blood stream where they assist in regulating growth, brain development and metabolism (Figure 1.8) (Dunn 1985; Dunn and Dunn 1999; de Crombrugghe and Edelhoch 1966; Pit-Rivers 1976).

Figure 1.8 Thyroid hormone synthesis. The secretion of newly synthesised thyroglobulin from the follicular cell into the follicle lumen (top left). In the lumen, iodine is oxidised to molecular iodine and bound to tyrosyl residues of Tg (one or two per residue). Once iodinated, adjacent tyrosyl residues are conjugated (bottom left). The protein is then taken back into the cell where the hormones are cleaved from the polypeptide then secreted into the blood stream (bottom right; Walter 2003).

Until recently, no other physiological roles for Tg were known. It has been suggested that Tg has regulatory, suppressive roles for thyroid-specific gene expression, including its own expression (Luo et al. 2014; Suzuki et al. 1999) and that Tg may have additional extrathyroidal functions. Traditionally, Tg was considered to be exclusively produced and function within the thyroid gland, and serum Tg was only clinically important for the diagnosis of thyroid diseases, especially for detecting the reoccurrence and metastasis of thyroid cancer post thyroidectomy (Ahn et al. 2013). The observations that (i) putative Tg receptors are expressed in non-thyroidal cells (e.g. the asioglycoprotein receptor expressed in hepatocytes), and (ii) Tg binds to non-thyroid follicular

Chapter 1 cells (Luo et al. 2014), are consistent with Tg having physiological roles in other parts of the body. However, the concentration of Tg in serum is extremely low in healthy, euthyroid individuals (ranging between 1.6 – 30 ng/ml). The level of Tg found in serum is influenced by a person’s genetics, iodine intake, smoking status, pregnancy, and gender, resulting in a broad and constantly fluctuating range of concentrations (Wang et al. 2016a; Premawardhana et al. 1994; Bertelsen and Hegedus 1994; Knudsen et al. 2001).

It was recently suggested that Tg may have both intramolecular and EC activity. Intramolecular chaperones are encoded in the primary sequence of the protein as either an N-terminal (type I) or C-terminal (type II) sequence extension. This sequence is typically not part of the primary functional domain, however, it is essential for and facilitates the folding of the primary functional domain (Chen and Inouye 2008). Congenital hypothyroidism is a serious disease caused by mutations in the Tg gene affecting the ChEL domain of Tg (Lee et al. 2008; Macchia et al. 1998). Cellular responses seen in patients suffering from congenital hypothyroidism include dilation of the ER, induction of ER chaperone proteins, and activation of ER stress signalling pathways and ER-associated degradation of the mutated Tg (Baryshev et al. 2004; Kim et al. 1996; Menon et al. 2007). Lee et al expressed two recombinant Tg fragments (truncated Tg missing the ChEL domain and a tagged ChEL domain peptide) in cells. When truncated Tg (only containing region I-II-III) was expressed, the protein did not fold properly and became trapped within the ER. However, when truncated Tg was co-expressed with the ChEL domain peptide, truncated Tg was folded and transited correctly through the ER and Golgi, suggesting that the ChEL domain of the Tg molecule may act as an intramolecular chaperone. These results indicate that the ChEL domain of Tg is required for the proper folding and secretory clearance of Tg, however, more research on the structure of Tg and the putative chaperone activity of the ChEL domain would be valuable.

It was first discovered that Tg might have EC activity while using it as a non-chaperone control protein in a protein aggregation assay at the University of Wollongong. Tg was incubated with the client protein creatine phosphokinase (CPK) at 43oC. In this assay Tg did not behave as expected, but instead decreased the aggregation of CPK. The experiment was subsequently repeated, using varying concentrations of Tg with a constant concentration of CPK. The results indicated a concentration-dependent inhibition of CPK aggregation by Tg (Constantinescu, unpublished data). Further study showed that Tg can inhibit the aggregation of a wide range of amorphously aggregating and amyloidogenic client proteins in a concentration-dependent manner. In addition, it was shown that Tg forms HMW complexes with aggregating citrate synthase (CS) and the complexes bind to the surfaces of two different macrophage cell lines (Frazier 2012). Like CLU,

Chapter 1

Hp, and α2M, Tg is a secreted glycoprotein linked via disulfide bonds and is the most abundant extracellular protein in the thyroid lumen. Previous results suggested that Tg has EC activity, however, further investigations were needed to clearly establish this. Interestingly, several diseases of the thyroid (amyloidosis of the thyroid, thyroid goitre, and TMC) are characterised by amyloid deposits within the thyroid and surrounding tissue (Erickson et al. 2015; Nessim and Tamilia 2005; Khurana et al. 2004; Aydin et al. 2016; Westermark 1975; Pai et al. 2010; Kimura et al. 1997). If the chaperone activity of Tg operates in vivo, this may be relevant to the future development of therapies for thyroid-associated PDDs.

1.6.2 Neuroserpin Neuroserpin (NS), also called protease inhibitor 12, or PI12 according to the accepted serine protease inhibitor (SERPIN) nomenclature, is a member of the SERPIN superfamily (Silverman et al. 2001). NS is a secreted glycoprotein predominantly expressed in the central and peripheral nervous systems during development (neurogenesis) and in the adult brain in areas where synaptic changes are associated with learning and memory (synaptic plasticity) (Galliciotti and Sonderegger 2006; Takehara et al. 2009). The protein is comprised of 410 amino acid residues and has a mass of 46 kDa (Schrimpf 1997; Miranda and Lomas 2006). Like the other SERPIN family members, NS shares the typical, conserved “SERPIN structure” with a tertiary structure characterised by three β-sheets (A, B, and C), nine main α-helices, and a long and flexible exposed reactive centre loop (RCL), which is exposed to the solvent and acts as a pseudo substrate for the protease tissue-type plasminogen activator (tPA) (Figure 1.9) (Fabbro and Seeds 2009; Belorgey et al. 2007; Ricagno et al. 2009; Miranda and Lomas 2006). Once tPA binds to the RCL of NS, the protease cleaves the RCL at the P1–P1′ peptide bond (predicted to be between residues Arg362 and Met363) (Hastings et al. 1997; Srikeerthana and de Causmaechker 2010).

Chapter 1

P1 P1’

RCL

s3A

s2A

s6A s5A s2A

Figure 1.9 Ribbon representation of NS structure. β-Sheet A is depicted in yellow, Sheet B in blue and Sheet C in cyan; α-helices are red, and loops are green. The individual strands of β-Sheet A are labelled as s1A, s2A, s3A, s5A and s6A. The P1-P1’ tPA cleavage site is indicated on the RCL (Ricagno et al. 2009).

Cleavage of the RCL results in significant conformational changes within the NS molecule: (i) the RCL upstream of the cleaved peptide bond (the P1’ strand) is inserted between strands 3 and 5 of β-sheet A as the “new” strand 4 (Figure 1.10), a process known as β-sheet expansion (Gooptu and Lomas 2009; Huntington et al. 2000; Huntington 2011) and (ii) tPA, which is covalently bound to the P1 residue of the RCL, is translocated from the proximal to the distal end of the SERPIN (~70 Å away), opposite to the original location of the intact RCL and resulting in the formation of a covalently-linked acyl-enzyme intermediate (Ricagno et al. 2009; Takehara et al. 2009; Saga et al. 2016; Knaupp and Bottomley 2009; Devlin and Bottomley 2005).

Chapter 1

s4A s3A s2A

s6A s1A s5A

Figure 1.10 Ribbon representation of cleaved NS. β-Sheet A is depicted in yellow, β-sheet B in blue and β-sheet C in cyan; α-helices are red, and loops are green. The individual strands of β-sheet A are labelled as s1A, s2A, s3A, s5A and s6A. Once tPA binds to NS the RCL is cleaved at the P1-P1’ peptide bond. tPA remains covalently bound to the P1 residue (not shown in the figure) while the remaining RCL (starting with the P1’ residue) is inserted into β-sheet A between s3A and s5A as the new fourth strand (depicted by the green strand labelled s4A in the figure; Ricagno et al. 2009).

These significant conformational changes in the NS molecule require considerable structural flexibility and result in the inhibition of tPA through deformation of its catalytic triad; as a result of translocation tPA loses ~ 30% of its structure (Huntington et al. 2000; Gooptu and Lomas 2009). For other serpins, deacylation (the dissociation of the SERPIN-protease complex) typically takes weeks, however, in the case of NS and tPA, the acyl-enzyme intermediate is relatively unstable and short-lived. Several studies report that NS-tPA complexes dissociate within minutes in vitro (Barker-Carlson et al. 2002; Ricagno et al. 2009; Lee et al. 2015a; Lee et al. 2015b). Upon dissociation of the NS-tPA complex, active tPA is released, however, NS permanently remains in a stable, loop-inserted “cleaved” conformer, subsequently unable to interact with tPA again.

Chapter 1

Interestingly, like other SERPINs, the native NS structure is considered to be metastable and sometimes referred to as a “folding intermediate” on the pathway to a more thermodynamically favourable conformation (Whisstock and Bottomley 2006). It is only upon the insertion of the RCL into β-sheet A that a significant decrease in free energy occurs and a more thermodynamically stable structure is achieved. Taken together, the intrinsic structural flexibility and inherent metastability of the NS native conformation render the protein susceptible to mutations that result in the formation of dysfunctionally folded NS (Saga et al 2016; Knaupp and Bottomley 2009). Site mutations in SERPIN proteins can lead to diseases, collectively known as “serpinopathies”, and are characterised by the formation of either latent SERPINs or SERPIN polymers (both of which are inactive as protease inhibitors due to the inaccessibility of the RCL) (Ricagno et al. 2010). Latent serpins form when the intact RCL inserts itself as the fourth strand of β-sheet A (as described above), creating a stable, inactive conformer. Although latent SERPINs are implicated in some serpinopathies, SERPIN polymers are the more prominent species known to cause disease (Knaupp and Bottomley 2009).

The initiation of SERPIN polymerisation, mechanisms of elongation, and the structural organisation of SERPIN polymers are still matters of debate and not fully understood (Chiou et al. 2009; Yamasaki et al. 2008; Takehara et al. 2010; Krishnan and Gierasch 2011). Some studies report that the RCL of one SERPIN molecule is simply inserted into the β-sheet A of a neighbouring SERPIN molecule, whilst others report a more complex domain-swapping mechanism (Noto et al. 2015a; Ricagno et al. 2010; Wisstock and Bottomley 2006; Knaupp and Bottomley 2009). A better understanding of the mechanisms behind SERPIN polymerisation (both the initiating step and polymer elongation) may prove useful in developing therapies that prevent serpinopathies characterised by the deposition of SERPIN polymers.

There are currently several known pathological mutations in the NS gene (SERPINI1) that lead to a dementia-like serpinopathy known as familial encephalopathy with NS inclusion bodies (FENIB). These mutations include: Ser49Pro, Ser52Arg, His338Arg, and Gly392Glu (Silverman et al. 2010; Belorgey et al. 2007), with each mutated variant having a different propensity to polymerise (and the degree of polymerisation is directly related to disease severity). In vitro, these mutations lead to the development and accumulation of both intra- and extracellular deposits of NS polymers (Noto et al. 2015a; Davis et al. 2002; Davies et al. 2009; Giampietro et al. 2017). Although it is clear that NS polymers are associated with FENIB, it is not known what effects polymerisation have on the function of secreted NS. While NS polymerisation is most often due to mutations in the NS gene, it should be noted that wild type NS can also polymerise under

Chapter 1 conditions of thermal stress (most studies induce NS polymerisation at 50°C) and under acidic conditions (Miranda and Lomas 2006; Noto et al. 2012; Takehara et al. 2010; Santangelo et al. 2012; Ricagno et al. 2009).

The overall physiological role of NS in the human nervous system is still unclear. Based on in vitro models, it has been proposed that NS (i) plays an important role in axonogenesis and synaptogenesis during development and synaptic plasticity in adults (Ricagno et al. 2009; Parmar et al. 2002), (ii) has neuroprotective effects against ischemic stroke (Cole et al. 2007), (iii) lowers the cytotoxicity of Aβ aggregates in the AD brain (Kinghorn et al. 2006), and (iv) regulates the spreading of kainic acid-induced seizures (Yepes et al. 2002). Most of these suggested functions (except NS binding Aβ aggregates) appear to be linked with its ability to bind and inhibit tPA (i.e. regulation of protease activity in the neurovascular system) (Cole et al. 2007). Although these proposals are supported by experimental evidence, it is important to note that the in vivo mechanism by which NS preferentially complexes with tPA, and the overall effects of complex formation, are not entirely understood (Galliciotti and Sonderegger 2006). More research is required in order to fully understand the physiological role of NS in the human nervous system.

It was shown that NS binds to Aβ, preventing maturation of fibrils and lowering their cytotoxicity on rat pheochromocytoma (PC12) cells and E14 rat embryonic cortical neurons in a concentration- dependent manner (Kinghorn et al. 2006). Though Kinghorn showed that NS can bind Aβ1-42 at the N-terminus or middle parts of the peptide, the mechanism by which this complex forms is still not known. This was an interesting discovery and led to the investigation of NS as a putative EC. A series of aggregation assays at the University of Wollongong revealed that NS has in vitro chaperone activity and dose-dependently reduces amyloid formation by several proteins, including coiled-coil β (ccβ) and Aβ1-42 (Whiten 2011). Whiten also showed that NS was able to inhibit the amorphous aggregation of CS and alcohol dehydrogenase (ADH), however it was significantly less effective than CLU at the same molar concentration.

1.6.3 Transthyretin Human TTR (hTTR), previously named thyroxin binding prealbumin, is a highly conserved, 55 kDa homotetrameric protein comprised of four identical monomeric subunits (A, B, A’, & B’). Each monomer consists of 127 amino acid residues (after proteolytic cleavage of the 20-residue N-terminal signal sequence) and is organised into a “β-sandwich conformation” (Kanda et al. 1974; Buxbaum and Johansson 2017). This includes an inner β-sheet (strands DAGH), an outer β- sheet (strands BCEF), and an α-helix (consisting of nine residues) (Figure 1.11). The inner and

Chapter 1 outer β-sheets lie orthogonal to one another and approximately 10 Å apart. Two monomers interact to form a dimer via hydrogen bonding and two dimers interact to form the tetramer via additional hydrogen bonds and hydrophobic interactions (Prapunpoj and Leelawatwattana 2009; Vieira and Saraiva 2014; Buxbaum and Reixach 2009). The inner β-sheets between two dimers form the two T4 binding pockets within the TTR molecule (Wang et al. 2007; Myung et al. 2013).

Monomer A

α-helix A

T4 Binding T4 Binding Pocket Pocket

H G α-helix A’ D A α-helix B’

E B C Monomer A’ Monomer B’

A B

Figure 1.11 Ribbon representation of homotetrameric TTR structure. A) TTR is made up of four identical monomers (A, B, A’ and B’) shown in blue, yellow, green and red respectively. The two T4 binding channels that form between the inner β-sheets of the AB and A’B’ dimers are also indicated by the grey shaded area. B) Depicts the same molecule rotated 90° in the horizontal plane. The β-strands of each monomer are named A-H (labelled on the green A’ monomer), with the inner β-sheet comprised of strands DAGH and the outer β-sheet being comprised of strands CBEF. The alpha helices associated with each monomer are also indicated (adapted from Wang et al. 2007).

hTTR is mainly synthesised in the liver, however, additional sites of synthesis include cells of the blood–CSF barrier, the choroid plexus of the brain, retinal pigment epithelium, the pancreas, and neurons under stress (Vieira and Saraiva 2014; Liz et al. 2010; Wang et al. 2014). TTR is highly multifunctional (Prapunpoj and Leelawatwattana 2009) and is best known for distributing T3 and, to a lesser extent T4, to the bloodstream and CSF of vertebrates and indirectly transporting vitamin A (retinol) by binding retinol-binding protein. TTR is the only known distributor and regulator of T4 within the CSF. T4 enters the brain by either diffusing across the blood-brain barrier or the blood-CSF barrier and is sequestered by TTR (two T4 binding sites per TTR molecule) (Feldt- Rasmussen and Rasmussen 2007). TTR carries only one T4 molecule per TTR tetramer and when T4 is bound to TTR, it cannot be accessed by tissues (Wirth et al. 2014; Liz et al. 2010). 31

Chapter 1

Several other functions for TTR, specifically in the nervous system, have recently emerged. Owing to its function in thyroid hormone and retinol distribution (Vieira and Saraiva 2014), TTR has been found to have important, indirect roles in processes relating to behaviour (Sullivan et al. 1999; Sullivan et al. 2006), cognition (Sousa et al. 2007), neuropeptide maturation (Mar et al. 2009), neurogenesis, nerve regeneration, axonal growth (Fleming et al. 2007; Fleming et al. 2009) and metabolism (Ingenbleek and Young 2002; Wolf 1995). TTR also has several important clinical roles. TTR serum concentration has been utilized as a marker of nutritional and inflammatory status in a variety of conditions (Roma et al. 2012; Ding et al. 2014; Baeten et al. 2006).

TTR is also implicated in a series of diseases characterised by the deposition of TTR aggregates (in the form of amyloid fibrils) in various tissues. There are four classes of TTR amyloidoses: (i) senile systemic amyloidosis (sometimes referred to as senile cardiac amyloidosis, SSA), (ii) familial amyloidotic polyneuropathy (FAP), (iii) familial amyloidotic cardiomyopathy (FAC), and (iv) central nervous system selective amyloidosis (CNSA). In all four cases, the amyloid fibrils are formed from monomeric TTR species, either from the sporadic dissociation of the wild type TTR tetramer (in the case of SSA), or due to genetic mutations that cause thermal or kinetic destabilisation of the tetramer (linked to the development of FAP, FAC, and CNSA) (Johnson et al. 2005; Buxbaum and Johansson 2017; Li and Buxbaum 2011; Hershman Babbes et al. 2008). SSA is the most common TTR-associated amyloidosis and is characterised by the sporadic deposition of wild type TTR fibrils in the myocardium. The exact cause for disease onset and the dissociation of wild type TTR tetramers remains elusive, yet it appears to be related to the aging process (Pinney et al. 2013; Ruberg and Berk 2012; Hershman Babbes et al. 2008). It has been shown that proteolysis of TTR leads to the generation of TTR amyloid, which could cause the onset of disease (Suhr et al. 2017; Ihse et al. 2011). SSA mainly affects males over the age of 60 and leads to cardiac dysfunction and ultimately death from cardiac arrest (Westermark et al. 2003). There are currently more than 120 mutations responsible for the development of the tissue-specific diseases FAP, FAC, and CNSA (Lai et al. 2015; Yamamoto et al. 2018). These diseases are typically more severe than SSA and the onset of disease does not necessarily correlate with age. FAP and FAC are characterised by the deposition of mutant amyloidogenic TTR (secreted by the liver) in the peripheral and autonomic nerves and heart (Li and Buxbaum 2011). CNSA is a rare disease characterised by deposition of mutant amyloidogenic TTR (secreted by the choroid plexus) in the central nervous system (CNS) (Faria et al. 2015; Sekijima et al. 2005). If left untreated, these diseases lead to organ dysfunction and death, often within ten years of disease onset (Johnson et al. 2005). Additionally, TTR amyloid deposits have also been found associated with conditions

Chapter 1 such as preeclampsia, carpel tunnel, osteoarthritis, and spinal stenosis (Buxbaum and Reixach 2009; Chen and Zhang 2011; Westermark et al. 2014; Yanagisawa et al. 2015).

Interestingly, although TTR has a propensity to form amyloid itself, it has also been shown to bind to Aβ and inhibit Aβ fibril formation, ultimately decreasing Aβ cytotoxicity in vitro. Several studies have investigated the interaction between TTR and Aβ (Schwarzman et al. 1994; Costa et al. 2008; Du and Murphy 2010), however, the means by which TTR supresses Aβ amyloid formation is still not entirely clear. One possibility is that TTR has substrate-specific proteolytic activity towards Aβ. Several studies have suggested that TTR may inhibit Aβ fibrillogensesis by both direct binding to Aβ (Buxbaum et al. 2008) and via a proteolytic activity that can cleave Aβ in several positions (Liz et al. 2004; Liz et al. 2009; Liz et al. 2010; Geneste et al. 2014), and the ability of TTR to cleave Aβ is inhibited by Kunitz protease inhibitor (Costa 2009). However, the evidence for the proteolytic activity of TTR is limited and may reflect the presence of a protease that co-purifies from human plasma with the TTR used in these studies. As TTR can inhibit Aβ aggregation, it would be interesting to investigate whether it can inhibit the aggregation of other proteins, thereby exhibiting broad-spectrum chaperone activity.

1.7 A model for the clearance of extracellular protein aggregates The involvement of receptor-mediated endocytosis in the removal and intracellular degradation of extracellular proteins, other macromolecules, and cellular debris is a fundamental biological process and has been under investigation since the late 1970s (Goldstein et al. 1979). There are many receptors, expressed by various cells types and tissues that assist in the removal of unwanted extracellular molecules, such as the clearance of Aβ from the brain by LRP1 (Sagare et al. 2007). Three well-established ECs (CLU, α2M, and Hp) are known to bind specific endocytic cell-surface receptors (Table 1.2) and have been implicated in receptor-mediated endocytosis (Wilson and Easterbrook-Smith 2000; Djakiew et al. 1984; Narita et al. 1997; Madsen et al. 2001). This provides a possible mechanism for ECs to clear misfolded proteins from body fluids. The current theory is that ECs bind to hydrophobic regions of stress-induced, misfolded proteins to form stable, soluble, HMW complexes (currently there is only evidence to support this theory for amorphously aggregating client proteins) (Wyatt and Wilson 2010; Wyatt and Wilson 2013). These complexes can then be directed to specific cell-surface receptors for internalisation and degradation, thus preventing the accumulation of protein aggregates in the extracellular space (Yerbury et al. 2005a; Yerbury and Wilson 2010). It has been shown that once internalised, clusterin-client and Hp-client complexes are targeted to lysosomes for proteolytic degradation (Wyatt et al. 2011; Sultan et al. 2013). Although the receptors listed below are known to bind ECs, their involvement in the 33

Chapter 1 chaperone-mediated clearance of misfolded proteins from body fluids has not been thoroughly investigated previously. It is still unknown how ECs interact with misfolded proteins to form complexes and which specific receptors are involved in their clearance. Lakins et al. 2002 report data suggesting that CLU may have separate binding sites for interactions between misfolded proteins and receptors.

Table 1.2 Examples of cell surface receptors that bind known ECs. The receptors listed may play a role in clearing EC-stressed protein complexes. * Denotes putative ECs.

EC Receptor Reference Megalin (LRP2 or Glycoprotein 330) Wilson and Easterbrook-Smith 2000 Very low-density lipoprotein receptor (VLDLR) Leeb et al. 2014 Apolipoprotein E receptor 2 (ApoER2) Leeb et al. 2014 CLU Plexin A4 Kang et al. 2016 Triggering receptor on myeloid cells 2 (TREM2) Yeh et al. 2016 Unidentified scavenger receptors Wyatt et al. 2011 α2M LRP1 Kounnas et al. 1992; Barcelona et al. 2011 Hp CD163 Madsen et al. 2001 Tg* Megalin Marino et al. 2000; Lisi et al. 2006 LRP1 Alemi et al. 2016 TTR* Megalin Sousa et al. 2000; Gomes et al. 2016

Whether or not the receptors listed in Table 1.2 are actually involved in clearing EC-client protein complexes from body fluids remains to be established. Megalin, LRP1, ApoER2, and VLDLR are all members of the low-density lipoprotein (LDL) receptor family. CLU has been shown to bind all of these receptors (except LRP1) and is internalised via LRP2 (Kounnas et al. 1995; Leeb et al. 2014; Riaz et al. 2017; Calero et al. 1999; Bartl et al. 2001; Lakins et al. 2002), LRP1 binds and internalises protease-activated α2M (Kounnas et al. 1992; Barcelona et al. 2011; Mikhailenko et al. 2001; French et al. 2008), and Tg and TTR (putative ECs) both bind and undergo endocytosis via megalin (Marino et al. 2000; Sousa et al. 2000; Gomes et al. 2016; Lisi et al. 2006; Fleming et al. 2009). Several studies also show that CLU-Aβ complexes bind megalin (Zlokovic et al. 1996) and that CLU, α2M, and TTR can facilitate the clearance of Aβ by forming an EC-Aβ complex and binding to megalin (CLU-Aβ) and LRP1 (α2M-Aβ and TTR-Aβ) (Hammad et al. 1997; Narita et al. 1997; Alemi et al. 2016). In addition, Bajari et al. 2003 demonstrated that CLU and CLU-leptin complexes bind to ApoER2 and VLDLR, and Byun et al. 2014 showed that CLU- leptin complexes bind and are internalised via LRP2.

Chapter 1

Other receptors of interest include (currently unidentified) scavenger receptors, plexin A4, and TREM2. In 2011, Wyatt et al. found that pre-incubation of cells with fucoidan, a known inhibitor of scavenger receptors, significantly inhibited the binding of several CLU-client protein complexes, thus indicating that some interactions between CLU-client complexes and cells were via scavenger receptors. Hp, another established EC, is known to bind the scavenger receptor CD163 (Madsen et al. 2001). Separate studies show that (i) Hp binds and is internalised via CD163 (Graversen et al. 2002), and (ii) Hp forms soluble complexes with misfolded β2M (an amyloidogenic protein), which then bind to the surface of macrophage cells, and are subsequently internalised and degraded in lysosomes (Sultan et al. 2013). There is, however, currently no direct evidence to establish that Hp-misfolded protein complexes bind to, and are cleared, via CD163. Plexin A4 and TREM2 have recently been described as novel receptors for CLU (Kang et al. 2016; Yeh et al. 2016) and mutations in both plexin A4 and TREM2 have been linked with the development of AD (Zhou et al. 2018; Ulrich et al. 2017; Jin et al. 2014; Wang et al. 2016b; Xiang et al. 2016). Kang et al. 2016 showed that CLU binds plexin A4 in vitro and that there is an inverse correlation between CLU concentration in the CSF and the expression of plexin A4. Brain tissue from patients with AD showed an elevated concentration of CLU, but significantly decreased levels of plexin A4 (by about 50%). It was also observed that plexin A4 knockdown mice showed both behavioural and memory deficits, raising the possibility that memory loss in AD may be associated with a decrease in plexin A4 expression. It is also possible that CLU uses plexin A4 as a conduit to clear misfolded proteins (especially Aβ) from the extracellular space. However, further work is needed to test this hypothesis, as currently it is not known whether CLU-client protein complexes bind to and are internalised by plexin A4. TREM2 is another potential candidate receptor for clearing CLU-client protein complexes from the extracellular environment. One study showed that TREM2 facilitates the cellular uptake of CLU-Aβ complexes, suggesting that TREM2 and CLU may together work to clear toxic Aβ from the brain, however, these results must be interpreted with caution as this study used lipidated CLU in their binding assays and could not detect unlipidated CLU binding to TREM2 (Yeh et al. 2016).

These findings suggest possible EC and receptor combinations that may work together to remove toxic misfolded proteins from body fluids. There are many different cell surface receptors and the expression of these changes with cell type (Murdoch and Finn 2000), cell growth stage (Sallusto et al. 1998), physiological location (or the microenvironment surrounding the cell), cell function, and during various physiological stresses (e.g. oxidative stress (Nie et al. 1998; Keidar et al. 2002), or inflammation (Wu et al. 2012)). Due to the variability and transient nature of cell surface receptor expression, identification of all receptors involved in the uptake of EC-client protein

Chapter 1 complexes will be challenging and take some time. Nevertheless, this information is likely to play an important role in the eventual development of new therapies for PDDs. Identification of the specific receptors involved in receptor-mediated endocytosis of EC-stressed client complexes will progress our understanding of extracellular proteostasis and identify the critical cells and tissues involved in this process. Once these have been identified, targeted manipulation of receptor expression could form part of a viable therapy for PDDs.

1.8 Experimental objectives The focus of this project is to identify new ECs and investigate their roles in inhibiting protein aggregation. In addition, this study will focus on the hypothesis that EC-stressed client protein HMW complexes are directed to specific cell surface receptors and internalised to the lysosome for degradation. The specific aims of this study are:

(i) Characterise the chaperone activity of Tg, NS and TTR. (ii) Investigate the ability of the putative ECs to form HMW complexes with amorphously aggregating clients. (iii) Investigate the ability of EC-client protein complexes to interact with specific cell surface receptors.

Current drug therapies for PDDs, such as AD, are limited to alleviating symptoms of disease rather than targeting the direct causes of the disease (Wyatt et al. 2012). Further understanding of the cellular and molecular mechanisms causing the onset of disease, as well as continued investigation of the structures, roles and mechanisms of ECs could lead to innovative therapies for disease prevention and control. Understanding of the chaperone activities of Tg, NS, TTR, and other ECs could prove to be useful in the future design of disease therapies.

Chapter 2

Chapter Two General Materials & Methods

2.1 Materials Chemicals: All chemicals were of analytical grade and prepared in MilliQ water (Millipore; Billerica, MA, USA) unless otherwise stated. Methanol, ethanol, and NaCl were purchased from Univar Ltd. (Richmond, BC, Canada). Novagen’s Overnight Express™ Instant Terrific Broth (TB) autoinduction media and BugBuster® Master Mix Protein Extraction Reagent glycerol and

KH2PO4 were from Merck (Baywater, VIC, Australia). cOmplete and cOmplete ethylenediaminetetraacetic acid (EDTA)-free protease inhibitor cocktail tablets, Bacto-tryptone, yeast extract, agar, and ampicillin were from Roche (Dee Why, NSW, Australia). Glacial acetic acid was from Panreac (Barcelona, Spain). 1-O-n-Octyl-β-D-Glucopyranoside (β-Dog) was from AG Scientific, Inc. (San Diego, CA, United States). All other chemicals and reagents were purchased from Sigma Aldrich (St. Louis, MO, USA).

Bacterial Cell Lines: Library Efficiency® DH5α™ E. coli and BL21(DE3) chemically competent E. coli were purchased from Thermo Fisher Scientific (Scoresby, VIC, Australia). NiCo21(DE3) competent E. coli were purchased from New England BioLabs Inc. (Ipswich, MA, USA).

Proteins: Alpha-lactalbumin (α-Lac), bovine serum albumin (BSA), CS, CPK, lysozyme, ovalbumin (Oval), ovotransferrin (OT), bovine Tg (bTg) and thrombin were purchased from

Sigma Aldrich. Aβ1-42 and Cal were purchased from ChinaPeptides (Shanghai, China). Purified recombinant human α-syn and Coiled-coil beta “w” (ccβw), a modified version of the ccβ peptide that has an additional tryptophan at its N terminus (as described in Kammerer et al. 2004), was purchased from GenScript (Piscataway, NJ, United States). Purified human tPA was a kind gift from Associate Professor Marie Ranson (Illawarra Health & Medical Research Institute, University of Wollongong, Australia). Purified SOD1 was a kind gift from Dr. Luke McAlary (Illawarra Health & Medical Research Institute, University of Wollongong, Australia). hTg was purchased from CALBIOCHEM (subsidiary of Merk, Bayswater, VIC, Australia). Both hTTR and monomeric TTR (mTTR), a stable monomeric mutant of hTTR that carries F87M and L110M mutations and is unable to form a tetramer, were a kind gift from Professor Buxbaum (The Scripps Research Institute, CA, United States). Synthetic peptides corresponding to a region on NS and TTR (and a scrambled control peptide) were purchased from GenicBio Limited (Shanghai, China). The purification of CLU, glutathione-S-transferase (GST) tagged receptor-associated protein

Chapter 2

(RAP) fusion protein (RAP-GST), and mutant Cys-less chloride intracellular channel 1 (Clic1) is described below.

Antibodies: All antibodies used in this study are described within the text, with the exception of G7 and 41D hybridoma cell supernatants (mouse, monoclonal anti-human clusterin) and DNP9 hybridoma cell supernatant (a species matched negative control for G7 and 41D). The G7 mouse hybridoma cell line was a kind gift from Dr. B. Murphey (St. Vincent’s Hospital, Melbourne, VIC, Australia) and the mouse hybridoma cell lines 41D and DNP9 were designed and produced by Senior Professor Mark Wilson as described in (Humphreys et al. 1999). G7, 41D and DNP9 were cultured in Dulbecco’s modified eagle medium (DMEM):F-12 (Thermo Fisher Scientific) supplemented with 10% heat-inactivated fetal bovine serum (FBS) (Bovagen; SD, United States) and maintained in an incubator at 37°C with humidified air containing 5% (v/v) CO2. Thawed FBS was heat-inactivated by incubation at 56°C for 30 min and then stored at 4°C. The cultures were expanded by routinely adding culture medium (DMEM:F12 + 10% (v/v) FBS) until the desired volume (approximately 250 mL) was attained. The cultures were subsequently left in the incubator until the majority of the cells had died to maximise antibody yield. Cell cultures were then centrifuged at 300 x g for 5 minutes at room temperature to remove cells and cellular debris and 0.1% (w/v) sodium azide was added to the cell supernatants prior to filtration through a MILLEXÒGV 0.22 μm DuraporeÒ PVDF membrane (Merk). The filtered cell supernatants were then stored at 4°C.

2.2 General methods 2.2.1 Estimation of protein concentration Protein concentration was estimated by measuring the absorbance at 280 nm (A280) in a quartz cuvette using a SpectroMax Plus spectrophotometer (Molecular Devices; San Jose, California, United States) and the respective extinction coefficient for each protein. Alternatively, estimation of protein concentration was determined by a bicinchoninic acid (BCA) assay using methods previously described (Smith et al. 1985).

2.2.2 SDS-PAGE analysis Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was conducted using a Hoefer SE 260 Mighty Small II gel system connected to a BioRad Powepac 300. All SDS gels were comprised of a stacking and resolving gel, 5% and 10% (w/v) bis-acrylamide, respectively. Gels were either short (8.5 x 8.5 cm) or tall (8.5 x 10.5 cm). Samples were prepared under non- reducing conditions by incubating with 1x sample buffer (50 mM Tris-HCl (pH 6.8), 10% (w/v) 38

Chapter 2 glycerol, 2% (w/v) SDS, and 0.01% (w/v) bromophenol blue) or reducing conditions (1x sample buffer with the addition of 5% (v/v) 2-mercaptoethanol) and boiled for 5 min. After denaturation, the samples were loaded onto the gel and electrophoresed at 90V in SDS running buffer (0.025 M Tris, 0.192 M glycine, 0.1% (w/v) SDS, pH 8.5) until the samples crossed into the resolving gel (approximately 20 min), after which, the voltage was increased to 120V and the gel was run until the dye front reached the bottom of the gel. Gels were then stained in 0.1% (w/v) Coomassie™ Brilliant Blue, 40% methanol, and 10% acetic acid and destained with the same solution without Coomassie™ Brilliant Blue (unless otherwise specified). All gels were run with Precision Plus Proteinä Dual Color Standards (BioRad) unless otherwise stated.

2.2.3 Western blot analysis After separation by SDS-PAGE, proteins (1 μg or less) were transferred to BioTraceä NT nitrocellulose membranes (PALL Life Sciences) using a Mini Trans-Blot Cell Western blotting apparatus (Bio-Rad) in Western transfer buffer (0.025 M Tris, 0.192 M glycine, 20% (v/v) methanol, pH 8.5) at 90V for 120 min at 4°C. Afterwards, the membrane was blocked with phosphate buffered saline (PBS; 2.7 mM KCl, 1.75 mM KH2PO4, 135 mM NaCl, 10 mM

Na2HPO4, pH 7.4) containing 5% (w/v) skim milk (PBS/skim milk) for 1 h at room temperature or overnight at 4°C. Primary antibodies at the manufacturer’s recommended dilution were incubated with the membrane in PBS/skim milk for 1 h at room temperature. The membrane was then washed 3 times with PBS containing 0.1% (v/v) Triton X-100 and once with PBS prior to incubation with an appropriate horseradish peroxidase (HRP) conjugated secondary antibody in PBS/skim milk for 1 h at room temperature. The membrane was then washed as described above and proteins were detected using an enhanced chemiluminescence (ECL) SuperSignal Western pico substrate kit (Pierce Biotechnology; Rockford, IL, USA) as per the manufacturer’s protocol. The membrane was imaged using an Amersham imager 600 (GE Healthcare).

2.2.4 Dot blot analysis The methods used for dot blot analysis are identical to those described for Western blot analysis with the exception that purified proteins (0.25 – 0.5 μg) were pipetted directly onto the nitrocellulose membrane instead of being transferred from a gel. The membrane was then treated as described in the previous section.

2.2.5 Protein aggregation assays In vitro protein aggregation assays are commonly used to investigate the ability of putative chaperones to inhibit the aggregation of a wide range of client proteins (Bhuwan et al. 2017; 39

Chapter 2

Wilson and Easterbrook-Smith 2000; Yerbury et al. 2007; French et al. 2008). The methodology for inducing the in vitro aggregation of many amorphously aggregating and amyloidogenic client proteins is well established in the literature. Protein precipitation assays were carried out in clear, non-treated, flat-bottom 384-well microplates (Greiner; Kremsmunster, Austria), unless otherwise stated, using a POLARStar Optima microplate reader (BMG Labtech; Offenburg Germany). For amorphously aggregating client proteins, changes in turbidity (as an indicator of protein aggregation) were monitored by measuring the absorbance at 360 nm (A360) over time. Aggregation of all amorphously aggregating client proteins (except that of Clic1 and BSA) was induced by heat stress; 43°C for CS and CPK, 55°C for GST and 60°C for OT. Clic1 was the only amorphously aggregating client protein to aggregate at 37°C without the use of oxidising/reducing reagents, however, a significantly higher molar concentration of Clic1 was required when compared to other client proteins. BSA was induced to aggregate under reducing conditions with 20 mM dithiothreitol (DTT). Aggregation of amyloidogenic client proteins was monitored by measuring changes in Thioflavin T (ThT) fluorescence intensity in arbitrary fluorescent units (AFU) over time. All amyloidogenic client proteins were incubated at 37°C with the exception of Ab, which was incubated at 30°C.

In all protein aggregation assays, the client was induced to aggregate alone, and together with non- chaperone control proteins or CLU. Lastly, various molar ratios of putative chaperone:client were tested to establish whether or not the putative chaperone had a concentration-dependent effect on client protein aggregation. Typically, a number of non-chaperone control proteins were tested for non-specific effects on protein aggregation under the assay conditions, and one (or more) that showed minimal non-specific effects was selected for further work. Relative to the control proteins, genuine chaperones show a clear and pronounced dose-dependent inhibition of protein aggregation. The samples were loaded onto the microplate (surrounded by a buffer “moat” to ensure even heating of all samples) and then placed into the plate reader (set to room temperature). The plate reader thermostat was then set to the desired temperature, allowing the plate and samples to equilibrate (for approximately 10 min) prior to starting the assay, thus decreasing the risk of condensation forming on the plate seal during the assay. Immediately before starting the assay, the plate was covered using a SealPlateÒ microplate film. All assays were performed in triplicate with 50 μL/well unless otherwise stated.

2.2.6 Mammalian cell culture The following cell lines were obtained from the American Type Culture Collection (ATCC; Manassas, VA, United States) and were cultured as needed: HEK293 (human embryonic kidney 40

Chapter 2 endothelial), LADMAC (murine bone marrow lymphoblast), EOC13.31 (murine microglial), PC- 12 (rat adrenal pheochromocytoma), RAW 264.7 (Abelson murine leukemia virus-induced tumour macrophage), HepG2 (human liver epithelial), and SH-SY5Y (human bone marrow neuroblastoma). The human thyroid follicular epithelial cell line (Nthy-ori 3-1) was purchased from Cellbank Australia (Westmead, NSW, Australia). All cell lines were cultured in media supplemented with heat-inactivated FBS; HEK293, RAW 264.7, and SH-SY5Y were cultured in DMEM:F-12 supplemented with 10% (v/v) FBS and LADMAC, HepG2, and Nthy-ori 3-1 were cultured in Roswell Park Memorial Institute (RPMI)-1640 medium (Thermo Fisher) supplemented with 10% (v/v) FBS. PC-12 cells were cultured in RPMI-1640 containing 5% (v/v) FBS and EOC13.31 cells were cultured in RPMI-1640 supplemented with 10% (v/v) FBS and 10% (v/v) LADMAC-conditioned medium. LADMAC-conditioned medium was prepared as described by the ATCC. All cell lines were maintained in an incubator at 37°C with humidified air containing

5% (v/v) CO2. HEK293 and SH-SY5Y cells were routinely passaged with Trypsin-EDTA containing 0.05% (w/v) trypsin and 0.02% (w/v) EDTA, pH 7 (Gibco) at 37°C for 5 – 10 min. HepG2 and Nthy-ori 3-1 cells were passaged using Trypsin-EDTA containing 0.25% (w/v) trypsin and 0.02% (w/v) EDTA, pH 7 (Sigma-Aldrich) at 37°C for 5 – 10 min. RAW 264.7 and EOC13.31 cells were routinely passaged by scraping in serum-free media and the suspended cells (LADMAC and PC-12) were passaged by decanting the cells and diluting them with culture medium to a suitable density. For all cell-binding experiments, cells were removed from the flask by either gentle scraping (RAW264.7 and EOC13.31) or with Acutase (Sigma-Aldrich) (HepG2) according to the manufacturer’s instructions. Acutase is a cell detachment solution containing proteolytic and collagenolytic enzymes, which are neither mammalian nor bacterially derived. Acutase was used in place of trypsin/EDTA to avoid damaging cell surface receptors during the detachment process. Transfected HEK293 were detached from the surface of plates or flasks using 0.5 strength PBS containing 5 mM EDTA.

For long term storage, cells were resuspended in cold freeze-down medium (50% (v/v) FBS, 10% (v/v) dimethyl sulfoxide (DMSO), 40% (v/v) DMEM:F-12) to a final cell density of 5 x 106 cells/mL. Aliquots (1 mL) of the cell suspension were transferred to sterile cryovials and then placed in a Cryo 1°C freezing container (Nalgene; Sydney, NSW, Australia). The freezing container was stored at -80°C overnight to achieve a freezing rate of -1°C/min and then the vials transferred to a liquid nitrogen storage facility.

Chapter 2

2.2.7 CLU purification CLU was purified from human plasma kindly donated by the Wollongong Public Hospital (Wollongong, Australia). Human blood containing 10 mM sodium citrate (pH 7) was collected from the hospital and centrifuged at 1300 x g for 30 min at 4°C to separate the plasma layer, which was subsequently stored at -20°C. CLU was purified from ~ 0.5 L of human plasma by immunoaffinity chromatography as previously described (Wilson and Easterbrook-Smith 1992). Briefly, frozen plasma was thawed overnight in the presence of a Roche cOmplete protease inhibitor cocktail (per the manufacturer’s recommended dose) and 5 mM EDTA (to prevent clotting) at 4°C. Thawed plasma was first filtered through a Whatmanä glass microfiber GF/Cä filter (GE Healthcare) to remove large debris, followed by filtration through a 0.45 μm cellulose nitrate membrane (Sartorius). The cleared filtrate was then passed through a 15 mL G7 (anti-CLU)- Sepharose column that had been equilibrated in PBS containing 0.05% (w/v) sodium azide (PBS/az), at 0.5 mL/min at 4°C. The column was then washed with 100 mL of PBS/az, followed by 100 mL of PBS containing 0.5% (v/v) Triton X-100 for the removal of lipids. The column was pre-eluted with 100 mL of 0.2 M sodium acetate, 0.5 M NaCl, pH 5 to remove any non-specifically bound proteins. CLU was then eluted with 2 M guanidine hydrochloride in PBS, pH 7.4 and a UV trace monitor was used to collect a fraction containing CLU, which was immediately dialysed against PBS. Post dialysis, the concentration of clusterin was determined spectrophotometrically (A280) using a theoretical extinction coefficient of 0.848 for a 0.1% (w/v) solution (calculated using ExPASy) and CLU purity was checked by SDS-PAGE. Purified CLU was then aliquoted and stored at -20°C. Approximately 40 mg of CLU was obtained from 0.5 L of plasma.

2.2.8 Purification and restriction digestion of the pGEX-2T(GST-RAP) plasmid for the purification of recombinant Schistosoma japonicum GST-tagged human RAP Glycerol stocks of Library Efficiency® DH5α™ cells containing the plasmid, pGEX-2T(RAP- GST), a kind gift from Dr. Yonghe Li (Washington University School of Medicine, WA, United States), were grown on sterile Luria broth (LB) agar plates containing 100 μg/mL ampicillin. A starter culture was then prepared by adding a single bacterial colony to 3 mL of sterile LB containing 100 μg/mL ampicillin and incubating for 8 h at 37°C, shaking at 180 rpm. The starter culture was then added to 120 mL of sterile LB containing 100 μg/mL ampicillin and grown overnight under the same conditions as the starter culture. Post-incubation, the cells were centrifuged at 5,000 x g for 10 min at room temperature. The bacterial cells were then lysed and plasmid DNA was purified using a Promega PureYield™ MaxiPrep System kit, in accordance with the manufacturer’s protocol. DNA concentration and purity were determined spectrophotometrically (DNA concentration, in μg/mL, was determined by multiplying the 42

Chapter 2 absorbance value at 260 nm by 50 and DNA purity was calculated by dividing the absorbance value at 260 nm by the absorbance vale at 280 nm, with an acceptable A260/A280 value ranging between 1.8 and 2).

To confirm that the pGEX-2T plasmid contained the RAP-GST insert, the purified DNA was digested using the restriction enzyme EcoR1 or doubly digested using the restriction enzymes EcoR1 and BamH1 following the manufacturer’s protocol. The plasmid DNA was then analysed on a 1% (w/v) agarose gel, electrophoresed in TAE buffer (40 mM Tris, 20 mM acetic acid, 1 mM EDTA, pH 8) at 80 V for approximately 1 h. The gel was then stained with 1 μg/mL ethidium bromide for 30 min and de-stained in MilliQ water for 15 min prior to being imaged.

2.2.9 Transformation, expression, and purification of recombinant RAP-GST For RAP-GST expression, 50 μL of BL21(DE3) chemically competent E. coli cells were transformed with either 50 ng of the pGEX-2T(RAP-GST) plasmid or 5 μL of sterile water as a negative control. After mixing the competent cells with the plasmid DNA (or water) they were placed on ice for 30 min, heat-shocked at 42°C for 45 s, and placed back on ice for an additional 2 min. Subsequently, 0.3 mL of pre-warmed LB was added to each tube and the cultures were incubated at 37°C for 1 h with shaking at 180 rpm. After incubation, each tube was aliquoted onto pre-warmed LB agar plates (50 μL, 100μL, & 200 μL per plate) containing 100 μg/mL ampicillin. The samples were spread evenly and allowed to dry for 25 min prior to an overnight incubation at 37°C. The plates were then stored at 4°C (for up to 1 month).

For the expression of RAP-GST, a starter culture was prepared by inoculating a single colony of pGEX-2T(RAP-GST)-transformed cells in 15 mL of pre-warmed LB containing 100 μg/mL ampicillin. The starter culture was incubated at 37°C overnight with shaking at 180 rpm. Subsequently, the starter culture was added to 1 L of LB containing 100 μg/mL ampicillin and the bacterial culture was incubated at 37°C with shaking at 180 rpm until the optical density (OD600) reached between 0.6 – 0.7 (approximately 3.5 h). Protein expression was then induced with 0.4 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) and the culture was incubated for an additional 4 h under the same conditions. The cells were then harvested by centrifugation at 5,000 x g for 10 min at 4°C. The supernatant was removed, and the tubes were inverted for 5 min to drain any remaining media prior to weighing the cell pellets. The cell pellets were then stored at -20°C until further use.

Chapter 2

To purify RAP-GST, cell pellets were resuspended in 3 mL of Tris buffered saline (TBS; 0.5 M Tris, 1.5 M NaCl, pH 8) per one gram of cells containing 0.1 % Triton-X 100 (TBST), lysozyme (80 mg/g cells), Roche EDTA-free cOmplete protease inhibitor cocktail (per the manufacturer’s instructions), DNase (200 μg/mL), and MgCl (1 mg). For lysis, the cells were subjected to 3 cycles of snap freezing (liquid nitrogen) and immediate thawing (37°C water bath) and then centrifuged at 33,000 x g for 40 min at 4°C. Lysate was then filtered through a 0.45 μm cellulose nitrate membrane (Sartorius) before being passed over a 5 mL glutathione Sepharose® 4B column (GE Healthcare; Pittsburgh, PA, USA), equilibrated in TBST buffer using an NGC™ Liquid Chromatography System (Bio-Rad). The column was then washed with 25 mL of TBST buffer, followed by 25 mL of TBS buffer. Bound RAP-GST was eluted with 25 mL of 15 mM glutathione in TBS and 5 mL fractions were collected. The fractions were assessed via 10% SDS-PAGE and samples containing RAP-GST were pooled and dialysed against 50 mM Tris, 20 mM NaCl, pH 8 for subsequent thrombin digestion. Proteolysis was performed using approximately 50 U of thrombin, in the presence of 2.5 mM CaCl2, for 4 h at room temperature. The sample was centrifuged to remove aggregates that formed during the digestion and then passed over the glutathione Sepharose® 4B column again, which had been equilibrated in 50 mM Tris, 20 mM NaCl, pH 8. The flow through, containing RAP and trace amounts of thrombin, was collected in 5 mL fractions. Bound GST was then eluted with 15 mM glutathione in 20 mM Tris, 20 mM NaCl, pH 8 and 2 mL fractions were collected. Fractions containing RAP and GST were analysed by SDS-PAGE and the samples containing purified protein were pooled and dialysed against PBS/az. Post dialysis, the concentration of GST was determined spectrophotometrically (A280) using a theoretical extinction coefficient of 1.69 for a 0.1% (w/v) solution (calculated using ExPASy). RAP concentration was determined by BCA assay as described in section 2.2.1. Purified GST and RAP were both stored at 4°C.

2.2.10 Purification of recombinant Clic1 Construction of the pET28a plasmid containing the mutant Cys-less Clic1 (simply referred to as Clic1 in this study) cDNA insert is described in Goodchild et al. 2011. For the expression of Clic1, a glycerol stock of BL21(DE3) chemically competent E. coli cells (a kind gift from Associate Professor Heath Ecroyd (Illawarra Health & Medical Research Institute, University of Wollongong, Australia)), transformed with the pET28a(Clic1) plasmid, were streaked onto a pre- warmed LB agar plate, containing (50 μg/mL kanamycin) and incubated at 37°C overnight. The plate was stored at 4°C the following day (for up to 1 month). A starter culture was prepared by inoculating a single colony of pET28a(Clic1)-transformed cells in 100 mL of pre-warmed LB containing 50 μg/mL kanamycin. The starter culture was incubated at 37°C overnight with shaking

Chapter 2 at 180 rpm. Subsequently, the starter culture was added to 950 mL of LB containing 50 μg/mL kanamycin and the bacterial culture was incubated at 37°C with shaking at 180 rpm until the optical density (OD600) reached between 0.6 – 0.7 (approximately 2 hours). Protein expression was then induced with 0.25 mM IPTG and the culture was incubated for an additional 4 h under the same conditions. The cells were then harvested by centrifugation at 5,000 x g for 10 min at 4°C. The supernatant was removed and the tubes were inverted for 5 min to drain any remaining media prior to weighing the cell pellets. The cell pellets were then stored at -20°C until further use.

To purify Clic1, cell pellets were resuspended in 3 mL of cell lysis buffer (50 mM Tris, 100 mM NaCl, 0.1 % Triton-X 100, pH 8) per 1 gram of cells containing lysozyme (0.8 mg/g cells), cOmplete EDTA-free protease inhibitor cocktail tablets (following the manufacturer’s instructions), DNase (200 μg/mL), and MgCl (1 mg). For lysis, the cells were subjected to 3 cycles of snap freezing (liquid nitrogen) and immediate thawing (37°C water bath) and then centrifuged at 33,000 x g for 40 min at 4°C. Lysate was then filtered through a 0.45 μm cellulose nitrate membrane (Sartorius) before being passed over a 5 mL HisTrap HP column (GE Healthcare), equilibrated in filtered buffer comprised of 50 mM Tris, 300 mM NaCl and 5 mM imidazole (pH 8) using an NGC™ Liquid Chromatography System (Bio-Rad). Filtered lysate was then loaded onto the column at 1 mL/min. Bound Clic1 was eluted using an imidazole gradient from 5 mM to 500 mM imidazole in 50 mM Tris and 300 mM NaCl (pH 8) and 1 mL fractions were collected. The eluted fractions were analysed by 10% SDS-PAGE and samples containing purified Clic1 were pooled and dialysed against PBS/az. Post dialysis, the concentration of Clic1 was determined spectrophotometrically (A280) using a theoretical extinction coefficient of 0.647 for a 0.1% (w/v) solution (calculated using ExPASy); purified Clic1 was stored at 4°C.

2.2.11 Expression and purification of recombinant NS Purified pQE-81L plasmid containing the human NS cDNA insert was a kind gift from Professor David Lomas (University of Cambridge, UK). The expression and purification of recombinant NS is described in detail in section 3.3.1.

Chapter 3

Chapter Three NS and TTR: Potential New ECs

3.1 Introduction 3.1.1 NS NS is a secreted SERPIN family member that is widely expressed in the CNS and peripheral nervous system. It is comprised of 410 amino acids and has a mass of ~ 46 kDa. NS is expressed during development, predominantly during the late states of neuronal development, during neuronal migration, axogenesis, synaptogenesis, and rearrangement of synaptic connections and throughout adulthood in regions of high plasticity, particularly, regions associated with learning and memory (Reumann et al. 2017; Ali et al. 2017; Noto et al. 2015a; Noto et al. 2015b; Teesalu et al. 2004).

NS exerts its only known physiological role by binding and inhibiting tPA (a protease that activates the plasminogen system) (Yepes et al. 2000; Yepes and Lawrence 2004). Interestingly, however, NS has been found associated with Aβ deposits in brain tissue from patients with AD. It was also shown that NS could bind and stabilise Aβ, in turn reducing Aβ cytotoxicity in vitro (Kinghorn et al. 2006). These findings suggest that NS may play an important neuroprotective role in AD by interacting with Aβ. One study also reports that NS expression is upregulated in the brain of patients with AD (Subhadra et al. 2013). If NS does play a neuroprotective role in AD, an increase in NS expression should be beneficial, however, it has been shown that increased levels of NS lead to a decrease in brain tPA activity and an increased level of NS-tPA complexes, which may actually progress disease pathology (Fabbro and Seeds 2009). As an inhibitor of tPA, NS may impair the plasmin-mediated degradation and clearance of Aβ from the brain (Gupta et al. 2015). However, NS-tPA complexes are quite unstable and fall apart within minutes in vitro, resulting in cleaved (inactive) NS and the release of active tPA (Barker-Carlson et al. 2002; Ricagno et al. 2009; Lee et al. 2015a). Interestingly, Lee et al. 2015b suggest that perhaps it is tPA that regulates NS activity (by way of NS cleavage) rather than NS merely inhibiting tPA function. A deeper understanding of the physiological roles of NS in the human body will shed more light on the functional relationship between NS and tPA, and the implications that this relationship may have in AD.

Chapter 3

3.1.2 TTR TTR is a 55 kDa homotetrameric protein and is predominantly expressed by the liver and choroid plexus, which secrete TTR into the blood stream and CSF respectively (Quintas et al. 1999; Palha 2002). Several studies also report that TTR is expressed and secreted by the retina (Zhang et al. 2013; Cavallaro et al. 1990; Bui et al. 2001). In plasma, TTR is the secondary carrier of T4 and the only known transporter of retinol (vitamin A), which it binds indirectly via retinol binding protein (Schreiber 2002). In the CSF and brain, TTR is the only known protein to bind and regulate T4 (Wirth et al. 2014). In the mammalian eye, TTR plays a crucial role in the biochemistry of the visual process by delivering retinol to photoreceptors (Cavallaro et al. 1990; Bui et al. 2001; Eichenbaum and Zheng 2000).

TTR may also have an important neuroprotective role in AD. Several studies report that TTR facilitates the unidirectional transport of Aβ from the brain across the blood-brain barrier to the blood stream, and that TTR expression levels are decreased in patients with AD (Alemi et al. 2016; Li and Buxbaum 2011). TTR has also been found associated with Aβ plaques in AD tissue, and TTR-Aβ complexes have been co-immunoprecipitated from cortical lysates of APP23 mice (transgenic mice with AD pathology) and from human brain tissues of some AD patients (Li et al. 2013). In APP23 mouse models, it was observed that the overexpression of the wild type human TTR (hTTR) transgene significantly decreased AD pathology (i.e. a significant reduction in Aβ plaque formation and AD-associated behaviour), and silencing of the endogenous murine TTR gene progressed disease pathology with an increase in Aβ brain lesions (Buxbaum and Johansson 2017; Buxbaum et al. 2008). Several studies have also shown that TTR prevents Aβ aggregation and inhibits Aβ-induced cytotoxicity in vitro (Du and Murphy 2010). However, one important observation is that the interaction between TTR and Aβ appears to be highly dependent upon the quaternary structure of TTR. While many studies show that wild type hTTR does bind to soluble Aβ monomers and oligomers and can significantly decrease cytotoxicity, it should be noted that mTTR is significantly more potent at inhibiting Aβ aggregation in vitro compared to hTTR (Li et al. 2013; Mangrolia et al. 2016; Li and Buxbaum 2011). One study specifically reports that wild type hTTR preferentially binds Aβ oligomers (but not fibrils), whereas mTTR has a much stronger binding affinity for Aβ monomers (Du and Murphy 2010). However, another study reported the opposite, stating that tetrameric TTR preferentially binds Aβ monomers and mTTR does not interact with monomeric Aβ (Garai et al. 2018). Taken together, these data demonstrate complex interactions between TTR and Aβ that are dependent on the structure of TTR and suggest that TTR may have an important neuroprotective role in AD pathology in vivo. It is very intriguing, however,

Chapter 3 that wild type hTTR appears capable of protecting cells from cytotoxic Aβ (demonstrated by multiple studies in vitro), yet in in vitro protein aggregation assays, it has little to no inhibitory effect on Aβ aggregation, compared to mTTR (which appears to be significantly more efficient at inhibiting amyloid formation). The physiological relevance of comparing wild type hTTR with mTTR in these studies is open to question since hTTR is by far the predominant species found in vivo, with concentrations being 1000-fold higher than that of mTTR (Buxbaum and Johansson 2017). Furthermore, mTTR species are typically associated with amyloidogenic diseases in vivo (described below). One study addresses this by showing that the tetrameric hTTR structure is destabilised and dissociates upon hTTR binding to soluble Aβ oligomers in vitro (Yang et al. 2013). This study hypothesised that the hTTR molecule acts as a bait and trap, where tetrameric TTR recognises and binds to accumulating Aβ oligomers, then dissociates into monomers. Liberated mTTR can then bind to excess monomeric Aβ to prevent further incorporation of Aβ into aggregating fibrils. This data suggests that an in vivo Aβ interaction-induced dissociation of wild type hTTR may exist and that investigating the chaperone activity of mTTR could be physiologically relevant, however, if Aβ interactions do in fact induce the dissociation of hTTR into monomers, a significant difference in the ability of hTTR versus mTTR to inhibit Aβ aggregation in vitro would not be expected.

Interestingly, Aβ is not the only amyloidogenic protein found to interact with TTR. Cascella et al. 2013 showed that hTTR and mTTR significantly decreased the cytotoxicity of oligomeric Aβ1-

42 and HypF-N (N-terminal domain of E. coli hydrogenase maturation protein) relative to controls. These results were obtained in vitro using MTT assays and by measuring the influx of Ca2+ ions into cells, which generate ROS (a sign of cellular stress) (Ojha et al. 2011; Sengupta et al. 2016; Deas et al. 2016; Drews et al. 2016). Despite mTTR reducing the cytotoxicity of Aβ and HypF- N, when cells were incubated with mTTR alone (but not wild type TTR alone) a ~20% decrease in MTT reduction was measured, indicating that mTTR was itself toxic. A more recent study (Jain et al. 2017) reported that both hTTR and mTTR inhibit the in vitro amyloid aggregation of CsgA (a protein precursor subunit which, in conjunction with CsgB, forms the amyloid curli fibers found in Enterobacteriacea biofilms). However, wild type hTTR did not significantly inhibit amyloid formation by CsgA at any of the concentrations tested, but rather slightly slowed the kinetics of CsgA aggregation. In agreement with other studies, the mTTR species used by (Jain et al. 2017) significantly inhibited CsgA aggregation.

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3.1.3 NS and TTR are putative ECs Although NS and TTR are unrelated proteins, it has been reported that both proteins are found associated with Aβ plaques in AD brain tissue. It was also shown that NS and TTR both bind and stabilize Aβ, thereby reducing Aβ aggregation and cytotoxicity in vitro. It has also been reported that TTR has an important role in clearing Aβ from the brain. In addition to the described interactions with Aβ, TTR has also been reported to inhibit the aggregation of CsgA (mTTR only) and protect cells from the cytotoxicity of HypF-N oligomers. These characteristics (observed for other ECs) suggest that NS and TTR may have promiscuous chaperone activity and warrant their investigation as potential new ECs. Discovering new ECs that reside in the brain and CNS is particularly important, as these organs are susceptible to many (currently) untreatable PDDs. Treating PDDs by drug delivery to the brain is challenging due to restricted passage of molecules through the blood-brain barrier and the complexity of tissue permeability in the brain (Krol 2012), however, utilising resident proteins may prove to be a viable technique for developing therapies for PDDs of the CNS and brain. This study aims to (i) determine whether NS and TTR have “generic” chaperone activity (i.e. the ability to inhibit the aggregation of a variety of amorphously- aggregating and amyloidogenic client proteins) and (ii) compare the sequence and structural characteristics of NS and TTR using a bioinformatics approach.

3.2 Materials As described in Section 2.1.

3.3 Methods 3.3.1 Optimised expression & purification of recombinant NS Previously, recombinant His-tagged NS was expressed using a BL21*(DE3) IPTG-inducible expression system at 37°C (Whiten 2011). After expression, NS was purified using (i) immobilised metal ion affinity chromatography, followed by (ii) anion exchange chromatography. Whiten followed published methodology that lead to very low yields of recombinant NS, either being expressed in the form of inclusion bodies (0.7 mg of NS per 1 L of cultured cells), which required re-solubilisation, or as soluble, cytosolic protein (1.4 mg of NS per 1 L of cultured cells). For this reason, the pQE-81L(NS) plasmid (Figure 3.1) was sent to the Protein Production Unit (PPU) (Monash University, Australia) for the development of an optimised NS expression protocol that would result in a greater yield of protein. All NS used in this study was purified as per the methods developed by the PPU, which are described below.

Chapter 3

NS cDNA insert

pQE-81L 4.8 kbp

Figure 3.1 Plasmid map of pQE-81L(NS) containing human NS cDNA, used to transform chemically competent NiCo21(DE3) E. coli. The red arrow indicates the position of the NS cDNA insert. The plasmid contains an ampicillin resistance gene (Ampicillin), colicin E1 (Col E1), lactose repressor (lac lq), lactose operon (lac O), multiple cloning site (MCS), T5 promoter (PT5), and ribosome binding site (RBS). Plasmid map was sourced from Addgene.

NiCo21(DE3) E. coli were aseptically transformed with the pQE-81L(NS) plasmid encoding N- terminally his-tagged NS. Briefly, a 50 μL suspension of chemically competent NiCo21(DE3) cells was incubated with plasmid DNA (~ 100 ng) on ice for 30 min prior to being heat shocked in a 42°C water bath for 45 s. The cells were then incubated on ice for an additional 5 min. The transformed cells were subsequently incubated with 500 μL of super optimal broth with catabolite repression (SOC) media and incubated at 37°C for 1 h while shaking at 250 rpm. The cells were then plated onto LB agar containing 100 μg/mL ampicillin. The plates were allowed to dry for 15 min before an overnight incubation at 37°C. After the overnight incubation, plates were stored at 4°C for up to 1 month.

For the expression of recombinant NS, a single colony of transformed NiCo21(DE3) cells was used to inoculate 50 mL of LB media containing 100 μg/mL ampicillin. The starter culture was incubated at 37°C overnight while shaking at 200 rpm. The starter culture was then added to 950 mL of Overnight Express™ Instant TB autoinduction media (prepared as per the manufacturer’s instructions) containing 100 μg/mL ampicillin. The expression culture was then incubated at 28°C overnight. Bacterial cells were then harvested by centrifugation at 6,000 x g for 15 min at 4°C. The supernatant was removed, and the cell pellets were either used immediately or stored at -20°C for up to 1 month. To extract recombinant NS, the bacterial pellet was lysed using BugBuster® Master Mix according to the manufacturer’s protocol. Briefly, bacterial cells were resuspended in 5 mL of BugBuster® Master Mix per 1 g of pelleted cells, containing Roche cOmplete protease inhibitor cocktail (as per the manufacturer’s recommended dosage). The cell suspension was then 50

Chapter 3 incubated on a rocking platform at room temperature for 20 min. Once lysed, cellular debris was removed by centrifugation at 16,000 x g for 20 min at 4°C. Cleared cell lysate was then passed through 0.22 μm MILLEXÒGV 0.22 μm DuraporeÒ PVDF syringe filter (Merk) in preparation for chromatographic purification.

To purify NS, the filtered lysate was loaded at 0.5 mL/min onto a HisTrap nickel affinity column (5 mL bed volume; GE Healthcare) that had been pre-equilibrated in filtered 50 mM Tris, 0.3 M NaCl, 5 mM imidazole (pH 8) using an NGC™ Liquid Chromatography System (Bio-Rad). The column was then washed with 5 column volumes of the aforementioned buffer. Bound protein was eluted at 1 mL/min using an imidazole gradient of 5 – 125 mM imidazole in 50 mM Tris, 0.3 M NaCl (pH 8) over the course of 100 min and 50 x 2 mL fractions were collected. The column was then washed with 500 mM imidazole in 50 mM Tris, 0.3 M NaCl (pH 8) to elute any remaining bound proteins from the column. All collected samples were then analysed by non-reducing 10% (w/v) SDS-PAGE (described in section 2.2.2). Samples containing purified NS were pooled and dialysed against PBS. NS concentration was determined spectrophotometrically (A280) using an extinction coefficient of 0.803 for a 0.1% (w/v) solution (Noto et al. 2015a) and the samples were then stored at -80°C. Approximately 8 mg of NS was obtained from 1 L of cell culture.

3.3.2 Immunoblotting of recombinant NS To confirm that the purified protein was indeed NS, 1 μg of the protein was first loaded onto a 10% SDS polyacrylamide gel and electrophoresed as described in section 2.2.2. After electrophoresis, the protein was transferred onto a nitrocellulose membrane as described in Section 2.2.3. Subsequently, the membrane was blocked for 1 h at room temperature with PBS/skim milk and then probed using a polyclonal rabbit anti-NS antibody (Abcam: ab16171) diluted 1:500 in PBS/skim milk. Unbound antibody was removed by washing the membrane three times with PBS containing 0.1% Triton X-100 followed by two washes with PBS. The membrane was then incubated with a polyclonal goat anti-rabbit Ig-HRP conjugate (Dako: P0448) diluted 1:5000 in PBS/skim milk for 1 h at room temperature. Lastly, the membrane was washed as before, and bound antibodies were detected using an ECL SuperSignal Western pico substrate kit (Pierce Biotechnology) following the manufacturer’s protocol. The membrane was imaged using an Amersham imager 600 (GE Healthcare).

Chapter 3

3.3.3 Characterisation of recombinant NS structure and activity Native PAGE To confirm that the purified NS was monomeric, aliquots of NS in PBS (at 32.6 μM = 1.5 mg/mL) were either freshly thawed or heated to 37°C and 43°C for 6 h and 4 h, respectively. NS was also incubated at 50°C for 30 min to induce polymerisation. 20 μg of each NS sample was then incubated with 1x non-denaturing sample buffer (100 mM Tris-HCl, 10% glycerol and 0.0025% (w/v) bromophenol blue, pH 8.6) and analysed by native PAGE using a Hoefer SE 260 Mighty Small II gel system connected to a BioRad Powerpac 300. The 8.5 x 10.5 cm2 gel was comprised of a stacking gel (5% (w/v) bis-acrylamide mix, 125 mM Tris-HCl (pH 6.8), 0.1% (w/v) ammonium persulfate (APS), 0.1% (w/v) N,N,N’,N’-Tetramethylethylenediamine (TEMED)) and a resolving gel (8% (w/v) bis-acrylamide mix, 375 mM Tris-HCl (pH 8.8), 0.1% (w/v) APS and 0.06% (w/v) TEMED). The samples were loaded onto the gel and electrophoresed in running buffer (25 mM Tris-base and 192 mM glycine pH 8.3) at 90V until the dye front reached the bottom of the gel (~ 4 h). Gels were then stained with InstantBlue™ Protein Stain (Expedeon) according to the manufacturer’s instructions before imaging. tPA-inhibition The functional activity of recombinant NS was assessed by non-reducing SDS PAGE analysis to detect the formation of SDS-stable NS-tPA complexes and the production of cleaved NS (Belorgey et al. 2002). NS and tPA were incubated together at a molar ratio of NS:tPA = 1:1.5 for 5 min at room temperature. Active tPA was inhibited by the addition of SDS sample buffer (to a final concentration of 1x) and the reaction was stopped by boiling the samples for 5 min. For comparison, samples containing NS and tPA alone were also treated as described above. The samples were analysed by non-reducing 10% SDS-PAGE, transferred to nitrocellulose membrane and then probed for NS as described in section 3.3.2.

3.3.4 Protein aggregation assays The chaperone activity of NS, hTTR, and mTTR was tested using a variety of amorphously- aggregating client proteins (BSA, CS, CPK, Clic1) and the amyloidogenic clients (ccβw and α- syn). A list of the client proteins utilised in this study, and a brief description of the assay conditions used for each client, is provided in Table 3.1.

Chapter 3

Table 3.1 Summary of the client proteins tested and the conditions used to induce aggregation. Asterisks (*) indicate the client proteins that form amyloid fibrils; other proteins all generate amorphous aggregates.

Client Concentration Temperature Protein (μM) (°C) Buffer Composition α-syn * 25 37 PBS BSA 20 37 PBS, 20 mM DTT

ccβw * 75 37 100 mM Na2HPO4, pH 7.8 Clic1 60 37 PBS CPK 15 43 PBS 50 mM Tris-HCl, 5 mM HEPES, pH 8 (TH CS 1.3 or 1.8 43 buffer)

A POLARstar OMEGA microplate reader (BMG Labtech) was used to measure amorphous protein aggregation (as changes in turbidity, A360) and amyloid fibril formation (as in increase in ThT fluorescence). The concentration of ThT used in each assay is described below and fluorescence intensity was measured using a 440 + 10 nm excitation filter and a 480 + 10 nm emission filter. To determine whether the effects of NS and TTR were specific, CLU was used as a positive control chaperone protein and BSA, α-Lac, OT, Oval or SOD1 were used as non- chaperone negative control proteins. It was confirmed that, when incubated alone, NS, TTR, or the control proteins, did not generate turbidity or ThT fluorescence under the assay conditions used (data not shown unless turbidity was generated). Control proteins (except for CLU) were used at the highest molar ratios of the putative EC:client tested. Buffer alone was also used as a control in all assays. Samples were incubated in the wells of a Greiner 384-well microplate (50 μL/well) in triplicate and all assays were repeated at least twice, unless otherwise specified.

NS aggregation assays CS (1.8 μM, 0.15 mg/mL), with or without NS, was incubated in TH buffer at 43°C with 20 s shaking (double orbital, 100 rpm) before each measurement for a minimum of 4 h. The molar ratios of NS:CS tested varied between 1:5 (0.55 mg/mL NS) and 10:1 (27.6 mg/mL NS). Clic1 (60 μM, 1.62 mg/mL), with or without NS, was incubated in PBS at 37°C with 20 s shaking (as above) before each measurement for a minimum of 6 h. The molar ratios of NS:Clic1 tested varied between 1:5 (0.55 mg/mL NS) and 5:1 (13.8 mg/mL NS). ccβw was prepared by first dissolving 1 mg of the peptide in 25 μL of DMSO. The ccβw solution was then added to 975 μL of room temperature 100 mM Na2HPO4 (pH 8) to a final ccβw concentration of 1 mg/mL (2.5% DMSO) and immediately vortexed to prevent protein precipitation. The final concentration of ccβw was determined spectrophotometrically (A280) nm using a theoretical extinction coefficient

Chapter 3

(absorbance of a 0.1% (w/v) solution = 2.46; calculated using ExPASy). ccβw (75 μM, 0.17 mg/mL), with or without NS, was incubated in 100 mM Na2HPO4 containing 25 μM ThT (pH 7.8) at 37°C with 20 s shaking (double orbital, 600 rpm) before each measurement for a minimum of 12 h. The molar ratios of NS:ccβw tested varied between 1:100 (0.035 mg/mL NS) and 1:10 (0.345 mg/mL NS). α-syn (25 μM, 0.36 mg/mL) containing 1.25 μM α-syn “seed” fibrils, with or without NS, was incubated in PBS containing 20 μM ThT at 37°C (without shaking) for a minimum of 24 h. α-syn seed fibrils were prepared by first incubating monomeric α-syn (1.25 μM) in a 42°C water bath with gentle stirring for 24 h (to initiate fibril formation), followed by 15 cycles of sonication (cycle time = 1.5 s with 30% amp) to break up the newly formed fibrillar material. This process was repeated once more before storing the fibril “seeds” at -80°C. The molar ratios of NS:α-syn tested varied between 1:200 (0.006 mg/mL NS) and 1:10 (0.12 mg/mL NS). All α-syn assays were performed by Dr. Megan Kelly (Illawarra Health & Medical Research Institute, University of Wollongong, Australia). Control proteins were used as described above. hTTR and mTTR aggregation assays BSA (20 μM, 1.33 mg/mL), with or without hTTR, was incubated in PBS containing 20 mM DTT at 37°C with 20 s shaking (double orbital, 100 rpm) before each measurement for a minimum of 8 h. The molar ratios of hTTR:BSA tested varied between 1:40 (0.028 mg/mL hTTR) and 1:1 (1.11 mg/mL hTTR). This assay was performed only once. CPK (15 μM, 1.2 mg/mL), with or without hTTR or mTTR, was incubated in PBS at 43°C with 10 s shaking (double orbital, 100 rpm) before each measurement for a minimum of 4 h. The molar ratios of hTTR:CPK tested varied between 1:50 (0.017 mg/mL hTTR) and 3:1 (2.5 mg/mL hTTR), and for mTTR:CPK between 1:50 (0.004 mg/mL mTTR) and 5:1 (1.04 mg/mL mTTR). CS (1.3 μM, 0.112 mg/mL), with or without hTTR or mTTR, was incubated in TH buffer at 43°C with 20 s shaking (as above) before each measurement for a minimum of 4 h. The molar ratios of hTTR:CS tested varied between 1:20 (0.003 mg/mL hTTR) and 10:1 (0.722 mg/mL hTTR), and for mTTR:CS between 1:20 (0.002 mg/mL mTTR) and 10:1 (0.18 mg/mL mTTR). ccβw (75 μM, 0.17 mg/mL), with or without hTTR or mTTR, was incubated in 100 mM Na2HPO4 containing 25 μM ThT (pH 7.8) at 37°C with 20 s shaking (double orbital, 600 rpm) before each measurement for a minimum of 12 h. The molar ratios of hTTR:ccβw tested varied between 1:20 (0.22 mg/mL hTTR) and 1:2 (2.2 mg/mL hTTR), while mTTR was tested at molar ratios of mTTR:ccβw between 1:100 (0.01 mg/ml mTTR) and 1:10 (0.11 mg/ml mTTR). α-syn (25 μM, 0.36 mg/mL) containing 1.25 μM α-syn “seed” fibrils, with or without hTTR or mTTR, was incubated in PBS containing 20 μM ThT at 37°C (without shaking) for a minimum of 24 h. The molar ratios of hTTR:α-syn tested varied between 1:10 (0.14 mg/mL hTTR) and 1:1 (1.38 mg/mL hTTR), while mTTR was tested at molar ratios of mTTR:α-

Chapter 3 syn between 1:50 (0.007 mg/ml mTTR) and 1:1 (0.35 mg/ml mTTR). All α-syn assays were performed by Dr. Megan Kelly (Illawarra Health & Medical Research Institute, University of Wollongong, Australia). Control proteins were used as described above.

3.3.5 Bioinformatic analysis of NS and TTR sequences To investigate whether or not NS and TTR share any sequence homology, a pairwise sequence alignment was performed using the National Center for Biotechnology Information (NCBI) Align Sequences Protein BLAST tool; default settings and algorithm parameters were used for all pairwise analyses. The NCBI Standard Protein BLAST tool was also used to determine if NS or TTR sequences are conserved in other proteins throughout the entire NCBI protein database; adjustments of settings and algorithm parameters were required for this analysis and these are described in Table 3.2. All protein sequences were obtained from UniProtKB/Swiss-Prot reviewed entries unless otherwise specified and included a signal sequence (residues 1 – 16 for NS and 1 – 20 for TTR). The PyMOL Molecular Graphics System (Version 1.2.8 Schrodinger, LLC.) was used to visualise the three-dimensional structures of NS and TTR with structural data obtained from the Research Collaboratory for Structural Bioinformatics (RCSB) Protein Data Bank (PDB); the PDB IDs for NS and TTR used in this study were 3F5N (Ricagno et al. 2009) and 4N85 (Yokoyama et al. 2014), respectively.

Table 3.2 Algorithm parameter settings used in NCBI’s Standard Protein BLAST tool. It was necessary to make several changes to the default settings and algorithm parameters when utilising the NCBI Standard Protein BLAST tool. These changes were made to reduce the stringency of the search, to identify matches with lower identity scores that might still be of potential interest. The left column describes the parameter that was altered, the middle column lists the program’s default setting and the right column indicates the setting that was selected for use in this study. Default settings were used for all other parameters not listed in the table.

Setting/Algorithm Default Setting Changed To: Parameter Database Non-redundant Protein Sequences Reference Proteins Organism (no entry) Homo sapiens (taxid:9606) Max Target Sequences 100 20,000 Expect Threshold 10 100,000 Gap Costs Existence: 11 Extension: 1 Existence: 11 Extension: 2 Conditional compositional score Compositional Adjustments No adjustments matrix adjustment Low complexity Regions (check Low complexity Regions Filter box unticked) (check box ticked)

Chapter 3

3.3.6 Synthetic peptides corresponding to NS residues 300 – 313 in aggregation assays The effects on ccβw amyloid formation of a synthetic peptide corresponding to a region of interest on NS (PNS; NS residues 300-313, EIDLKDVLKALGIT), and a negative control peptide comprised of the same residues randomly scrambled (PCTRL; KGEVDLAIDLTKIL), were tested to investigate whether the PNS region may be involved in the binding of NS to amyloidogenic proteins. ccβw was induced to aggregate as described in Section 3.3.4. The molar ratios of PNS or PCTRL to ccβw tested varied between 1:20 (0.006 mg/mL PNS or PCTRL) and 10:1 (1.15 mg/mL PNS or PCTRL).

3.4 Results 3.4.1 A more efficient method for the expression and purification of recombinant NS The NS used in this study was purified using the methods developed by the PPU (as described in section 3.3.1). In short, soluble, cytosolic his-tagged NS was expressed in NiCo21(DE3) E. coli at 28°C and purified using a HisTrap nickel affinity column. Many endogenously expressed bacterial proteins also bind to nickel columns, albeit at a lower affinity. To overcome this, a low concentration of imidazole (5 mM) was included in the binding buffer to minimise non-specific binding. Additionally, the protein was eluted from the HisTrap column using an imidazole gradient (5 – 125 mM imidazole), which produced two distinct peaks in absorbance at 280 nm (Fig. 3.2 A), eluting at 10 – 20 mL (peak 1) and 25 – 38 mL (peak 2).

The fractions corresponding to each elution peak were analysed by 10% SDS-PAGE to identify the protein species present (Figure 3.2 B). For comparison, purified NS (Lane 1) was also electrophoresed. The fractions corresponding to peak 1 were comprised of many unidentified contaminating proteins (lanes 2 – 5), none of which appeared at the expected size of NS (46 kDa). Peak 2 (lanes 6 – 8) contained a ~ 46 kDa protein (red arrow) at the expected size of NS and significantly lesser amounts of a protein of ~ 40 kDa (green arrow). The apparent size of the ~ 40 kDa protein, and the fact that it co-elutes with NS, suggests that this protein may be proteolytically cleaved NS (Belorgey et al. 2002). The fractions containing the purified ~ 46 kDa and ~ 40 kDa proteins were pooled yielding ~ 8 mg of protein.

Chapter 3

A 1500 Absorbance Peak 2 125 Imidazole 1200 100

900 Peak 1 75

600 50

300 25 Absorbance 280 nm (mAU) 280 nm Absorbance

0 0 (mM) Concentration Imidazole 0 10 20 30 40 50 Elution Volume (mL) B

Markers (kDa) 1 2 3 4 5 6 7 8

100

50 NS Cleaved NS 37

Figure 3.2 Imidazole elution of NS from a HisTrap column. A) An example chromatogram produced during the imidazole gradient elution of NS from a 5 mL HisTrap column. Bound protein was eluted in two separate peaks (peaks 1 & 2). B) SDS PAGE analysis of eluted fractions. Molecular weight markers are indicated on the left side of the image and lane 1 contains 5 μg of purified NS. Lanes 2 – 5 contain eluted fractions between 10 and 18 mL (peak 1), which contain a variety of unidentified bacterial proteins. Lanes 6 – 8 contain eluted fractions between 26 – 36 mL (peak 2), which contained two distinct bands corresponding to the masses of NS (46 kDa; red arrow) and putative proteolytically cleaved NS (40 kDa; green arrow).

To confirm that the two purified proteins were in fact NS and cleaved NS, 2 μg of the sample was electrophoresed on an SDS polyacrylamide gel and then analysed by immunoblotting (as described in section 3.3.2). The anti-NS antibody bound to both bands at ~ 46 kDa (red arrow) and ~ 40 kDa (green arrow), but not to any of the molecular marker proteins or tPA (data not shown) (Figure 3.3). These results confirm that the purified proteins are NS and cleaved NS.

Chapter 3

Markers (kDa) 50 NS Cleaved NS 37

Figure 3.3 Immunoblotting of purified NS. The polyclonal rabbit anti-NS antibody bound to both of the proteins present in the sample, however did not interact with any of the molecular marker proteins or tPA (data not shown).

3.4.2 Structural and functional characterisation of recombinant NS After confirming that the purified protein was NS, the next step was to confirm that the recombinant NS was monomeric and functional as a protease inhibitor. It is currently unknown how NS polymerisation will affect the protein’s ability to bind and stabilise aggregating proteins (such as Aβ), therefore, it was imperative to confirm that NS was monomeric when assessing its putative chaperone activity.

Native PAGE (pH 8) was used to determine if recombinant NS had polymerised during the purification process; for comparison, 20 μg of NS was either untreated or heated to 50°C for 30 min (to induce polymerisation; Figure 3.4). The extent of NS polymerisation was also investigated after 6 h and 4 h incubations at 37°C and 43°C, respectively, as these were conditions to be used in protein aggregation assays. Native PAGE analysis indicated that the recombinant NS purified in this study was predominantly monomeric. The results also showed that heating purified NS to 37°C for 6 h did not induce NS polymerisation, however, a small amount of NS polymers may have formed when NS was heated to 43°C. These results suggest that protein aggregation assays performed at 37°C should not induce significant NS polymerisation and are, therefore, suitable for testing NS for chaperone activity. Results from protein aggregation assays performed at 43°C should be interpreted with caution as the assay conditions may induce some limited polymerisation of NS, which could affect how NS interacts with aggregating client proteins. Incubation of NS at 50°C for 30 min produced various sized oligomers and significant levels of polymeric NS (~ 24% of the total NS loaded aggregated). Densitometric analysis was performed using ImageJ software (version I.x; National Institute of Health).

Chapter 3

°C for 6 h °C for 4 h °C for 30 min As purified 37 43 50

NS Polymers

NS Oligomers

Monomeric NS

Figure 3.4 Native PAGE analysis of heated and unheated NS. Purified recombinant NS was examined for polymerisation by native PAGE. The identity of samples is indicated above each lane (molecular markers were not used). Untreated purified NS was predominantly monomeric and contained negligible levels of oligomers. Heating NS to 37°C (6 h) did not induce detectable polymerisation, however, incubation at 43°C (4 h) may have produced small amounts of NS polymers (visible as a small increase in stained material sitting at the bottom of the sample well). Heating at 50°C for 30 min generated several oligomer species and larger polymers (labelled). This is representative of two individual experiments. Gel densitometry was performed to determine the extent of NS polymerisation relative to the total amount of protein loaded into each well (monomeric NS + polymerised NS).

It was also necessary to confirm that the purified NS was functionally active and could inhibit tPA (to form covalently-linked NS-tPA complexes). Latent NS is inactive as a tPA inhibitor (due to the inaccessibility of the RCL) and it is not known what effects NS latency might have on the protein’s putative chaperone activity. To confirm its functionality, NS was incubated with a molar excess of tPA (NS:tPA = 1:1.5) followed by SDS PAGE (Figure 3.5 A). Equivalent amounts of NS (red arrow) and tPA (blue arrow) alone were also included for comparison. Incubating NS with tPA produced two high molecular weight species; one migrating at just under 100 kDa (purple arrow) and one migrating between the 50 and 75 kDa markers (yellow arrow). These two species are likely to represent NS-tPA complexes produced by the interactions of NS with single-chain tPA (purple arrow) or two-chain tPA (yellow arrow) (similar results are described in Tsang et al.

Chapter 3

2014). tPA is produced as a single-chain protein, however, upon exposure to proteases, tPA is cleaved into a two-chain species that is held together via a single disulfide bond (Bertrand et al. 2015; Boose et al. 1998). Despite incubating NS with a molar excess of tPA, a small amount of NS was not incorporated into a complex. Two possible explanations for this are (i) it may be that some of the recombinant NS is inactive as a protease-inhibitor (i.e. latent) or (ii) the incubation time was not long enough for all of the NS to become incorporated into NS-tPA complexes. Proteolytically cleaved NS was also detected at ~ 40 kDa (green arrow) and is known to be produced following the spontaneous dissociation of unstable NS-tPA complexes (Belorgey et al. 2002).

To confirm that the higher molecular weight species were NS-tPA complexes, the proteins were subsequently transferred to a nitrocellulose membrane for immunoblotting analysis (Figure 3.5 B). The anti-NS antibody bound to both NS (red arrow) and cleaved NS (green arrow), however, it did not interact with any of the molecular marker proteins or tPA. The anti-NS antibody also bound to both high molecular weight species (purple and yellow arrows), confirming that these did contain NS and only form when NS is incubated with tPA.

NS-tPA NS-tPA

tPA

NS NS Cleaved Cleaved NS NS

A B

Figure 3.5 Detection of covalently linked NS-tPA complexes. A) 10% SDS-PAGE analysis of the covalently linked complex formed between NS and tPA. Molecular markers are indicated on the left side of the image and the sample loaded into each well is indicated at the top of the image; see keys for sample IDs. For comparison, NS and tPA alone were also analysed. When NS is incubated with tPA, two high molecular weight, SDS-stable species formed and an increase in cleaved NS was also apparent. B) Immunoblotting confirmed that the high molecular weight species contained NS and only form when NS is incubated with tPA. Two different sizes of NS-tPA complexes were produced, corresponding to those containing single-chain tPA (purple arrow) and one with two-chain tPA (yellow arrow).

Chapter 3

3.4.3 The effects of NS in protein aggregation assays This study examined NS for “generic” chaperone activity using two amorphously-aggregating proteins (CS and Clic1) and two proteins known to form amyloid (ccβw and α-syn). Incubating recombinant NS for 6 h at 37°C did not induce its polymerization, therefore, Clic1 (currently the only known client protein to readily form amorphous aggregates at 37°C without the use of an oxidizing/reducing agents) was selected as an appropriate client protein to assess NS for chaperone activity. As only a very low level of NS polymers formed after incubating the protein at 43°C for 4 h, CS (which readily forms amorphous aggregates at 43°C) was also used to assess NS for chaperone activity. Both ccβw and α-syn form amyloid fibrils under physiological conditions (at 37°C in phosphate buffer). The assay conditions used for each client protein are briefly described in Table 3.1.

NS did not completely inhibit CS precipitation at any of the concentrations used in this study (Figure 3.6 A). Even when tested at a substantial molar excess (NS:CS = 5:1), at the assay endpoint, NS could only inhibit ~ 47% of CS aggregation. At molar ratios of NS:CS = 1:1 and 1:5, NS only reduced endpoint CS aggregation by about 7%. The non-chaperone control protein BSA (tested at a molar ratio of BSA:CS = 5:1) slightly slowed the kinetics of CS aggregation, however, it did not inhibit the level of endpoint CS aggregation, confirming that the effects of NS were specific. The CLU control for this particular assay is not shown due to condensation problems in the corresponding wells. However, typically at a molar ratio of CLU:CS = 1:1, CLU inhibits ~ 95% of CS aggregation (see Figure 3.6 B).

NS did not inhibit Clic1 aggregation at any of the concentrations tested in this study (Figure 3.6 B). NS alone also showed a significant increase in turbidity, indicating that the sample had aggregated under the assay conditions. When used in molar excess (NS:Clic1 = 3:1 and 5:1), NS significantly increased Clic1 aggregation compared to the Clic1 alone sample (data for NS:Clic1 is not shown). The negative control protein BSA had no significant effect on Clic1 aggregation and the chaperone CLU inhibited endpoint Clic1 aggregation by ~ 94%. The erratic fluctuations in absorbance recorded in this assay typically result from condensation forming within the microplate, however, this assay was performed twice with similar results and condensation was not observed in either experiment.

Chapter 3

1.6

1.4

1.2 1 CS 1.8 uM

BSA:CS = 5:1 0.8 A360 NS:CS = 5:1 0.6 NS:CS = 1:1 NS:CS = 1:5 0.4

0.2

0 0 1 2 3 Time (h)

7 Clic1 60 uM

6 CLU:Clic1 = 1:2 BSA:Clic1 = 3:1 5 NS 180 uM 4 NS:Clic1 = 3:1

A360 3 NS:Clic1 = 1:1 NS:Clic1 = 1:3 2 NS:Clic1 = 1:5

0 0 1 2 3 4 5 Time (h)

Figure 3.6 NS is an inefficient inhibitor of amorphous CS and Clic1 aggregation. A) CS (1.8 µM) was incubated at 43°C for 4 h to induce aggregation and B) Clic1 (60 µM) was incubated at 37°C for 6 h to induce aggregation. In both cases, client aggregation was measured as turbidity at 360 nm (y-axis) as a function of time in h (x-axis). The keys indicate the molar ratios of NS:client or control:client used. Data points plotted are means ± SEM (n = 3) and the lines connecting each data point indicate the trends in the data. The results shown are representative of one experiment (CS) and two independent experiments (Clic1).

Chapter 3

In contrast, NS concentration-dependently inhibited the aggregation of ccβw (Figure 3.7 A). A molar ratio of NS:ccβw = 1:10 completely inhibited the formation of ThT-reactive material until approximately 8 h into the assay, at which point the ThT fluorescence of the sample gradually increased to ~ 28% of that measured for ccβw alone at assay endpoint. At lower ratios of NS:ccβw, the effects were less pronounced. NS:ccβw = 1:20 decreased endpoint ThT fluorescence by only ~ 25% (and delayed the onset of ccβw aggregation by about 4 h), and at NS:ccβw = 1:50 and 1:100 there was no inhibition of ccβw aggregation at the assay endpoint (although NS:ccβw = 1:50 did appear to delay the onset of aggregation by ~ 1 h). When used at a molar ratio of Oval:ccβw = 1:10, the non-chaperone control protein Oval did not significantly inhibit ccβw aggregation, confirming that the effects of NS are specific. Tested at the same molar ratio (chaperone:ccβw = 1:10), NS performed similarly to CLU until ~ 8 h into the assay, when the ThT fluorescence of wells containing ccβw and NS began to increase while those containing ccβw and CLU sample did not. An initial decrease in ThT fluorescence may be observed in some amyloid aggregation assays, which is possibly due to changes in temperature at the beginning of the assay as the absorbance and emission of ThT fluorescence decreases with an increase in temperature (Malmos et al. 2017).

NS also showed a concentration-dependent inhibition of α-syn aggregation (Figure 3.7 B). Molar ratios of NS:α-syn = 1:200, 1:100, 1:50, 1:25, and 1:10 reduced endpoint ThT fluorescence of wells containing α-syn by ~ 45%, 61%, 74%, 86%, and 97% respectively. When the non- chaperone control protein BSA was incubated with α-syn at a molar ratio of BSA:α-syn = 1:10, there was no significant inhibition of ThT fluorescence, suggesting that the effects of NS are specific. Although NS significantly and specifically inhibited α-syn aggregation, it was outperformed by CLU, which at a 1:50 molar ratio of CLU:α-syn reduced endpoint ThT fluorescence by ~ 99% relative to α-syn alone.

Chapter 3

100000 90000 ccβw 75 uM 80000 CLU:ccβw = 1:10 70000 Oval:ccβw = 1:10 60000 NS:ccβw = 1:10 50000 NS:ccβw = 1:20 AFU 40000 NS:ccβw = 1:50 30000 NS:ccβw = 1:100 20000 10000 0 0 1 2 3 4 5 6 7 8 9 10 11 12 Time (h)

B 32000

28000

24000 ⍺-syn 25 uM 20000 CLU:⍺-syn = 1:50 16000 BSA:⍺-syn = 1:10 AFU NS:⍺-syn = 1:10 12000 NS:⍺-syn = 1:25 NS:⍺-syn = 1:50 8000 NS:⍺-syn = 1:100 4000 NS:⍺-syn = 1:200

0 0 5 10 15 20 25 Time (h)

Figure 3.7 NS inhibits amyloid formation by ccβw and α-syn. A) ccβw (75 µM) and B) α-syn (25 µM) were incubated at 37°C for 12 h and 24 h, respectively, to induce amyloid formation. In both cases aggregation was measured as ThT fluorescence (in arbitrary fluorescence units (AFU); y-axis) as a function of time in h (x-axis). The keys indicate the molar ratios of NS:client or control:client used in each assay. Data points plotted are means ± SEM (n = 3 for ccβw assays or n = 5 for α-syn assays). The figure is representative of two individual experiments. The data shown in these figures was produced by Dr. Megan Kelly (Illawarra Health & Medical Research Institute, University of Wollongong, Australia).

Chapter 3

3.4.4 The effects of hTTR and mTTR in protein aggregation assays To test TTR for “generic” chaperone activity, tetrameric hTTR and mTTR were incubated with three amorphously-aggregating client proteins (BSA (hTTR only), CPK, and CS) and two amyloidogenic clients (ccβw and α-syn). BSA was selected because TTR has only one cysteine residue per monomer (Cys10) and no intramolecular disulfide bonds (Kingsbury et al. 2008). As it contains no disulfide bonds, TTR should not be structurally altered by reducing agents (such as DTT), which is required to induce the aggregation of BSA at 37°C. As TTR is thermally stable below temperatures of 101.7°C (Saelices et al. 2015), CS and CPK were also selected as appropriate clients, whose amorphous aggregation is induced by heat-stress at 43°C. Both ccβw and α-syn produce amyloid fibrils under physiological conditions (at 37°C in phosphate buffer).

At all concentrations tested, hTTR did not inhibit the reduction-induced aggregation of BSA. In contrast, under the same conditions, a molar ratio of CLU:BSA = 1:1 inhibited nearly 90% of the endpoint level of aggregation of BSA (Figure 3.8). The non-chaperone control protein SOD1 slowed the kinetics of BSA aggregation to some extent but did not significantly reduce the endpoint level of BSA aggregation.

0.6

0.5 BSA 20 uM 0.4 CLU:BSA = 1:1 A360 SOD1:BSA = 1:1 0.3 hTTR:BSA = 1:1 hTTR:BSA = 1:10 0.2 hTTR:BSA = 1:20 hTTR:BSA = 1:40 0.1

0 0 5 10 15 20 Time (h)

Figure 3.8 Effects of hTTR on the DTT-induced aggregation of BSA. BSA (20 µM) was incubated with 20 mM DTT for 18 h at 37°C to induce aggregation. BSA aggregation was measured as turbidity at 360 nm (y-axis) as a function of time in h (x-axis). The key indicates the molar ratios of chaperone:client or non-chaperone control:client used. Data points plotted are means ± SEM (n = 3). The figure shows data from a single experiment.

Chapter 3

Neither hTTR nor mTTR showed clear dose-dependent inhibition of the aggregation of CPK (Figure 3.9 A and B). Relative to CPK alone, endpoint aggregation of CPK was reduced at molar ratios of hTTR:CPK = 2:1, 1:1, 1:5, and 1:10 by ~ 31%, 21%, 27%, and 21% respectively, and at molar ratios of mTTR:CPK = 5:1, 1:1, 1:5, and 1:10 by ~ 19%, 14%, 37%, and 23% respectively. When incubated with a molar excess of mTTR (mTTR:CPK = 5:1), CPK aggregation was delayed, however, the level of endpoint aggregation was not significantly reduced. At a molar ratio of CLU:CPK = 1:1, CLU reduced the endpoint level of CPK aggregation by ~ 75%.

Chapter 3

A 0.9 0.8

0.7 CPK 15 μM 0.6 CLU:CPK = 1:1 0.5 α-Lac:CPK = 5:1

hTTR:CPK = 2:1 A360 0.4 hTTR:CPK = 1:1 0.3 hTTR:CPK = 1:5 0.2 hTTR:CPK = 1:10 0.1 0 0 1 2 3 4 5 6 7 Time (h)

B 0.9

0.8

0.7 CPK 15 μM 0.6 CLU:CPK = 1:1 0.5 α-Lac:CPK = 5:1

A360 0.4 mTTR:CPK = 5:1 0.3 mTTR:CPK = 1:1 0.2 mTTR:CPK = 1:5 mTTR:CPK = 1:10 0.1 0 0 1 2 3 4 5 6 7 Time (h)

Figure 3.9 Effects of hTTR and mTTR on the heat-induced aggregation of CPK. CPK (15 µM) was incubated at 43°C for 7 h to induce aggregation, which was measured as turbidity (A360, y-axis) as a function of time in h (x-axis). The keys indicate the molar ratios of TTR:CPK used in each assay A) hTTR:CPK and B) mTTR:CPK. Data points plotted are means ± SEM (n = 3). The data shown is representative of two independent experiments.

Chapter 3

Neither hTTR nor mTTR inhibited the aggregation of CS (Figure 3.10 A and B), however, the highest concentrations of hTTR and mTTR used (TTR:CS = 10:1) did appear to reduce the “settling” of CS aggregates in the plate (noted by a decrease in turbidity after 2 h). Aggregate “settling” is a common observation in protein precipitation assays. Relative to CS alone, a molar ratio of CLU:CS = 1:1 reduced the level of endpoint CS aggregation by ~ 88%. Under the conditions tested, the non-chaperone control proteins (α-Lac and SOD1) had no significant effect on the level of endpoint aggregation of CPK or CS, however, like TTR (at a molar ratio of TTR:CS = 10:1), SOD1 also reduced the observed “settling” of CS aggregates.

Chapter 3

A 0.6

0.5 CS 1.3 µM CLU:CS = 1:1 0.4 SOD1:CS = 10:1 hTTR:CS = 10:1 0.3 A360 hTTR:CS = 5:1

0.2 hTTR:CS = 1:1 hTTR:CS = 1:5 0.1 hTTR:CS = 1:10

0 1 2 3 Time (h)

B 0.6

0.5 CS 1.3 µM

0.4 CLU:CS = 1:1 SOD1:CS = 10:1 0.3 mTTR:CS = 10:1 A360 mTTR:CS = 5:1 0.2 mTTR:CS = 1:1

mTTR:CS = 1:5 0.1 mTTR:CS = 1:10

0 0 1 2 3

Time (h)

Figure 3.10 Effects of hTTR and mTTR on the heat-induced aggregation of CS. CS (1.3 µM) was incubated at 43°C for 3 h to induce aggregation, which was measured as turbidity (A360, y-axis) as a function of time in h (x-axis). The keys indicate the molar ratios of A) hTTR:CS or B) mTTR:CS and the control:client used in each assay. Data points plotted are means ± SEM (n = 3). The data shown is representative of two independent experiments.

Chapter 3

The effects of hTTR and mTTR on amyloid formation was also tested. Molar ratios of hTTR:ccβw = 1:10, 1:5, 1:4, 1:3, and 1:2 reduced endpoint ccβw aggregation by ~ 13%, 20%, 35%, 49%, and 63% respectively. However, when used at similar concentrations, the non-chaperone control protein Oval (1:2 molar ratio shown) completely inhibited the formation of ThT-reactive ccβw aggregates at the assay endpoint (Figure 3.11), suggesting that the inhibitory effect observed for hTTR is not specific.

35000

30000

ccβw75 uM 25000 Oval:ccβw = 1:2 20000 hTTR:ccβw = 1:2 hTTR:ccβw = 1:3 AFU 15000 hTTR:ccβw = 1:4 hTTR:ccβw = 1:5 10000 hTTR:ccβw = 1:10

5000

0 0 10 20 30 40 50 60 Time (h)

Figure 3.11 Effects of hTTR on amyloid formation by ccβw. ccβw (75 µM) was incubated at 37°C for 60 h to induce aggregation. Aggregation was measured as ThT fluorescence in AFU (y-axis) over time in h (x-axis). The key indicates the molar ratios of hTTR:ccβw or Oval:ccβw used in the assay. Data points plotted are means ± SEM (n = 3). The figure is representative of two individual experiments.

To further test whether or not hTTR had any ability to specifically inhibit the aggregation of ccβw, a panel of six negative control proteins (Figure 3.12 A (BSA), B (SOD1), C (OT), D (GST), E (a-Lac) and F (apo-transferrin (aTF)) were incubated with ccβw as described previously at control:ccβw ratios ranging between 1:1 and 1:10. The results from these assays confirmed that, under the conditions tested, many of the non-chaperone control proteins non-specifically interact with ccβw to inhibit the formation of ThT-reactive aggregates.

Chapter 3

28000 ccβw 28000 A B ccβw 24000 BSA 24000 SOD 20000 1:1 20000 1:1 1:5 1:5

16000 1:10 16000 1:10 12000 12000

8000 8000 4000 4000 0 0 0 11 22 33 44 55 66 0 11 22 33 44 55 66

28000 C ccβw 28000 D ccβw 24000 OT 24000 GST

20000 1:1 20000 1:1 1:5 1:5 16000 16000 1:10 1:10 AFU 12000 12000 8000 8000 4000 4000

0 0

0 11 22 33 44 55 66 0 11 22 33 44 55 66

28000 ccβw 28000 ccβw E F 24000 α-Lac 24000 aTF 1:1 1:1 20000 20000 1:5 1:5 16000 1:10 16000 1:10 12000 12000 8000 8000 4000 4000 0 0 0 11 22 33 44 55 66 0 11 22 33 44 55 66

Time (h) Figure 3.12 Effects of non-chaperone control proteins on amyloid formation by ccβw. ccβw (75 µM) was incubated with A) BSA, B) SOD1, C) OT, D) GST, E) α-Lac, and F) aTF at 37°C for 65 h. Aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). The key indicates the molar ratios of control:ccβw used in the assay. Data points plotted are means ± SEM (n = 3). The figure is representative of one experiment.

Chapter 3

Relative to a-syn alone, molar ratios of hTTR:α-syn = 1:10, 1:5, and 1:1 reduced endpoint ThT fluorescence by ~ 13%, 36%, and 67% respectively and this effect is hTTR specific (Figure 3.13). Relative to a-syn alone, a 1:1 molar ratio of BSA:a-syn, did not significantly reduce the endpoint ThT fluorescence. Although hTTR can significantly and specifically inhibit α-syn aggregation, its chaperone action is much less potent than that of CLU. At a molar ratio of CLU:a-syn = 1:50, CLU reduced the endpoint ThT fluorescence by ~ 87%. Compared with CLU, a 50 times higher molar concentration of hTTR was needed to reduce endpoint ThT fluorescence by ~ 67%.

7000 6000 α-syn 25 uM 5000 CLU:α-syn = 1:50 4000 BSA:α-syn = 1:1 AFU 3000 hTTR:α-syn = 1:1 2000 hTTR:α-syn = 1:5 hTTR:α-syn = 1:10 1000 0 0 5 10 15 20 25 Time (h)

Figure 3.13 Effects of hTTR on amyloid formation by α-syn. α-syn (25 μM) was incubated at 37°C for 24 h to induce aggregation. Aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). The keys indicate the molar ratios of hTTR:α-syn or control:α-syn used in each assay. Data points plotted are means ± SEM (n = 3). The figure is representative of two individual experiments. The data shown in this figure was produced by Dr. Megan Kelly (Illawarra Health & Medical Research Institute, University of Wollongong, Australia).

mTTR reduced endpoint ccβw aggregation (Figure 3.14 A). Relative to ccβw alone, molar ratios of mTTR:ccβw = 1:100, 1:50, 1:20, and 1:10 reduced endpoint ThT fluorescence by ~ 15%, 33%, 38%, and 88% respectively (data for mTTR:ccβw = 1:50 is not shown). By comparison, at TTR:ccβw = 1:10, hTTR only reduced endpoint ThT fluorescence by ~ 12%, confirming that mTTR is a significantly more potent chaperone than hTTR. Relative to ccβw alone, a 1:10 ratio of Oval:ccβw had no significant effect on the endpoint ThT fluorescence. When used at the same molar ratio (CLU/mTTR:ccβw = 1:10), mTTR outperformed CLU (which reduced endpoint levels of ThT fluorescence by ~ 68%), indicating that mTTR is a very efficient inhibitor of ccβw aggregation.

Chapter 3 mTTR also inhibited α-syn amyloid formation in a concentration-dependent manner (Figure 3.14 B). Relative to α-syn alone, molar ratios of mTTR:α-syn = 1:50, 1:20, 1:10, and 1:2 reduced endpoint ThT fluorescence by ~ 43%, 56%, 73%, and 78% respectively (data for mTTR:α-syn = 1:50 is not shown). Under the same conditions, hTTR (at a molar ratio of hTTR:α-syn = 1:5) only reduced endpoint ThT fluorescence by ~ 44%, which is less than that achieved at mTTR:α-syn = 1:50. Thus, in this assay, mTTR has a significantly more potent chaperone activity than hTTR. In contrast, a molar ratio of BSA:α-syn = 1:2 had no significant effect on the level of endpoint ThT fluorescence, showing that the effects of mTTR (and hTTR) are specific. At CLU:α-syn = 1:50, CLU reduced the endpoint ThT fluorescence by ~ 88%, showing that while effective, in this assay, mTTR is a less potent chaperone.

Chapter 3

A 50000 45000 ccβw 75 uM 40000 CLU:ccβw = 1:10 35000 Oval:ccβw = 1:10 30000 mTTR:ccβw = 1:10 25000 AFU mTTR:ccβw = 1:20 20000 mTTR:ccβw = 1:100 15000 hTTR:ccβw = 1:10 10000 5000 0 0 10 20 30 40 Time (h)

B 6000

5000 α-syn 25 uM CLU:α-syn = 1:50 4000 BSA:α-syn = 1:2 3000 mTTR:α-syn = 1:2

AFU mTTR:α-syn = 1:10 2000 mTTR:α-syn = 1:20 hTTR:α-syn = 1:5 1000

0 0 5 10 15 20 25 Time (h)

Figure 3.14 Effects of mTTR on amyloid formation by ccβw and α-syn. A) ccβw (80 µM) and B) α-syn (25 μM) were incubated at 37°C for 40 and 24 h, respectively, to induce aggregation. For both client proteins, aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). The keys indicate the molar ratios of TTR:client or control:client used in each assay. Data points plotted are means ± SEM (n = 3). The data shown represents two independent experiments for each client. The data shown for α-syn was produced by Dr. Megan Kelly (Illawarra Health & Medical Research Institute, University of Wollongong, Australia).

Chapter 3

3.4.5 NS and TTR share sequence homology To determine whether NS and TTR share any regions of sequence homology, a pairwise sequence alignment was performed using the NCBI Align Sequences Protein BLAST tool. NS and TTR sequences were obtained from UniProtKB/Swiss-Prot. This analysis revealed a homologous 14- residue sequence, calculated as being 64% identical (71% similar) and corresponding to an α- helical structure on both NS (residues 300 – 313) and TTR (residues 92 – 105; Figure 3.15 A). The specific amino acid residues corresponding to the α-helix on NS (LKDVLKAL; obtained from UniProtKB/Swiss-Prot) and TTR (TKSYWKA; obtained from Cho et al. 2014) are underlined in each sequence in Figure 3.15 A. PyMOL was used to generate three-dimensional structures of both NS and TTR and the 14 residues corresponding to the homologous region are highlighted in red on each structure (Figure 3.15 B).

A NS (Q99574): 300 E I D L K D V L K A L G I T 313 E I D K K A L G I + TTR (P02766): 92 E I D T K S Y W K A L G I S 105

NS TTR

Figure 3.15 Short conserved region shared between NS and TTR. NS and TTR were assessed in a pairwise alignment, which identified one homologous region as being 64% identical and 71% similar. A) The sequences for NS (upper) and TTR (lower) corresponding to this region are depicted. Identical residues are highlighted in red and the specific residues corresponding to the α-helix have been underlined. The C- terminal residue was not identical between NS (threonine) and TTR (serine), however these residues have similar properties (indicated by the +). Residue numbering shown includes the N-terminal signal sequences for both NS and TTR. B) PyMOL-generated structures of NS and TTR highlighting the 14-residue homologous region, which corresponds to an α-helix (depicted in red) on both NS (indicated by the single red arrow) and TTR (indicated by the four red arrows; one α-helix per identical monomeric subunit).

Chapter 3

To investigate whether the NS-TTR 14-residue homologous region is also conserved amongst other proteins (in particular chaperones, or proteins that are known to interact with amyloid), the 14-residue sequence was “blasted” against the NCBI protein database (section 3.3.5). This analysis identified many proteins of interest and these are listed in Table 3.3. Pairwise alignments (as described above) were used to compare either the 14-residue NS sequence or TTR sequence with the sequences of each protein listed in Table 3.3 and the identity/similarity scores for each alignment are provided. Only proteins of interest (such as known chaperones, proteins known to bind amyloid, proteins that share large contiguous regions of similarity, or proteins in the same family as NS or TTR) are listed in Table 3.3 as the number of matches obtained from the BLAST alignment was too large to include all of them.

Chapter 3

Table 3.3 Pairwise alignment results for the 14-residue sequences in NS and TTR compared with corresponding sequences in other proteins. Proteins of interest are listed below (first column). Identical residues are shown in red and the identity and similarity scores for each alignment are shown. Only regions of similarity containing four or more similar residues are shown here, and – denotes no similarity found. Proteins that are known chaperones or proteins that have been shown to interact with amyloid are indicated by a superscripted number, which corresponds to a reference with this information. References are listed directly after the table on the following page.

% % UniProt Aligned with NS Sequence: Aligned with TTR Sequence: Protein Residues Iden/Sim Residues Iden/Sim ID EIDLKDVLKALGIT EIDTKSYWKALGIS to NS to TTR 303-314 DLKEPLKVLGIT 75/83 Glia-derived nexin P07093 303-314 DLKEPLKVLGIT 50/58 92-99 LKKINKA 57/71 122-129 LKETLKAL 75/87 470-473 EIDT 100/100 Coiled-coil domain-containing protein 73 Q6ZRK6 213-218 LKKELK 67/67 342-345 KALG 100/100 133-143 EIDLKDVLKGL 91/91 DnaJ (Candidatus Jorgensenbacteria) 1 A0A2H0NEY4 - 3-11 KDYYKILGV 56/66 DnaJ Homolog Subfamily B Member 11 2 Q9UBS4 39-44 KDIKKA 67/83 - 1480-1484 EISTK 80/80 6908-6912 ALSIS 80/80 3557-3565 LKNVLKDLG 78/88 7737-7740 WMAL 75/75 Dystonin Q03001 3143-3149 DVKDILK 71/85 5593-5598 EIQEKS 67/67 1460-1466 LKDAEKA 71/71 2690-2695 EIAGKS 67/67 1238-1243 KAKAIS 67/67 Serine protease inhibitor I2 O75830 296-308 VDFKDVLYSLNIT 62/69 - Pleckstrin homology domain-containing family A member 2 3 Q9HB19 244-249 LKDVLK 100/100 - Beta-2-adrenergic receptor 4 P07550 271-277 ALKTLGI 71/71 - Alpha-1-antitrypsin P01009 322-333 DLKSVLGQLGIT 75/75 - Talin-1 Q9Y490 2085-2092 KDVAKALG 88/88 2089-2092 KALG 100/100 3158-3163 LKEFLK 67/83 Apolipoprotein B-100 5 P04114 3220-3223 TKSY 100/100 2292-2304 IDVRVLLDQLGTT 46/46 Apolipoprotein L6 Q9BWW8 49-54 LKEDLK 67/83 - Apolipoprotein D 6 P05090 38-44 DVNKYLG 71/71 - Apolipoprotein A-1 7 P02647 44-47 DVLK 100/100 - Amyloid protein-binding protein 2 8 Q92624 69-73 VLRAL 80/80 - Formyl peptide receptor 2 9 P25090 95-100 VLGVLG 67/67 - Ephrin type-A receptor 4 P54764 832-836 DVIKA 80/80 - Insulin receptor 10 P06213 128-132 LKELG 80/80 - Low-density lipoprotein receptor-related protein 1 11 Q07954 - 577-581 DTTSY 80/80 434-437 KALG 100/100 434-437 KALG 100/100 Low-density lipoprotein receptor-related protein 4 12 O75096 448-454 IDIRQVL 57/57 594-599 DTHLFW 50/67 Amyloid-beta A4 precursor protein-binding family B member 1 13 O00213 - 239-244 DTDSFW 67/83 DnaJ homolog subfamily A member 3, mitochondrial 14 Q96EY1 - 172-175 SYWK 100/100 Serine protease inhibitor H1 15 P50454 324-327 LGLT 75/75 300-305 KAVAIS 67/67 Serine protease inhibitor A12 Q8IW75 - 298-295 WKTL 75/75

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% % UniProt Aligned with NS Sequence: Aligned with TTR Sequence: Protein Residues Iden/Sim Residues Iden/Sim ID EIDLKDVLKALGIT EIDTKSYWKALGIS to NS to TTR 760-767 DTITEWKA 63/63 Alpha-2-macroglobulin 16 P01023 - 778-781 LGIS 100/100 Heat shock 105 kDa protein 17 Q92598 220-224 LKVLG 80/80 784-788 EIKTK 80/80 Heat shock 70 kDa protein 12B 18 Q96MM6 340-346 LKELYKA 57/71 405-409 ALNIS 80/80 Fibrinogen alpha chain 19 P02671 191-196 EVDLKD 83/100 248-251 WKAL 100/100 291-294 EIDT 100/100 BAG family molecular chaperone regulator 1 20 Q99933 204-209 LSALGI 83/83 206-209 ALGI 100/100 346-352 IDLKEAL 71/85 60/60 BAG family molecular chaperone regulator 5 21 Q9UL15 370-374 WNVLG 199-203 VLIAL 80/80 Leukocyte immunoglobulin-like receptor subfamily B member 2 22 Q8N423 7-12 VLICLG 67/67 - Advanced glycosylation end product-specific receptor 23 Q15109 344-347 ALGI 100/100 344-347 ALGI 100/100 216-223 LKSIMKAM 50/87 581-585 VLRAL 80/80 Voltage-dependent P/Q-type calcium channel subunit alpha-1A 24 O00555 - 1297-1303 IDLGLVL 71/71 1445-1450 NVLWAL 67/83 Toll-like receptor 2 25 O60603 82-88 VLTSNGI 57/57 - Glutamate receptor ionotropic, NMDA 2A 26 Q12870 11-15 VLPAL 80/80 -

1. DnaJ is a molecular chaperone (Huang et al. 2001; Cuellar et al. 2013; Wild et al. 1996; Zarouchlioti et al. 2018). 2. DnaJ B11 is an ER resident co-chaperone to Hsp40 (Lindberg et al. 2015; Chen et al. 2017), however, activation of the UPR can trigger the secretion of ER DnaJ B11 to act as an EC in the extracellular environment (Genereux et al. 2015). 3. Pleckstrin homology domains associate with the amyloid region of Aβ precursor protein (APP; Jin et al. 2007). 4. The beta-2-adrenergic receptor binds Aβ (Wang et al. 2010), however, this study suggests that Aβ binds to the N-terminal region of the receptor, which does not correspond to the region shown in Table 3.3. 5. AD senile plaques and neurofibrillary tangles immunostain for apolipoprotein B (Namba et al. 1992) and apolipoprotein B-100 has been shown to bind to Aβ (Shih et al. 2014). 6. AD neurofibrillary tangles and compact Aβ plaques immunostain for apolipoprotein D (Elliott et al. 2010). It has also been hypothesized that apolipoprotein D may be involved in the aggregation and/or clearance of Aβ from the brain (Desai et al. 2005). 7. Apolipoprotein A-1-Aβ complexes are detected in patients with AD, and apolipoprotein A-1 reduced Aβ cytotoxicity in vitro (Paula-Lima et al. 2009). 8. Amyloid protein-binding protein 2 is known to bind to APP (Senol et al. 2015). 9. Formyl peptide receptor 2 is known to bind amyloidogenic proteins, such as Aβ and the prion protein (Cattaneo et al. 2013). 10. Aβ has been shown to compete with insulin for binding to the insulin receptor (Xie et al. 2002; Tharp et al. 2016). 11. The low-density lipoprotein receptor-related protein 1 binds to secreted APP and Aβ and likely plays an important role in clearing Aβ from the brain (Kanekiyo and Bu 2014; Goto and Tanzi 2002). 12. The low-density lipoprotein receptor-related protein 4 has been shown to interact with APP (Choi et al. 2013). 13. Aβ A4 precursor protein-binding family B member 1 binds to APP (Nensa et al. 2014). 14. DnaJ A3 is a mitochondrial co-chaperone (Elwi et al. 2012; Momiuchi et al. 2015; Kakkar et al. 2016). 15. Serine protease inhibitor H1 (also known as Hsp47) is a known collagen-specific chaperone (Ishida and Naqata 2011), however, it has also been reported to interact with APP and co-localize with amyloid deposits in some patients with AD (Bianchi et al. 2011).

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16. α2M is a well characterised EC (French et al. 2008; Wyatt et al. 2013; Seddighi et al. 2018). 17. Hsp105 is a known molecular chaperone (Yamagishi et al. 2011) and has been shown to depolymerise amyloid in a manner similar to some sHSPs (Torrente and Shorter 2013). 18. Hsp70s are well documented molecular chaperones, however there is little specific information available regarding the 12B family member (Radons 2016; Mayer and Bukau 2005). 19. Fibrinogen is a characterised EC (Fonseca et al. 2016; Tang et al. 2009a) and the alpha-chain may be responsible for some of the chaperone activity (Tang et al. 2009b). 20. BAG-1 has been shown to directly bind both Aβ and tau precursor proteins (Elliot et al. 2009). 21. BAG5 enhances Hsp70 protein refolding activity by acting as a nucleotide exchange factor (Arakawa et al. 2010) and has also been shown to rescue neuronal cells from Aβ cytotoxicity in vitro (Guo et ai. 2015). 22. Leukocyte immunoglobulin-like receptor B2 binds oligomeric Aβ and the interaction is thought to occur within the first or second Ig-like domains (Kim et al. 2013b). 23. The advanced glycosylation end product-specific receptor is a cell surface receptor for Aβ (Yan et al. 1997) and has been shown to also interact with β-sheet fibrils formed by amyloid A, the prion peptide and amylin (Schmidt et al. 2000). 24. Oligomeric (but not monomeric) Aβ has been shown to regulate the α1A subunit of the P/Q-type calcium channel (Mezler et al. 2012). 25. Toll-like receptor 2 interacts with bacterial curli fibrils (Rapsinski et al. 2015) and Richard et al. 2008 suggest that toll-like receptor 2 is an endogenous receptor involved in clearing toxic Aβ from the brain. 26. Oligomeric Aβ has been shown to interact with glutamatergic receptors of the NMDA type (Danysz and Parsons 2012)

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Interestingly, many of the proteins listed in Table 3.3 are known to interact with amyloid and eight of these are known chaperones (see above). When aligning the 14-residue NS sequence with the proteins listed in Table 3.3, an interesting “pattern” emerged with many proteins following NS’s

KXXXK arrangement, where residues indicated by “X” are not necessarily identical to the NS sequence. There are also a number of protein sequences that do not follow this pattern, some of which align with residues towards the N-terminal or C-terminal end of the 14-residue NS sequence instead. The NCBI protein database BLAST yielded far fewer proteins of interest that share sequence homology with the 14-residue TTR sequence. Not only were there fewer proteins that share sequence homology, there also does not appear to be an obvious pattern of similarity shared by majority of the proteins analysed: only one protein (glia-derived nexin) shares a large region of homology (12 residues), 12 proteins share sequence similarity with the N-terminal region of the TTR 14-residue sequence and 14 proteins share sequence similarity with the C-terminal region. The significance of these alignments is not yet known. However, a better understanding of the physiological roles (in particular possible chaperone activity) of the proteins listed in Table 3.3, and identification of the regions responsible for recognising and binding unfolded proteins (in particular amyloid) may help identify and characterise the regions of NS and TTR responsible for their chaperone activities.

One very interesting protein to be identified in Table 3.3 is the coiled-coil domain-containing protein 73. Coiled-coil domains typically contain a repeated pattern of seven residues “hxxhcxc” (known as a heptad), where “h” represents a hydrophobic residue (often isoleucine, leucine, or valine) and “c” represents a charged residue (Mason and Arndt 2004). When this particular order of residues folds into an α-helix, the hydrophobic residues are arranged to one side of the helix, with hydrophilic residues on the opposite side, creating an amphipathic structure (Walshaw et al. 2001), however the “heptad” reside arrangement is not an exclusive requirement to generate an amphipathic α-helix. This is particularly interesting because, not only do amphipathic α-helices play important biological roles in protein-protein interactions, it is possible that these structures are also involved in the binding of some chaperones to misfolded or aggregating proteins. This predicted binding mechanism would require that the chaperone have a surface-exposed amphipathic α-helix, whose hydrophobic surface would selectively bind to regions of exposed hydrophobicity on unfolded proteins in the surrounding environment. Harndahl et al. 2001 showed that the small heat shock protein Hsp21 lost all chaperone activity when residues that fell within a surface-exposed amphipathic α-helix were mutated, suggesting that this particular amphipathic α- helix is involved in the chaperone action of Hsp21. It should be noted however, that the mutations

Chapter 3 described in this study also lead to significant conformational changes in Hsp21, which in itself could be responsible for the protein’s loss of chaperone activity.

To predict whether or not the corresponding α-helices on NS and TTR were amphipathic in nature, or contain obvious hydrophobic regions, the α-helical region of each 14-mer sequence was first analysed for the coiled-coil heptad pattern. The coiled-coil heptad pattern was not observed in either the NS or TTR sequence. To further investigate whether the homologous α-helices might be amphipathic, predicted α-helical pinwheel structures were also generated (Figure 3.16 A) using the Helical Wheel Projections tool created by Don Armstrong and Raphael Zidovetzki (Version: Id: wheel.pl,v 1.4 2009-10-20 21:23:36 don Exp; University of California, Riverside, United States; all default settings were used). The pinwheel projection tool predicts the arrangement of each residue around the central axis of the α-helix. The α-helix corresponding to NS appears to be amphipathic with five hydrophobic residues (L5, L1, L8, V4, and A7) arranged on one side of the helix. The remainder of the helix is comprised of charged residues. In contrast, the α-helix corresponding to TTR does not appear to be amphipathic and has no obvious distribution of hydrophobic residues in a single region of the helix. Lastly PyMOL was used to visualise the hydrophobic residues (highlighted yellow) on each α-helix (shown in red) in Figure 3.19 B. For NS, the PyMOL-generated α-helix looks quite comparable to the pinwheel helix generated using the Helical Wheel Projections tool, with all hydrophobic residues distributed on side of the helix. Similarly, the arrangement of hydrophobic residues on the TTR α-helix generated in PyMOL appears similar when compared to the predicted α-helix generated using the Helical Wheel Projections tool. It is interesting that both NS and TTR share such a strikingly homologous sequence corresponding to an α-helix, and that this particular sequence (or parts thereof) have also been identified in other proteins (including chaperones) known to interact with amyloid. These findings warrant further investigation of the homologous region in NS and TTR to determine whether this region is involved in binding to amyloid structures.

Chapter 3

T1 L8 L1 W5 L5 Y4 V4

NS TTR K2 K2 A7 A7 K6 K6 D3 S3

NS TTR Figure 3.16 Amphipathicity of predicted NS and TTR α-helices. A) Helix pinwheel projections were generated to show predicted regions of hydrophobicity on the homologous predicted α-helices corresponding to NS (left) and TTR (right). Hydrophobic residues are depicted in yellow, charged residues in blue, and polar residues in red. B) PyMOL images were also generated to examine the arrangement of hydrophobic residues and visualise the predicted α-helices within the three-dimensional structure of each protein. The homologous predicted α-helices are depicted in red, with hydrophobic residues shown in yellow.

Chapter 3

3.4.6 PNS does not inhibit the formation of ccbw amyloid As a first approach to test whether or not the NS/TTR homologous region is involved in the binding of NS and TTR to amyloid, the synthetic peptides PNS and PCTRL were generated and their effects on the formation of ccβw amyloid were tested. Neither PNS nor PCTRL inhibited endpoint ccβw aggregation (Figure 3.17 A and B). Relative to ccβw alone, molar ratios of PNS significantly increased endpoint ThT fluorescence in a mostly concentration-dependent manner (PNS:ccβw = 1:20, 1:10, 1:5, 1:1, and 10:1 increased ThT fluorescence by ~ 173%, 186%, 214%, 288%, and 272% respectively). By comparison, PCTRL also increased endpoint ThT fluorescence, however, this does not appear to be concentration-dependent effect. Neither peptide increased ThT fluorescence when incubated alone under the assay conditions, confirming that they do not form amyloid species. A non-chaperone, negative control protein was not used in this assay. When used at a molar ratio of CLU:ccβw = 1:10, CLU completely reduced endpoint ThT fluorescenc.

Chapter 3

15000

12000 ccβw 75 uM CLU:ccβw = 1:10 9000 PNS

AFU PNS:ccβw = 10:1 PNS:ccβw = 1:1 6000 PNS:ccβw = 1:5 PNS:ccβw = 1:10 3000 PNS:ccβw = 1:20

0 0 20 40 60 Time (h)

B 15000

12000 ccβw 75 uM CLU:ccβw = 1:10 9000 PCTRL PCTRL:ccβw = 10:1 AFU 6000 PCTRL:ccβw = 1:1 PCTRL:ccβw = 1:5 PCTRL:ccβw = 1:10 3000 PCTRL:ccβw = 1:20

0 0 20 40 60 Time (h)

Figure 3.17 Effects of PNS and PCTRL on amyloid formation by ccβw. ccβw (75 µM) was incubated at 37°C for 60 h to induce aggregation. Amyloid formation was measured as ThT fluorescence (AFU; y- axis) as a function of time (h, x-axis). The keys indicate the molar ratios of A) PNS:ccβw or B) PCTRL:ccβw and CLU:ccβw used. Data points plotted are means ± SEM (n = 3). The data shown represents two independent experiments.

Chapter 3

3.5 Discussion Prior to this project, one of the biggest challenges of working with recombinant NS was that its expression and purification was time consuming and ultimately resulted in very low yields. Unfortunately, there appears to be no standard NS expression protocol published in literature, for example, Kreuger et al. 1997 expressed recombinant mouse NS in E. coli DH5α cells, Hill et al. 2001 expressed recombinant human NS is Drosophila S2 cells, Ricagno et al. 2009 expressed recombinant human NS in E. coli Rosetta (DE3) pLys cells and Belorgey et al. 2002 do not specify the strain of E. coli used for their NS expression. In addition, publications rarely specify the yields of recombinant NS they obtain, making it impossible to directly compare between different methods.

The experiments performed in this study required substantial quantities of NS. To develop a more efficient NS expression and purification protocol, the pQE-81L plasmid containing the human NS cDNA insert was sent to the PPU at Monash University (Melbourne, Australia). There, the PPU assessed the expression levels of NS in various bacterial strains using expression at both 28°C (overnight) and 20°C (overnight, followed by 24 h at 18°C) (Table 3.4).

Table 3.4 Expression levels of recombinant NS in various bacterial strains at 28°C and 20°C. The symbol key is as follows: expression not detected (-), poor expression (+), low expression (++), moderate expression (+++), and high expression (++++) (PPU unpublished report).

Cell Line 28°C 20°C BL21(DE3) Gold - - BL21(DE3) C43 - - BL21(DE3) C41 + + BL21(DE3) RP ++ +++ BL21(DE3) RIL + + Rosetta-gami (DE3) - - Rosetta-gami B (DE3) - - Rosetta 2 (DE3) - - Rosetta Blue (DE3) ++ ++++ BLR (DE3) ++ + NEB NiCo21(DE3) ++++ ++ BL21*(DE3) ++ ++++

The cells were lysed and recombinant NS was purified as described above. The NS purified from each bacterial strain was then analysed by SDS-PAGE analysis and it was found that NEB NiCo21(DE3) cells incubated at 28°C produced the largest yield of soluble, uncleaved recombinant NS (PPU unpublished report; data not shown). The PPU provided a detailed

Chapter 3 expression and purification protocol for recombinant NS (described in section 3.3.1) and the methodology was easily followed to obtain similar yields.

There are several mutations that lead to the self-polymerisation of NS (Miranda et al. 2004; Roussel et al. 2013), a characteristic of the disease FENIB. Several publications show, however, that wild type NS is also capable of polymerising under acidic conditions (Belgorey et al. 2010) and under thermal stress (temperatures at or above 45°C; Noto et al. 2015; Noto et al. 2012; Ricagno et al. 2010). In 2011, Whiten compared the chaperone activity of native (monomeric) NS and NS that had been induced to polymerise. These results showed that NS polymers are less potent at inhibiting protein aggregation for some client proteins (Whiten 2011). It was, therefore, necessary to confirm that the recombinant NS used in this study was of monomeric form prior to testing the protein for chaperone activity. This was done using non-denaturing native-PAGE at pH ~ 8 (NS has a pI of 4.1 and would be negatively charged at pH 8). Unheated recombinant NS and NS heated to 50°C were compared on the gel. NS was also heated to the same temperatures used in the aggregation assays performed in this study (37°C and 43°C for 6 h and 4 h respectively) to determine if these conditions would induce NS polymerisation. The results indicated that the recombinant NS produced in this study was predominately monomeric and incubation at 37°C for 6 h did not induce NS polymerisation. However, incubation at 43°C for 4 h may generate very low levels of NS polymers.

Previous work used two amyloidogenic client proteins (Aβ1-42 and ccβ) and two amorphously- aggregating client proteins (ADH and CS) to compare the chaperone activity of monomeric NS with that of NS that had been heat-treated to induce polymerisation (Whiten 2011). Whiten’s results showed that for some client proteins, polymeric NS had potent chaperone activity and performed similarly to (or in the case of CS, better than) monomeric NS. However, in other assays, polymeric NS was significantly less effective at inhibiting protein aggregation than monomeric NS. Despite there being no consistent pattern for how NS polymerisation affects its chaperone activity, it is obvious that there are differences between the interactions of monomeric and polymeric NS with different client proteins and that caution must be taken when testing NS for chaperone activity under assay conditions that might induce its polymerisation.

Once confirmed that the recombinant NS was monomeric, it was next necessary to show that it was active as a protease inhibitor. The ability of NS to bind tPA in an SDS-stable complex was tested to assess the function of NS. Figure 3.8 clearly shows the formation of a two high molecular weight complexes only when NS and tPA are incubated together. Coincident with the formation

Chapter 3 of NS:tPA complexes, the level of native NS detected is drastically reduced and a smaller, proteolytically-cleaved NS species is produced. The western blot in Figure 3.8 B confirmed that NS is a component of the high molecular weight complexes. Following confirmation that recombinant NS was monomeric and functional as a tPA inhibitor, it was tested for chaperone activity. Most amorphously aggregating clients will not aggregate at physiologically relevant temperatures within an experimentally convenient timeframe. For this reason, the in vitro aggregation of most amorphously aggregating clients is induced with heat (temperatures typically ranging between 43°C – 60°C) or chemical stress (such as reagents that cause oxidation or reduction) (Bucciantini et al. 2002). Day et al. 2002 showed that although heat stress does increase the rate of client protein aggregation, it does not alter the aggregation pathway. Unlike most amorphously-aggregating clients, many amyloidogenic client proteins can be induced to aggregate under physiological conditions and at 37°C, without the use of chemical stress. Furthermore, Bucciantini et al. 2002 showed that the in vitro-induced aggregation of several non-disease associated proteins (such as the SH3 domain from bovine phosphatidyl-inositol-3’-kinase and HypF-N) produced fibrils that were structurally identical to aggregates found in vivo. Additionally, oligomeric species produced from the in vitro aggregation of non-disease associated proteins have cytotoxic effects similar to those produced by oligomers formed from disease-associated proteins (Rymer and Good 2001; Kayed et al. 2003; Pike et al. 1991).

As mentioned above, selecting appropriate client proteins to assess the chaperone activity of NS was important as wild type NS is susceptible to polymerisation at temperatures above 45°C (Yamasaki et al. 2008) and oxidation induces conformational changes in the NS molecule which can impair its protease inhibitor activity (Mohsenifar et al. 2007). Two amorphously aggregating client proteins were selected in this study: CS and Clic1. CS had been used previously to assess the chaperone activity of NS and was used in this study for comparison. The results obtained using CS in the current study were comparable to those previously obtained (Whiten 2011), therefore, this assay was performed only once. Under the conditions tested, NS did not completely inhibit CS aggregation. At the highest molar ratio of NS:CS tested (NS:CS = 5:1), NS only inhibited CS aggregation by ~ 47% at the assay endpoint. At molar ratios of NS:CS = 1:1 and 1:5, NS was less effective and only reduced CS aggregation by ~ 7%. Although the inhibitory effects observed for NS were specific to NS, when compared to CLU, which typically requires a CLU:CS molar ratio of 1:1 to inhibit CS aggregation by approximately 95%, NS was significantly less effective at inhibiting CS aggregation. As shown in Figure 3.4, incubating NS under the conditions used to induce CS aggregation (4 h at 43°C) resulted in negligible NS polymerisation (the vast majority

Chapter 3 of NS remained monomeric), suggesting that polymerisation is an unlikely explanation for the poor ability of NS to inhibit CS aggregation.

The ability of NS to inhibit amorphous protein aggregation was further investigated using Clic1 as a client protein, which can aggregate at only 37°C. The Clic1 used in this study is a mutant that has had four of its six native cysteine residues mutated to alanine: C89A, C178A, C191A, and C223A (Goodchild et al. 2011; Littler et al. 2004). Removal of these cysteine residues significantly destabilises Clic1, which allows the protein to readily form amorphous aggregates within 6 h when incubated at 37°C in phosphate buffer (without the use of oxidising/reducing agents). NS did not significantly reduce endpoint Clic1 aggregation at any of the molar ratios tested. However, when used at NS:Clic1 molar ratios greater than 1:1, it instead increased the level of endpoint turbidity relative to Clic1 alone. Despite the Clic1 assay being performed at 37°C (a temperature that should not induce any structural changes in the NS molecule), NS alone also showed an increase in turbidity. The buffer used in the Clic1 assay (PBS) did not contain any unusual additives and aggregation of NS alone was not observed in α-syn assays (which were also carried out in PBS at 37°C). One possible explanation for the increase in turbidity measured for NS alone in the Clic1 assays is that high concentrations of Clic1 (60 µM = 1.62 mg/ml) are required to achieve measurable Clic1 aggregation. Using NS at a molar excess in these assays could result in concentrations that exceed the solubility of NS (at a NS:Clic1 = 3:1 ratio, NS is at a concentration of 180 µM or 8.3 mg/ml). This hypothesis could also explain the observed increase in turbidity for samples containing molar ratios of NS:Clic1 greater than 1:1. At a CLU:Clic1 molar ratio of 1:2, CLU reduced endpoint Clic1 aggregation by ~ 94%, confirming that CLU is a significantly more potent chaperone in this assay. Previous work also showed that NS does not inhibit the DTT-induced aggregation of BSA or α-Lac (Whiten 2011). ccβw is a synthetic peptide designed to quickly form amyloid fibrils when incubated at 37 °C (Kammerer et al. 2004). NS concentration-dependently inhibited the increase in ThT fluorescence associated with the formation of amyloid by ccβw. Molar ratios of NS:ccβw = 1:50 and 1:100 had no effect on the end-point aggregation of ccβw. NS performed similarly to CLU when each were tested at a molar ratio of chaperone:client = 1:10, however the sample containing NS did start to show an increase in ThT reactive fluorescence around 8 h into the assay, whereas the CLU sample remained relatively constant until the assay endpoint. These results are nearly identical to those reported in (Whiten, 2011). NS also dose-dependently inhibited the generation of ThT fluorescence associated with the formation of amyloid by α-syn. Relative to α-syn alone, molar ratios of NS:α-syn = 1:200, 1:100, 1:50, 1:25, and 1:10 reduced endpoint ThT fluorescence by ~

Chapter 3

45%, 61%, 74%, 86%, and 97% respectively. Although NS significantly and specifically inhibits α-syn aggregation, five times more NS (NS:α-syn = 1:10) was required to nearly abolish α-syn amyloid formation compared to CLU, which only required CLU:α-syn = 1:50 to achieve approximately the same level of inhibition.

This study also investigated TTR as a putative new EC. TTR has been shown to bind Aβ, and thereby inhibit fibril formation and reduce Aβ cytotoxicity, suggesting that it may possess EC activity. TTR is very thermodynamically stable (up to 100°C) and contains no disulfide bonds, suggesting that the use of heat (below 100°C) or reducing agents (like DTT) in aggregation assays is unlikely to affect TTR structure (Saelices et al. 2015). Given these considerations, CS and CPK (induced to aggregate at 43°C) and BSA (induced to aggregate using 20 mM DTT) were selected as appropriate clients to test the ability of TTR to inhibit amorphous protein aggregation. At the molar ratios tested, neither hTTR or mTTR inhibited the aggregation of amorphously aggregating clients in a concentration-dependent manner. TTR had no effect on the aggregation of BSA or CS, even when present at a large molar excess with CS (TTR:CS = 10:1). Interestingly, both hTTR and mTTR did have some effect on CPK aggregation; at molar ratios of hTTR:CPK of 1:5 to 1:20, TTR reduced CPK end point aggregation by approximately 25%, but there was no apparent progressive increase in inhibition at higher ratios of TTR:CPK. At molar ratios of hTTR:CPK = 1:5 to 1:20, TTR reduced endpoint CPK aggregation by ~ 25%, but there was no apparent progressive increase in inhibition at higher ratios of TTR:CPK. When mTTR was present at a molar excess (mTTR:CPK = 5:1), it slowed CPK aggregation, however, no reduction in the level of endpoint aggregation was measured. On a molar basis, relative to either hTTR or mTTR, CLU was a far more potent inhibitor of the amorphous aggregation of all three client proteins tested.

This study also investigated the ability of hTTR and mTTR to inhibit amyloid formation by ccβw and α-syn. mTTR was significantly more effective at inhibiting amyloid formation by ccβw and α-syn than hTTR. Inhibition of ccβw aggregation by hTTR was not specific because when tested at the same high molar ratios required to measure effects for hTTR, the non-chaperone control proteins tested also significantly reduced ccβw aggregation. In contrast, mTTR specifically and dose-dependently inhibited ccβw aggregation. A molar ratio of mTTR:ccβw = 1:10 reduced endpoint ccβw aggregation by ~ 88%, whereas, at the same molar ratio, Oval (a non-chaperone control protein) did no significant effect on ccβw aggregation. Although hTTR did specifically and dose-dependently inhibit α-syn amyloid formation, it was significantly outperformed by mTTR. At a molar ratio of mTTR:α-syn = 1:10, mTTR inhibited endpoint α-syn ThT fluorescence by 73%. At the same molar ratio, hTTR only reduced endpoint ThT fluorescence by ~ 13%. These

Chapter 3 results are in agreement with published data that shows that, relative to mTTR, hTTR is not a very efficient inhibitor of amyloid formation.

As mentioned above, hTTR is unable to specifically inhibit ccβw aggregation. It is able, however, to specifically inhibit the aggregation of α-syn. It may appear curious that hTTR specifically inhibits the aggregation of only specific amyloidogenic clients, however, one possible explanation for this observation is that hTTR has been shown to interact preferentially with amyloid seeds (or oligomers) compared to monomeric forms (Du and Murphy 2010). Yang et al. 2013 showed in an ELISA that soluble Aβ aggregates that had been pre-aggregated bound to immobilised TTR with a significantly higher affinity compared to freshly prepared Aβ. As mentioned in the methods section above, α-syn aggregation assays were conducted using α-syn pre-aggregated “seeds”, whereas ccβw assays were prepared with monomeric ccβw. If hTTR preferentially binds protein oligomers, this could explain why hTTR was able to specifically, and more efficiently inhibit fibril formation by α-syn (which was induced to aggregate using pre-aggregated α-syn seeds) but not ccβw.

Collectively, these results indicate that on a molar basis CLU is a more efficient EC than NS or TTR, which is not surprising as it is the most potent EC described to date. Despite NS and TTR being very inefficient inhibitors of amorphous protein aggregation, NS and mTTR are reasonably efficient inhibitors of amyloid formation. The idea that chaperones can be “amyloid-specific” is a new concept. CLU, α2M, and Hp are classified as “generic” or broad-spectrum ECs because they have been shown to dose-dependently inhibit both amorphous and amyloid aggregation of proteins. Many PDD-associated proteins are amyloidogenic, and therefore, amyloid-forming clients are the most commonly used when assessing a protein for chaperone activity. However, amorphously aggregating proteins are also associated with many physiological disorders and disease. Correctly classifying chaperones as “generic” or “amyloid-specific” may prove to be important when investigating the mechanism by which proteins exert their protective chaperone effects. Many chaperones are thought to target unfolded proteins by recognising and binding to exposed regions of hydrophobicity that are otherwise hidden within the structure of native conformations. This hypothesis provides a mechanism for “generic” chaperone activity, however, it does not provide a mechanistic explanation for chaperones that only inhibit amyloid formation.

Until now, a structural and functional relationship between NS and TTR has never been investigated. In the current literature, NS and TTR are considered to be structurally and functionally distinct in their known physiological roles, despite multiple publications reporting

Chapter 3 that both proteins bind and stabilise Aβ, to inhibit its aggregation and cytotoxicity. The current study shows that both NS and TTR specifically and concentration-dependently inhibit amyloid formation by ccβw and α-syn in vitro, NS and mTTR being more efficient than hTTR for all amyloidogenic clients tested. NS and TTR were also incubated with a wide range of amorphously aggregating client proteins and results from these assays showed that neither hTTR nor mTTR significantly inhibited the aggregation of BSA (hTTR only), CS or CPK at the concentrations tested in this study. Although NS did partially inhibit the amorphous aggregation of CS, NS was substantively less efficient compared to CLU. This suggests that NS and TTR may be amyloid- specific chaperones and are acting via a similar mechanism (which may be different than that used by other generic ECs). The important question is: what feature(s) make NS and TTR “amyloid- specific” chaperones? It is unlikely that they simply target molecules that have regions of exposed hydrophobicity as this is the mechanism by which generic ECs are thought to exert their chaperone action. An obvious hypothesis is that there is a common structural element shared between NS and TTR responsible for their recognition and interaction with highly ordered and conserved amyloid structures.

Analyses using the NCBI Align Sequences Protein BLAST tool revealed a homologous region shared by NS and TTR that is 64% identical and 71% similar. Crystallographic structures of NS and TTR revealed that in both proteins this region corresponds to an a-helix. When looking at the crystallographic structures of NS and TTR, both of these α-helices are exposed at the surface of the protein, however the TTR α-helix is slightly sheltered by other structural components of the protein when in the tetrameric form. When TTR dissociates into monomers, the α-helix is more exposed, which could account for mTTR having a greater chaperone action compared to hTTR. Identification of the exact Aβ binding site(s) on TTR has been the focus of many studies. Li et al. 2013 and Jain et al. 2017 suggest that the TTR T4 binding pocket (residues 13 – 27, 107 – 110, and 120 – 122) are involved in the binding of Aβ or CsgA (respectively) to TTR and that binding occurs due to a hydrophobic-hydrophobic interaction (Li et al. 2016). Interestingly, however, several studies provide evidence suggesting that the TTR α-helix is involved in the binding of TTR to Aβ. Du and Murphy 2010 were the first to make this suggestion after analysing TTR-Aβ fragments via tandem mass spectrometry. Later, Du et al. 2012 showed that a mutation in the TTR α-helix, specifically L102A (L82A if lacking the signal peptide sequence) significantly reduced the binding affinity of TTR for Aβ and Yang et al. 2013 showed that the same mutation significantly reduced the ability of TTR to protect cells from cytotoxic Aβ in vitro. These data suggest that this α-helical region of TTR is involved in its ability to interact with Aβ and could also mediate interactions with other amyloid-forming proteins. As there are inconsistencies in the 91

Chapter 3 identified Aβ-binding site(s) in TTR, it is worthwhile to mention that the a-helix on NS, though strikingly similar in sequence to the TTR a-helix, may have different functional role.

NCBI BLAST analyses revealed many other proteins sharing partial homology with the NS-TTR sequence (Table 3.3). It is quite interesting that, despite NS and TTR having such high sequence similarity in this 14-residue α-helix, there are some obvious differences when looking at the pairwise alignments between NS or TTR and the proteins listed in Table 3.3. The 14-residues corresponding to NS shared significant sequence similarity with more proteins than the TTR sequence, and the homologous regions shared between NS and the other proteins, in general, encompassed a larger number of residues compared to the TTR sequence alignments. While TTR does share this particular pattern (KXXXK; where X represents a residue that may vary between individual proteins), not a single pairwise alignment between TTR and the proteins listed in Table 3.3 resulted in a region of similarity that included this entire region. The significance of this 14- residue homologous region, and in particular the “KXXXK” pattern, is not yet known, but further investigation into the properties of the amino acid residues in this region (hydrophobicity, polarity, charge, etc.) and the arrangement of the residues may identify a pattern that could (in part) explain how these proteins interact with amyloidogenic proteins.

As mentioned above, hydrophobicity is a common topic when it comes to discussing how chaperones recognise and interact with misfolded proteins. Although the exact mechanisms by which chaperones identify their target protein are not yet fully understood, surface-exposed amphipathic α-helices on chaperones may be involved in this mechanism. Amphipathic α-helices are naturally occurring structures with hydrophobic residues arranged to one side of the helix and hydrophilic residues arranged on the other. Despite sharing high sequence similarity, the NS and TTR α-helices appear to be quite different in the physical arrangement of their hydrophobic residues. The sequence corresponding to the NS α-helix appears to produce a highly amphipathic structure. In contrast, the TTR sequence predicts an α-helix that is mainly hydrophilic with a few hydrophobic residues at the end of the sequence, leading into a loop structure.

If these 14-residue regions, or more specifically the α-helices that they correspond to, are involved in the amyloid-specific chaperone activity of NS and TTR, it is likely that they interact with a specific amyloid structural element (whether that be charge, shape, size, or a combination of physical properties) rather than simply recognising regions of exposed hydrophobicity on a misfolded protein.

Chapter 3

One approach trialled to determine whether the homologous NS-TTR regions are involved in their amyloid specificity was to synthesise peptides corresponding to these regions. The peptides were then tested in protein aggregation assays to determine their effects on amyloid formation. In fact, this same approach was utilised by Cho et al. 2014 to investigate whether various TTR sequences were involved in the binding of TTR to Aβ. Cho et al. synthesised a peptide that corresponds to the TTR α-helix, however they were not able to show any interaction between the synthetic peptide and Aβ, and it was therefore not tested in any protein aggregation assays. Their conclusions stated that the synthetic peptide corresponding to the TTR α-helix did not form a helical structure, which may be required for Aβ interaction.

The current study employed a similar approach using the 14-residue sequence corresponding to the NS α-helix to determine if this region interacts with amyloidogenic proteins and inhibits fibril formation. A control sequence-scrambled peptide was also used, to test whether or not the specific sequence was required to affect amyloid formation. These results showed that both PNS (especially) and PCTRL increased ccβw amyloid formation; for PNS only, this increase was concentration-dependent. These findings suggest that PNS and PCTRL are, in some way, interacting with ccβw. When PNS or PCTRL were incubated alone, they did not generate ThT- reactive species, suggesting that they do not form amyloid on their own. One explanation for the dose-dependent enhancement of ccβw aggregation measured for PNS is that it binds to and stabilises an otherwise unstable intermediate species produced during amyloid formation, increasing the net production of ThT-reactive material (Figure 3.18) (Wilson et al. 2008; Yerbury et al. 2007). Therefore, although PNS did not inhibit ccβw amyloid formation, it did specifically and does-dependently affect the process, suggesting that there may be some inherent specificity imbedded in the 14-mer sequence for amyloid structures. Other factors that may have impacted on the results obtained include: (i) PNS may not have formed a helical structure in solution (as it would in the intact NS molecule), which may have affected its ability to interact with amyloid species, and (ii) more than one region on the NS molecule may be required to inhibit amyloid formation and the 14-residue sequence alone is insufficient.

Chapter 3

Figure 3.18 Predicted effects of ECs on amyloid formation. The kinetics of amyloid formation is often depicted as a sigmoidal curve to show the three distinct phases of amyloid assembly: lag phase, elongation phase, and lastly the production of mature fibrils (represented by the orange curve). When ECs are incubated with amyloidogenic clients, they can have varying effects on the formation of amyloid in vitro. When ECs are incubated with amyloidogenic clients at high but still sub-stoichiometric concentrations, they are able to completely inhibit amyloid formation (represented by the green arrow). However, when the concentration of the EC is much lower than that of the amyloid-forming client, ECs may stabilise normally unstable intermediate species, resulting in increased amyloid formation (represented by the red curve). Although PNS and PCTRL were incubated with ccβw at a molar excess, their structure (or lack thereof) may result in similar mechanisms of enhanced aggregation. Modified from Wilson et al. 2008.

Another obvious approach to this question is to generate mutant forms of NS and TTR that have mutations in the region of interest. As described above, this has been done for TTR. Several publications have identified L102 on the TTR EF loop (immediately adjacent to the EF α-helix) and L130, which corresponds to a residue on strand G of the inner β-sheet, as being involved in binding to Aβ (Du et al. 2012; Cho et al. 2014). Compared to wild type TTR, L102A and L130A

Chapter 3

TTR mutants exhibited weak binding to Aβ and a reduced ability to protect cells from Aβ cytotoxicity (Du et al. 2012; Yang et al. 2014; Li et al. 2013). Experiments such as these have not yet been performed with NS, however, in collaboration with researchers from the University of Cambridge, our lab group has designed several NS mutants to determine which residues in the NS α-helix, if any, are involved in its amyloid-specific chaperone activity. The ability of each NS mutant to inhibit amyloid formation and cytotoxicity will be compared with that of wild type NS.

A third potential approach to identify the NS and TTR regions involved in interacting with amyloidogenic proteins would be to design complementary antibodies that sterically shield the specific regions of interest. For example, a complementary antibody that targets the NS α-helix sequence could be generated and incubated with NS. The effects of shielding particular regions of the NS molecule can be tested in amyloid forming protein aggregation assays. If the region shielded by the antibody is involved in the chaperone action of NS, then the protein would likely lose some (or all) of its chaperone activity. This could be a good approach as modifications to the native NS and TTR structure would not be required, however, it may be very costly and time- consuming to generate the complementary antibodies, therefore, it was more practical to first use one of the alternative approaches described above to test whether the NS and TTR α-helices identified in this study are directly involved in the amyloid-specific chaperone activity.

Taken together, the results of this study provide, for the first time, an established NS expression and purification protocol that utilises a single chromatography step for purification and results in a relatively high yield of soluble recombinant NS. This study also examined the chaperone activity of NS and two forms of TTR (hTTR and mTTR). It was found that NS and TTR (mTTR specifically) are potent amyloid-specific ECs, however, they are much less effective (or ineffective in the case of TTR) at inhibiting amorphous protein aggregation. It was shown that NS and TTR share sequence homology over a 14-residue region, with the majority of this conserved sequence corresponding to a surface-exposed α-helix. This region was also found to share limited sequence homology with other proteins (some of which are known chaperones or known to interact with amyloidogenic proteins). The functional significance of this 14-residue region is not fully understood, however, it may be directly involved in the binding of NS and TTR to amyloidogenic proteins (published data already suggests that L102 on the TTR α-helix is involved in TTR binding to Aβ). Identifying and characterising the residues or regions of NS and TTR that specifically interact with amyloidogenic proteins to prevent aggregation and cytotoxicity could further our understanding of the mechanism by which these chaperones interact with various amyloid species (monomers, oligomers, or fibrils) and may prove useful in the future development of therapeutic

Chapter 3 drugs to treat diseases characterised by the formation of amyloid deposits. In any case, these findings suggest that NS and TTR are likely to play important protective roles in the human body, especially in the brain and CSF, which warrants further investigation into how they interact with PDD-relevant amyloidogenic proteins.

Chapter 4

Chapter Four Tg is a Newly Identified EC

4.1 Introduction Tg is a 660 kDa homodimeric glycoprotein secreted exclusively by thyroid follicular cells and is stored within the lumen of the thyroid. There are currently no crystallographic structures for Tg, however, its primary structure amino acid sequence is known and is described in Chapter 1. Tg plays a crucial role in the physiological storage of iodine (Ahad and Ganie 2010) and in thyroid hormone synthesis by acting as the protein precursor to T3 and T4. When thyroid hormone is required by the body, iodinated Tg is endocytosed from the thyroid lumen by follicular cells and trafficked to lysosomes for proteolytic cleavage and hormone release. In addition to its essential role in thyroid hormone synthesis, Tg also plays a role in the negative feedback regulation of thyroid functional gene expression and acts as an inducer of thyrocyte growth (Luo et al. 2014). Tg also exhibits intramolecular chaperone activity in that a domain within Tg assists in holo-Tg folding prior to its translocation from the ER (Lee et al. 2008). Furthermore, this study presents evidence of a novel physiological role for Tg as an EC which has not previously been described.

Discovering Tg’s putative EC activity was purely coincidental. Whilst investigating the chaperone activity of a2M, bTg was one of several proteins being trialled as a non-chaperone control protein in an assay measuring the amorphous aggregation of CPK. However, bTg was shown to significantly inhibit the aggregation of CPK and preliminary data indicated that this effect was dose-dependent (P. Constantinescu, unpublished data). It was subsequently shown that (i) bTg could dose-dependently and specifically inhibit the aggregation of several other amorphously aggregating client proteins (including CS and OT) and the amyloidogenic client peptide ccbw, (ii) bTg forms stable, soluble HMW complexes with CS, and (iii) FITC-labelled Tg-CS complexes bind to the surface of RAW 264.7 macrophages (Frazier 2012). The ability of bTg to specifically and dose-dependently inhibit the aggregation of a wide range of proteins (both amorphously- aggregating and amyloidogenic) and subsequently form stable, soluble HMW complexes with amorphously-aggregating client proteins are characteristics shared by other known ECs, such as CLU (Wyatt et al. 2009c).

Co-localisation with amyloid deposits in vivo is another feature shared by other known ECs. Although there is currently no published data showing the co-localisation of Tg with amyloid in vivo, there are several PDDs that lead to the deposition of amyloid in the thyroid and surrounding

Chapter 4 tissues, which warrant further investigation of Tg as an EC. TMC is a cancer that originates in the parafollicular C cells of the thyroid (Khurana et al. 2004). TMC tumours oversecrete the hormone calcitonin, which leads to calcitonin amyloid formation and deposition in surrounding tissues (Erickson et al. 2015). In addition to TMC, patients suffering from systemic amyloidosis (primary or secondary – though more common in the latter) can also exhibit amyloid deposition in the thyroid gland. Occasionally, though rare, systemic amyloidosis of the thyroid can lead to thyroid amyloid goitre, which is clinically defined by the apparent enlargement of the thyroid gland due to amyloid deposition (Yildiz et al. 2009). Surprisingly, many studies report that patients exhibiting amyloid goitre of the thyroid have normal thyroid function, however, this data is mostly restricted to medical case studies (rather than peer-reviewed scientific publications). Additionally, these studies typically assess thyroid function by measuring thyroid hormone levels, and not Tg directly (Kimura et al. 1997). Determining whether or not amyloid formation is linked to a decrease in Tg expression, or if Tg interacts with amyloid in vivo, would help to establish a physiological role for Tg as an EC.

Although no such role has been previously described for Tg, preliminary data suggests that Tg has promiscuous chaperone activity. Discovering new ECs, especially ones that reside in specific organs, may prove to be useful in developing therapies that target organ-specific PDDs, such as TMC. This study aims to (i) confirm that both bTg and hTg have the ability to inhibit the aggregation of a variety of amorphously-aggregating and amyloidogenic client proteins (all previous experiments were performed with only bTg), (ii) confirm that bTg forms HMW complexes with amorphously aggregating clients, (iii) determine if bTg can protect cells from cytotoxic amyloid in vitro, and (iv) compare the sequence of hTg with other known ECs to identify any sequence similarities using a bioinformatic approach.

4.2 Materials As described in Section 2.1.

4.3 Methods 4.3.1 Tg in protein aggregation assays The chaperone activity of Tg was tested using several amorphously-aggregating client proteins (CS and OT) and the amyloidogenic clients (Aβ and Cal). A list of the client proteins utilised in this study, and a brief description of the assay conditions used for each, is provided in Table 4.1.

Chapter 4

Table 4.1 Summary of the client proteins and conditions used to induce aggregation in this study. The client protein concentration and the assay conditions (temperature and specific buffer) have been listed for each client protein; * indicates the client proteins that form amyloid fibrils, with all other proteins generating amorphous aggregates.

Client Concentration Temperature Buffer Composition Protein (μM) (°C) Aβ * 20 30 PBS Cal * 10 37 PBS CS 1.8 43 TH buffer OT 12.5 60 PBS

A POLARstar OMEGA microplate reader (BMG Labtech) was used to measure amorphous protein aggregation (as changes in turbidity, A360) and amyloid fibril formation (as an increase in ThT fluorescence). The concentration of ThT used in each assay is described below and fluorescence intensity was measured with excitation at 440 + 10 nm and emission collected at 490 + 10 nm. To confirm that the effect of Tg was specific, CLU was used as a positive chaperone control and BSA, α-Lac, OT, Oval or SOD1 were used as non-chaperone control proteins. It was confirmed that, when incubated alone, Tg and the control proteins did not generate turbidity or ThT fluorescence under the assay conditions used. Control proteins (except for CLU) were used at the highest molar ratios of Tg:client tested. Buffer alone was also used as a control in all assays. Immediately prior to placement in the microplate reader, the sample wells were covered using SealPlate. Samples were incubated in the wells of a Greiner 384-well microplate (50 μL/well) in triplicate and all assays were repeated at least twice, unless otherwise specified.

Ab1-42 peptide (simply referred to as Ab in this study) was dissolved in 10 mM NaOH to a concentration of 1 mg/mL, aliquoted into 1.5 mL Eppendorf tubes (50 μL/tube), immediately snap- frozen in liquid nitrogen, and then stored at -80°C. Ab (20 μM, 0.135 mg/mL), with or without bTg, was incubated in PBS containing 50 μM ThT at 30°C with 30 s shaking (double orbital, 200 rpm) before each measurement for a minimum of 24 h. The molar ratios of bTg:Ab tested varied between 1:5 (2.64 mg/mL bTg) and 1:600 (0.022 mg/mL bTg). Cal was prepared as described by Avidan-Shpalter and Gazit 2006. Briefly, 1 mg of Cal was dissolved in 1 mL of 1,1,1,3,3,3- hexafluoro-2-propanol (HFIP) and sonicated in a water bath for 5 min at room temperature to remove any pre-formed aggregates. The peptide was then aliquoted into 2 mL (round-bottom) Eppendorf tubes (100 μL/tube). Remaining HFIP was evaporated off using nitrogen gas to produce a Cal film, which was then stored at -20°C. Immediately prior to use, Cal films were dissolved in PBS to a working concentration of 1 mg/mL. Cal (10 μM, 0.034 mg/mL), with or without bTg, was incubated in PBS containing 25 μM ThT at 37°C with 20 s shaking (double orbital, 100 rpm) 99

Chapter 4 before each measurement for a minimum of 100 h. The molar ratios of Tg:Cal tested varied between 1:50 (0.132 mg/mL Tg) and 1:500 (0.013 mg/mL Tg). CS (1.8 μM, 0.15 mg/mL), with or without bTg or hTg, was incubated in TH buffer at 43°C with 20 s shaking (double orbital, 100 rpm) before each measurement for a minimum of 4 h. The molar ratios of Tg:CS tested varied between 1:1 (1.19 mg/mL Tg) and 1:4 (0.297 mg/mL bTg) or 1:6 (0.198 mg/mL hTg). OT (12.5 μM, 0.95 mg/mL), with or without bTg or hTg, was incubated in PBS at 60°C with 10 s shaking (double orbital, 100 rpm) before each measurement for a minimum of 3 h. The molar ratios of Tg:OT tested varied between 1:5 (1.65 mg/mL bTg) and 1:20 (0.413 mg/mL bTg) for bTg and between 1:10 (0.825 mg/mL hTg) and 1:40 (0.206 mg/mL Tg) for hTg. Control proteins were used as described above.

4.3.2 Investigating Tg structure using native PAGE To investigate the structural integrity of Tg at 60°C, aliquots of freshly prepared bTg and hTg in PBS (at 5 mg/mL) were either stored at 4°C for 3 h or heated to 60°C for between 5 min and 3 h. 20 μg of each sample in non-denaturing sample buffer (100 mM Tris-HCl, 10% (v/v) glycerol and 0.0025% (w/v) bromophenol blue, pH 8.6) was analysed by native PAGE using a Hoefer SE 260 Mighty Small II gel system connected to a BioRad Powerpac 300. The 8 x 5 cm2 gel was comprised of a stacking gel (5% (w/v) bis-acrylamide mix, 125 mM Tris-HCl (pH 6.8), 0.1% (w/v) APS, 0.1% (w/v) TEMED) and a resolving gel (8% (w/v) bis-acrylamide mix, 375 mM Tris-HCl (pH 8.8), 0.1% (w/v) APS and 0.06% (w/v) TEMED). The samples were loaded onto the gel and electrophoresed in running buffer (25 mM Tris-base and 192 mM glycine pH 8.3) at 90V until the dye front reached the bottom of the gel (~ 4 h). Gels were then stained with InstantBlue™ Protein Stain (Expedeon) according to the manufacturer’s instructions and imaged using an Amersham A1600 imager.

4.3.3 Measuring regions of solvent-exposed hydrophobicity on Tg with bisANS To investigate whether or not thermal stress induced changes in the hydrophobicity of the Tg molecule, freshly prepared bTg and hTg at 300 μg/mL in PBS were either stored at 4°C for 4 h or heated to 60°C for 4 h. BSA at 300 μg/mL in PBS was used as a positive control (as it is very hydrophobic) and PBS alone was used for blank-correcting all other samples. All samples were loaded into the wells of a Greiner clear, flat-bottomed 96-well microplate (50 μL/well) in triplicate. Stocks of 4,4’-dianilino-1,1’-binaphthyl-5,5’-disulfonic acid, dipotassium salt (bisANS) (Thermo Fisher Scientific) were originally prepared by dissolving the salt in DMSO to a concentration of 564 μg/mL (838 μM) followed by storage at -20°C. For each assay, a stock of bisANS was thawed and then diluted in PBS to a concentration of 10 μM. To each well containing protein or PBS, 50

100

Chapter 4

μL of the diluted bisANS stock was added to achieve a final protein concentration of 150 μM and 5 μM bisANS. The plate was then incubated at 28°C for 15 min before being read in a POLARstar OMEGA microplate reader (BMG Labtech). BisANS fluorescence intensity was measured with excitation at 360 + 10 nm and emission collected at 490 + 10 nm.

4.3.4 Tg-client protein complexes The ability of Tg to form stable, soluble HMW complexes with the amorphously-aggregating client protein, CS, was assessed. Samples were prepared in Greiner 384-well microplates (50 μL/well) and the extent of CS aggregation was monitored using a POLARstar OMEGA microplate reader as described previously.

Formation of Tg-client complexes: Tg-client complexes were formed by incubating CS with Tg under the assay conditions described above. Briefly, CS (1.8 μM) was incubated with or without Tg in TH buffer at 43°C for a minimum of 4 h at a 1:1 molar ratio of Tg:CS. CS was also incubated under the same conditions with or without Tg at 4°C. Tg was also incubated alone at 43°C to confirm that Tg alone did not form HMW species under these conditions. Samples were then collected from the microplates and pooled in preparation for size exclusion chromatography (SEC).

Detection and purification of Tg-CS complexes using SEC: Pooled samples were concentrated to ~ 200 μL using a Vivaspin 500 (MWCO 30) microcentrifuge concentrator (GE Healthcare, Parramatta, NSW, Australia) according to the manufacturer’s instructions. The samples were then centrifuged at 16,000 x g for 5 min to remove any insoluble material (i.e. protein aggregates) before being passed through a Superoseä 6 10/300 SEC column (24 mL packed bed volume) connected to an NGC™ Liquid Chromatography System (Bio-Rad) equilibrated in filtered PBS/az at 0.3 mL/min and collected in 0.5 mL fractions. The UV trace (A280) was used to monitor the elution of proteins and a minimum of 24 mL of PBS/az eluate was collected for each sample to ensure no proteins remained in the column. Fractions containing the putative Tg-CS complexes were then pooled and concentrated using EMD Milliporeä Amiconä Ultra-15 centrifugal filter units (Merk-Millipore). An approximate concentration was determined (estimated by A280) using a SpectroMax Plus 384 spectrophotometer (Molecular Devices, San Jose, California, USA); the theoretical extinction coefficients of Tg (E0.1% = 1.104) and CS (E0.1% = 1.55) were averaged for these calculations. Theoretical extinction coefficients were calculated using the Expasy ProtParam tool. The putative Tg-CS complexes were stored at 4°C in PBS/az.

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Detection of Tg-CS complexes using SDS-PAGE: To confirm that a HMW complex had formed, and that the complex contained both Tg and CS, 5 μg each of Tg and CS and 10 μg of the putative Tg-CS complex were loaded onto a 8.5 x 10.5 8% (w/v) SDS polyacrylamide gel and electrophoresed as described in section 2.7, with the exception that the samples were heated to 99°C for 3 h to induce the dissociation of the complex prior to electrophoresis.

Detection of Tg-CS complexes using sandwich ELISA: To confirm that the HMW Tg-CS complexes were comprised of physically-associated Tg and CS, the complexes were assessed by sandwich ELISA (see Figure 4.1). Samples were prepared in quadruplicate in a Greiner 96-well ELISA High Binding microplate. The wells containing the “sandwich” were first coated with 50 μL of a polyclonal rabbit anti-CS antibody (LifeSpan BioSciences, Inc..: LS-C147385) at 3 μg/mL (diluted in 20 mM Tris-HCl, 5 mM HEPES, pH 8) at 37°C for 1 h. The wells were then washed 3 times with PBS and subsequently blocked with 100 μL of 1% (w/v) HDC heat-denatured casein in PBS containing 0.01% (w/v) thimerosal (HDC) for 30 min at 37°C. These wells were then washed as above and incubated with 50 μL of the Tg-CS complex at 20 μg/mL (diluted in HDC) for 1 h at 37°C. The plate was then washed as described above and subsequently incubated with 50 µL of a mouse monoclonal anti-Tg antibody (GeneTex: GTX21984) at 3 µg/mL (diluted in HDC) for 1 h at 37°C. The plate was again washed as described above and incubated with 50 µL of anti-mouse IgG-HRP conjugated antibody (Abcam: 205719) diluted 1:4000 in HDC for 1 h at 37°C. The plate was again washed and bound anti-mouse IgG-HRP antibody was detected by incubating each well with 100 µL of ELISA buffer (0.05 M citric acid, 0.1 M Na2HPO4, pH 5, containing 1 µL/mL 30% H2O2 and 2.5 mg/mL o-phenylenediamine dihydrochloride OPD) at room temperature until a bright, orange colour developed (approximately 2.5 min). The reaction was stopped with 1 M HCl (50 µL/well) and the absorbance of the wells at 490 nm was measured in a SpectraMax microplate reader.

To confirm that the antibodies used in this ELISA were binding specifically, various controls were used. First, Tg and CS (5 µg/mL) were used alone in place of the Tg-CS complex (50 µL/well) to show that both Tg and CS must be present and physically associated for the “sandwich” ELISA to produce a significant absorbance value. To confirm that the anti-CS and anti-Tg antibodies were specifically reacting with the proteins in the complex, the Tg-CS complex was captured by a specific antibody (anti-CS or anti-Tg) and a species-matched, non-specific antibody (polyclonal rabbit anti-a2M antibody (Abcam: ab58703) or a mouse monoclonal anti-Ub antibody (Santa Cruz Biotechnology, Inc.: SC-8017) were used in place of the rabbit anti-CS or mouse anti-Tg antibodies, respectively). Non-specific antibody controls were used at the same concentrations as 102

Chapter 4 the anti-Tg and anti-CS antibodies. To verify that the anti-mouse IgG-HRP conjugate did not bind non-specifically, the wells of the plate were first blocked with HDC and washed as described above, followed by incubation with the anti-mouse IgG-HRP conjugate. The wells were again washed and detected as described above. Finally, as positive controls, selected wells were coated with Tg or CS (50 µL/well) at 5 µg/mL (diluted in 20 mM Tris-HCl, 5 mM HEPES, pH 8), washed and blocked as described above, and then incubated with mouse anti-Tg or rabbit anti-CS antibodies respectively. The wells were then washed and incubated with an anti-mouse IgG-HRP conjugate (as above) and goat anti-rabbit IgG-HRP conjugate (1:2000) (Chemicon: AP307P), respectively. The wells were then washed, and bound antibodies were detected as described above.

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Figure 4.1 Schematic diagram showing the different configurations of reagents used in a sandwich ELISA to specifically detect Tg-client protein complexes. The figure shows when molecules should interact (depicted by a physical association) and predicts which molecules should not interact with each other (shown by a gap separating molecules). The figure also indicates for each sample set whether or not an HRP signal is predicted to develop. The key at the top indicates the colour/shape representing each molecule in the ELISA.

Size analysis of Tg-client complexes: Dynamic light scattering (DLS) and nanoparticle tracking analysis (NTA) were used to determine the average size and size distribution of bTg, CS, and Tg- CS complexes. For DLS analysis, solutions of bTg and CS (1 mg/mL) and purified Tg-CS 104

Chapter 4 complexes (0.1 mg/mL) in 0.22 µm filtered PBS were loaded into the wells of a Greiner 96-well microplate (100 µL/well) in triplicate and analysed using a Zetasizer Auto Plate Sampler (Malvern Instruments, Worcestershire, UK). Average particle diameters were recorded as a frequency distribution curve and the average diameter (averaged peak of 3 normally distributed curves) was reported for each well for a total of nine technical replicates for each sample. For NTA, solutions of bTg, CS, and purified Tg-CS complexes at 8 µg/mL in 0.22 µm filtered PBS were individually passed through the cell chamber of a NanoSight LM10 (Malvern Instruments) and the particle sizes were recorded as a frequency distribution curve (n = 3 for each sample).

4.3.5 Cytotoxicity assays To determine if Tg was capable of rescuing cells from cytotoxic amyloid, it was first necessary to analyse the toxicity of the amyloid species produced in this study. Cal (80 – 200 μM) and Ab (100 μM) were prepared and aggregated under sterile conditions. Microplates and film covers were incubated under UV light for 2 h in a laminar-flow biosafety cabinet and all buffers and protein solutions were syringe-filtered using Millexâ GV 0.22 μm filters (Millipore). Cal and Ab, prepared as described in section 4.3.1, were loaded into the wells of a Greiner 96-well microplate (100 μL/well) and the plate was sealed with a SealPlateâ (Excel Scientific, Inc.) plate film prior to being removed from the biosafety cabinet. The peptides were aggregated under the same conditions outlined in section 4.3.1, except that samples to be added to cells did not contain ThT. For each peptide, one well containing ThT was used to monitor amyloid formation during the assay using a POLARstar OMEGA microplate reader. At various time points, aliquots of each peptide were removed from the plate under sterile conditions (to ensure a variety of amyloid species were obtained), snap-frozen in liquid nitrogen, and stored at -80°C.

Nthy-ori 3-1, PC12, HEK293, and SH-SY5Y cells were cultured as described in section 2.11. Nthy-ori 3-1 and PC12 cells (in RPMI containing 1% FCS) and HEK293 and SH-SY5Y (in DMEM-F12 containing 1% FCS) were seeded at 10,000 cells per well (except for Nthy-ori 3-1, which were seeded at 12,000 cells per well) in a Greiner 96-well sterile tissue culture-grade microplate and allowed to adhere overnight in a humidified incubator at 37°C with 5% (v/v) CO2. The media was then replaced with serum-free RPMI-1640 or DMEM-F12 (100 μL/well) with or without additives. Frozen peptide stocks were thawed immediately before addition to cells and used at final working concentrations of 20 – 50 μM (Cal) and 30 μM (Ab). Serum-free media, sterile-filtered BSA (at the same concentration as Cal or Ab) and PBS were used as non-toxic controls and all proteins were incubated with cells for a minimum of 48 h. Additives constituted no more than 25% of the total volume in each well. During optimisation, it was confirmed that 105

Chapter 4 media containing 25% PBS did not affect cell viability (data not shown). Immediately prior to starting the viability assays, media for the positive “cell death” control was replaced with 20 μL of 100% ethanol (EtOH) and the plate was allowed to sit in the biosafety cabinet for 10 min to allow for the ethanol to evaporate. 100 μL of serum-free media was then added back to these samples and cell viability was assessed. Cell viability was analysed using either (i) MTS (3-(4,5- dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) assay or (ii) calcein-AM (calcein-acetoxymethyl (AM) ester) assay. In both assays, results were adjusted for background absorbance or fluorescence by subtracting the absorbance or fluorescence of wells that contained only media + PBS (no cells).

In MTS assays, metabolically active cells reduce MTS to produce the dark purple molecule formazan, which is soluble in cell culture media. Formazan absorbance can then be measured spectrophotometrically and used as an indirect measurement of cell viability. For these assays, CellTiter 96â Aqueous One Solution (Promega, Madison, WI, USA) was thawed immediately prior to use in a 37°C water bath for 10 min and 20 μL of this was added to each well. The plate was wrapped in foil and returned to the incubator and monitored hourly for a maximum of 5 h. When sufficient colour had developed, the absorbance of wells at 492 nm was measured using a SpectraMax microplate reader.

Calcein-AM is membrane permeable and diffuses into cells. In the cytoplasm of viable cells calcein-AM is cleaved to release fluorescent calcein, which remains trapped inside the cell. Fluorescent intensity is used as a measure of cell viability. For calcein-AM assays, calcein AM (Invitrogenä, Thermo Fisher Scientific, North Ryde, NSW Australia) was added to each well to a final concentration of 1 μM and the plate was covered in foil and returned to the incubator for 30 min. The media containing excess calcein AM was carefully removed and replaced with PBS. The plate was read in a POLARstar OMEGA microplate reader using excitation and emission band pass filters of 485 + 12 nm and 520 + 10 nm, respectively.

4.3.6 Bioinformatic analysis comparing Tg and other known ECs To investigate whether or not Tg shares sequence homology with other known ECs, pairwise sequence alignments were performed using the NCBI Align Sequences Protein BLAST tool; default settings and algorithm parameters were used for all pairwise analyses. The NCBI Standard Protein BLAST tool was also used to determine if any parts of the Tg sequence are conserved in other proteins throughout the entire NCBI protein database; adjustments of settings and algorithm parameters were required for this analysis and these are described in Table 4.2. Only human 106

Chapter 4 sequences were used in bioinformatical analyses and all protein sequence information was obtained from UniProtKB/Swiss-Prot reviewed entries unless otherwise specified.

Table 4.2 Algorithm parameter settings used in NCBI’s Standard Protein BLAST tool. It was necessary to make several changes to the default settings and algorithm parameters when utilising the NCBI Standard Protein BLAST tool. These changes were made to reduce the stringency of the search, to identify matches with lower identity scores that might still be of potential interest. The left column describes the parameter that was altered, the middle column lists the program’s default setting and the right column indicates the setting that was selected for use in this study. Default settings were used for all other parameters not listed in the table.

4.4 Results 4.4.1 Tg inhibits the aggregation of a wide range of proteins This study examined bTg and hTg for general chaperone activity using two amorphously- aggregating proteins (CS and OT) and two proteins known to form amyloid (Aβ and Cal). The assay conditions used for each client protein are briefly described in Table 4.1. bTg (only) has previously been shown to inhibit the aggregation of CPK, CS, OT and ccβw (Frazier 2012).

Tg inhibited Aβ amyloid formation (Figure 4.2). Relative to Aβ alone, molar ratios of Tg:Aβ = 1:200, 1:10, and 1:5 inhibited endpoint ThT fluorescence by ~ 40%, 83%, and 83% respectively (data for bTg:Aβ = 1:5 is not shown). Very low concentrations of bTg were less efficient at inhibiting Aβ aggregation with a molar ratio of bTg:Aβ = 1:400 having no inhibitory effect and Aβ:bTg = 1:600 actually increasing Aβ amyloid formation. Relative to Aβ alone, a 1:5 ratio of the non-chaperone control proteins SOD1 and Oval had no inhibitory effect on Aβ aggregation (Oval actually increased the endpoint ThT fluorescence of Aβ by ~ 17%). When used at the same molar concentration, bTg performed similar to CLU (CLU:Aβ = 1:5), which inhibited endpoint ThT- reactive Aβ by ~ 83%, indicating that, like CLU, Tg is a very efficient inhibitor of Aβ aggregation.

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50000

45000

40000 Aβ 20 µM 35000 CLU:Aβ = 1:5 30000 SOD1:Aβ = 1:5 Oval:Aβ = 1:5 25000

AFU bTg:Aβ = 1:10 20000 bTg:Aβ = 1:200 15000 bTg:Aβ = 1:400 10000 bTg:Aβ = 1:600 5000

0 0 2 4 6 8 10 12 Time (h) Figure 4.2 bTg is an efficient inhibitor of Ab aggregation. Ab (20 µM) was incubated at 30°C for nearly 12 h to induce amyloid formation. Aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). The key indicates the molar ratios of bTg:Ab or control:Ab used. Data points plotted are means ± SEM (n = 4). The figure is representative of two independent experiments.

Although bTg did inhibit Cal aggregation in a concentration dependent manner, the negative control, BSA, completely inhibited Cal aggregation when used at BSA:Cal = 1:5 molar ratio. As a molar ratio of bTg:Cal = 1:100 was sufficient to almost completely inhibit the formation of ThT- reactive Cal amyloid, several negative control proteins were incubated with Cal at the same molar ratio to determine whether they had an inhibitory effect. At a control:Cal molar ratio = 1:100, Oval, a-Lac, OT, and GST inhibited Cal amyloid formation between 80 – 90% (data not shown). As all negative controls used in this study also inhibited Cal aggregation, it could not be concluded that bTg inhibits Cal aggregation specifically.

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140000

120000

Cal 10 µM 100000 CLU:Cal = 1:1

80000 BSA:Cal = 1:5

bTg:Cal = 1:5 AFU 60000 bTg:Cal = 1:100

40000 bTg:Cal = 1:250 bTg:Cal = 1:400 20000

0 15 30 45

Time (h)

Figure 4.3 bTg does not specifically inhibit Cal aggregation. Cal (10 µM) was incubated at 37°C for nearly 45 h to induce amyloid formation. Aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). The key indicates the molar ratios of bTg:Cal or control:Cal used. Data points plotted are means ± SEM (n = 3). The figure is representative of one independent experiments.

Both bTg (Figure 4.4 A) and hTg (Figure 4.4 B) dose-dependently inhibited the aggregation of CS. A molar ratio of Tg:CS = 1:1 inhibited CS aggregation by ~ 93% (bTg) and ~ 88% (hTg). At lower ratios of Tg:CS, the effects were less pronounced. At bTg:CS = 1:2, endpoint CS aggregation was decreased by only ~ 33% and at bTg:CS = 1:4 CS aggregation actually increased by ~ 39%. hTg was slightly better at inhibiting endpoint CS aggregation with hTg:CS = 1:2 and 1:4 reducing this by ~ 68% and 44%, respectively. When used at a molar ratio of hTg:CS = 1:6, endpoint CS aggregation increased by ~ 34%. At a 1:1 molar ratio, the non-chaperone control protein BSA did not inhibit endpoint CS aggregation, but rather increased it by ~ 90%, confirming that the inhibitory effects of both bTg and hTg are specific. Tested at the same molar ratio (chaperone:CS = 1:1), both bTg and hTg outperformed CLU, which inhibited CS endpoint aggregation by ~ 56%.

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A 1.2

0.8 CS 1.8 µM CLU:CS = 1:1 0.6 BSA:CS = 1:1

A360 nm 0.4 bTg:CS = 1:1

bTg:CS = 1:2 0.2 bTg:CS = 1:4 0 0 0.5 1 1.5 2 2.5 3 3.5 Time (h)

1.2

CS 1.8 µM 0.8 CLU:CS = 1:1 0.6 BSA:CS = 1:1

A360 nm hTg:CS = 1:1 0.4 hTg:CS = 1:2 0.2 hTg:CS = 1:4 hTg:CS = 1:6 0 0 0.5 1 1.5 2 2.5 3 3.5 Time (h) Figure 4.4 bTg and hTg are efficient inhibitors of CS aggregation. CS (1.8 µM) was incubated at 43°C for 4 h to induce aggregation. Protein aggregation was measured as turbidity at 360 nm (y-axis) as a function of time in h (x-axis). CS was incubated with either A) bTg, or B) hTg and the keys indicate the molar ratios of Tg:CS or control:CS used. Data points plotted are means ± SEM (n = 3) and each result shown is representative of two independent experiments.

Both bTg (Figure 4.5 A) and hTg (Figure 4.5 B) dose-dependently inhibited the aggregation of OT. For OT assays, hTg appeared to be a more potent EC than bTg (hTg only required a molar ratio of hTg:OT = 1:10 to inhibit endpoint OT aggregation by ~ 97%, whereas a 1:5 molar ratio of bTg:OT was required to inhibit ~ 94% of endpoint OT aggregation). However, both forms of Tg were efficient inhibitors of OT aggregation. Lower ratios of Tg:OT were still very effective at inhibiting OT aggregation. At bTg:CS = 1:15 and 1:20 endpoint OT aggregation was decreased 110

Chapter 4 by ~ 73% and 15%, respectively. hTg was slightly better at inhibiting endpoint OT aggregation with hTg:CS = 1:20, 1:30, and 1:40 inhibiting ~ 90%, 67%, and 36%, respectively. At 1:1 and 1:10 molar ratios, the non-chaperone control proteins SOD1 and a-Lac did not inhibit OT aggregation, with a-Lac actually increasing endpoint OT aggregation by ~ 31%, confirming that the inhibitory effects of both bTg and hTg are specific. CLU (at a 1:1 molar ratio) was used as a positive chaperone control in both assays. Both bTg and hTg were tested alone and neither protein generated turbidity under these assay conditions (data not shown).

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A 0.45 0.4 0.35 OT 12.5 µM 0.3 CLU:OT = 1:1 0.25 SOD1:OT: 1:1 0.2

A360 nm bTg:OT = 1:5 0.15 bTg:OT = 1:15 0.1 bTg:OT = 1:20 0.05 0 0 50 100 150 200 Time (min) B 1.2

1 OT 12.5 µM 0.8 CLU:OT = 1:1

0.6 ⍺-Lac:OT = 1:10 hTg:OT = 1:10 A360 nm 0.4 hTg:OT = 1:20 0.2 hTg:OT = 1:30 hTg:OT = 1:40 0 0 100 200 300 Time (min) Figure 4.5 bTg and hTg are efficient inhibitors of OT aggregation. OT (12.5 µM) was incubated at 60°C for up to 5 h to induce aggregation. OT aggregation was measured as turbidity at 360 nm (y-axis) as a function of time in min (x-axis). OT was incubated with either A) bTg or B) hTg and the keys indicate the molar ratios of Tg:OT or control:OT used. Data points plotted are means ± SEM (n = 4 for bTg assays and n = 3 for hTg assays) and the results shown are each representative of two individual experiments.

4.4.2 Thermal stress induced structural changes in the Tg molecule As described above, OT was induced to aggregate at 60°C. It is well known that Tg loses structural stability and dissociates at temperatures above 55°C (Metzger and Edelhoch 1961; Edelhoch and Metzger 1961), however, it is not known whether these heat-induced structural changes affect the chaperone activity of Tg. To investigate the structural changes that occur in bTg and hTg, native PAGE was used to compare unheated Tg and Tg that had been heated at 60°C for various periods of time (Figure 4.6). A small amount of self-aggregated Tg is observed in the unheated Tg sample

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(more prominent in bTg compared to hTg). It is possible that this species corresponds to a 27s Tg polymer with a reported molecular mass of 1,320 kDa (4 x 330 kDa subunits) (Vecchio et al. 1966), though this was not confirmed with molecular markers in this study. Tg is heterogeneous in nature and known to self-aggregate (a requirement for storage within the thyroid lumen) and several studies have reported difficulties in obtaining pure 660 kDa Tg dimer (Van Zyl et al. 1969; Baudry et al. 1998; Deshpande et al.1997) Incubating both bTg and hTg at 60°C for only 5 min was enough to induce significant structural changes in nearly all Tg in the sample. At this time point, the majority of the Tg had self-aggregated to form several HMW species (> 660 kDa). In addition to polymerisation, the dissociation of Tg into monomeric subunits (330 kDa) was also observed for both bTg and hTg within 5 min of heating. As the 60°C incubation time increased, no significant changes in the amount of polymerised Tg were observed, however, the abundance of monomeric Tg decreased over time, likely due to its aggregation.

Native 5 min10 min 20 min1 h 2 h 3 h Native 5 min 10 min20 min1 h 2 h 3 h

Tg Tg Aggregates Aggregates

Tg Tg (660 kDa) (660 kDa) Monomer Monomer

(330 kDa) (330 kDa)

bTg hTg

Figure 4.6 Images of InstantBlueä-stained native-PAGE gels showing heat-induced structural changes in Tg. 8% native-PAGE was used to separate 20 μg each of bTg (left) or hTg (right) that had been stored at 4°C for 4 h (Native) or heated at 60°C for various time points between 5 min and 3 h (indicated at the top of each gel). Molecular markers were not used in this PAGE experiment and these results are representative of three independent experiments for bTg (time points greater than 20 min only) and one experiment for hTg.

4.4.3 Thermal stress increased surface hydrophobicity of both bTg and hTg Once it had been confirmed that heating Tg to 60°C does induce significant structural changes, the next step was to determine whether or not these structural changes affected the surface hydrophobicity of Tg. Analysis of the surface hydrophobicity of bTg and hTg using bisANS showed a significant increase in hydrophobicity for both species after Tg had been incubated at 60°C for 4 h relative to native Tg (Figure 4.7). As expected, a high level of fluorescence intensity was measured for wells containing BSA. 113

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14000 12000 10000 8000 * * 6000 AFU 4000 2000 0 4°C 60°C 4°C 60°C BSA bTg hTg

Figure 4.7 BisANS assay shows a significant increase in solvent-exposed hydrophobicity on Tg following heating at 60°C. The y-axis is fluorescence intensity (AFU) and the x-axis identifies the sample. bTg (blue) and hTg (pink) were either stored at 4°C or incubated at 60°C for 4 h. BSA (yellow) was used as a positive control. The results shown are the average fluorescence intensities (n = 3 + SEM) and were adjusted for background fluorescence by subtracting the intensity of wells that contained only PBS (no protein). This figure is representative of two individual experiment and * indicates a significant increase in fluorescence intensity compared to the corresponding 4°C sample in each sample set (bTg or hTg) (student T-test, p < 0.05).

4.4.4 bTg forms stable, soluble HMW complexes with CS Several methods were used to confirm the formation of HMW Tg-CS complexes and to show that bTg and CS were physically associated in the complex. Particle size analysis was also carried out to determine the size of the Tg-CS complexes.

Detecting Tg-CS complexes by SEC: Co-incubation of bTg and CS at 43°C for 4 h produced a

7 species which eluted within the void volume (V0 > 4 x 10 Da) of a Superoseä 6 10/300 SEC column run at ~ 7 mL (Figure 4.8). A similar species was not observed when the native proteins were incubated for the same period of time at 4°C. Under the latter condition, the SEC profile observed matched the combined SEC profiles of native bTg (eluting at ~ 12 mL) and CS (eluting at ~ 17 mL). There were no differences between the elution profiles for Tg incubated at 4°C vs 43°C, however, the elution peak observed for native CS disappeared after heating the protein at 43°C for 4 h (indicating that nearly all CS precipitated out of solution under these conditions). Neither bTg nor CS produced HMW species when heated alone at 43°C. These results confirm that a HMW species only forms when bTg and CS are co-incubated at 43°C, conditions that induce CS aggregation.

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Putative HMW Tg:CS Complex bTg CS > 4 x 107 Da 660 kDa 85 kDa 2000 1800 1600 1400 Tg + CS 43°C 1200 Tg + CS 4°C 1000 800 CS 43°C

A280 nm CS 4°C 600 400 bTg 43°C 200 bTg 4°C 0 0 2 4 6 8 10 12 14 16 18 20 22 24 Volume (mL)

Figure 4.8 SEC profiles of native and heat-stressed bTg and CS. The y-axis indicates absorbance at 280 nm and the x-axis indicates the elution volume (mL) from a Superoseä 6 10/300 SEC column. CS and bTg alone (at 1.8 μM) and bTg with CS at a 1:1 molar ratio (at 1.8 μM) were incubated at 43°C for 4 h. The resulting SEC profiles were compared to the SEC profiles of the native proteins (incubated at 4°C for 4 h). Each trace is labelled on the right with the protein species present in the sample and its pre-SEC treatment. The protein species corresponding to each elution peak is indicated at the top of the figure. Heated and unheated bTg and CS were only analysed once, however, the unheated bTg + CS mixture was analysed twice and heated bTg + CS samples were analysed on at least three separate occasions; similar results were obtained in all cases.

Detecting Tg-CS complexes by SDS-PAGE: Reducing SDS-PAGE analysis confirmed that both bTg (blue arrow) and CS (pink arrow) were present in the putative HMW Tg-CS complex that had eluted in the void volume of the Superoseä 6 10/300 SEC column (Figure 4.9). Bands corresponding to both bTg and CS were also observed when the HMW complex was analysed under non-reducing conditions, however, these bands were comparatively faint, indicating that majority of the complex had not dissociated and did not enter the gel. For comparison, bTg and CS were also loaded onto the gel under non-reducing and reducing conditions.

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CS Molecular -CS - Markers

(kDa) bTg CS HMW Tg bTg CS HMW Tg

bTg 200 bTg

150

100

CS CS

Reduced Non-reduced Samples Samples Figure 4.9 Images of Coomassie-stained SDS-PAGE gels analysing SEC fractions. 8% SDS-PAGE was used to separate 5 μg each of bTg and CS, and 10 μg of putative HMW Tg-CS complex (purified by SEC) under reducing and non-reducing conditions (indicated below images). Precision Plus Proteinä Dual Color Standards (BioRad) were used (left) and the image was cropped to remove empty wells.

Detecting Tg-CS complexes with sandwich ELISA: Sandwich ELISA, in which plate bound rabbit polyclonal anti-CS antibody and a mouse monoclonal anti-Tg antibody were used, indicated 7 that co-incubation of bTg and CS for 4 h at 43°C produced a species (eluting at V0 > 4 x 10 Da) that contains physically associated bTg and CS (Figure 4.10; yellow sample). Compared to control samples (Figure 4.10; blue samples), SEC purified putative HMW Tg-CS complex showed a significantly higher absorbance (p < 0.05, Student’s t-test).

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* *

0.35 * 4 3.5 0.3 3 0.25

2.5 0.2 2

A490 A490 nm 0.15 A490 nm 1.5

0.1 1

0.05 0.5

0 0 a-1CS a-2CS a-3CS a-a42M a-5CS bTg1 CS2 Tg-CS bTg CS Tg-CS Tg-CS a-Tg a-CS a-Tg a-Tg a-Tg a-Tg a-Ub

Test Negative Controls Positive Controls

Figure 4.10 Sandwich ELISA confirms that bTg and CS are physically associated in Tg-CS complexes. The y-axis is absorbance (490 nm) and the x-axis identifies each sample; the molecules were added to the plate in the order in which they are listed, and the red text indicates which molecule was bound to the plate prior to blocking for each sample set. Bound antibody was detected with an appropriate HRP conjugated antibody. The results shown are the average A490 nm (n = 4 + SEM). This figure is representative of two individual experiments and * indicates significantly greater A490 nm compared to the indicated negative control (Student’s t-test, p < 0.05).

Determining the size of Tg-CS complexes: For DLS results, all values reported here correspond to the Z-averages obtained for each sample unless otherwise stated. DLS analysis of bTg and CS samples indicated normally distributed particle sizes with average diameters of approximately 45.6 nm and 8.5 nm, respectively (Figure 4.11). Although CS appeared to be a monodisperse solution, having a dispersity index (DI) score (+ standard error) of 0.24 + 0.007, this was not the case for bTg (DI score of 0.55 + 0.01). DLS indicated that the bTg sample contained molecules of two distinct sizes with mean diameters of 23.8 nm and 187.9 nm (these values were obtained from the peak distribution curve). The larger particle observed here may correspond to the Tg polymer described in section 4.4.2. DLS analysis also indicated that the SEC purified HMW Tg-CS complex had an average diameter of 82.9 nm, which is significantly larger than bTg or CS alone as determined by the Student’s t-test (p < 0.05).

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* 100 * 80 60 40 20 Diameter (nm) Diameter 0 bTg CS Tg-CS

Figure 4.11 DLS estimates of the mean diameters of HMW Tg-CS complexes and uncomplexed native bTg and CS. The y-axis is particle size (nm) and the x-axis identifies each sample. The results shown are the Z-averaged mean diameter of normally distributed curves; n = 9 + SEM for each sample. This figure is representative of one individual experiment and * indicates a significant difference in size when comparing the Tg-CS complex to either bTg or CS (Student’s t-test, p < 0.05).

Due to the well-known and documented limitations of using DLS to resolve polydisperse solutions (Pusey and Tough 1985; Stetefeld et al. 2016), NTA was also used in this study to estimate the particle size(s) of purified Tg-CS complexes. NTA analysis indicated that the CS solution was comparatively monodisperse (relative to bTg and Tg-CS) with the majority of the particles in the sample resolving with a mean diameter around 54.5 nm and a range between 21.5 – 106.5 nm (Figure 4.12; green trace). A small fraction of CS particles did resolve at larger sizes, which could be attributed to CS aggregation. NTA analysis of the bTg sample showed a more polydisperse solution compared to CS with three distinct mean particle sizes (Figure 4.12; blue trace: (i) the highest number of particles resolved with a mean diameter of 84.5 nm and a range between 57.5 – 113.5 nm, (ii) the next most abundant peak shows particles resolving with a mean diameter of 135.5 nm and a range between 113.5 – 168.5 nm, and (iii) a small number of particles resolved with a mean diameter of 194.5 nm and a range between 168.5 – 303.5 nm. As mentioned above, the bTg particles resolving at larger sizes (> 113.5 nm) likely correspond to bTg polymers that are known to be in this solution (as described in section 4.4.2). In addition to having the largest sized particles, the purified HMW Tg-CS complex sample was by far the most polydisperse sample tested (Figure 4.12; pink trace). Tg-CS had five distinct mean particle diameters; the largest number of Tg-CS particles resolved with a mean diameter around 214.5 nm and a range between 163.5 – 287.5 nm. There are two peaks indicating that some Tg-CS particles are smaller than 163.5 nm (mean diameters resolving at around 153.5 nm and 92.5 nm) and two peaks indicating that some Tg-CS particles are larger than 287.5 nm (mean diameters resolving at around 325.5 nm and 438.5 nm). 118

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18 16 14 )

6 12

10 bTg 8 CS 6 Tg-CS

Particle Count (10 Particle 4 2 0 0 100 200 300 400 500 600 Diameter (nm) Figure 4.12 NTA distribution curves for estimation of the mean diameters of HMW Tg-CS complexes and uncomplexed native bTg and CS. The y-axis indicates the number of particles counted (106) and the x-axis is particle size (nm). The results shown are the average of three normally distributed curves for each sample. This figure represents one individual experiment.

4.4.5 Assessing cytotoxicity of Cal and Ab To determine whether or not Tg can protect cells from cytotoxic amyloid, it was first necessary to confirm that the amyloid species produced in this study were cytotoxic. Cal was selected as it has previously been shown to produce cytotoxic amyloid species. Furthermore, Cal amyloid can accumulate within thyroid tissue in vivo in association with TMC, making its use physiologically relevant when testing the cytoprotective properties of Tg (a thyroid resident protein). Nthy-ori 3- 1, a euthyroid follicular cell line, was chosen (for physiological relevancy) as the first cell type to test.

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300000 4 250000 3 200000

150000 AFU

100000 2 1. monomers 2. early-stage oligomers 50000 3. late-state oligomers 1 4. amyloid fibrils 0 0 5 10 15 20 25 Time (h) Figure 4.13 ThT assay measuring amyloid formation by Cal. Cal (80 µM) was induced to aggregate at 37°C for 24 h Aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). The amyloid species expected to be present at each stage of aggregation is indicated: (1) predominately monomers, (2) early stage oligomeric species, (3) larger oligomers, and (4) mature amyloid fibrils. Data points plotted are each from a single well (n = 1). The results shown are representative of at least four independent experiments.

MTS assays were used to measure the viability of Nthy-ori 3-1cells that had been incubated with 20 μM Cal (sampled at various stages of aggregation) or BSA. MTS analysis indicated that Cal (at any of the collected timepoints tested in this study) did not significantly reduce Nthy-ori 3-1 viability when compared to cells that were treated with PBS only (Figure 4.14). In contrast, wells containing cells treated with 100% EtOH for 20 min showed a significantly reduced MTS absorbance (Student t-test, p < 0.05).

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1.6 1.4 1.2 1 0.8 * 0.6

A492 nm 0.4 0.2 0 PBS 0 h 3 h 5 h 8 h 12 h 30 h 100%EtOH EtOH 20 μM Cal taken at the indicated timepoint during aggregation

Figure 4.14 MTS assay to test Nthy-ori 3-1 viability after incubation with 20 μM Cal. The y-axis is absorbance (492 nm) and the x-axis identifies the sample incubated with cells for 48 h. The first samples (PBS) indicates cells that were treated with an equivalent volume of PBS in place of Cal. The results shown are the average A492 nm (n = 3 + SEM) and this figure is representative of two independent experiments; * indicates a significant reduction in A492 nm compared to the PBS control (Student’s t-test, p < 0.05).

As cytotoxicity may be cell-specific, three other cell types (PC12, HEK293, and SH-SY5Y) were incubated with PBS, Cal, or BSA for 48 h and their viability was assessed using MTS. For these assays, Cal (200 μM) was incubated for 24 h at 37°C under the same conditions as above (24 h timepoint aliquots were used for these assays) and the working concentration of Cal added to the cells was increased from 20 μM to 50 μM. Although MTS analysis did show a small decrease in A492 (18% for PC12, 7% for HEK293, and 14% for SH-SY5Y) for cells treated with 50 μM Cal compared to cells treated with only PBS, similar results were also observed for cells treated with 50 μM BSA, indicating that the decrease in cell viability was not specific to Cal (Figure 4.15). Wells containing cells treated with 100% EtOH showed a significant reduction in absorbance.

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A B

PC12 HEK293

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0.6 0.6 0.4 0.4 A492 A492 nm A492 A492 nm 0.2 0.2 * * 0 0 PBS 50 µM 50 µM EtOH PBS 50 µM 50 µM EtOH Cal BSA Cal BSA

C SH-SY5Y 0.8

0.6

0.4 A492 A492 nm 0.2 * 0 PBS 50 µM 50 µM EtOH Cal BSA

Figure 4.15 MTS assay to test the effects of 50 μM Cal on the viability of PC12, HEK293, and SH- SY5Y cells. The y-axis is absorbance (492 nm) and the x-axis identifies the sample incubated with A) PC12, B) HEK293, and C) SH-SY5Y cell lines for 48 h. The results shown are the average A492 nm (n = 4 + SEM) and are from a single experiment; * indicates a significant reduction in A492 nm compared to the PBS control (Student’s t-test, p < 0.05).

As bTg could not be shown to specifically inhibit Cal aggregation, nor could cytotoxic Cal amyloid be generated under the conditions tested here, cytotoxicity assays using Ab were trialled instead. Ab is the most extensively characterised amyloid-forming protein to date and most publications show that it is the oligomeric species that are the most cytotoxic. Although Ab amyloid is not found in thyroid tissue, most studies agree that amyloid structure is characteristically similar regardless of the identity of the native protein or amino acid sequence, and it is the structure itself that is inherently responsible for cytotoxicity (Serpell 2014; Shewmaker et al. 2011; Eisenberg and Jucker 2012; Schubert et al. 1995). bTg does inhibit Ab amyloid formation (Figure 4.2), suggesting that Ab is a suitable test protein for assessing the ability of Tg to protect against

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Chapter 4 cytotoxic amyloid. For these studies, only SH-SY5Y cells were used and viability was assessed using 1 μM calcein-AM. Ab (100 μM) was induced to aggregate under sterile conditions at 37°C in a 96-well microplate. One sample, containing ThT, was used to monitor the progression of Ab aggregation (Figure 4.16). In order to assess the toxicity of several amyloid species, Ab (without ThT) was removed from the microplate at various time points (0, 0.5, 1, 3, and 12 h) during the course of aggregation, snap-frozen and stored at -80°C until use.

120000

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60000 AFU 40000

20000

0 0 3 6 9 12 15

Time (h) Figure 4.16 ThT assay measuring Ab amyloid formation. Ab (100 µM) was induced to aggregate at 37°C for 14 h. Aggregation was measured as ThT fluorescence in AFU (y-axis) as a function of time in h (x-axis). Each data point plotted is from a single well (n = 1). The results shown are representative of two independent experiments.

Results from calcein-AM assays indicated that incubating SH-SY5Y with 30 μM Ab collected at 0, 0.5, and 1 h did not significantly reduce cell viability when compared to cells that were treated with an equivalent volume of PBS only (Figure 4.17). Although incubating cells with 30 μM Ab (collected at 3 and 12 h) did show a small reduction in calcein fluorescence (~ 5% and 12% respectively) compared to cells treated with only PBS, similar results were also observed for cells treated with 30 μM BSA (~ 17% reduction in calcein fluorescence), indicating that the decrease in cell viability was not specific to Ab. Cells treated with 100% EtOH for 20 min produced a cytotoxic effect as indicated by the significant reduction in calcein fluorescence (student T-test, p < 0.05).

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100000 90000 80000 70000 60000 50000 AFU 40000 30000 20000 10000 * 0 PBS 0 hr 0.5 hr 1 hr 3 hr 12 hr BSA 100%EtOH EtOH Figure 4.17 Calcein-AM assay to measure SH-SY5Y viability after incubation with 30 μM Ab. The y- axis is fluorescence intensity (AFU) and the x-axis identifies the sample incubated with SH-SY5Y cells for 48 h. PBS indicates cells that were treated with an equivalent volume of PBS in place of Ab or BSA. The results shown are the average fluorescence intensities (n = 5 + SEM) and this figure represents one individual experiment and * indicates a significant reduction in fluorescence compared to the PBS control (student T-test, p < 0.05).

4.4.6 Tg does not share sequence homology with other ECs In an attempt to identify potential regions responsible for the chaperone action of Tg, NCBI BLAST pairwise alignments were carried out to compare the amino acid sequence of Tg with that of CLU, a2M, Hp, NS and TTR. These alignments did not result in any large contiguous or significant regions of homology between the Tg sequence and the sequences of known ECs. The NCBI Standard Protein BLAST tool was also employed to search for other proteins of interest (known chaperones or proteins known to interact with amyloid) that share sequence homology with Tg. The results from this search did not yield any proteins of interest sharing significant sequence homology with Tg.

4.5 Discussion One aim of this study was to determine whether or not both bTg and hTg possess a general EC activity. In vitro protein aggregation assays can be used to measure the ability of a putative chaperone to inhibit the aggregation of a wide range of client proteins. Selecting suitable client proteins to test Tg in this way was challenging as the Tg structure is sensitive to reducing agents and temperature. Tg contains 122 cysteine residues, which are involved in forming 61 intrachain disulfide bonds (Lee et al. 2008) and Tg would lose its structural integrity under reducing conditions. Tg is also thermally destabilised at temperatures above ~ 55°C, which induces the dissociation of Tg dimers to monomers (Edelhoch and Metzger 1961; Metzger and Edelhoch

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1961), and it was unknown how this might affect the chaperone activity of Tg. Another challenge is the sheer mass of Tg at 660 kDa. In order to undergo amorphous aggregation in vitro on an experimentally convenient time scale, a relatively high concentration (> 20 μM) of client protein is often required. This can be problematic because, even when using low molar ratios of Tg:client, it can necessitate the use of very high mass concentrations of Tg. These challenges were therefore considered when selecting the client proteins to be used in assessing Tg for chaperone activity.

In this study, Ab and Cal were chosen as amyloidogenic clients. Both of these proteins generate amyloid at 37°C under non-reducing conditions and for most ECs, lower molar ratios of the chaperone:client are typically effective at inhibiting amyloid formation compared to the ratios required to inhibit amorphous protein aggregation (Yerbury et al. 2007; Wyatt et al. 2013). Aβ is a widely-characterised amyloidogenic peptide that quickly forms amyloid when heated to 30°C (Kammerer et al. 2004). Cal is a physiologically relevant, disease-associated protein of the thyroid, making it an ideal candidate to test Tg for EC activity. Although bTg did not specifically inhibit the aggregation of Cal, both bTg and hTg concentration-dependently inhibited the increase in ThT fluorescence associated with the formation of Ab. At molar ratios of bTg:Aβ = 1:5, 1:10, and 1:20, bTg reduced the level of end-point aggregation of Aβ. At lower concentrations, bTg either had no inhibitory effect on Aβ aggregation (bTg:Aβ = 1:400) or increased Aβ aggregation (bTg:Aβ = 1:600). As described in section 3.6 (page 92), it is not uncommon for ECs, when used at very low molar concentrations, to increase the aggregation of amyloidogenic proteins. bTg performed identically to CLU when used at the same molar ratio (CLU:Aβ = 1:5). Neither non-chaperone control (SOD1 or Oval) inhibited Aβ aggregation, confirming that the inhibitory action of Tg was specific. Previous work also tested the effects of bTg on the aggregation of ccβw, which showed a concentration-dependent inhibition of the formation of ThT-reactive amyloid (Frazier 2012).

In vitro protein aggregation assays were also performed with two amorphously-aggregating clients (CS and OT). CS was induced to aggregate at 43°C, well below the temperature required to destabilise Tg (~ 55°C). At the highest molar ratio of Tg:CS tested (Tg:CS = 1:1), both bTg and hTg inhibited CS aggregation by ~ 93% and 88% respectively at the assay endpoint. At a molar ratio of bTg:CS = 1:2 and hTg:CS = 1:2 and 1:4, Tg was less effective and only reduced endpoint CS aggregation by ~ 33%, 68%, and 44%, respectively. At the lowest molar ratios of bTg:CS and hTg:CS tested (1:4 and 1:6 respectively), endpoint CS aggregation actually increased by ~ 38% and 34%, respectively. In this assay, both bTg and hTg outperformed CLU, which only inhibited ~ 56% of endpoint CS aggregation when used at the same molar ratio (CLU:CS = 1:1). When comparing the effectiveness of bTg versus hTg in CS aggregation assays, bTg and hTg were 125

Chapter 4 similarly effective at the highest molar ratio tested (Tg:CS = 1:1), however, at lower molar ratios of Tg:CS, hTg appears to be more effective at inhibiting CS aggregation than bTg.

Both bTg and hTg concentration-dependently inhibited the aggregation of OT. Relative to OT alone, the molar ratios of bTg:OT tested (1:20, 1:15, and 1:5) reduced the level of endpoint OT aggregation by ~ 94%, 73%, and 15% respectively. The molar ratios of hTg:OT tested (1:10, 1:20, 1:30, and 1:40) reduced endpoint OT aggregation by ~ 97%, 90%, 67%, and 36%, respectively. CLU was not tested at molar ratios lower than CLU:OT = 1:1. OT aggregates at 60°C, a temperature known to induce structural changes within the Tg molecule. These structural changes are shown in Figure 4.6. Within 5 min of incubation at 60°C, both bTg and hTg dimers have dissociated into monomers and formed higher molecular weight polymers. Although the effects of these structural changes on the chaperone activity of Tg were unknown, incubating Tg at 60°C for up to 5 h did not abolish its chaperone activity. In fact, both bTg and hTg appeared to be more effective at inhibiting the aggregation of OT versus CS. These apparent differences in the chaperone efficiency of Tg may simply result from different specificities of interaction between Tg and individual client proteins, however the possibility that the chaperone activity of Tg is enhanced upon dissociation into dimers is worth investigating.

Interestingly, increased chaperone activity due to the dissociation of quaternary structures has been reported for other ECs, one example being a2M. The hydrophobicity of a2M increases when the native tetramer dissociates into dimers, and is associated with an increase in its chaperone efficiency (Wyatt et al. 2014). To determine whether or not heating Tg to 60°C had any effect on its surface hydrophobicity, bTg and hTg were either stored at 4°C or incubated at 60°C for 4 h followed by bisANS analysis. These results indicated a significant increase (~ 358% for bTg and ~ 244% for hTg) in the surface-exposed hydrophobicity of Tg relative to Tg that had been stored at 4°C. Heating Tg to temperatures > 55°C is not physiologically relevant in vivo, however, heat is not the only means of inducing changes in the Tg structure. In addition to heat, alkaline pH and low ionic strength can also induce the dissociation of Tg into monomeric subunits (Edelhoch and Metzger 1961; Metzger and Edelhoch 1961; Schneider and Edelhoch 1970). It has also been reported that interactions with Ca2+ ions induce structural changes in Tg (Gentile et al. 1998; Formisano et al. 1983). Though it has only been reported that interactions with Ca2+ ions promote the soluble polymerisation of Tg for colloidal storage (not the dissociation of Tg dimers into monomers), one study reports that the binding of Ca2+ to Tg does increase its surface hydrophobicity (Acquaviva et al. 1991). Oxidants could be another possible inducer of structural changes in Tg. 126

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Many PDDs are thought to be associated with high levels of oxidative stress, and induction of oxidative stress in cells by amyloid is one possible explanation for the cytotoxicity of amyloid. It was shown that exposure to physiologically relevant levels of hypochlorite (an oxidant produced by the immune system) induces the dissociation of the a2M tetramer (Wyatt et al. 2014). Although there are currently no reports of Tg dimers dissociating into monomers in vivo, the human thyroid harbours high levels of ROS, in particular hydrogen peroxide (H2O2), and ROS produced during the coupling of iodine to tyrosine residues by peroxidases during thyroid hormone synthesis (Karbownik-Lewinska and Kokoszko-Bilska 2012). Several disorders of the thyroid are also associated with oxidative stress, such as hypothyroidism and TMC (Chakrabarti et al. 2016; Hosseini-Zijoud et al. 2016). Determining whether or not oxidative stress, or more specifically oxidants, have any effect on the structure of Tg, and if that modified structure has a defined biological function (such as enhanced chaperone action) would further our understanding of the physiological role of Tg in the body and possibly lead to novel therapies for PDDs.

Once it was established that Tg could dose-dependently inhibit the aggregation of wide range of client proteins, it was next assessed for its ability to form HMW complexes with the amorphously- aggregating client protein CS. CS was selected because of the possible client proteins available, it required the lowest final concentration (1.8 μM), it aggregates within 3 h at 43°C (a temperature suitable for maintaining Tg structural integrity), and it does not require the use of reducing reagents.

SEC of bTg + heat-stressed CS sample revealed the formation of a new protein species (not present 7 in samples containing native, non-heated proteins) that eluted in the exclusion limit (V0 > 4 x 10 Da) of a Superoseä 6 10/300 column. This species only appeared when bTg was co-incubated with CS at 43°C and did not appear in the SEC profile of bTg or CS heated separately, or the mixture of the two proteins incubated at 4°C, suggesting the heat-induced formation of a HMW complex between bTg and CS. The purified, putative HMW complex was then assessed by 8% SDS-PAGE under reducing and non-reducing conditions to confirm that both bTg and CS were present in this sample. Non-reducing SDS-PAGE was not very effective at dissociating the complex (indicating a high level of stability). In contrast, reducing the sample dissociated the HMW complex and distinct bands corresponding to both reduced bTg and CS were detected. Although these results strongly suggest that bTg and CS are directly associated in a complex, it does not directly demonstrate a physical association between the two proteins. A “sandwich” ELISA approach was used to confirm that HMW complexes were formed as a result of a direct physical association between bTg and CS. This involved capturing the Tg-CS complex with an 127

Chapter 4 anti-CS antibody bound to an ELISA plate and then probing with an anti-Tg antibody. Results from this assay confirmed the presence of complexes containing both bTg and CS. The complexes were stored at 4°C in PBS/az.

Both DLS and NTA analyses of the purified HMW Tg-CS complexes indicated that the complexes were significantly larger than native bTg or CS alone. DLS analysis indicated smaller mean diameters (45.6 nm, 8.5 nm, and 82.9 nm) for bTg, CS and Tg-CS respectively compared to NTA analysis (the most frequent average mean diameters for bTg, CS and Tg-CS were 84.5 nm, 54.5 nm, and 214.5 nm respectively), and indicated that the Tg-CS solution was monodisperse, which contradicted NTA analysis. The limitations of DLS in resolving polydisperse solutions have been well documented and were considered when using DLS here. It was not surprising that the Tg-CS complexes were polydisperse as this is also the case with CLU-client complexes (Wyatt et al. 2009c). It is possible that the lack of organisation observed in amorphous protein aggregation could lead to the formation of aggregates that vary in both size and structure, which could influence how chaperones interact with them.

This study also aimed to determine if, like other ECs, Tg could protect cells from the cytoxicity of amyloid in vitro. These experiments first required the production of a cytotoxic amyloid species. Cal was selected on the basis that (i) it is a thyroid-resident peptide that is known to form amyloid in vivo, making it physiologically relevant for testing with Tg and (ii) several studies have shown that Cal is cytotoxic, although there is little information describing the exact species responsible for its cytoxicity. One group published two studies reporting that Cal oligomers are the most toxic species (Diociaiuti et al. 2014; Malchiodi-Albedi et al. 2010), which agrees with most literature published regarding the toxicity of other amyloid-forming peptides (such as Ab) (Verma et al. 2015; Kayed and Lasagna-Reeves 2013). Three separate studies also report that Cal amyloid is cytotoxic (Wang et al. 2005; Rymer and Good 2001; Schubert et al. 1995). Neither of these studies verified the composition of their Cal amyloid (oligomer vs mature fibril), however, the methodologies used to generate their Cal amyloid suggests that their sample likely contained mature fibrils.

As it was uncertain which Cal amyloid species would generate a measurable level of cytoxicity, aliquots of Cal were taken at six different timepoints (0, 3, 5, 8, 12, and 30 h) during the aggregation process to ensure a wide range of amyloid species could be tested. Preliminary cytotoxicity assays used Nthy-Ori 3-1 euthyroid follicular cells, however none of the Cal amyloid samples were cytotoxic towards these cells when measured by MTS assays. As cytoxicity may be 128

Chapter 4 cell-specific, a panel of three other cell types (HEK293, PC12 and SH-SY5Y; the latter two are commonly used to assess amyloid toxicity) were tested. To conserve limited stocks of Cal, the cytotoxicity of only one aggregation time point (30 h) was tested with the cells. Again, however, no significant reduction in MTS was observed for cells incubated with Cal when compared to the non-toxic control protein BSA, indicating that the Cal produced in this study was not detectably cytotoxic. As MTS measures metabolic activity (an indirect indicator of cell viability), it could be that this assay was not sensitive enough to measure cytotoxicity in cells that had been starved (no FCS) for nearly three days.

Ab has been extensively used in cell cytoxicity assays and numerous studies have shown that Ab oligomers are more toxic to cells in vitro compared to fibrillar Ab. This study examined the effects of Ab (collected at 0, 0.5, 1, 3 and 12 h aggregation time points) on the viability of SH-SY5Y cells using calcein-AM. The results from this experiment indicated that none of the Ab samples were cytotoxic relative to cells treated with BSA. As none of the aggregates generated in this study were shown to be cytotoxic, the ability of Tg to protect cells from amyloid toxicity was not assessed.

Like many other ECs, Tg is a secreted glycoprotein that can inhibit the aggregation of a wide range of both amorphously aggregating and amyloidogenic client proteins. Like CLU, Tg can form stable, soluble, HMW complexes with amorphously aggregating proteins. Tg is secreted into the thyroid lumen, rather than the blood stream like other ECs, and is historically thought to remain there until needed for thyroid hormone synthesis, however, multiple reports suggest that that is not the case. Although significantly lower than other ECs, serum concentrations of Tg in euthyroid individuals can vary between 10 – 30 ng/mL (within the thyroid lumen, Tg concentrations can reach between 100 – 800 mg/mL) (Wang et al. 2016a; Premawardhana et al. 1994; Bertelsen and Hegedus 1994; Knudsen et al. 2001; Druetta et al. 1998; Feldt-Rasmussen 1978; Herzog et al. 1992; Marino and McCluskey 2000), indicating that Tg is released from the follicular lumen. Other reports suggesting that Tg may have an extrathyroidal role include a description of the renal expression of “thyroid-specific” genes (e.g. thyrotropin, Tg, and Pax8) (Sellitti et al. 2000; Plachov et al. 1990), the expression of putative Tg receptors (including the asialoglycoprotein receptor (ASGPR)) in non-thyroidal cells (hepatocytes and glomerular mesangial of the kidneys), and that fact that circulating plasma Tg is taken up by liver macrophages (where it is degraded) (Luo et al. 2014; Brix et al. 1997). The fact that Tg uptake in vivo occurs in the liver is particularly interesting because radioactively-labelled HMW CLU-client protein complexes injected into rats have also been shown to localise to the liver, where they are subsequently cleared from circulation and degraded within lysosomes (Wyatt et al. 2011). 129

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Thus, although an established extrathyroidal role for Tg is not yet confirmed, there is plenty of evidence to suggest that Tg is not restricted to the thyroid lumen. Interesting questions relevant to this study are what conditions or stimuli trigger the release of Tg into the bloodstream, and could this be used as a therapeutic approach for the selective release of an EC? Future investigations of the mechanisms by which Tg is released into the bloodstream may provide valuable answers.

The intracellular uptake and degradation of Tg by thyroid follicular cells must be highly regulated to ensure appropriate amounts of thyroid hormone are released. Endocytosis of Tg from the lumen typically occurs by way of micropinocytosis, which can result from either fluid-phase pinocytosis and/or receptor-mediated endocytosis (Figure 4.13). Both forms of micropinocytosis result in the invagination of the plasma membrane to form small vesicles (derived from coated pits), which then fuse with endosomes. Fluid-phase non-specific pinocytosis leads to the trafficking of Tg to lysosomes for proteolytic cleavage, however, receptor-mediated endocytosis of Tg can direct the protein into several different intracellular trafficking pathways: (i) Tg is trafficked to the lysosome for proteolytic cleavage, (ii) immature forms of Tg are trafficked to the Golgi for maturation and then recycled back to the colloid, or (iii) Tg bypasses lysosomal degradation and is secreted into the bloodstream (Marino and McCluskey 2000; Vono-Toniolo et al. 2005). The binding of Tg to receptors appears to be regulated by the extent of Tg iodination and glycosylation (in particular, N-acetylglucosamine residues located on the B carbohydrate chain) (Shifrin et al. 1982; Consiglio et al. 1981; Palumbo and Ambrosio 1982; Mezgrhani et al. 1997).

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Figure 4.18 Schematic representation of thyroidal endocytosis of Tg and associated trafficking pathways. The key on the left identifies structural components and the four possible pathways have been labelled. 1) Tg is trafficked to lysosomes for proteolytic cleavage and hormone secretion. 2) Endocytosis occurs via a low-affinity Tg receptor and the protein is trafficked to lysosomes for cleavage and hormone secretion. 3) Immature Tg is internalised by a specific receptor and recycled back to the colloid lumen in the mature form. 4) Tg is internalised by megalin and transported to the basolateral surface for exocytosis into the bloodstream. This process is known as transcytosis and allows Tg to by-pass lysosomal degradation (Marino and McCluskey 2000).

Several receptors have been reported to be involved in the intracellular uptake of Tg (Consiglio et al. 1981; Consiglio et al. 1979; Marino and McCluskey 2000), however, the receptor of interest for this study is megalin. Megalin has been shown to mediate the internalisation and transcytosis of Tg for release into the bloodstream (Marino et al. 2000; Lisi et al. 2006). This process may be involved in the regulation of thyroid hormone production as it allows Tg to by-pass lysosomal degradation. It was also shown that megalin expression in thyrocytes is dependent upon thyroid- stimulating hormone (TSH; Marino et al. 1999; Herzog 1983). This trafficking mechanism may be a viable route to retrieve Tg from the thyroid lumen if an imbalance in proteostasis should occur. Tg has also been shown to bind to the asialoglycoprotein receptor and it has been proposed that this receptor may be involved in the uptake and recycling of immature Tg molecules (Rousset et al. 2015). A better understanding of the mechanisms that mediate the release of Tg into plasma could provide evidence to establish an extrathyroidal role for Tg as an EC.

This study is the first to show that both bTg and hTg have efficient chaperone activity in vitro. As the hTg and bTg sequences are 77% identical and 85% similar (see Appendix I), it was not surprising that both species of Tg are active as chaperones. This study demonstrated that Tg possesses several EC characteristics, however, it would be an important to establish whether or not Tg can protect cells from cytotoxic amyloid. As amyloidoses of the thyroid do exist, a better understanding of Tg’s physiological role as an EC could prove valuable when developing future therapies to treat PDDs in both the thyroid and other regions of the body. 131

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Chapter Five Investigating the Binding and Uptake of HMW Chaperone-client Complexes by Cell Surface Receptors

5.1 Introduction ECs are crucial members of the extracellular protein quality control network in vivo and play an essential role in preventing the aggregation of unfolded proteins associated with many PDDs. The current model suggests that ECs prevent protein aggregation by forming stable soluble, complexes with aggregating proteins and facilitate their removal from the extracellular environment by receptor-mediated endocytosis and intracellular degradation. Several studies provide evidence to support the formation of soluble HMW complexes formed between ECs (e.g. CLU, a2M, and Hp) and stressed, amorphously-aggregating client proteins (Wyatt et al. 2009c; French 2008; Wyatt and Wilson 2013; Yerbury et al. 2005b). In further support of this model, it was also shown that biotinylated CLU-client complexes bind to (currently unknown) cell surface receptors on human blood monocytes and isolated rat liver cells, and, in the latter cell type were internalised and degraded within lysosomes (Wyatt et al. 2011) (Figure 5.1). Although there is strong evidence to support a role for ECs in the removal of toxic protein aggregates from the extracellular environment, little is known regarding which cell surface receptors are involved in the internalisation of chaperone-client complexes.

There are currently seven receptors known to bind established (and putative) ECs that could be involved in the cellular uptake of EC-client complexes, which include megalin (binds CLU, Tg, and TTR), LRP1 (binds a2M and TTR), CD163 (binds Hp), and VLDLR, ApoER2, plexin A4, and TREM2 (which have all been shown to bind CLU) (Table 1.2). Currently, unidentified scavenger receptors have also been implicated in the binding of HMW CLU-client complexes to human blood monocytes and isolated rat liver cells (Wyatt et al. 2011). Wyatt et al. also showed that pre-incubation of human monocytes and rat liver cells with fucoidan inhibited the binding of biotinylated HMW CLU-client complexes to the cell surface by 50 – 75%. Fucoidan is a known antagonistic ligand and inhibitor of class A scavenger receptors (Iesaki et al. 2014), class E scavenger receptors (Oka et al. 1998), and class F scavenger receptors (Berwin et al. 2004). This data suggests that CLU-client HMW complexes bind to scavenger receptors, and likely other receptor types as fucoidan did not inhibit 100% of the binding of the CLU-complexes to cells.

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1. ECs (blue) recognise and bind to 2. The HMW complex binds to one or more misfolded client proteins (black) (currently unknown) cell surface receptors to form HMW complexes. (purple) Cell Membrane

Cell Surface Receptor Lysosome

3. The HMW complex is then internalised 4. The HMW complex is degraded within via receptor-mediated endocytosis and lysosomes trafficked to lysosomes

Figure 5.1 Schematic drawings showing the formation, internalisation, and degradation of HMW chaperone-client complexes. The current accepted model for the chaperone-mediated clearance of misfolded proteins from the extracellular environment is based on data that shows (1) ECs form stable, soluble HMW complexes with misfolded proteins (clients) in vitro (this has only been reported for amorphously aggregating clients), (2) purified HMW chaperone-client complexes bind to various (unidentified) cell surface receptors, (3) the complexes are internalised by cells and are shown to co-localise with lysosomes, and (4) the HMW complexes are proteolytically degraded (Wyatt et al. 2013).

The current study aimed to (i) confirm that HMW CLU-CS, CLU-GST, and Tg-CS complexes bind to various cell surfaces, (ii) identify specific receptors involved in the binding of HMW chaperone-client complexes to cell surfaces, and (iii) determine whether or not chaperone-client complexes are internalised and trafficked to lysosomes for degradation. As all previous work investigating the binding of CLU-client complexes to cell surfaces was conducted using either biotinylated or fluorophore-labelled HMW CLU-client complexes (Wyatt et al. 2011; Dabbs 2013; Frazier 2012), the initial work in this study followed a similar approach (using biotinylated chaperone-client complexes), however, it was later discovered that biotinylation of the complexes led to structural changes (described below), enhancing their binding to cell surfaces. As biotinylation of the chaperone-client complexes was found to be unsuitable for investigating their interactions with cell surfaces, methodology had to be changed. Owing to time limitations, much 133

Chapter 5 of the work described that was conducted with non-biotinylated HMW complexes is preliminary in nature and some work performed using biotinylated HMW complexes could not be repeated with unlabelled complexes.

5.2 Materials As described in Section 2.1, Table 5.1 lists all antibodies used in this study. Mouse IgG 2a control was from Silenus (987720005), streptavidin-Alexa-488 (SA-488) was from Molecular Probes, and streptavidin-HRP was from Sigma-Aldrich (S5512). DNP9 was used as the species-matched negative control antibody for G7 and was produced by Mark Wilson as described in Humphreys et al. 1999.

Table 5.1 List of antibodies and conjugates used in this study. The below table lists the antibodies used in this study and relevant information pertaining to each one, such as host species, conjugation (if applicable), and supplier information. Superscripted numbers were used to differentiate between multiple antibodies against the same antigen.

Antigen Host Species Conjugation Supplier

2,4-dinitrophenyl mouse DNP9; described in Section 2.1 ApoER2 mouse Abcam: ab58216 CLU mouse G7; described in Section 2.1 CS1 rabbit Abcam: ab182888 CS2 rabbit Sapphire Biosciences: LS-C147385 GST1 rabbit Alexa-488 Thermo Fisher Scientific: A-11131 GST2 rabbit Millipore: AB3282 HB rabbit Dako: A0118 Mouse IgG goat Alexa-488 Abcam: ab150117 Mouse IgG goat CF-488 Biotium: 20010 Mouse IgG goat HRP Dako: P0447 Rabbit IgG goat Alexa-488 Life Technologies: A11008 Rabbit IgG goat HRP Dako: P0448 Tg mouse Sapphire Bioscience: 000-13993 VLDLR mouse Abcam: ab75591

5.3 Methods 5.3.1 Mammalian cell culture Mammalian cells were cultured as described in Section 2.2.6. Information regarding each cell line can also be found in Section 2.2.6. For binding experiments, RAW 264.7 cells were harvested between 60 – 70% confluency.

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5.3.2 Expression and purification of GST-tagged RAP The RAP-GST fusion protein was expressed and purified as described in Sections 2.2.8 and 2.2.9.

5.3.3 Formation of HMW chaperone-client complexes The ability of CLU and bTg to form stable, soluble HMW complexes with the amorphously- aggregating client protein CS, and CLU with GST was assessed. All HMW complexes were prepared in Greiner 384-well microplates (50 μL/well). bTg-CS complexes were prepared and purified as described in Section 4.3.4 and CLU-GST complexes were prepared as described in Wyatt et al. 2009c. Briefly, GST (20 μM) was incubated with CLU in PBS at 60°C for a minimum of 2 h at a 1:10 molar ratio of CLU:GST. CLU-CS complexes were prepared under the same assay conditions used to generate bTg-CS complexes. Briefly, CS (1.8 μM) was incubated with CLU (at a 1:1 molar ratio of CLU:CS) in TH buffer at 43°C for a minimum of 4 h. Samples were then collected from the microplates and passed through a SEC column as described in section 4.3.4. The SEC profiles of CLU-GST and CLU-CS are shown in Appendix II. The SEC profiles of both heated and unheated CLU, GST, and CS are described in Wyatt et al. 2009c and were not re- analysed in the current study. Purified HMW chaperone-client complexes were collected and their concentration was estimated using a SpectroMax Plus 384 spectrophotometer (Molecular Devices; measuring A280 in a quartz cuvette). The theoretical extinction coefficients of the chaperone and client in each complex were averaged for these calculations. Theoretical extinction coefficients were calculated using the Expasy ProtParam tool as: bTg E0.1% = 1.104, CLU E0.1% = 0.84, GST E0.1% = 1.69, and CS E0.1% = 1.55. The HMW chaperone-client complexes were stored at 4°C in PBS/az.

5.3.4 Biotinylation of HMW complexes and control proteins Proteins (including HMW chaperone:client complexes; at between 1 – 3 mg/mL) were first dialysed against PBS (pH 8.5). To each solution, 0.25 mg of EZ-Link NHS-Biotin (Pierce Biotechnology; freshly prepared at 40 mg/mL in DMSO) was added per mg of protein. The samples were then incubated with gentle shaking for a minimum of 2 h at room temperature followed by dialysis against PBS/az to remove unconjugated biotin. Biotinylated proteins and HMW complexes were stored at 4°C.

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5.3.5 Transient transfection of HEK293 cells HEK293 cells in DMEM-F12 containing 10% (v/v) FBS were seeded into the wells of Greiner tissue culture-treated 6-well microplates (2 mL/well) and allowed to adhere overnight at 37°C in a humidified incubator containing 5% (v/v) CO2. When cells reached between 60 – 70% confluency, they were transfected using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol. Briefly, the culture media was removed and replaced with 1.5 mL of fresh DMEM-F12 containing 10% (v/v) FBS. Sterile DNA-lipofectamine complexes were pre- formed by incubating 4 µg of DNA in 250 µL with an equal volume of Lipofectamine 2000 diluted 1:25 in serum-free DMEM-F12 for 20 min at room temperature prior to adding the mixture to each well (a total volume of 500 µL/well was added dropwise to each well). Cells were returned to the incubator and left for an additional 48 h. Transfection efficiency was monitored by examining the expression levels of enhanced green fluorescent protein in a control well with a Leica DC500 epifluorescence microscope or by flow cytometry. Table 5.2 lists the specific plasmids used in this study.

Table 5.2 List of plasmids used in this study. The below table lists the plasmids used in this study for transient transfections, the specific protein expressed by each plasmid, and the source from which the plasmid was obtained.

Protein Plasmid Sourced from: Expressed ApoER2 pcDNA3.1(-)(ApoER2) Dr. Yonghe Li EGFP pEGFP-N1 Clontech Laboratories, Inc. SRA-1 pEBS/PL(SRA-1) Dr. Yves Delneste as described in Delneste et al. 2002 SREC-1 pcDNA3.1(SREC-1) Dr. Yves Delneste as described in Jeannin et al. 2005 VLDLR pcDL-SRa296(VLDLR) Dr. Yonghe Li

5.3.6 Cell binding assays using biotinylated HMW chaperone-client complexes To determine whether biotinylated CLU-CS, CLU-GST, and Tg-CS complexes bound to cell surfaces, a panel of cell types, and cells transfected to express various receptors were screened. Cells were washed free of culture medium with ice cold PBS containing 0.01% (w/v) sodium azide prior to detachment. RAW 264.7 and EOC13.31 cells were detached by scraping in PBS containing 0.01% (w/v) sodium azide before being centrifuged at 350 x g for 5 min at 4°C. HepG2 were detached using Acutase (according to the manufacturer’s protocol) and SRA-1 or SREC-1 transfected HEK293 cells (transfected as described in Section 5.3.5) were detached with half- strength PBS containing 5 mM EDTA, pH 7.4. Cells were then washed twice in ice cold PBS containing 0.01% (w/v) sodium azide by centrifugation at 350 x g for 5 min at 4°C. After washing,

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Chapter 5 cells were resuspended in blocking buffer (1% (w/v) BSA in PBS containing 0.01% (w/v) sodium azide) at 1 x 106 cells/mL and approximately 1 x 106 cells/tube were aliquoted into 15 mL polystyrene tubes. Cells were blocked for 1 h on ice and then pelleted by centrifugation (as above). Cells were then resuspended with 100 µL of biotinylated proteins (HMW chaperone-client complexes or negative control proteins) at 100 µg/mL (unless otherwise stated) in blocking buffer and incubated for 1 h on ice. After washing in 15 mL of PBS containing 0.01% (w/v) sodium azide the cells were resuspended in 100 µL blocking buffer containing 2.5 µg/mL SA-488 and incubated on ice for 30 min. Cells were then washed as above and resuspended in 500 µL of cold PBS before analysis by flow cytometry as described in Section 5.3.11. The level of cell-associated background fluorescence was measured by only incubating cells with 2.5 µg/mL SA-488.

5.3.7 Inhibition of HMW chaperone-client complexes binding to cell surface receptors Inhibition of the binding of HMW complexes to cells was trialled using fucoidan, a known inhibitor of scavenger receptors. This study looked at the effects of fucoidan on the binding of biotinylated CLU-GST and non-biotinylated Tg-CS complexes to RAW 264.7 macrophages and both non-biotinylated and biotinylated CLU-GST complexes to EOC13.31 microglia. Cells were detached and blocked as described in Section 5.3.6. After blocking, cells were incubated with or without 500 µg/mL fucoidan for 30 min on ice. Next, cells were washed once by centrifugation for 5 min at 350 x g at 4°C in cold PBS containing 0.01% (w/v) sodium azide and then resuspended in blocking buffer containing biotinylated or non-biotinylated HMW complexes and detected as described in Sections 5.3.6 or 5.3.10, respectively.

5.3.8 Affinity adsorption to identify cell surface receptors for HMW CLU-CS complexes Once it was confirmed that biotinylated HMW chaperone-client complexes bound to the surfaces of the cells tested in this study, isolation of the specific receptors involved was attempted. This approach involved three main steps: (i) incubating cells with or without the HMW complexes, (ii) lysing cells to release membrane-bound proteins (e.g. receptors), and (iii) capturing the complex (in theory) still bound to the receptor, as shown in Figure 5.2. Bound proteins were then released from the beads and assessed by SDS-PAGE to compare the eluates corresponding to cells incubated with and without the complex. If proteins were recovered from the lysate of cells that had been incubated with the complex, but not in the lysate prepared from cells that had not been incubated with complex, these bands would be excised from the gel and identified by mass spectrometry.

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1. 2. 3. Cell Membrane

Cell Surface Cytosol Receptor

HMW Chaperone- client Complex

Figure 5.2 Schematic depiction of receptor affinity adsorption assay methodology. Receptor pull-down assays employed three main steps: 1) HMW chaperone-client complexes were incubated with cells and allowed to bind to cell surface receptors, 2) cell membranes were disrupted to release membrane bound proteins, and 3) cell lysates were incubated with resin that interacts with a specific component of the HMW complex (i.e. either the biotin tag or the individual proteins making up the HMW complex). Some experiments used batch adsorption, whilst others used a home-made affinity column.

As biotinylated CLU-CS, CLU-GST, and Tg-CS were all shown to bind RAW 267.4 cells, this cell line was selected for these experiments. Biotinylated CLU-CS HMW complexes were chosen for preliminary tests as a large amount of the complex was required for these experiments and preparing CLU-CS complexes was more efficient and cost-effective compared to preparing CLU- GST and Tg-CS complexes (which require larger masses of proteins to be used).

Affinity adsorption to streptavidin-Sepharose Cells were detached, blocked, and incubated with or without biotinylated CLU-CS as described in Section 5.3.6, except that the number of cells per sample used was increased from 1 x 106 cells to 10 x 106 cells per sample and the volumes of buffers used were increased accordingly. After incubation with or without the biotinylated complex, 2 x 106 cells from each sample were removed, washed, and incubated with SA-488 for analysis by flow cytometry as described in Section 5.3.11 to confirm that CLU-CS complex had bound to the cell surface. The remainder of the cells were washed thrice in PBS containing 1% (w/v) BSA and 0.01% (w/v) sodium azide before being incubated with 1 mL PBS containing 1% (v/v) Tergitol-type nonyl phenoxypolyethoxylethanol (NP-40), 2 mM EDTA, and a Roche cOmplete protease inhibitor cocktail tablet (according to the manufacturer’s instructions) for 30 min on ice to disrupt the cell membrane. Cellular debris and nuclei were removed by centrifugation at 350 x g for 5 min at 4°C and cell lysates were transferred to 1.5 mL Eppendorf tubes containing 100 µL of a streptavidin Sepharose bead slurry (Invitrogen) that had been washed once in PBS by centrifuging at 1200 x g for 5 min and discarding the

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Chapter 5 supernatant. Cell lysates were incubated with the streptavidin beads at 4°C with gentle shaking for a minimum of 2 h. The beads were then washed thrice with 1.5 mL of PBS containing 0.1% (v/v) Triton X-100 and then twice with 1.5 mL of PBS. The beads were finally resuspended in 100 µL of 1 x SDS sample buffer containing 5% (v/v) 2-mercaptoethanol, boiled for 5 min, and eluted proteins analysed by 10% SDS-PAGE as described in Section 2.2.2. For comparison, purified CLU-CS complex, CLU, and CS were also electrophoresed under reducing conditions.

This approach was unsuccessful due to the interference of many endogenous proteins binding to the streptavidin beads from all lysate samples. The large number of bands that appeared on the gel made it impossible to (i) identify which bands corresponded to the CLU-CS complex without immunostaining (it was not confirmed that the beads actually captured the complex), and (ii) make meaningful comparisons between samples prepared from cells incubated or not with CLU-CS (data not shown).

Immunoaffinity adsorption using anti-CS antibody coupled to Sepharose As streptavidin beads were found to bind many proteins non-specifically from cell lysates, an alternative affinity matrix was produced by coupling polyclonal anti-CS antibody (CS1 in Table 5.1) to CNBr-activated Sepharose beads (GE Healthcare). To conjugate anti-CS to CNBr-activated

Sepharose, anti-CS was first dialysed into coupling buffer (0.1 M NaHCO3, 0.5 M NaCl, pH 8.5) at 4°C with three buffer changes. After dialysis, the A280 of the antibody was taken. The CNBr- activated Sepharose was prepared by first swelling 0.344 g of dry CNBr-activated Sepharose (swells to ~ 1.2 mL when hydrated) in 1 mM HCl for 15 min at room temperature. The bead slurry was then transferred to a sintered glass funnel and washed with 200 mL of 1 mM HCl, followed by 5 mL of coupling buffer. The beads were then swiftly transferred to a 5 mL tube containing 1 mL of anti-CS at 10 mg/mL in coupling buffer. The mixture was placed on a rotating shaker at room temperature for 2 h, and then at 4°C to rotate overnight. The mixture was then centrifuged at 1200 x g for 1 min and the A280 of the supernatant was measured to confirm that the majority of the antibody had been conjugated to the beads (the A280 was ~ 10-fold lower than the original A280). The beads were next washed thrice in coupling buffer by centrifugation at 1200 x g for 5 min at 4°C, followed by a 2 h incubation with filtered blocking buffer (1 M ethanolamine in coupling buffer, passed through a MILLEXÒGV 0.22 μm DuraporeÒ PVDF syringe filter, Merk) at room temperature. The beads were then washed 4 times by centrifugation at 1200 x g at 4°C, alternating between a low pH wash buffer (0.1 M acetic acid, 0.5 M NaCl, filtered as above) and filtered coupling buffer. The beads were then washed twice in filtered PBS/az before being stored in this buffer at 4°C. 139

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It was next confirmed that the newly conjugated anti-CS-CNBr Sepharose could capture the HMW CLU-CS complex by incubating 20 μL of packed beads (containing ~ 20 μg of anti-CS) with 20 μg of CLU-CS at 1 mg/mL overnight on a rotating shaker at 4°C. As a negative control, 20 μg of CLU-CS was also incubated with 20 μL of unconjugated Sepharose 4 beads (GE Healthcare). Both sets of beads were then pelleted, and the supernatant of the anti-CS beads was collected for analysis (to analyse what did not bind to the beads). The beads were next washed thrice with 1.5 mL of PBS containing 0.1% (v/v) Triton X-100 and then twice with 1.5 mL of PBS as described above. Next, the beads were resuspended in 50 μL of 1x SDS sample buffer and boiled for 5 min to release the proteins captured by the beads. The beads were then pelleted by centrifugation at 1200 x g for 5 min. The supernatants were transferred to clean 1.5 mL Eppendorf tubes and 5% (v/v) 2- mercaptoethanol was added to reduce the samples. The supernatants were boiled for an additional 5 min and assessed by 10% SDS-PAGE as described in Section 2.2.2. For comparison, 10 μg of purified CLU-CS complex and 5 μg each of purified CLU and CS were also electrophoresed under reducing conditions.

After establishing that the anti-CS Sepharose did capture the CLU-CS complex, a small homemade column was assembled using a glass Pasteur pipette packed at the bottom with ~ 5 mm of absorbent cotton (to retain the beads). Tubing was attached to the pipette tip so that clamping could be used to stop the buffer flow. The anti-CS beads (~ 0.5 mL) were carefully transferred to the column and washed with 10 mL of filtered PBS/az. The column was stored in filtered PBS containing 0.01% (w/v) sodium azide at 4°C.

In an attempt to use anti-CS-Sepharose to capture CLU-CS (and potentially bound receptors), RAW 264.7 cells and non-biotinylated CLU-CS complexes were used. Cells were detached, blocked, and incubated with or without non-biotinylated CLU-CS complex as described in Section 5.3.6, except that 150 x 106 cells per sample were used. After incubating cells with or without non-biotinylated CLU-CS, cells were washed thrice in PBS containing 1% (w/v) BSA and 0.01% (w/v) sodium azide before being incubated with 15 mL PBS containing 1% (v/v) NP-40, 2 mM EDTA, and a Roche cOmplete protease inhibitor cocktail tablet (according to the manufacturer’s instructions) for 30 min on ice. Cellular debris and nuclei were removed by centrifugation at 350 x g for 5 min at 4°C and cell lysates were transferred to the homemade anti-CS-CNBr column. Prior to addition of the lysates, the column had been prepared by first washing with elution buffer (0.1 M glycine containing 1.2% (w/v) b-Dog (AG Scientific), pH 2.4). The column was then equilibrated into PBS containing 1.2% (w/v) b-Dog. All column wash and elution buffers contained a detergent to prevent the precipitation of membrane-bound proteins (i.e. receptors). b- 140

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Dog was specifically chosen because it is non-ionic (can be used at a low pH) and it forms micelles small enough to flow through a 3 kDa VivaSpin concentrator (Millipore). Each sample was passed over the column four consecutive times to ensure the sample had adequate time to bind. The column was then washed with 5 mL of PBS containing 1.2% (w/v) b-Dog, pH 7.4, followed by 2.7 mL of elution buffer to elute bound proteins. The eluent was collected in a 1.5 mL Eppendorf tube containing 300 μL of 1 M Tris, pH 8, to immediately increase the sample pH. The resulting sample pH was checked using litmus strips to confirm that it was between 7 – 8 (1 M Tris was added if required). The samples were then concentrated using a 3 kDa VivaSpin concentrator to ~ 120 μL. To each sample, 40 µL of 4 x SDS sample buffer and 5% (v/v) 2-mercaptoethanol were added and the samples were boiled for 5 min before being analysed by 15% SDS-PAGE as described in Section 2.2.2. 1 μg each of purified CLU and CS, and 5 μg of purified CLU-CS were also electrophoresed under the same conditions. The gel was first stained as described in Section 2.2.2, followed by silver staining using a Pierceä silver staining kit (Thermo Fisher).

In addition to the adsorption affinity assay, to confirm that the complex had bound to the cell surface, 2 x 106 cells from each sample (prior to cell lysis) were immunostained (described in Section 5.3.10) with undiluted G7 or DNP9 hybridoma supernatant and analysed by flow cytometry as described in Section 5.3.11.

5.3.9 Analysis of non-biotinylated and biotinylated HMW chaperone-client complexes 5.3.9.1 SDS-PAGE analyses To determine whether there were structural differences between biotinylated and non-biotinylated chaperone-client complexes, 15 μg each of the non-biotinylated and biotinylated complex was assessed by 10% SDS-PAGE (as described in Section 2.2.2) under non-reducing and reducing conditions. For comparison, 5 μg of the individual proteins making up the complex were also included in these analyses.

5.3.9.2 Dot blot analyses Dot blot analysis was also used to examine whether there were any significant structural differences between non-biotinylated and biotinylated chaperone-client complexes. As antibodies recognise specific motifs on proteins, both the non-biotinylated and biotinylated versions of the complex should be detectable if they are structurally similar. For dot blot analysis of CLU-CS complexes, 1, 0.5, 0.25, and 0.125 μg each of non-biotinylated and biotinylated CLU-CS was dotted onto a membrane. To ensure the antibodies bound specifically, 1, 0.5, 0.25, and 0.125 μg each of positive and negative control proteins were also included in the analysis. For dot blot 141

Chapter 5 analysis of CLU-GST complexes, 5 μg each of non-biotinylated and biotinylated CLU-GST, and 1 μg each of positive and negative control proteins were dotted onto membranes. Membranes were blocked, washed, and imaged as described in Section 2.2.4. Blots containing CLU-CS complexes were detected using 4 methods: (i) using 0.5 μg/mL of anti-CS1 and a goat anti-rabbit IgG-HRP conjugate at 0.125 μg/mL, (ii) using 0.5 μg/mL of anti-CS2 and a goat anti-rabbit IgG- HRP conjugate at 0.125 μg/mL, (iii) using undiluted G7 supernatant and a goat anti-mouse IgG- HRP conjugate at 0.25 μg/mL, and (iv) using 2 μg/mL of conjugated streptavidin-HRP. Blots containing CLU-GST complexes were detected using 1 μg/mL of either anti-GST1 or anti-GST2 and a goat anti-rabbit IgG-HRP conjugate at 0.125 μg/mL.

5.3.9.3 NTA analyses To compare the mass of non-biotinylated versus biotinylated CLU-CS HMW complexes, NTA was used as described in Section 4.3.4. NTA was used as a third confirmatory approach to examine whether there were structural differences between the non-biotinylated and biotinylated complexes.

5.3.9.4 Binding to RAW 264.7 cells The binding to RAW 264.7 cells of non-biotinylated and biotinylated CLU-CS complexes was compared. The cells were incubated with the different complexes as described in Section 5.3.10, and then incubated with a mixture of anti-CS1 and anti-CS2 at 2.5 µg/mL. Bound anti-CS antibody was detected with 2.5 µg/mL goat anti-rabbit IgG-Alexa-488. Cells were analysed by flow cytometry as described in Section 5.3.11.

5.3.10 Cell binding assays using non-biotinylated HMW chaperone-client complexes To determine whether non-biotinylated HMW complexes bind to RAW 264.7, EOC13.31, and transfected HEK293 cells (transfected as described in Section 5.3.5) expressing ApoER2 and VLDLR, cells were detached, washed, and blocked as described in Section 5.3.6. After washing in 15 mL of PBS/az the cells were resuspended in 100 µL of primary antibody diluted in blocking buffer (with the exception of G7 and DNP9 supernatants, which were used undiluted) and incubated on ice for 30 min. Cells were then washed as above before resuspension in 100 µL of blocking buffer containing an appropriate fluorophore-conjugated secondary antibody and again incubated on ice for 30 min. Cells were then washed and resuspended in 500 µL of cold PBS before being analysed by flow cytometry as described in Section 5.3.11. In each case, the level of cell-associated background fluorescence was measured by substituting a species-matched negative control primary antibody for the specific primary antibody. The specific antibodies, and the 142

Chapter 5 concentrations used for each test are described in the results (Section 5.4.6). Table 5.1 lists the antigen, species, conjugate, and supplier for all antibodies used in this study.

5.3.11 Flow cytometry analysis For flow cytometry experiments, cells were resuspended in 500 µL of cold PBS and held on ice. Propidium iodide (PI) (Sigma Aldrich) was added at a final concentration of 1 µg/mL to each tube immediately prior to analysis using a Fortessa LSR II (BD Biosciences) flow cytometer. Discrete populations (i.e. cellular debris, single cells, and cell aggregates) were gated based on forward scatter and side scatter and viable cells were gated based on PI exclusion. The acquired data was analysed using FlowJo version 9 software (TreeStar Inc.).

5.4 Results Initial work in this study used biotinylated complexes (as used by previous members in our laboratory), however, during the project it became apparent that biotinylation of the complexes induced structural changes that affected their binding to cells. Thus all subsequent experiments were performed with non-biotinylated HMW complexes.

5.4.1 Neither biotinylated CLU-CS nor biotinylated Tg-CS bind to SRA-1 or SREC-1 transfected HEK293 cells To investigate the binding of biotinylated CLU-CS and biotinylated Tg-CS binding to SRA-1 and SREC-1 scavenger receptors, non-transfected HEK293 cells and cells that had been transfected to express SRA-1 and SREC-1, were incubated with 10 μg/mL of an acetylated Dil-labelled LDL probe (Molecular Probes) or 50 μg/mL of biotinylated complexes. Acetylated LDL preferentially binds to scavenger receptors, rather than LDL receptors, and was used to probe for the expression of SRA-1 and SREC-1. Both SRA-1 (pink curve) and SREC-1 (green line) transfected cells showed an increase in Dil fluorescence after being incubated with acetylated LDL compared to non-transfected cells that had been incubated with the probe (black line), indicating that the level of scavenger receptor expression had increased following transfection (Figure 5.3 A). Using antibodies that specifically recognise SRA-1 and SREC-1 would confirm the expression of these particular receptors, however, these were not used in preliminary tests and investigations using these receptors were not continued as neither biotinylated CLU-CS (Figure 5.3 B) nor biotinylated Tg-CS (Figure 5.3 C) bound to the transfected cells (SA-488 fluorescence did not increase compared to non-transfected cells incubated with the complexes).

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% Max of

Dil Fluorescence

Non-transfected + Dil-LDL Probe c SRA-1 transfected + Dil-LDL Probe SREC-1 transfected + Dil-LDL Probe

B C

% Max of % Max of

SA-488 Fluorescence SA-488 Fluorescence

c Non-transfected + biotinylated CLU-CS c Non-transfected + biotinylated bTg-CS

SRA-1 transfected + biotinylated CLU-CS SRA-1 transfected + biotinylated bTg-CS

SREC-1 transfected + biotinylated CLU-CS SREC-1 transfected + biotinylated bTg-CS

Figure 5.3 Flow cytometry analysis of transfected HEK293 cells expressing SRA-1 and SREC-1. Cells were incubated with A) an acetylated Dil-labelled LDL probe, B) biotinylated CLU-CS, or C) biotinylated Tg-CS. The x-axes represent fluorescence intensities, the y-axes indicate the frequency of events (%), and the keys identify each sample. This experiment was performed only once.

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5.4.2 Biotinylated chaperone-client complexes bind to the surface of RAW 264.7 macrophages Biotinylated CLU-CS bound to RAW 264.7 cells, indicated by the large increase in fluorescence of cells incubated with the biotinylated complex and detected with SA-488 (pink line) compared to cells that had only been incubated with SA-488 (black line) (Figure 5.4). The biotinylated negative control proteins BSA, Oval, and a-Lac, did not bind significantly to the cells.

No protein

+ biotinylated CLU-CS + biotinylated BSA

% Max of + biotinylated Oval

+ biotinylated a-Lac

SA-488 Fluorescence

Figure 5.4 Flow cytometry analysis of biotinylated CLU-CS and control proteins binding to RAW 264.7 cells. Cells were incubated with or without biotinylated proteins as described in the key and bound proteins were detected with 2.5 µg/mL SA-488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on forward scatter and side scatter alone. The results shown are representative of two independent experiments.

Biotinylated Tg-CS also bound to the surface of RAW 264.7 macrophages, shown by the large increase in fluorescence of cells that had been incubated with the complex and detected with SA- 488 (green line) versus cells only incubated with SA-488 (black line) (Figure 5.5). Previous work showed that the level of binding of FITC-labelled BSA, CLU, Tg, and CS to RAW 264.7 cells is significantly lower than that of FITC-labelled Tg-CS (Frazier 2012) (see Appendix III), therefore, these controls were not included in the current study.

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No protein

% Max of + biotinylated Tg-CS

SA-488 Fluorescence

Figure 5.5 Flow cytometry analysis of biotinylated Tg-CS binding to RAW 264.7 cells. Cells were incubated with or without biotinylated Tg-CS as indicated in the key and bound protein were detected with 2.5 µg/mL SA-488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Viable cells in this experiment were gated based on PI exclusion. The results shown are representative of two independent experiments.

Biotinylated CLU-GST also bound to RAW 264.7 cells, indicated by the large increase in SA-488 fluorescence of cells incubated with the complex (green line) versus cells that had not been incubated with CLU-GST (black line) (Figure 5.6). The binding of biotinylated GST was not assessed in preliminary tests. Pre-incubation of the macrophages with fucoidan inhibited CLU- GST binding by more than 50% (cyan line), indicating that biotinylated CLU-GST complexes bind, at least in part, to scavenger receptors on the surface of these cells. Before other inhibitors could be tested, results were obtained suggesting that non-biotinylated chaperone-client complexes were not binding to RAW 264.7 (discussed in Sections 5.4.6). This required further investigation before any additional work could be performed using biotinylated chaperone-client complexes.

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No protein + biotinylated CLU-GST

% Max of + fucoidan + biotinylated CLU-GST

SA-488 Fluorescence

Figure 5.6 Flow cytometry analysis of biotinylated CLU-GST binding to RAW 264.7 cells. Cells were incubated with or without biotinylated CLU-GST as indicated in the key and bound protein was detected with 2.5 µg/mL SA-488 (see key). The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Viable cells in this experiment were gated based on PI exclusion. The results shown are representative of two independent experiments.

5.4.3 Biotinylated CLU-GST and biotinylated Tg-CS bind to EOC13.31 microglial cells Flow cytometry analysis of EOC13.31 cells that had been incubated with biotinylated CLU-GST showed a significant increase in fluorescence compared to cells that had been incubated with SA- 488 only, indicating that the biotinylated complex binds to the surface of these cells (Figure 5.7 A; green vs black lines). The extent of binding of biotinylated Tg-CS to the surface of EOC13.31 cells was significantly lower than that of biotinylated CLU-GST (Figure 5.7 B). Appropriate negative controls would need to be tested to confirm that the binding of biotinylated Tg-CS is specific, however, these controls were not tested in the current study. Nor did this study test the binding of biotinylated CLU-CS to EOC13.31 cells.

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A B

% Max of % Max of

SA-488 Fluorescence SA-488 Fluorescence

c No protein No protein + biotinylated CLU-GST + biotinylated Tg-CS

Figure 5.7 Flow cytometry analysis of biotinylated CLU-GST and Tg-CS binding to EOC13.31 cells. A) Cells were incubated with or without biotinylated CLU-GST and bound protein was detected with 2.5 µg/mL SA-488. B) Cells were incubated with or without biotinylated Tg-CS and bound protein was detected with 2.5 µg/mL SA-488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Viable cells in this experiment were gated based on PI exclusion. The results shown represent a single experiment.

5.4.4. Biotinylated CLU-GST, but not biotinylated Tg-CS, binds to HepG2 liver cells Flow cytometry detected biotinylated CLU-GST complexes bound to the surface of HepG2 liver cells, noted by the large increase in fluorescence for cells incubated with the biotinylated complex versus cells incubated with SA-488 only (Figure 5.8 A, green vs black line). Biotinylated Tg-CS did not bind to this cell type (Figure 5.8 B).

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A B

% Max of % Max of

SA-488 Fluorescence SA-488 Fluorescence

No protein No protein + biotinylated CLU-GST + biotinylated Tg-CS

Figure 5.8 Flow cytometry analysis of biotinylated CLU-GST and Tg-CS binding to HepG2 cells. A) Cells were incubated with or without biotinylated CLU-GST and bound protein was detected with 2.5 µg/mL SA-488. B) Cells were incubated with or without biotinylated Tg-CS and bound protein was detected with 2.5 µg/mL SA-488 (see keys). The x-axis is fluorescence intensity, the y-axis indicates the frequency of events (%). Viable cells in this experiment were gated based on PI exclusion. This experiment was performed twice with CLU-GST complexes and once with Tg-CS complexes.

5.4.5 Anti-CS recognised and captured CLU-CS complexes As mentioned in Section 5.3.8, streptavidin-Sepharose beads could not be used to capture biotinylated CLU-CS complexes due to the high level of non-specific binding (data not shown). To circumvent this issue, an affinity column was designed using anti-CS antibody that, in theory, would specifically bind CS contained in CLU-CS complexes.

It was first necessary to confirm that anti-CS could bind to CS complexed with CLU and pull the complex out of solution. Purified CLU, CS, and CLU-CS, and eluates from anti-CS-Sepharose and unconjugated Sepharose 4B that had been incubated with CLU-CS, were analysed by reducing 10% SDS-PAGE (Figure 5.9). After incubation with CLU-CS, the anti-CS beads were pelleted and the supernatant was collected for SDS-PAGE analysis to determine if any CLU-CS did not bind to the beads. Due to the possibility of bead saturation, a small amount of CLU-CS may not have bound to the beads (a small amount of CS can be seen in the last sample on the gel). These results indicated that the anti-CS beads were able to capture the CLU-CS complex; bands corresponding to the approximate sizes of CLU and CS appear in this sample but were not detected in the corresponding Sepharose 4B eluate. Another possibility is that some of the anti-CS- 149

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Sepharose beads were accidentally collected when removing the supernatant from this sample, and following denaturation, bound CLU-CS was released from the beads. Three bands (yellow arrows) did appear in both the anti-CS beads incubated with CLU-CS sample and in the supernatant sample which do not correspond to CLU or CS. These unidentified bands may correspond to eluted anti- CS antibody as no other proteins were present in the sample, however this was not confirmed.

-CS -CS CNBr 1 - CNBr 1 - -CS

Molecular -CS CS CS - Anti Markers - Anti Sepharose 4B Sepharose + CLU Supernatant (kDa) CLU CS CLU Sepharose + +CLU CLU

200 150

100 75

50 Anti-CS

37 CLU

Anti-CS

Anti-CS 25

Figure 5.9 Reducing 10% SDS-PAGE analysis confirms that anti-CS-CNBr Sepharose captures the CLU-CS complex. Molecular makers are indicated at the left of the figure and sample identities are indicated at the top. For comparison, 5 μg each of CLU (pink arrow) and CS (blue arrow) and 10 μg of CLU-CS complex were also electrophoresed. Yellow arrows indicate samples that may correspond to eluted anti-CS antibody. The gel was stained with Coomassieä Brilliant Blue as described in Section 2.2.2. This gel represents a single experiment.

5.4.6 Anti-CS affinity adsorption assays did not capture receptor-bound CLU-CS Once confirmed that anti-CS-Sepharose could bind and capture CLU-CS complexes from solution, affinity adsorption was attempted. Analysis by reducing 15% SDS-PAGE showed no difference in the protein composition of eluates from affinity beads after incubation of the beads with lysates of cells that had been incubated with or without CLU-CS (Figure 5.10). Although several

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Chapter 5 unidentified proteins did appear in both eluate samples, there is no obvious difference in the protein composition of these eluates.

Molecular -CS Lysates Markers -CS U (kDa) CLU CS Lysates + CLU CL

200 150 100 75

CS 37 CLU

Figure 5.10 Reducing 15% SDS-PAGE analysis reveals no difference in the protein composition of affinity adsorption eluates from lysates prepared from cells incubated with or without the CLU-CS complex. Molecular makers are indicated at the left of the figure and sample identities are indicated at the top. For comparison, 1 μg each of CLU (pink arrow) and CS (blue arrow) and 5 μg of CLU-CS complex were also electrophoresed. The gel was stained using a Pierceä silver staining kit according to the manufacturer’s instructions. Results shown represent a single experiment.

There are two plausible reasons as to why bands corresponding to CLU or CS did not appear in the affinity adsorption eluates corresponding to RAW264.7 cells incubated with the CLU-CS complex: (i) only a small amount of the complex binds to the cell surface and not enough sample was loaded onto the gel to detect it (despite silver staining being highly sensitive), or (ii) the complex did not bind to the cells in this experiment. To address the latter possibility, a simultaneous flow cytometry experiment was conducted to confirm that the CLU-CS complex actually bound to cells (Figure 5.11). The results indicated that CLU-CS did not bind to the RAW 264.7 cells used in this experiment.

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+ CLU-CS + DNP9 + CLU-CS + G7

% Max of

Alexa-488 Fluorescence

Figure 5.11 Flow cytometry analysis to detect CLU-CS binding to RAW 264.7 cells. Cells were incubated with CLU-CS and then undiluted G7 or DNP9 supernatant as described in the key. Bound antibodies were detected with 2.5 µg/mL goat anti-mouse IgG-Alexa-488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Viable cells were gated based on PI exclusion and this figure is representative of two independent experiments.

As biotinylated CLU-CS had bound to RAW 264.7 cells, it was initially surprising that binding of non-biotinylated CLU-CS was not detected. To avoid the possibility that G7 supernatant could not bind to CLU in the complex, the above experiment was repeated, using 2.5 µg/mL each of two different anti-CS antibodies (Table 5.1) Bound primary antibody was detected with 2.5 µg/mL goat anti-rabbit IgG-Alexa-488. Cells were also incubated with or without biotinylated CLU-CS as a positive binding control. Results from this experiment were similar to those shown in Figure 5.11 for the detection of CLU-CS with G7 supernatant; i.e. bound CLU-CS was not detected with G7 (Figure 5.12 A). In addition, binding of CLU-CS to RAW 264.7 cells was not detected using either of the anti-CS antibodies (Figure 5.12 B & C). In contrast, binding of biotinylated CLU-CS to RAW 264.7 cells was detected with SA-488 (Figure 5.12 D). These results indicated that either (i) the antibodies being used were not capable of detecting the bound complex, or (ii) biotinylated CLU-CS binds to RAW 264.7 cells, but non-biotinylated CLU-CS does not (suggesting that the complexes are somehow different).

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A B

% Max of % Max of

Alexa-488 Fluorescence Alexa-488 Fluorescence

1 + CLU-CS + DNP9 + anti-CS + CLU-CS + G7 + CLU-CS + anti-CS1

C D

% Max of % Max of

Alexa-488 Fluorescence SA-488 Fluorescence

+ anti-CS2 No protein

+ CLU-CS + anti-CS2 + biotinylated CLU-CS

Figure 5.12 Flow cytometry analysis showing that non-biotinylated CLU-CS is not detected on the surface of RAW 264.7 cells with anti-CLU or anti-CS antibodies. Cells were incubated with or without CLU-CS or biotinylated CLU-CS. Bound complexes were then detected with A) undiluted G7 or DNP9 supernatants and 2.5 µg/mL goat anti-mouse IgG-Alexa-488, B) 5 µg/mL anti-CS1 (Table 5.1) and 2.5 µg/mL goat anti-rabbit IgG-Alexa-488, C) 5 µg/mL anti-CS2 (Table 5.1) and 2.5 µg/mL goat anti-rabbit IgG-Alexa-488, or D) SA-488. The keys identify each sample, the x-axis is fluorescence intensity, and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on forward and side scatter only and the results shown are representative of two independent experiments.

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5.4.7 The biotinylation of HMW chaperone-client complexes modifies their structure SDS-PAGE analysis indicated that there were structural differences between biotinylated and non- biotinylated chaperone-client complexes. Both the complex, and the individual proteins making up the complex were analysed under non-reducing and reducing conditions on the same gel. Under non-reducing conditions, some CLU (pink and orange arrows) dissociated from the non- biotinylated CLU-CS complex, however, no bands corresponding to CS (blue arrow) were observed and much of the complex remained trapped at the bottom of the sample well (Figure 5.13). It should be noted that purified human CLU typically contains HMW species that react with anti-CLU antibodies. The exact identities of the HMW CLU species are not known, but they are likely to be either oligomerised CLU or complexes formed between CLU and other proteins in plasma. Biotinylated CLU-CS did not detectably dissociate into its CLU and CS components; most of it remained in HMW form that did not enter the gel matrix. Under reducing conditions, most of the non-biotinylated CLU-CS complex dissociated into CLU and CS, but the biotinylated complex again remained mostly trapped at the bottom of the sample well. Two distinct bands did appear when the biotinylated CLU-CS complex was reduced that are not detected in reduced CLU-CS; one band migrating above 200 kDa (orange arrow), which may represent a HMW CLU species, and one band migrating between 37 and 50 kDa, which is just above where CS migrates on the gel (probably representing biotinylated CS). The additional mass of conjugated biotin (~ 244 Da per biotin group) may result in CS running slightly slower than normal, though this was not confirmed.

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CS -CS -

Molecular

CS Markers -CS - (kDa) CLU CS CLU Biotinylated CLU CLU CS CLU Biotinylated CLU

HMW HMW CLU 200 CLU 150 100 75 CLU

50 CS CS 37 CLU

Non-reduced Reduced Figure 5.13 SDS-PAGE analysis of CLU-CS and biotinylated CLU-CS complexes. 5 µg each of CLU, CS, and 15 µg each of CLU-CS and biotinylated CLU-CS were assessed under non-reducing (left) and reducing conditions (right). Molecular markers are indicated on the left, samples are identified at the top of the figure, and labelled arrows identify specific proteins on each gel. The gel was stained with Coomassieä Brilliant Blue as described in Section 2.2.2. Images were cropped to remove empty wells and this gel represents a single experiment.

Under reducing conditions, most of the non-biotinylated CLU-CS complex dissociated into CLU and CS, but the biotinylated complex again remained mostly trapped at the bottom of the sample well. Two distinct bands did appear when the biotinylated CLU-CS complex was reduced that are not detected in reduced CLU-CS; one band migrating above 200 kDa (orange arrow), which may represent a HMW CLU species, and one band migrating between 37 and 50 kDa, which is just above where CS migrates on the gel (probably representing biotinylated CS). The additional mass of conjugated biotin (~ 244 Da per biotin group) may result in CS running slightly slower than normal, though this was not confirmed.

SDS-PAGE analysis of CLU-GST also indicated differences in the structure of the non- biotinylated and biotinylated complexes (Figure 5.14). Similar to CLU-CS, under non-reducing conditions CLU-GST partially dissociated into CLU and GST, and some of the complex remained too large to enter the gel. Under these conditions, biotinylated CLU-GST did not dissociate into monomeric CLU and GST, but instead remained trapped at the bottom of the sample well. A small

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Chapter 5 amount of HMW CLU (orange arrow) was detected in both the non-biotinylated and biotinylated CLU-GST samples.

-GST -GST

Molecular GST GST Markers - - (kDa) CLU GST CLU Biotinylated CLU CLU GST CLU Biotinylated CLU

HMW 200 CLU 150 Lane 100 Smearing 75 CLU

37 CLU

GST

25 GST Unknown

20 Protein Non-reduced Reduced

Figure 5.14 SDS-PAGE analysis of CLU-GST and biotinylated CLU-GST. CLU, GST, CLU-GST, and biotinylated CLU-GST were assessed under non-reducing (left) and reducing (right) conditions. Molecular markers are indicated on the left, samples are identified at the top of the figure, and labelled arrows identify specific proteins on each gel. The gel was stained with Coomassieä Brilliant Blue as described in Section 2.2.2. Images were cropped to remove empty wells and this gel represents a single experiment.

Interestingly, both CLU-GST and biotinylated CLU-GST complexes dissociated into free CLU and GST under reducing conditions. The CLU and GST that had dissociated from biotinylated CLU-GST complexes migrated slightly more slowly than CLU and GST from non-biotinylated complexes, which is most likely due to the mass of their conjugated biotin groups. There also appears to be some band smearing through the top half of the gel in the biotinylated CLU-GST sample that is not observed for non-biotinylated CLU-GST. When reduced, an unidentified protein that migrates between 20 and 25 kDa (yellow arrow) appears in both CLU-GST and biotinylated CLU-GST (slightly higher molecular weight) but does not correspond to either CLU or GST alone. The identity of this species in unknown.

Comparisons of Tg-CS and biotinylated Tg-CS by non-reducing SDS PAGE were difficult as there was little protein visible on the gel in these samples due to neither form of the complex dissociating, however, Tg-CS does appear to be structurally different to biotinylated Tg-CS 156

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(Figure 5.15). Upon close examination, a small amount of dissociated Tg (pink arrows) and CS (blue arrow) can be seen in the non-biotinylated Tg-CS sample, however, most of the complex remained too large to enter the gel matrix. Compared to Tg-CS, an even smaller amount of Tg dissociated from biotinylated Tg-CS, and there was no free CS detected in this sample. Making comparisons between such small amounts of protein was done with caution, as there are other possible explanations for the subtle differences observed here (e.g. one sample containing slightly more total protein than the other). Increasing the total amount of protein loaded for each sample and/or using a more sensitive stain may have allowed for more accurate conclusions. The same amounts of Tg-CS and biotinylated Tg-CS were also assessed under reducing conditions and free Tg and CS were easily detected in the non-biotinylated complex sample. CS was detected in the biotinylated Tg-CS sample, running at a slightly higher molecular weight compared to the corresponding band in the non-biotinylated sample. Under reducing conditions, Tg dissociates into multiple bands, ranging in size from 100 to >200 kDa. Bands corresponding to reduced Tg were detected in both reduced Tg-CS and biotinylated Tg-CS samples, however, a significantly greater amount of one of the reduced Tg bands (running at >200 kDa) (yellow arrow) was detected in reduced biotinylated Tg-CS compared to reduced Tg or Tg-CS.

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-CS

Molecular

Markers tinylated Tg -CS -CS (kDa) Tg CS Tg Bio Tg CS Tg

> 200 kDa Tg Tg 200 Tg 150

100 75

50 CS CS

37 Non-reduced Reduced Figure 5.15 SDS-PAGE analysis of Tg-CS and biotinylated Tg-CS complexes. Tg, CS, Tg-CS, and biotinylated Tg-CS were assessed under non-reducing (left) and reducing (right) conditions. Molecular markers are indicated on the left, samples are identified at the top of the figure, and labelled arrows identify specific proteins on each gel. The gel was stained with Coomassieä Brilliant Blue as described in Section 2.2.2. Images were cropped to remove empty wells and this gel is representative of one experiment.

Dot blot analysis also showed differences in the ability of antibodies to recognise non-biotinylated versus biotinylated chaperone-client complexes. Dot blot analysis confirmed that two different anti-CS antibodies (Table 5.1) specifically recognised CS present in non-biotinylated CLU-CS complexes but not in biotinylated CLU-CS complexes (Figure 5.16). The chemiluminescence signal from anti-CS1 detecting purified CS is quite strong compared to detection of the CLU-CS complex, which is not surprising due to the possibility of CS molecules being partially buried within the complex and inaccessible to the antibody. There was no detectable difference in anti- CS2 recognising purified CS versus non-biotinylated CLU-CS complexes, suggesting that the CS motif recognised by anti-CS2 was readily available and not obscured within the structure of the BSA complex. The negative control proteins CLU or BSA did not generate a detectable chemiluminescence signal when incubated with anti-CS1 or anti-CS2, indicating that both antibodies specifically recognised CS. One surprising observation was that neither anti-CS1, nor anti-CS2 bound to biotinylated CLU-CS complexes, further suggesting that the biotinylated complexes were structurally different to the non-biotinylated complex.

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μg μg

1 0.5 0.25 0.125 1 0.5 0.25 0.125

CLU-CS CLU-CS

Biotinylated Biotinylated CLU-CS CLU-CS CS CS

CLU CLU BSA BSA

Detection with anti-CS1 Detection with anti-CS2

Figure 5.16 Dot blot detection of CLU-CS complexes with anti-CS antibodies. Anti-CS antibodies (Table 5.1) were used at 0.5 μg/mL to probe membranes containing 1, 0.5, 0.25, and 0.125 μg each of CLU- CS, biotinylated CLU-CS, CS, CLU, and BSA. Samples are indicated at the left of the figure and the amount of protein (μg) dotted onto the membrane is indicated at the top of the figure. These dot blots were performed once.

In similar assays, G7 supernatant specifically recognised CLU present in non-biotinylated CLU- CS complexes but was less effective at detecting CLU present in the corresponding biotinylated complexes (Figure 5.17). A weaker chemiluminescence signal was detected for G7 bound to 1 μg of biotinylated CLU-CS versus 1 μg of non-biotinylated complex, and no signal was detected for smaller amounts of biotinylated CLU-CS. G7 did not bind to CS or BSA, indicating that the antibody was specifically interacting with CLU on the membrane. Detection of streptavidin-HRP bound to biotinylated BSA and biotinylated CLU-CS confirmed the successful biotinylation of these proteins (Figure 5.17).

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μg

1 0.5 0.25 μg CLU-CS 1 0.5 0.25 0.125 Biotinylated CLU CLU-CS

CLU CLU-CS

Biotinylated CS CLU-CS

CS BSA

BSA Biotinylated BSA

Detection with G7 Supernatant Detection with streptavidin-HRP

Figure 5.17 Dot blot detection of CLU-CS complexes with G7 anti-CLU and streptavidin-HRP. Undiluted G7 hybridoma supernatant (left) or 2 μg/mL streptavidin-HRP (right) were used to probe 1, 0.5, 0.25, and 0.125 μg each of membrane-immobilised CLU, CLU-CS, biotinylated CLU-CS, CS, and BSA. Samples are indicated to the left of the blot images and the amount of protein dotted onto the membrane is indicated at the top of each. These dot blots were performed once.

Detection of CLU-GST complexes using anti-GST antibodies showed that two different anti-GST antibodies detected GST present in both non-biotinylated and biotinylated CLU-GST complexes (Figure 5.18), although in both cases the detection was significantly weaker for GST present in biotinylated CLU-GST complexes. Both antibodies specifically recognised and bound to GST, with no non-specific binding of either antibody to BSA.

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CLU-GST CLU-GST

Biotinylated Biotinylated CLU-GST CLU-GST

BSA BSA

GST GST

Detection with anti-GST1 Detection with anti-GST2

Figure 5.18 Dot blot detection of CLU-GST complexes with two different anti-GST antibodies. Anti- GST antibodies (Table 5.1) were used at 1 μg/mL to probe membranes on which 5 μg each of CLU-GST and biotinylated CLU-GST, and 1 μg each of BSA and GST had been immobilised. Each sample was dotted onto the membrane in triplicate (n = 3) and samples are identified at the left of the images. These dot blots were performed once.

SDS-PAGE and dot blot analyses of HMW chaperone-client complexes formed with 3 different client proteins indicated that the biotinylation process is likely to induce structural changes in the complexes, but it was still unclear what those changes might be, and how they could affect the binding of the complexes to cell surfaces. NTA size analysis confirmed that biotinylated CLU- CS was, on average, larger than CLU-CS (Figure 5.19). Unsurprisingly, both CLU-CS (pink trace) and biotinylated CLU-CS (green trace) appear to be polydisperse solutions with particles ranging in size from ~ 20 – 300 nm. For CLU-CS there were three main size populations (P1, P2, and P3) at ~ 44, 60, and 84 nm respectively, and the single major biotinylated CLU-CS peak (P4) resolved at ~ 92 nm. The size distribution of the biotinylated complexes indicates, on average, larger particles compared to the non-biotinylated complexes, which contained smaller particles not detected in analyses of biotinylated complexes. It was therefore concluded that biotinylation of the chaperone-client complexes could induce an increase in size, and this in turn affected their binding to cell surfaces (possibly causing them to non-specifically interact with cells). NTA analyses of other chaperone-client complexes were not performed.

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P2 P3 25 P1 ) 6 20 P4 Clu-CSCLU-CS Biotinylated CLU Clu-CS-CS 15

10 Particle Count (x10 Particle

0 0 50 100 150 200 250 300 350 400 Diameter (nm) Figure 5.19 NTA distribution curves estimating the mean diameters of CLU-CS and biotinylated CLU-CS complexes. The x-axis is particle size (nm), the y-axis indicates the number of particles counted (106), and the key identifies each sample. The results shown are from one normally distributed curve for each sample. This figure represents two independent experiments.

Flow cytometry analysis detected binding of biotinylated CLU-CS to RAW264.7 cells, but not binding of non-biotinylated CLU-CS (Figure 5.20). Interestingly, however, the binding of non- biotinylated CLU-CS could not be detected with anti-CS antibodies (Figure 5.20 B). These results definitively confirmed that biotinylation of HMW chaperone-client complexes can alter their binding to cell surfaces and that biotinylated complexes should not be used for investigating such interactions. At this point in this study, the use of biotinylated chaperone-client complexes was discontinued.

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A B

% Max of % Max of

SA-488 Fluorescence Alexa-488 Fluorescence

No protein + anti-CS mix

+ Biotinylated CLU-CS + CLU-CS + anti-CS mix

+ Biotinylated CLU-CS + anti-CS mix

Figure 5.20 Flow cytometry analysis comparing the binding of CLU-CS and biotinylated CLU-CS to RAW 264.7 cells. Cells were incubated without the complex (no protein), with biotinylated CLU-CS, or with non-biotinylated CLU-CS. Bound complexes were then detected with A) 2.5 µg/mL SA-488, or B) 2.5 µg/mL each of anti-CS1 and anti-CS2 (Table 5.1) followed by 2.5 µg/mL goat anti-rabbit IgG-Alexa- 488. The x-axis shows fluorescence intensity, and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on PI exclusion and the experiment was performed only once.

5.4.8 Non-biotinylated Tg-CS and CLU-GST complexes do not bind specifically to RAW 264.7 macrophages As described above, cell binding assays using biotinylated HMW CLU-client protein complexes were discontinued, therefore, cells were re-tested for the binding of non-biotinylated complexes. Flow cytometry results showed that non-biotinylated Tg-CS complexes did not bind specifically to RAW 264.7 cells (Figure 5.21 A & B). There was also no specific binding of Tg to RAW 264.7 cells (Figure 5.21 C).

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% Max of

Alexa-488 Fluorescence

+ Tg-CS + anti-HB + Tg-CS + anti-CS2 + fucoidan + Tg-CS + anti-CS2

B C

% Max of % Max of

Alexa-488 Fluorescence Alexa-488 Fluorescence

+ CS + anti-HB + Tg + mouse IgG 2a

+ CS + anti-CS2 + Tg + anti-Tg

+ Tg-CS + anti-HB

+ Tg-CS + anti-CS2

Figure 5.21 Flow cytometry analysis of non-biotinylated Tg-CS binding to RAW 264.7 cells. A) Cells were incubated with Tg-CS and 5 µg/mL of either anti-CS2 or anti-HB (control antibody), or pre-treated with fucoidan prior to incubation with the non-biotinylated complex and 5 µg/mL anti-CS2. Bound antibodies were detected with 2.5 µg/mL goat anti-rabbit IgG-Alexa-488. B) Cells were incubated with either CS or Tg-CS and detected with 5 µg/mL of either anti-CS2 or anti-HB. Bound antibodies were detected with 2.5 µg/mL goat anti-rabbit IgG-Alexa-488. C) Cells were incubated with Tg and 5 µg/mL of either anti-Tg or mouse IgG 2a negative control antibody. Bound antibodies were detected with 2.5 µg/mL goat anti-mouse IgG-Alexa-488. The x-axis shows fluorescence intensity and the y-axis indicates the frequency of events (%). Viable cells were gated based on PI exclusion. The experiment was performed twice, each in triplicate (n = 3).

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Flow cytometry analysis indicated that non-biotinylated CLU-GST complexes specifically bound to RAW 264.7 cells (Figure 5.22 A). However, both CLU and GST were also shown to bind to RAW 264.7 cells.

% Max of

Alexa-488 Fluorescence

+ CLU-GST + DNP9

+ CLU-GST + G7

B C

% Max of % Max of

Alexa-488 Fluorescence Alexa-488 Fluorescence

+ CLU + DNP9 + GST + anti-HB

+ CLU + G7 + GST + anti-GST2

Figure 5.22 Flow cytometry analysis of non-biotinylated CLU-GST binding to RAW 264.7 cells. A) Cells were incubated with CLU-GST and either undiluted G7 anti-CLU or DNP9 (see key), and bound antibodies were detected with goat anti-mouse IgG-Alexa-488. B) Cells were incubated with CLU and either G7 anti-CLU or DNP9, and bound antibodies were detected with goat anti-mouse IgG-Alexa-488. C) Cells were incubated with GST and either 5 µg/mL anti-GST2 or anti-HB, and bound antibodies were detected with goat anti-rabbit IgG-Alexa-488. The x-axis is fluorescence intensity, the y-axis indicates the frequency of events (%), and the keys identify each sample. Cells in this experiment were gated based on PI exclusion. The experiment was performed twice, each in triplicate (n = 3). 165

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5.4.9 Non-biotinylated CLU-GST complexes bind to the surface of EOC13.31 cells Biotinylated CLU-GST was previously shown to bind to the surface of EOC13.31 cells (Figure 5.7), however due to the possibility of biotinylated complexes binding to cell surfaces non- specifically, this experiment was repeated and the ability of non-biotinylated CLU-GST to bind to EOC13.31 cells was also tested. Biotinylated CLU-GST was detected with SA-488 and non- biotinylated CLU-GST complexes were detected using G7 anti-CLU or DNP9 hybridoma supernatants and goat anti-mouse IgG-CF488. Flow cytometric analysis confirmed that biotinylated CLU-GST complexes bind to EOC13.31 cells (Figure 5.23 A). Specific binding of non-biotinylated CLU-GST complexes to EOC cells was also detected using G7 antibody (Figure 5.23 B). Pre-incubating EOC13.31 cells with fucoidan did not inhibit the binding of either biotinylated or non-biotinylated CLU-GST to the cells, suggesting that the complexes are not binding to scavenger receptors. Although the binding of non-biotinylated CLU-GST to EOC13.31 does appear to be significant, this experiment would need to be repeated with additional negative controls (e.g. cells incubated with non-complexed CLU and GST) to confirm that the binding of CLU-GST is specific to the complex.

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A B

% Max of % Max of

SA-488 Fluorescence CF-488 Fluorescence

No protein + CLU-GST + DNP9

+ biotinylated CLU-GST + CLU-GST + G7

+ fucoidan + biotinylated CLU-GST + fucoidan + CLU-GST + G7

Figure 5.23 Flow cytometry analysis of the binding of biotinylated and non-biotinylated CLU-GST complexes to EOC13.31 cells. A) Cells were either incubated with or without (no protein) biotinylated CLU-GST, or pre-treated with fucoidan before incubation with biotinylated CLU-GST. Bound complexes were detected with 2.5 µg/mL SA-488. B) Cells were incubated with non-biotinylated CLU-GST and either G7 or DNP9, or pre-treated with fucoidan prior to incubation with non-biotinylated CLU-GST and then G7. Bound antibodies were detected with 5 µg/mL goat anti-mouse IgG-CF-488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on PI exclusion. The results shown are representative of two independent experiments.

5.4.10 Non-biotinylated CLU-GST complexes do not bind to the surface of HepG2 liver cells Biotinylated CLU-GST complexes were confirmed to bind to HepG2 cells (Figure 5.24 A), however, binding of non-biotinylated CLU-GST was not detected with either anti-GST1 or anti- GST2 (Table 5.1; Figure 5.24 B and C). In the same experiment, the binding of biotinylated CLU- GST complexes to HepG2 cells was also assessed using anti-GST1, which did not detect their binding. These results suggest that non-biotinylated CLU-GST may bind to HepG2 cells, but the antibodies being used here cannot detect the complex bound to the cell surface, and further testing with different antibodies is warranted. Detection of bound CLU-GST complex with G7 was trialled, however, HepG2 cells produce large quantities of endogenous human CLU, which made it impossible to distinguish between bound endogenous CLU and bound CLU-GST (data not shown).

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% Max of

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+ CLU-GST + anti-HB No protein + CLU-GST + anti-GST2 + biotinylated CLU-GST + CLU-GST Figure 5.24 Flow cytometry analysis of biotinylated and non-biotinylated CLU-GST binding to HepG2 cells. A) Cells were incubated with or without (no protein) biotinylated CLU-GST and bound complex was detected with 2.5 µg/mL SA-488. B) Cells were incubated with non-biotinylated CLU-GST and 10 µg/mL of either anti-GST2 or anti-HB (control antibody). Bound primary antibody was detected with 2.5 µg/mL goat anti-rabbit IgG-Alexa-488. C) Cells were incubated with or without (no protein) CLU- GST or biotinylated CLU-GST. Bound complexes were detected with 10 µg/mL anti-GST1-488. The x- axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on PI exclusion. The experiment was performed twice, each in triplicate (n = 3).

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5.4.11 Non-biotinylated CLU-GST complexes bind to transfected HEK293 cells expressing ApoER2 and VLDLR ApoER2 and VLDLR are LDL-family receptors and are both known to bind CLU (Leeb et al. 2014). To investigate whether CLU-client complexes also bind to these receptors, non-transfected HEK293 cells, and HEK293 cells transfected to express ApoER2 or VLDLR, were incubated with CLU-GST and bound complexes were then detected with G7. As a positive control, transfected cells were also incubated with 50 µg/mL CLU. The binding of CLU to non-transfected cells was not tested in this study, though this control would be required to definitely conclude that CLU is binding to ApoER2 or VLDLR specifically. Successful expression of ApoER2 and VLDLR was confirmed using anti-ApoER2 or anti-VLDLR antibodies.

Detection with anti-ApoER2 antibody confirmed that HEK293 cells were successfully transfected with pcDNA3.1(1)(ApoER2) (Figure 5.25 A). Preliminary testing revealed that CLU-GST binds strongly to transfected HEK293 cells expressing ApoER2 (Figure 5.25 B). Relatively little CLU- GST bound to non-transfected HEK293 cells (Figure 5.25 C). As expected, CLU bound to transfected cells expressing ApoER2, although the level of binding detected was less than that for the HMW CLU-GST complex (Figure 5.25 D). The binding of GST alone to ApoER2 was not tested in this study, however, this control would be required to confirm that ApoER2 specifically binds HMW CLU-GST complexes.

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CF-488 Fluorescence CF-488 Fluorescence Non-transfected + CLU-GST + DNP9 Transfected + CLU + DNP9 Non-transfected + CLU-GST + G7 Transfected + CLU + G7

Figure 5.25 Flow cytometry analysis showing that CLU-GST binds to transfected cells expressing ApoER2. A) Non-transfected and pcDNA3.1(-)(ApoER2) transfected HEK293 cells were incubated with 2.5 µg/mL anti-ApoER2 (the key identifies each sample). B) pcDNA3.1(-)(ApoER2) transfected cells were incubated with CLU-GST and either G7 or DNP9. C) Non-transfected cells were incubated with CLU-GST and either G7 or DNP9. D) pcDNA3.1(-)(ApoER2) transfected cells were incubated with CLU and either G7 or DNP9. In all cases, bound primary antibodies were detected with 5 µg/mL goat anti-mouse IgG-CF- 488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on PI exclusion and the results shown are representative of two independent experiments.

Similar results were obtained for HEK293 cells transfected to express VLDLR, though the transfection efficiency appeared to be slightly lower compared to that of transfected ApoER2- expressing cells (Figure 5.26 A). Binding of CLU-GST complexes to transfected VLDLR- 170

Chapter 5 expressing cells, but not to untransfected cells, was detected (Figure 5.26). Binding of CLU to transfected VLDLR-expressing cells was also detected (Figure 5.26).

A B

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CF-488 Fluorescence CF-488 Fluorescence Non-transfected + CLU-GST + DNP9 Transfected + CLU + DNP9 Non-transfected + CLU-GST + G7 Transfected + CLU + G7

Figure 5.26 Flow cytometry analysis showing that CLU-GST complexes bind to transfected cells expressing VLDLR. A) Non-transfected and pcDL-Sra296(VLDLR) transfected HEK293 cells were incubated with 2.5 µg/mL anti-VLDLR (the key identifies each sample). B) pcDL-Sra296(VLDLR) transfected cells were incubated with CLU-GST and either G7 or DNP9. C) Non-transfected cells were incubated with CLU-GST and either G7 or DNP9. D) pcDL-Sra296(VLDLR) transfected cells were incubated with CLU and either G7 or DNP9. In all cases, bound primary antibodies were detected with 2.5 µg/mL goat anti-mouse IgG-CF-488. The x-axis is fluorescence intensity and the y-axis indicates the frequency of events (%). Cells in this experiment were gated based on PI exclusion and the experiment was performed once.

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5.5 Discussion The binding of chaperone-client complexes to a range of various cell types was assessed in this study with the initial aim of identifying the receptor type(s) involved in the binding of complexes to cells. The murine bone-derived macrophage line, RAW 264.7, and the murine microglial line, EOC13.31, were chosen as both macrophages and microglia are known to express scavenger receptors (Gough and Gordon 2000; Harb et al. 2009; de Vries et al. 1999; Campa et al. 2005; Fitzgerald et al. 2000; Coller and Paulnock 2001; Wilkinson and El Khoury 2012) which are thought to play a role in the binding of CLU and CLU-client complexes to cells. Microglia are also known for their role in tissue maintenance, injury response, and pathogen defence, and have been shown to clear Ab from the brain via scavenger receptor-mediated endocytosis (Zhao et al. 2017: Hansen et al. 2018). The binding of HMW chaperone-client complexes to the human hepatoblastoma cell line, HepG2 was also tested as Wyatt et al. 2011 showed that (i) radiolabelled CLU-client complexes injected into rats were localised to the liver and, (ii) that biotinylated CLU- client complexes bind to the surface of primary hepatocytes isolated from rat livers. The binding of chaperone-client complexes to specific scavenger receptors (SRA-1 and SREC-1) and LDL receptors (ApoER2 and VLDLR) was also investigated using transfected HEK293 cells. Results pertaining to experiments that used biotinylated HMW complexes will be discussed first, followed by the results of experiments that used non-biotinylated complexes.

Initial work in this study used biotinylated HMW chaperone-client complexes in cell binding assays for several reasons: (i) biotinylation of proteins is easy and relatively inexpensive for labelling large amounts of protein, (ii) detection of biotin tags with fluorescently-labelled streptavidin is highly specific and efficient, and (iii) previous work in our lab also used biotinylated CLU-client complexes to investigate their binding to cells and there was no prior knowledge that biotinylating the complexes might induce structural or functional changes in them.

The specific scavenger receptors involved in the binding of chaperone-client complexes to cell surfaces are not known, however, some molecular chaperones have been reported to interact with various scavenger receptors (Table 5.3). Furthermore, it has been shown that fucoidan inhibits (i) the binding of biotinylated CLU-client complexes to primary rat hepatocytes and the trafficking of radiolabelled CLU-client complexes to the liver (Wyatt et al. 2011), and (ii) the binding of Alexa fluorophore-labelled CLU-OT complexes to primary murine bone-derived macrophages and human white blood cells (granulocytes, lymphocytes, and monocytes) (Dabbs 2013). Fucoidan is known to inhibit the binding of ligands to class A, C, E, F, and H scavenger receptors, which include SRA-1, SRA-II (Platt et al. 1996), LOX-1 (Oka et al. 1998), SREC-1 (Berwin et 172

Chapter 5 al. 2004), and FEEL-1 (Tamura et al. 2003), but not class B scavenger receptors (Peiser and Gordon 2001; Gough and Gordon 2000; Fluiter and van Berkel 1997; Acton et al. 1994).

Table 5.3 Brief details of scavenger receptors. The below table lists the major scavenger receptors, their expression profiles (* indicates not expressed in that cell type), whether they have been reported to bind chaperones, and whether or not ligand binding to them is inhibited by fucoidan (adapted from Dabbs 2013).

Expression Binding of Fucoidan Class Receptor Reference Profile Chaperones Inhibited Dendritic cells HSP gp96 and Endothelial cells Hughes et al. 1995; de Winther calreticulin are Macrophages et al. 2000; Berwin et al. 2003; internalised by SRA-1 A SRA-1/11 Mast cells Yes Berwin et al. 2004; Delneste et

Monocytes al. 2007; Greaves and Gordon HSP70 does not bind Smooth muscle cells 2009; Stephen et al. 2010 SRA-1 Neutrophils * B cells Granucci et al. 2003; Delneste Dendritic cells A MARCO HSP70 does not bind Yes et al. 2007; Zhu et al. 2009; Macrophages Stephen et al. 2010 Microglia Adipocytes Fluiter and van Berkel 1997; B cells van Oosten et al. 1998; B SRB-1 Hepatocytes HSP70 does not bind No Delneste et al. 2007; Zhu et al. Macrophages 2009; Stephen et al. 2010 Monocytes Adipocytes B cells Dendritic cells Fluiter and van Berkel 1997; Epithelial cells Panjwani et al. 2000; Radsak et B CD36 Endothelial cells HSP70 does not bind No al. 2003; Berwin et al. 2004; Macrophages Delneste et al. 2007; Zhu et al. Monocytes 2009; Fischer et al. 2010 Platelets Neutrophils * Binds calreticulin, Moriwaki et al. 1998; Yoshida B cells HSP 60, and HSP70 et al. 1998; Delneste et al. Endothelial cells 2002; Theriault et al. 2006; Macrophages Internalises HSP Schaeffer et al. 2009; Fischer E LOX-1 Neutrophils gp96-peptide Yes et al. 2010; Matsutake et al. Platelets complexes into 2010; Stephen et al. 2010; Xie Smooth muscle cells dendritic cells and B et al. 2010; Yoshimoto et al. Monocytes * cells 2011; Stephen and Ponnambalam 2012 Adachi et al. 1997; Berwin et Binds calreticulin, al. 2004; Theriault et al. 2006; Endothelial cells HSP70-peptide Calderwood et al. 2007; Gong F SREC-1 Macrophages complexes, and Yes et al. 2008; Gong et al. 2009; Monocytes * HSP90-Oval Fischer et al. 2010; Murshid et complexes al. 2010; Stephen et al. 2010 B cells Dendritic cells Endothelial cells Shimaoka et al. 2000; Berwin G SR-PSOX None reported No Macrophages et al. 2004; Stephen et al. 2010 Smooth muscle cells T cells Endothelial cells Tamura et al. 2003; Theriault H FEEL-1 Macrophages Internalises HSP70 Yes et al. 2006; Stephen et al. 2010 Monocytes

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The results indicated that neither biotinylated CLU-CS or biotinylated Tg-CS bound to transfected cells expressing SRA-1 or SREC-1 in this study (Figure 5.3). However, it cannot be definitely concluded that HMW chaperone-client complexes do not bind to SRA-1 and SREC-1 scavenger receptors as the identities of the expressed receptors were not confirmed with specific antibodies.

Rather than trialling cell binding assays with cells expressing individual scavenger receptors, which would require the purchasing of many expensive plasmids and antibodies, a different approach was explored in an attempt to identify the specific receptors involved in HMW chaperone-client complexes binding to cell surfaces. First, various cell types known to express scavenger receptors (RAW 264.7 and EOC13.31), and a liver cell line expected to interact with HMW CLU-client complexes (HepG2) were selected and incubated with biotinylated HMW CLU- CS, Tg-CS, and CLU-GST to determine which cell types the complexes bind to; a summary of these results is shown in Table 5.4.

Table 5.4 A brief summary of results from cell binding experiments using biotinylated HMW chaperone-client complexes. The below table lists the biotinylated HMW complexes used in cell binding assays, the cell types trialled in this study, and which cell types each complex was shown to bind. N/A is written for complex/cell type combinations not tested in this study and the corresponding figure number for each result is shown; indicates that the complex bound and indicates no binding was detected.

Transfected Transfected HEK293 cells HEK293 cells RAW 264.7 EOC13.31 HepG2 expressing SRA-1 expressing SREC-1 CLU-CS Figure 5.3 B Figure 5.3 B Figure 5.4 N/A N/A CLU-GST N/A N/A Figure 5.6 Figure 5.7 A Figure 5.8 A Tg-CS Figure 5.3 C Figure 5.3 C Figure 5.5 Figure 5.7 B Figure 5.8 B

Here, it was shown that biotinylated CLU-CS, biotinylated Tg-CS, and biotinylated CLU-GST all bind to the surface of RAW 264.7 macrophages. Fucoidan was shown to inhibit the binding of biotinylated CLU-CST complexes to RAW 264.7 macrophages by more than 50%. These results implicate scavenger receptors as being involved in the binding of chaperone-client complexes to the surface of RAW 264.7 macrophages. This was not at all surprising as macrophages are known to express a wide range of scavenger receptors and as described above, scavenger receptors have been shown to bind molecular chaperones and other CLU-client complexes. The binding of individual biotinylated control proteins (e.g. CLU, Tg, CS, and GST) was not assessed in the current study, however, previous work showed that the binding of biotinylated CLU, Tg, and CS to the surface of RAW 264.7 cells was significantly lower than that of biotinylated Tg-CS (Frazier 2012), and biotinylated Tg-CS binds to a lesser extent than CLU-CS (see Appendix III). It was

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Chapter 5 also shown that biotinylated CLU-GST and Tg-CS bind to the surface of EOC13.31 cells, and that biotinylated CLU-GST (but not Tg-CS) binds to the surface of HepG2 liver cells.

After showing that all three biotinylated chaperone-client complexes bind to RAW 264.7 macrophages, this cell line was chosen for affinity adsorption assays, in an attempt to identify the specific receptors that bind the complexes. Two main methods were trialled here. The first approach, batch adsorption, required the incubation of cell lysates from cells that had been incubated with biotinylated CLU-CS with streptavidin conjugated Sepharose beads in a tube. It was not surprising that the streptavidin beads captured a significant number of proteins other than biotinylated CLU-CS as biotin is an endogenous molecule and a co-factor for many enzymes (Tytgat et al. 2015). Several other studies have reported experimental complications resulting from the interference of endogenous biotin in eukaryotic cells (Chen et al. 2012; Horling et al. 2012; McKay et al. 2008). Although SA-488 did not detect any endogenous biotin on the cell surface of RAW 264.7 cells, lysing of the cell membrane would have released intracellular biotinylated proteins.

The second approach, affinity adsorption, passed the lysates of cells that had been incubated with CLU-CS over a hand-made affinity column packed with anti-CS conjugated Sepharose. G7 conjugated Sepharose could have been trialled as well, however, anti-CS was tested first due to large quantities of this antibody being available and inexpensive. SDS-PAGE analysis confirmed that incubating the anti-CS beads with CLU-CS could indeed purify the complex as bands corresponding to both CLU and CS were observed in this sample, but not in the control sample (Figure 5.9). Three contaminating bands appeared in the sample containing CLU-CS eluted from the anti-CS beads and in the supernatant collected immediately after incubation of the complex with the anti-CS beads. Once the anti-CS beads had been mixed with SDS sample buffer (containing bromophenol blue), it was difficult to distinguish between the bead pellet and supernatant, and some beads may have been collected when removing the supernatant. If so, the subsequent incubation with 2-mercaptoethanol would have dissociated anti-CS antibody chains from the Sepharose. If this experiment were to be repeated, preparation of a colourless SDS sample buffer may help avoid this issue. This approach was unsuccessful at capturing CLU-CS (Figure 5.10), and therefore identifying the receptor(s) involved in binding CLU-CS to RAW 264.7 cells. A side-by-side flow cytometry experiment did not detect CLU-CS bound to the surface of the cells (Figure 5.11) and the most obvious explanation for these results was that CLU-CS did not bind to the cell surfaces in this particular experiment.

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To test whether or not non-biotinylated CLU-CS binds to RAW 264.7 cells, the cells were incubated with or without CLU-CS and biotinylated CLU-CS (as a positive binding control) in the same experiment. In this test, biotinylated CLU-CS was detected bound to the surface of the cells, however, the non-biotinylated complex was not (Figure 5.12). These results were the first indication that CLU-CS may be, in some way, structurally different to biotinylated CLU-CS. Before cell-binding and affinity adsorption assays could be continued, it was necessary to establish whether or not the biotinylation process changed the structure of the complex and caused it to bind to cells non-specifically.

Four different methods were used to compare biotinylated and non-biotinylated HMW chaperone- client complexes. SDS-PAGE results showed clear differences in the dissociation of non- biotinylated and biotinylated CLU-CS (Figure 5.13), CLU-GST (Figure 5.14), and Tg-CS (Figure 5.15) under non-reducing and reducing conditions, which clearly imply that biotinylated complexes are structurally different to their non-biotinylated counterpart. Dot blot analyses also showed differences in the ability of various antibodies to recognise non-biotinylated versus biotinylated CLU-CS (Figures 5.16 and 5.17) and CLU-GST (Figure 5.18). The most plausible explanation for the observed differences is that the changes induced by biotinylating the chaperone-client complexes altered the protein structure in such a way that the antibodies could no longer recognise their respective motifs on the proteins.

To investigate whether biotinylated CLU-CS complexes were larger in size than the corresponding non-biotinylated complex, NTA size analysis was used. NTA showed that both non-biotinylated and biotinylated CLU-CS samples were polydisperse solutions, with most particles resolving between 20 – 180 nm for CLU-CS and 25 – 220 nm for biotinylated CLU-CS (Figure 5.19). NTA confirmed that, on average, biotinylated CLU-CS contained larger molecules than the non- biotinylated complex; the most striking difference was the lack of smaller particles in the biotinylated sample.

Taken together, SDS-PAGE, dot blot analysis, and NTA particle sizing clearly indicate that non- biotinylated and biotinylated chaperone-client complexes are not equivalent, they differed in their abilities to dissociate under reducing conditions, and to be recognised by antibodies, and in the case of CLU-CS complexes, they differed in size. Collectively, these results demonstrate that biotinylation of chaperone-client complexes can alter their structure and change their biophysical properties. It was next necessary to determine whether there was a difference in the ability of non- biotinylated versus biotinylated complexes to bind to cells. A flow cytometry experiment was

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Chapter 5 conducted where non-biotinylated and biotinylated complexes were incubated with RAW 264.7 macrophages. Results from this experiment showed that non-biotinylated CLU-CS was not detectable on the cell surface, however, biotinylated CLU-CS was detected bound to RAW 264.7 cells using both SA-488 and the anti-CS antibody mixture (Figure 5.20). These results suggest that the binding of biotinylated CLU-CS to RAW 264.7 cells is mediated by a method other than the cells specifically recognising the CLU-CS complex.

Biotinylation is a commonly practiced method in laboratory research for the labelling, purification, and detection of proteins. The biotin molecule is quite small, only weighing ~ 244 Da and does not typically interfere with the normal structure or function of most proteins (Tytgat et al. 2015). Biotin has been shown to affect enzyme activity if it binds to a lysine residue in or near the enzyme active site, preventing substrate interactions (Kay et al. 2009), however that was not considered to be an issue in this study as HMW chaperone-client complexes do not have enzymatic activity. The exact reason as to why or how biotinylation structurally altered the HMW chaperone-client complexes used in this study is not known, and answering these questions was not an aim of this project. It is possible that labelling the complexes with biotin increased the hydrophobicity of these already “sticky” proteins causing them to interact with each other to form larger soluble aggregates. Regardless of how the structural changes came about, the fact remained that these complexes do not exhibit the same biophysical and functional properties as their non-biotinylated counterparts. It was therefore decided to discontinue their use in cell binding assays to avoid misleading results.

As the results obtained from cell binding assays using biotinylated chaperone-client complexes were unusable, repetition of these assays with non-biotinylated complexes was required. RAW 264.7 macrophages were incubated with non-biotinylated Tg-CS (Figure 5.21) and CLU-GST (Figure 5.22). Neither Tg-CS nor CLU-GST bound to these cells specifically. Pre-incubation of RAW 264.7 cells with fucoidan had no effect on the binding of non-biotinylated Tg-CS. It was also shown that neither fucoidan, nor polyinosinic acid (another inhibitor of scavenger receptors), inhibited the binding of Alexa-fluorophore-labelled CLU-Oval complexes to these cells (Dabbs 2013). Although the expression of specific scavenger receptors was not confirmed in the current study, the binding of the acetylated LDL probe to the cell surface suggests that scavenger receptors were present. As scavenger receptors are thought to play a role in the binding of chaperone-client complexes to cells, it was not expected that none of the complexes tested in this study would specifically bind to RAW 264.7 cells. It is possible that non-biotinylated chaperone-client complexes do not bind to the specific scavenger receptors expressed on this murine cell line.

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It was shown that non-biotinylated and biotinylated CLU-GST bind to EOC13.31 microglia (Figure 5.23). To confirm that CLU-GST binds to EOC13.31 cells specifically, this experiment would need to be repeated and the binding of CLU and GST alone also assessed, however time restrictions did not allow for this to be completed in this study. Fucoidan did not inhibit the binding of CLU-GST or biotinylated CLU-GST, suggesting that the complexes may be binding to something other than scavenger receptors. Microglia are the major phagocytic cells of the brain and in addition to scavenger receptors, they also express several LDL receptors, including LDLR, LRP1, and VLDLR, but not ApoER2 (Pocivavsek et al. 2009). The LDL receptors ApoER2 and VLDLR have been shown to bind CLU (Leeb et al. 2014; Dlugosz and Nimpf 2018) and LRP1 has been shown to bind a2M (Barcelona et al. 2011; Koummas et al. 1992), suggesting these receptors could be involved in the cellular uptake of chaperone-client complexes. RAP is known to inhibit LRP1, LRP2, and VLDLR (Kasza et al. 1997; Marino et al. 1999; Andersen et al. 2000; Herz and Strickland 2001; Yakovlev et al. 2012). If it can be established that non-biotinylated CLU-GST complexes specifically bind to EOC13.31 cells, it would be worthwhile to pre-incubate the cells with RAP to establish whether or not LDL receptors are involved in the binding of CLU- client complexes to microglial cells.

Binding of non-biotinylated CLU-GST to HepG2 cells was not detectable with either anti-GST1 or anti-GST2 (Figure 5.24). Cells were also incubated with biotinylated CLU-GST in this experiment as a positive binding control. SA-488 detection confirmed that biotinylated CLU-GST did bind to the surface of HepG2 cells, however, binding of the biotinylated complex was not detected using anti-GST1. This data suggests that anti-GST1, and possibly anti-GST2, may not be suitable for detecting HMW CLU-GST complexes in flow cytometry experiments, however this would be unlikely as these antibodies are both polyclonal. It would be worth repeating this assay using an anti-GST antibody that can detect both non-biotinylated and biotinylated CLU-GST bound to the cell surface to conclusively establish whether or not the non-biotinylated complex binds to HepG2 cells. Liver cells express a variety of receptor types required for their role in detoxifying blood plasma, including scavenger receptors (Terpstra and van Berkel 2000), LDL receptors (Soutar et al. 1986; Pal 1996), and the asioglycoprotein receptor. More specifically, HepG2 cells have been shown to endogenously express class B scavenger receptors (Rhainds et al. 1999; Sakai-Kato et al. 2017) and LRP1 (Moon et al. 2011), but not VLDLR or ApoER2 (Romagnuolo et al. 2017). If non-biotinylated HMW CLU-client complexes do not bind to HepG2 cells, other sources of liver cells should be trialled, such as primary rat hepatocytes (as described in Wyatt et al. 2011). Using a non-human cell line would also allow for the binding of CLU to be

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CLU has been shown to bind ApoER2 and VLDLR (Leeb et al. 2014; Dlugosz and Nimpf 2018), therefore, it seemed plausible that these receptors could be involved in the cellular uptake of CLU- client complexes. Flow cytometry analyses of HEK293 cells transfected to express ApoER2 and VLDLR showed that non-biotinylated CLU-GST complexes preferentially bind to both of these receptors (Figures 5.25 and 5.26). CLU bound to the surface of both ApoER2 and VLDLR expressing HEK293 cells, however the binding of CLU to non-transfected cells was not assessed to confirm that this binding was specific. In the case of ApoER2 transfected cells, the binding of CLU-GST was significantly greater than that of CLU alone, suggesting that ApoER2 may selectively bind CLU-client complexes. To confirm that the binding of CLU-GST to ApoER2 and VLDLR is specific, the binding of GST alone to ApoER2 will need to be assessed, however these are very interesting preliminary results and are the first to suggest a specific receptor involved in the binding of CLU-client complexes to cells. It would also be interesting to test the binding of Tg-client complexes to ApoER2 and VLDLR to determine whether these receptors bind chaperone-client complexes promiscuously or if they selectively target CLU-client complexes only.

Collectively, this study showed that (i) the biotinylation of HMW chaperone-client complexes can induce structural changes, which induce the binding of the complexes to cells, (ii) RAW 264.7 cells are not a suitable model for investigating the binding of chaperone-client complexes to cells, and (iii) ApoER2 and VLDLR bind CLU-GST complexes, and in the case of ApoER2 to a significantly greater extent than CLU alone. This study also showed that EOC13.31 microglia and HepG2 hepatocytes may specifically bind CLU-GST complexes, although further testing would be required to definitely confirm this. Biotinylation is such a common and widely used tool for protein labelling and it was very unexpected that this process would structurally alter the chaperone-client complexes used in this study. If the structural changes observed in this study had not been detected, the project would have likely continued using biotinylated complexes that we now know have altered cell binding properties. Although many of the cell-binding assays in the current study using non-biotinylated chaperone-client complexes are preliminary, this work has laid the foundation for future studies.

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Chapter Six Conclusions

PDDs are physically debilitating and life-threatening disorders that affect millions of people world-wide each year. In addition to decreasing the quality of life, PDDs also carry a significant economic burden for patients, their families, and healthcare systems. According to Alzheimer’s Disease International, there are approximately 44 million people living with AD, or a related dementia, worldwide. In the United States, approximately 10% of the population aged 65 and older have been diagnosed with AD (Hebert et al. 2013), with healthcare-associated costs of around $200 billion US dollars per annum according to the Alzheimer’s Association (USA). According to the Parkinson’s Foundation (US), approximately 10 million people worldwide are living with PD and nearly 60,000 Americans are diagnosed with PD each year, costing around $25 billion US dollars. The American ALS Association states that approximately 30,000 Americans are living with ALS, with more than 5,000 new cases of ALS being diagnosed each year in the United States; worldwide it is estimated that nearly 450,000 people are living with ALS. One study investigated the disease-duration costs associated with ALS and found that an individual can accrue over $1.4 million in medical costs whilst living with this disease (Obermann and Lyon 2015). There are many other PDDs not mentioned here (Table 1.1) that would add to the cumulative costs associated with these diseases. Despite many years of research effort, cures for most PDDs have remained elusive and with an increase in the aging population (those most vulnerable to acquiring PDDs), the physical and economic burden to our global healthcare system is predicted to rise at an alarming rate.

Although the exact cause for the onset of many PDDs is currently unknown, disease progression is most likely due to the unrelenting accumulation of protein aggregates and an overwhelming of proteostasis machinery. This could be due to (i) limited levels of functional ECs, which are unable to manage an increasing accumulation of protein aggregates, (ii) that there are not enough free or functional cell surface receptors available to clear misfolded proteins from the extracellular environment, or (iii) a combination of these. Researchers have identified many genetic mutations that can predispose individuals to developing PDDs, however, until the exact cause(s) for disease onset and/or progression is identified, it is worthwhile to target all potential mechanisms involved in the maintenance of proteostasis when looking for future PDD therapies. It is unlikely that an overexpression of ECs alone would be a viable treatment option for PDDs as several studies have already shown that some ECs are already upregulated in various PDD models (discussed below). Therefore, in this context, any new information gained in regards to the mechanism by which 180

Chapter 6 extracellular protein quality control is maintained could prove useful in developing novel therapies for PDDs.

There are currently six identified ECs (described in Section 1.5.2), and the list continues to grow as research in the field of extracellular protein quality control progresses. The discovery of new ECs is quite important for several reasons. First, some proteins are more structurally characterised than others, and if the structure of a newly identified EC has been extensively characterised, it may be possible to use that data to identify key structures or regions likely to be involved in the chaperone action of the protein. Identifying the specific structures or regions involved in mediating EC activity could provide information useful in the development of therapeutic approaches such as “pharmacological chaperones” and “kinetic stabilizers” (Balch et al. 2008; Wyatt et al. 2009a). Pharmacological chaperones and kinetic stabilizers are small molecules designed to bind and stabilise specific proteins (often interacting with ligand-binding sites), thereby preventing aggregation and allowing the protein to carry out its natural function (Convertino et al. 2016; Banning et al. 2016; Small 2014).

Secondly, PDDs can affect many different regions of the body, such as the brain, liver, heart, thyroid, and eyes. Four of the six major ECs, CLU, a2M, Hp, and SAP are found in blood plasma and migrate throughout the human body in the circulatory system, however, tissue-specific ECs may provide an efficient means of combating PDDs that affect particular organs or areas of the body. For example, NS is only expressed within the CNS and Tg within the thyroid; if protein misfolding were to occur in the brain or thyroid, resident proteins like NS and Tg would likely be the first responders acting to prevent the accumulation of toxic protein aggregates. Lastly, the discovery of new ECs could help identify cell surface receptors involved in the clearance of ECs bound to toxic protein aggregates from the extracellular environment. It is likely that several different receptors or receptor types are involved in this process, which could include selective receptors that only bind specific ECs or promiscuous receptors that will endocytose any EC-client protein complex regardless of the chaperone involved. The discovery of new ECs could also identify potential new receptors to investigate in this context if the EC is already known to interact with specific receptors. The work presented in this thesis has (i) identified three new ECs (Tg, NS, and TTR), the latter two of which are the first to be classified as “amyloid-specific” ECs, (ii) showed that, like CLU, Tg forms stable, soluble HMW complexes with amorphously aggregating CS, (iii) showed that heating Tg to 60°C leads to the dissociation of the Tg monomer and increases the surface hydrophobicity of the protein, (iv) showed that biotinylation alters the structure of HMW EC-client complexes and should not be used when examining their interactions with cells, 181

Chapter 6 and (v) identified ApoER2 and VLDLR as candidate receptors for the clearance and intracellular disposal of CLU-client protein complexes.

The first step in characterising the chaperone activity of Tg, NS, and TTR utilised in vitro protein aggregation assays. In vivo, protein misfolding is believed to occur due to the combined effects of intermittent stresses that occur over time, such as elevated temperatures, shear stress, and oxidation. It is also likely that in vivo protein misfolding is influenced by the large number of other macromolecules present in biological fluids, an effect known as macromolecular crowding. This study induced amorphous protein aggregation with shear stress (shaking) and either elevated temperatures (43 – 60°C) or reduction (DTT), whilst amyloid was formed by shaking the protein at 37°C. It is virtually impossible to replicate the complex conditions of the extracellular environment in in vitro protein aggregation assays. However, it has been shown that the mechanism of protein misfolding at physiological temperatures is the same as that occurring at higher temperatures - the rate at which the proteins misfold is simply increased at higher temperatures, which is experimentally more convenient (Day et al. 2002). The current study showed that, like CLU, Tg inhibits the aggregation of both amorphous and amyloidogenic client proteins (Section 4.4.1). It was also shown that NS and TTR efficiently inhibit amyloid aggregation but are much less effective at inhibiting amorphous protein aggregation (Sections 3.5.3 and 3.5.4).

The unique amyloid-specific chaperone activity shared by both NS and TTR lead to exploration of possible structural similarities shared between the two proteins. Structural comparisons identified an a-helix on both NS and TTR that contains high sequence homology (64% identical, 71% similar). Published research (in the case for TTR) implicated this a-helix in the binding of TTR to Ab (TTR has only one a-helix per each identical monomeric subunit). Due to the significant level of sequence similarity shared between the NS and TTR a-helices, it is possible that the NS a-helix is involved in its chaperone action as well (Section 3.5.5). Synthetic peptides corresponding to the NS a-helix (and a scrambled control) were generated and tested in an aggregation assay. The peptide corresponding to the NS a-helix did not inhibit amyloid formation by ccbw, but rather increased its aggregation (Figure 3.17). An independent study had also generated peptides corresponding to various regions of TTR, including the a-helix, however, they suggested that the synthetic peptide corresponding to the TTR a-helix lacked a helical structure, though it was not specified how this was measured in the article (Cho et al. 2014). The current study did not evaluate whether or not the synthetic peptides used in this study formed a-helices,

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Chapter 6 however circular dichroism spectroscopy could be used to do this in the future. If the synthetic peptide corresponding to the NS a-helix does not adopt an a-helical structure in solution, it is unlikely it would retain its native function in vitro. Another complementary approach for future work examining the role of these a-helices in the chaperone action of NS and TTR is site-directed mutagenesis (described in Section 3.6).

Future work examining the role of the conserved NS/TTR a-helix in chaperone activity is justified because a potential future approach to combat PDDs is the development of “therapeutic peptides”. The idea of peptide therapy has been around since the 1920s, when insulin was first used as a therapy for patients with diabetes (Rosenfeld 2002). Peptide therapy is an appealing strategy for several reasons: (i) compared to protein production, the synthesis of peptides is often quicker, easier, and more cost-efficient, (ii) peptides are relatively safe and well-tolerated due to their ability to mimic naturally occurring ligands, and (iii) peptides can be easily modified and manipulated, which is beneficial for increasing the specificity towards a particular target and for increasing plasma half-life (Fosgerau and Hoffmann 2015; Lau and Dunn 2018). One potential limitation to peptide therapy is that oral administration is typically not feasible due to the breakdown and destruction of the peptides during the digestive process, meaning most therapeutic peptides require subcutaneous injection. Although oral ingestion is often the preferred choice of administering drugs for the maximisation of patient compliance, the dosage frequency for many injectable therapeutic peptides has been lowered because the plasma half-life of the peptides has been increased, some requiring injection only once a month. There are currently over sixty approved therapeutic peptides in the United States (or other major markets) and over 150 peptide therapies being tested in clinical trials (Lau and Dunn 2018).

A defining characteristic of the established ECs is that they stabilise misfolded client proteins in solution by forming stable, soluble complexes with them to prevent their aggregation and precipitation. In vitro, HWM complexes between ECs and amorphously aggregating client proteins can be easily prepared and purified. The in vitro formation of complexes between ECs and endogenous plasma proteins would have required incubating CLU or Tg in plasma and then applying a form of stress (e.g. heat) to induce protein misfolding, however, this method would likely have generated a mixture of complexes that could not be easily separated and was, therefore, not employed in this study. Previous research has shown that the complexes formed between plasma proteins and CLU are of a similar size and structure compared to those produced between CLU and purified proteins used in in vitro aggregation assays, such as CS and GST (Wyatt 2009). Extensive work characterising CLU-client complexes has already been published (Wyatt et al. 183

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2009c). The current study further characterised the chaperone activity of Tg by confirming that bTg can form HMW complexes with CS (Section 4.4.4). Forming complexes between Tg and other client proteins was considered, however, due to the large size of Tg, and the large quantity of both Tg and clients that would have been required to purify an adequate amount of complex, it was not practical to pursue. For example, generating one batch of HMW complexes formed in a mixture of Tg and CPK at a 1:1 molar ratio would have required 242 mg of Tg and 31 mg of CPK (CPK required a concentration at or above 30 µM to generate a significant level of aggregation). Formation of complexes between Tg and client proteins that are induced to aggregate by reduction (such as BSA), or with heat above 50°C (such as OT and GST) was not possible as Tg loses its structural integrity under these conditions. Like with CLU-client complexes, bTg-CS complexes were found to be polydisperse in size, and on average, significantly larger than bTg or CS alone (Figures 4.11 and 4.12). This study did not investigate the ability of NS and TTR to form complexes with amorphously aggregating client proteins as neither EC was able to efficiently inhibit the aggregation of these client proteins.

Although there are no published reports of ECs forming HMW complexes with amyloidogenic client proteins, there is evidence to suggest that complexes between ECs and amyloid-forming proteins do form (Cascella et al. 2013; Garai et al. 2018). Several studies report that CLU forms stable complexes with Ab to prevent aggregation and clear the toxic protein form the extracellular space in AD animal models (Nelson et al. 2017; Hammad et al. 1997; Narayan et al. 2011). Furthermore, a recent study showed that (i) Hp prevents fibril formation of b2M by interacting with b2M oligomers, maintaining b2M in a soluble state, (ii) Hp protects RAW 264.7 cells against b2M fibril toxicity, and (iii) co-incubation of b2M with Hp promotes its lysosomal degradation, suggesting that a physical association between Hp and b2M facilitates the lysosomal degradation of b2M aggregates (Sultan et al. 2013). Although this paper lacks negative controls in their aggregation and cytoxicity assays (meaning the results must be interpreted with some caution), inhibition of protein aggregation and cytoprotective defence against amyloid toxicity are characteristics shared by other ECs. Lastly, a2M has also been shown to inhibit the aggregation of Ab (Narita et al. 1997) and b2M (Ozawa et al. 2011) by forming complexes with these proteins.

ECs accomplish two important tasks by forming complexes with misfolded proteins: (i) ECs bind and shield regions of hydrophobicity exposed on the misfolded protein to prevent aggregation, and (ii) ECs mediate the clearance of misfolded proteins from the extracellular environment by binding to specific cell surface receptors, which internalise the EC-client complexes for degradation

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(Wyatt et al. 2012). This study aimed to identify some of the receptors or receptor types involved in the chaperone-mediated endocytosis of EC-client complexes, however, due to complications arising with the methodology used in this study, only preliminary results were obtained. This study showed that biotinylating HMW chaperone-client complexes induces structural changes that caused the complexes to interact with cell surfaces non-specifically (Section 5.4.7). Two different approaches were used in this study in attempting to identify the receptors involved in the chaperone-mediated endocytosis of CLU-client and Tg-client complexes: (i) various cell types were screened for binding of the HMW complexes; in cases where it was shown that EC-client complexes bound to cells, lysates from these cells were used in attempts to affinity purify the complexes and to isolate and identify any associated receptors, and (ii) receptors of interest were overexpressed and the binding of HMW complexes to these receptors was assessed.

Interestingly, it was shown that the RAW 264.7 macrophages did not specifically bind HMW chaperone-client complexes (Figures 5.21 and 5.22), which was surprising as macrophages are known to express a wide range of scavenger receptors. Scavenger receptors have been implicated in the binding of CLU-client complexes by previous researchers in our lab (Wyatt et al. 2011; Dabbs et al. 2013). However, Wyatt et al. used biotinylated CLU-client complexes to assess binding to specific cells types, which was partially inhibited by fucoidan, suggesting the involvement of scavenger receptors. Although we now know that biotinylated chaperone-client complexes can bind to cell surfaces non-specifically, Wyatt et al. also showed that the uptake of radio-labelled CLU-client complexes injected into rats is delayed by pre-injecting the rats with fucoidan. As it is highly unlikely that radio-labelling (by iodination of tyrosine rings) would significantly alter the structure of chaperone-client complexes, it remains likely that scavenger receptors could be involved in the uptake of HMW EC-client complexes into cells. The RAW 264.7 cells used in this study may have lost the ability to express functional scavenger receptors, an artefact sometimes encountered in immortal cell lines that are at high passage numbers, which may explain why the complexes used here did not bind specifically.

Although specific receptors were not identified on EOC 13.31 microglia, it was shown that CLU- GST complexes bound to these cells, and that this binding is not inhibited by fucoidan (Figure 5.23). It would not be surprising if chaperone-client complexes specifically bind to microglia as these cells are known for their role in “cleaning up” debris and toxic molecules in the brain, however, it will be necessary in future to test the binding of both CLU and GST alone to EOC 13.31 cells to confirm that the complex is binding specifically. The hepatoblastoma cell line HepG2 was also used to investigate the binding of CLU-GST complexes. Initial results indicated

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Chapter 6 that CLU-GST complexes were not binding to the surface of these cells, however, it was later discovered that this may be an artefact related to the anti-GST antibodies used in this study (Figure 5.24). Although the manufacturer claims that their Alexa-488 labelled anti-GST antibody should be suitable for use in flow cytometry, there were no tests or references confirming its usability in this application. Furthermore, the non-labelled anti-GST antibody used in this study may only by suitable for use in Western blots and ELISAs (according to the manufacturer). Further investigations with a different anti-GST antibody are required to confirm whether or not CLU- client complexes bind to this cell line. Unfortunately, this study did not identify a cell type that specifically binds Tg-CS complexes. Further work will be required to determine whether Tg-client complexes are taken up by cells for the removal of toxic proteins. As Tg is a thyroid-specific protein, it is likely that Tg interacts with different receptors than CLU and may only bind to receptors found on thyroid cells (which could easily be tested).

This study identified two LDL receptors, ApoER2 and VLDLR, as potential candidate receptors that bind HMW CLU-GST complexes (Figures 5.25 and 5.26), however, the binding of GST alone was not tested in this study. Tbinding of GST must also be assessed to confirm that these receptors selectively bind HMW CLU-client complexes. Both ApoER2 and VLDLR have been shown to bind CLU (Leeb et al. 2014), and mutations in the genes encoding for ApoER2 and VLDLR have been identified as risk factors for the development of AD (Dlugosz and Nimpf 2018). ApoER2 is exclusively expressed in the CNS, placenta, and testis. Within the CNS, ApoER2 is found in the hippocampus (granule cells of the dentate gyrus and pyramidal cells), cerebellum (Purkinje cells), mitral cell layer of the olfactory bulb, neocortex (cell bodies and dendrites), and cortical neurons (Reddy et al. 2011). VLDLR is most abundantly expressed in the brain, with high levels of expression in the heart and skeletal muscle. VLDLR is expressed in nearly all regions of the brain, with the highest levels of mRNA found in the cortex and cerebellum; in particular, VLDLR is found on microglia, region-specific pyramidal neurons, neuroblasts, matrix cells, Cajal-Retzius cells, glioblasts, astrocytes, and oligodendrocytes (Reddy et al. 2011; Sakai et al. 2009). ApoER2 and VLDLR are involved in the correct positioning of newly generated neurons. Recently, it was shown that CLU is internalised by ApoER2 and VLDLR, activating the reelin pathway (Dlugosz and Nimpf 2018; Leeb et al. 2014) and another study reported that both receptors internalise CLU- leptin complexes for leptin clearance (Bajari et al. 2003). If these receptors internalise both uncomplexed CLU and CLU-leptin complexes, it is plausible that they may also play a significant role in clearing misfolded proteins (in the form of CLU-client complexes) from body fluids.

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Many PDDs result from the accumulation and aggregation of a single protein, however, it is important to note that protein aggregation may arise under vastly different scenarios, for which therapeutic treatment may differ. In some cases, protein aggregates may form due to genetic mutations (as is with familial AD), and in others due to an overwhelming of the extracellular protein quality control network during the aging process (in the case of sporadic AD). In order to develop effective therapies that prevent the progression of both familial and sporadic PDDs, we must first understand the cellular and molecular processes that drive protein quality control and identify the mechanism failures that contribute to disease. As stated above, for those PDDs involving extracellular proteins, an effective treatment option will likely require more than simply increasing the concentration of ECs available. Several studies actually report region-specific increases in CLU levels in the brains of patients with AD compared to healthy control patients, and that CLU expression increases in brain regions exhibiting Ab plaque development (Lidstrom et al. 1998; Baig et al. 2012; Miners et al. 2017). Another recent study showed that patients with pre-clinical AD (symptoms of mild cognitive impairment) and patients diagnosed with AD had significantly higher concentrations of serum a2M compared to healthy controls (Varma and Watts 2017). This data suggests that the body can preferentially overexpress ECs under the stress of PDD-associated protein aggregation and direct ECs to disease-impacted areas in an attempt to restore protein quality control. These changes, however, are insufficient to stop PDD progression. In fact, the overexpression of ECs may actually have adverse effects on other biological systems in the body; for example, the overexpression of CLU has been linked to the progression of cancer (Park et al. 2008). An increasing accumulation of soluble EC-client complexes in biological fluids may have toxic effects as well if there are a limited number of receptors available to efficiently clear them. It is possible that an upregulation of the receptors involved in clearing EC-client complexes would also be required if EC expression was increased. It should also be noted that an increase in the expression of some receptors can activate innate immune cells, such as macrophages, microglia, and neutrophils, which can lead to tissue inflammation (Deane et al. 2004; Mietus-Snyder et al. 1998; Horvai et al. 1995; Van Everbroeck et al. 2002; Ottonello et al. 2002).

Organ transplant and protein replacement therapy were once a popular approach for PDD treatment, however, organ transplant is invasive/high-risk and ineffective if multiple tissues produce the disease-associated protein. Further research has led to the development of small molecules which block amyloid formation in vitro, however, many of these act non-specifically, require a high concentration to be effective, and/or exhibit low tolerability in vivo (Wyatt et al. 2013; Bernier et al. 2004; Convertino et al. 2016). Peptide therapy (described above) may be an 187

Chapter 6 effective therapy alternative to small molecules as peptides are generally more tolerated by the body. Drug therapies that enhance chaperone activity may also be a viable option for treatment of some PDDs. The chaperone activity of a2M, and possibly Tg, is enhanced upon dissociation of the native protein (Wyatt et al. 2014). Drugs with low cytotoxic effects are currently being tested for their ability to induce the dissociation of a2M in vitro.

Other therapeutic approaches have been investigated and these primarily focus on the manipulation of various components of the proteostasis network with RNA interference (small interfering RNA (siRNA) and short-hairpin RNA (shRNA)) and cDNA as a means to treat PDDs (reviewed in Balch et al. 2008, Wyatt et al. 2009a, and Wyatt et al. 2013). Curbing the synthesis of disease-relevant proteins using RNA interference. The selective silencing of genes associated with the development and progression of PDDs (e.g. familial AD or ALS and Huntington’s disease) is a promising therapeutic approach currently under development (Aguiar et al. 2017; Miller et al. 2004; Chen et al. 2013; Zhang et al. 2018; Reddy and Miller 2015). The main limitations of RNA interference therapy include low target specificity (up- or downregulation of non-targeted genes), poor circulating stability (local administration and frequent dosages of the drug could be required), and both siRNA and shRNA have been linked to kidney and renal toxicity (Loisel et al.2001; Park et al. 2016; Tousignant et al. 2000; Castanotto and Rossi 2009), however, researchers are developing methods to overcome such limitations. In August 2018, the United States Food and Drug Administration approved the first-ever therapeutic drug based on RNA interference – patisiran. Patisiran treats hereditary TTR amyloidosis by silencing mutated TTR genes associated with disease (Adams et al. 2018).

Due to the complexity of the protein quality control network and our current lack of knowledge surrounding the onset and progression of PDDs, finding therapeutic targets for PDDs that act specifically and do not adversely affect other biological systems will be complicated and likely involve several aspects of the proteostasis network. A better understanding of how ECs interact with misfolded proteins and cell surface receptors could lay the foundation for novel and effective therapies to treat PDDs. This thesis identified and characterised three new ECs (two of which being the first classified as amyloid-specific), discovered a region on NS and TTR that is likely to be important in their chaperone action, and identified two receptors that bind CLU-GST complexes and that may be involved in the clearance of misfolded proteins from the extracellular environment. The data presented here contributes to the further understanding of extracellular protein quality control and future investigations could help identify suitable therapeutic options for the treatment of PDDs. Particularly, confirming if the NS or TTR a-helix is responsible for the 188

Chapter 6 amyloid-specific chaperone activity of these two proteins, and identifying what (if any) key biophysical properties are attributable to the a-helices’ amyloid-interactions, could provide valuable insights useful in the future development of peptide or small molecule therapies that inhibit amyloid formation associated with PDDs. Furthermore, a thorough understanding of the conditions required to induce the dissociation of Tg from dimer to monomer and establishing whether or not dimer dissociation improves the chaperone activity of Tg, could be useful in developing drugs that increase the potency of ECs. Lastly, identification of the specific receptors involved in clearing misfolded proteins from the extracellular environment, and the particular cells that are involved in this process, could lead to a therapeutic approach involving the upregulation of receptor expression to help efficiently clear misfolded proteins.

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Appendix I

The sequence for hTg (purple sequence) and bTg (magenta sequence) were aligned using the NCBI Align Sequence Protein BLAST tool (default settings). Identical residues are shown between the hTg and bTg sequences (black sequence). The sequences were found to be 77% identical and 85% similar; residues that are not identical, but have similar properties are indicated by + and gap insertions are indicated by -.

1 MALVLEIFTLLASICWVSANIFEYQVDAQPLRPCELQRETAFLKQADYVPQCAEDGSFQT 60 MAL L +F LL IC SANIFEYQVDAQPLRPCELQRE AFLK+ DYVPQCAEDGSFQT 1 MALALWVFGLLDLICLASANIFEYQVDAQPLRPCELQRERAFLKREDYVPQCAEDGSFQT 60

61 VQCQNDGRSCWCVGANGSEVLGSRQPGRPVACLSFCQLQKQQILLSGYINSTDTSYLPQC 120 VQC DG SCWCV A+G EV GSRQPGRP ACLSFCQLQKQQILLS YINST TSYLPQC 61 VQCGKDGASCWCVDADGREVPGSRQPGRPAACLSFCQLQKQQILLSSYINSTATSYLPQC 120

121 QDSGDYAPVQCDVQQVQCWCVDAEGMEVYGTRQLGRPKRCPRSCEIRNRRLLHGVGDKSP 180 QDSGDY+PVQCD+++ QCWCVDAEGMEVYGTRQ GRP RCPRSCEIRNRRLLHGVGD+SP 121 QDSGDYSPVQCDLRRRQCWCVDAEGMEVYGTRQQGRPARCPRSCEIRNRRLLHGVGDRSP 180

181 PQCSAEGEFMPVQCKFVNTTDMMIFDLVHSYNRFPDAFVTFSSFQRRFPEVSGYCHCADS 240 PQCS +G F PVQCK VNTTDMMIFDLVHSY+RFPDAFVTFSSF+ RFPEVSGYC+CADS 181 PQCSPDGAFRPVQCKLVNTTDMMIFDLVHSYSRFPDAFVTFSSFRSRFPEVSGYCYCADS 240

241 QGRELAETGLELLLDEIYDTIFAGLDLPSTFTETTLYRILQRRFLAVQSVISGRFRCPTK 300 QGRELAETGLELLLDEIYDTIFAGLDL STF ETTLYRILQRRFLAVQ VISGRFRCPTK 241 QGRELAETGLELLLDEIYDTIFAGLDLASTFAETTLYRILQRRFLAVQLVISGRFRCPTK 300

301 CEVERFTATSFGHPYVPSCRRNGDYQAVQCQTEGPCWCVDAQGKEMHGTRQQGEPPSCAE 360 CEVERF ATSF HPYVPSC +G+YQA QCQ GPCWCVD++G+E+ GTRQ+GEPPSCAE 301 CEVERFAATSFRHPYVPSCHPDGEYQAAQCQQGGPCWCVDSRGQEIPGTRQRGEPPSCAE 360

361 GQSCASERQQALSRLYFGTSGYFSQHDLFSSPEKRWASPRVARFATSCPPTIKELFVDSG 420 QSC SER++A SRL FG SGYFS+ L +PE+ S R ARF SCPP+IKELF+DSG 361 DQSCPSERRRAFSRLRFGPSGYFSRRSLLLAPEEGPVSQRFARFTASCPPSIKELFLDSG 420

421 LLRPMVEGQSQQFSVSENLLKEAIRAIFPSRGLARLALQFTTNPKRLQQNLFGGKFLVNV 480 + +PM++G+ +F E+L KEAIR +FPSR LARLALQFTTN KRLQQNLFGG+FLV V 421 IFQPMLQGRDTRFVAPESL-KEAIRGLFPSRELARLALQFTTNAKRLQQNLFGGRFLVKV 479

481 GQFNLSGALGTRGTFNFSQFFQQLGLASFLNGGRQEDLAKPLSVGLDSNSSTGTPEAAKK 540 GQFNLSGALGTRGTFNFS FFQQLGL F +G DLAKPLSVGL+SN ++ P+A+K 480 GQFNLSGALGTRGTFNFSHFFQQLGLPGFQDGRALADLAKPLSVGLNSNPASEAPKASKI 539

541 DGTMNKPTVGSFGFEINLQENQNALKFLASLLELPEFLLFLQHAISVPEDVARDLGDVME 600 D + KP VGSFGFE+NLQENQNAL+FL+S LELPEFLLFLQHAISVPED+ARDLGDVME 540 DVALRKPVVGSFGFEVNLQENQNALQFLSSFLELPEFLLFLQHAISVPEDIARDLGDVME 599

601 TVLSSQTCEQTPERLFVPSCTTEGSYEDVQCFSGECWCVNSWGKELPGSRVRGGQPRCPT 660 V SSQ C Q P LFVP+CT EGSYE+VQCF+G+CWCV++ G+EL GSRVRGG+PRCPT 600 MVFSSQGCGQAPGSLFVPACTAEGSYEEVQCFAGDCWCVDAQGRELAGSRVRGGRPRCPT 659

661 DCEKQRARMQSLMGSQPAGSTLFVPACTSEGHFLPVQCFNSECYCVDAEGQAIPGTRSAI 720 +CEKQRARMQSL+GSQPAGS+LFVPACTS+G+FLPVQCFNSECYCVD EGQ IPGTRSA+ 660 ECEKQRARMQSLLGSQPAGSSLFVPACTSKGNFLPVQCFNSECYCVDTEGQPIPGTRSAL 719

225

721 GKPKKCPTPCQLQSEQAFLRTVQALLSNSSMLPTLSDTYIPQCSTDGQWRQVQCNGPPEQ 780 G+PKKCP+PCQLQ+E+AFL TV+ L+SN S LP LS YIPQCS GQW VQC+GPPEQ 720 GEPKKCPSPCQLQAERAFLGTVRTLVSNPSTLPALSSIYIPQCSASGQWSPVQCDGPPEQ 779

781 VFELYQRWEAQNK-GQDLTPAKLLVKIMSYREAASGNFSLFIQSLYEAGQQDVFPVLSQY 839 FE Y+RWEAQN GQ LTPA+LL+KIMSYREAAS NF LFIQ+LYEAGQQ +FP L++Y 780 AFEWYERWEAQNSAGQALTPAELLMKIMSYREAASRNFRLFIQNLYEAGQQGIFPGLARY 839

840 PSLQDVPLAALEGKRPQPRENILLEPYLFWQILNGQLSQYPGSYSDFSTPLAHFDLRNCW 899 S QDVP++ LEG + QP N+ LEPYLFWQILNGQL +YPG YSDFS PLAHFDLR+CW 840 SSFQDVPVSVLEGNQTQPGGNVFLEPYLFWQILNGQLDRYPGPYSDFSAPLAHFDLRSCW 899

900 CVDEAGQELEGMRSEPSKLPTCPGSCEEAKLRVLQFIRETEEIVSASNSSRFPLGESFLV 959 CVDEAGQ+LEG R+EP+K+P CPGSCEE KLRVLQFIRE EEIV+ SNSSRFPLGESFL 900 CVDEAGQKLEGTRNEPNKVPACPGSCEEVKLRVLQFIREAEEIVTYSNSSRFPLGESFLA 959

960 AKGIRLRNEDLGLPPLFPPREAFAEQFLRGSDYAIRLAAQSTLSFYQRRRFSPDDSAGAS 1019 AKGIRL +E+L PPL P RE F E+FL GSDYAIRLAAQST FYQRR + +S A 960 AKGIRLTDEELAFPPLSPSRETFLEKFLSGSDYAIRLAAQSTFDFYQRRLVTLAESPRAP 1019

1020 ALLRSGPYMPQCDAFGSWEPVQCHAGTGHCWCVDEKGGFIPGSLTARSLQIPQCPTTCEK 1079 + + S Y+PQCDAFG WEPVQCHA TGHCWCVD KG ++P SLTARS QIPQCPT+CE+ 1020 SPVWSSAYLPQCDAFGGWEPVQCHAATGHCWCVDGKGEYVPTSLTARSRQIPQCPTSCER 1079

1080 SRTSGLLSSWKQARSQENPSPKDLFVPACLETGEYARLQASGAGTWCVDPASGEELRPGS 1139 R SGLLSSWKQA Q PSPKDLF+P CLETGE+ARLQAS AGTWCVDPASGE + PG+ 1080 LRASGLLSSWKQAGVQAEPSPKDLFIPTCLETGEFARLQASEAGTWCVDPASGEGVPPGT 1139

1140 SSSAQCPSLCNVLKSGVLSRRVSPGYVPACRAEDGGFSPVQCDQAQGSCWCVMDSGEEVP 1199 +SSAQCPSLC VL+SGV SRR SPGY PACRAEDGGFSPVQCD AQGSCWCV+ SGEEVP 1140 NSSAQCPSLCEVLQSGVPSRRTSPGYSPACRAEDGGFSPVQCDPAQGSCWCVLGSGEEVP 1199

1200 GTRVTGGQPACESPRCPLPFNASEVVGGTILCETISGPTGSAMQQCQLLCRQGSWSVFPP 1259 GTRV G QPACESP+CPLPF+ ++V GG ILCE SG +A Q+CQL C QG S FPP 1200 GTRVAGSQPACESPQCPLPFSVADVAGGAILCERASGLGAAAGQRCQLRCSQGYRSAFPP 1259

1260 GPLICSLESGRWESQLPQPRACQRPQLWQTIQTQGHFQLQLPPGKMCSADYADLLQTFQV 1319 PL+CS++ RWES+ PQPRACQRPQ WQT+QTQ FQL LP GK+CSADY+ LL FQV 1260 EPLLCSVQRRRWESRPPQPRACQRPQFWQTLQTQAQFQLLLPLGKVCSADYSGLLLAFQV 1319

1320 FILDELTARGFCQIQVKTFGTLVSIPVCNNSSVQVGCLTRERLGVNVTWKSRLEDIPVAS 1379 F+LDELTARGFCQIQVKT GT VSIPVC++SSV+V CL+RERLGVN+TWK +L D P AS 1320 FLLDELTARGFCQIQVKTAGTPVSIPVCDDSSVKVECLSRERLGVNITWKLQLVDAPPAS 1379

1380 LPDLHDIERALVGKDLLGRFTDLIQSGSFQLHLDSKTFPAET-IRFLQGDHFGTSPRTWF 1438 LPDL D+E AL GK L GRF DLIQSG+FQLHLDSKTF A+T IRFLQGD FGTSPRT F 1380 LPDLQDVEEALAGKYLAGRFADLIQSGTFQLHLDSKTFSADTSIRFLQGDRFGTSPRTQF 1439

1439 GCSEGFYQVLT-SEASQDGLGCVKCPEGSYSQDEECIPCPVGFYQEQAGSLACVPCPVGR 1497 GC EGF +V+ S+ASQD LGCVKCPEGSY QDE+CIPCP GFYQEQAGSLACVPCP GR 1440 GCLEGFGRVVAASDASQDALGCVKCPEGSYFQDEQCIPCPAGFYQEQAGSLACVPCPEGR 1499

1498 TTISAGAFSQTHCVTDCQRNEAGLQCDQNGQYRASQKDRGSGKAFCVDGEGRRLPWWETE 1557 TT+ AGAFSQTHCVTDCQ+NE GLQCDQ+ QYRASQ+DR SGKAFCVDGEGRRLPW E E 1500 TTVYAGAFSQTHCVTDCQKNEVGLQCDQDSQYRASQRDRTSGKAFCVDGEGRRLPWTEAE 1559

1558 APLEDSQCLMMQKFEKVPESKVIFDANAPVAVRSKVPDSEFPVMQCLTDCTEDEACSFFT 1617 APL D+QCL+M+KFEK+PESKVIF A+ V VRS+VP SE +MQCL DC DEAC F T 1560 APLVDAQCLVMRKFEKLPESKVIFSADVAVMVRSEVPGSESSLMQCLADCALDEACGFLT 1619

1618 VSTTEPEISCDFYAWTSDNVACMTSDQKRDALGNSKATSFGSLRCQVKVRSHGQDSPAVY 1677 VST E+SCDFYAW SD++AC TS + DALG S+ATSFGSL+CQVKVRS D AVY 1620 VSTAGSEVSCDFYAWASDSIACTTSGRSEDALGTSQATSFGSLQCQVKVRSREGDPLAVY 1679 226

1678 LKKGQGSTTTLQKRFEPTGFQNMLSGLYNPIVFSASGANLTDAHLFCLLACDRDLCCDGF 1737 LKKGQ T T QKRFE TGFQ+ LSG+Y+P+ FSASGA+L + HLFCLLACD D CCDGF 1680 LKKGQEFTITGQKRFEQTGFQSALSGMYSPVTFSASGASLAEVHLFCLLACDHDSCCDGF 1739

1738 VLTQVQGGAIICGLLSSPSVLLCNVKDWMDPSEAWANATCPGVTYDQESHQVILRLGDQE 1797 +L QVQGG ++CGLLSSP VLLC+V+DW DP+EA ANA+CPGVTYDQ+S QV LRLG QE 1740 ILVQVQGGPLLCGLLSSPDVLLCHVRDWRDPAEAQANASCPGVTYDQDSRQVTLRLGGQE 1799

1798 FIKSLTPLEGTQDTFTNFQQVYLWKDSDMGSRPESMGCRKDTVPRPASPTEAGLTTELFS 1857 I+ LTPLEGTQDT T+FQQVYLWKDSDMGSR ESMGCR+DT PRPASP+E LTT LFS 1800 -IRGLTPLEGTQDTLTSFQQVYLWKDSDMGSRSESMGCRRDTEPRPASPSETDLTTGLFS 1858

1858 PVDLNQVIVNGNQSLSSQKHWLFKHLFSAQQANLWCLSRCVQEHSFCQLAEITESASLYF 1917 PVDL QVIV+GN SL SQ+HWLFKHLFS QQANLWCLSRC E SFCQLAE+T+S LYF 1859 PVDLIQVIVDGNVSLPSQQHWLFKHLFSLQQANLWCLSRCAGEPSFCQLAEVTDSEPLYF 1918

1918 TCTLYPEAQVCDDIMESNAQGCRLILPQMPKALFRKKVILEDKVKNFYTRLPFQKLMGIS 1977 TCTLYPEAQVCDDI+ES+ +GCRLILP+ P AL+RKKV+L+D+VKNFY RLPFQKL GIS 1919 TCTLYPEAQVCDDILESSPKGCRLILPRRPSALYRKKVVLQDRVKNFYNRLPFQKLTGIS 1978

1978 IRNKVPMSEKSISNGFFECERRCDADPCCTGFGFLNVSQLKGGEVTCLTLNSLGIQMCSE 2037 IRNKVPMS+KSIS+GFFECER CD DPCCTGFGFLNVSQLKGGEVTCLTLNSLG+Q CSE 1979 IRNKVPMSDKSISSGFFECERLCDMDPCCTGFGFLNVSQLKGGEVTCLTLNSLGLQTCSE 2038

2038 ENGGAWRILDCGSPDIEVHTYPFGWYQKPIAQNNAPSFCPLVVLPSLTEKVSLDSWQSLA 2097 E GG WRILDCGSPD EV TYPFGWYQKP++ ++APSFCP V LP+LTE V+LDSWQSLA 2039 EYGGVWRILDCGSPDTEVRTYPFGWYQKPVSPSDAPSFCPSVALPALTENVALDSWQSLA 2098

2098 LSSVVVDPSIRHFDVAHVSTAATSNFSAVRDLCLSECSQHEACLITTLQTQPGAVRCMFY 2157 LSSV+VDPSIR+FDVAH+STAA NFSA RD CL ECS+H+ CL+TTLQTQPGAVRCMFY 2099 LSSVIVDPSIRNFDVAHISTAAVGNFSAARDRCLWECSRHQDCLVTTLQTQPGAVRCMFY 2158

2158 ADTQSCTHSLQGQNCRLLLREEATHIYRKPGISLLSYEASVPSVPISTHGRLLGRSQAIQ 2217 ADTQSCTHSLQ QNCRLLL EEAT+IYRKP I L + S PSVPI+THG+LLGRSQAIQ 2159 ADTQSCTHSLQAQNCRLLLHEEATYIYRKPNIPLPGFGTSSPSVPIATHGQLLGRSQAIQ 2218

2218 VGTSWKQVDQFLGVPYAAPPLAERRFQAPEPLNWTGSWDASKPRASCWQPGTRTSTSPGV 2277 VGTSWK VDQFLGVPYAAPPL E+RF+APE LNWTGSW+A+KPRA CWQPG RT T PGV 2219 VGTSWKPVDQFLGVPYAAPPLGEKRFRAPEHLNWTGSWEATKPRARCWQPGIRTPTPPGV 2278

2278 SEDCLYLNVFIPQNVAPNASVLVFFHNTMDREESEGWPAIDGSFLAAVGNLIVVTASYRV 2337 SEDCLYLNVF+PQN+APNASVLVFFHN + + S PA+DGSFLAAVGNLIVVTASYR 2279 SEDCLYLNVFVPQNMAPNASVLVFFHNAAEGKGSGDRPAVDGSFLAAVGNLIVVTASYRT 2338

2338 GVFGFLSSGSGEVSGNWGLLDQVAALTWVQTHIRGFGGDPRRVSLAADRGGADVASIHLL 2397 G+FGFLSSGS E+SGNWGLLDQV ALTWVQTHI+ FGGDPRRV+LAADRGGAD+ASIHL+ 2339 GIFGFLSSGSSELSGNWGLLDQVVALTWVQTHIQAFGGDPRRVTLAADRGGADIASIHLV 2398

2398 TARATNSQLFRRAVLMGGSALSPAAVISHERAQQQAIALAKEVSCPMSSSQEVVSCLRQK 2457 T RA NS+LFRRAVLMGGSALSPAAVI ERA+QQA ALAKEV CP SS QE+VSCLRQ+ 2399 TTRAANSRLFRRAVLMGGSALSPAAVIRPERARQQAAALAKEVGCPSSSVQEMVSCLRQE 2458

2458 PANVLNDAQTKLLAVSGPFHYWGPVIDGHFLREPPARALKRSLWVEVDLLIGSSQDDGLI 2517 PA +LNDAQTKLLAVSGPFHYWGPV+DG +LRE PAR L+R+ V+VDLLIGSSQDDGLI 2459 PARILNDAQTKLLAVSGPFHYWGPVVDGQYLRETPARVLQRAPRVKVDLLIGSSQDDGLI 2518

2518 NRAKAVKQFEESRGRTSSKTAFYQALQNSLGGEDSDARVEAAATWYYSLEHSTDDYASFS 2577 NRAKAVKQFEES+GRTSSKTAFYQALQNSLGGE +DA V+AAATWYYSLEH +DDYASFS 2519 NRAKAVKQFEESQGRTSSKTAFYQALQNSLGGEAADAGVQAAATWYYSLEHDSDDYASFS 2578

2578 RALENATRDYFIICPIIDMASAWAKRARGNVFMYHAPENYGHGSLELLADVQFALGLPFY 2637 RALE ATRDYFIICP+IDMAS WA+ RGNVFMYHAPE+Y H SLELL DV +A GLPFY 2579 RALEQATRDYFIICPVIDMASHWARTVRGNVFMYHAPESYSHSSLELLTDVLYAFGLPFY 2638 227

2638 PAYEGQFSLEEKSLSLKIMQYFSHFIRSGNPNYPYEFSRKVPTFATPWPDFVPRAGGENY 2697 PAYEGQF+LEEKSLSLKIMQYFS+FIRSGNPNYP+EFSR+ P FA PWPDFVPR G E+Y 2639 PAYEGQFTLEEKSLSLKIMQYFSNFIRSGNPNYPHEFSRRAPEFAAPWPDFVPRDGAESY 2698

2698 KEFSELLPNRQGLKKADCSFWSKYISSLKTSADGAKGGQSAESEEEELTAGSGLREDLLS 2757 KE S LLPNRQGLKKADCSFWSKYI SLK SAD K G SA+SEEE+ AGSGL EDLL 2699 KELSVLLPNRQGLKKADCSFWSKYIQSLKASADETKDGPSADSEEEDQPAGSGLTEDLLG 2758

2758 LQEPGSKTYSK 2768 L E SKTYSK 2759 LPELASKTYSK 2769

228

Appendix II

HMW CLU-CS and CLU-GST complexes were purified using SEC as described in Section 5.3.3. As described in Wyatt et al. 2009c, CLU-CS and CLU-GST complexes eluted in the void volume

7 (V0 > 4 x 10 Da) of a Superoseä 6 10/300 SEC column run at ~ 7 mL (Figure A1). Although CLU, CS, and GST were not individually passed over the column, the elution profiles of these proteins are similar to those shown in Wyatt et al. 2009c.

A 250 CLU-CS 200

150 CS

100 A280 nm CLU 50

0 0 4 8 12 16 20 24

Volume (mL)

1400 GST CLU-GST 1200 1000

800 600 CLU

A280 nm 400 200 0

0 4 8 12 16 20 24 Volume (mL)

Figure A1 SEC profiles of CLU-CS and CLU-GST. A) CLU-CS and B) CLU-GST eluted in the void volume of a Superoseä 6 10/300 SEC column The y-axis indicates absorbance at 280 nm and the x-axis indicates the elution volume (mL). The protein species corresponding to each elution peak is indicated at the top of the figure. The data here represents two independent experiments for CLU-CS and one experiment for CLU-GST. 229

Appendix III

FITC-labelled Tg-CS bound to the surface of RAW 264.7 macrophages, shown by the large increase in fluorescence of cells that had been incubated with the complex versus cells incubated with FITC- labelled BSA, CLU, Tg, or CS at the same concentration (Figure A2).

Number of Events

Raw 264.7 BSA CLU Tg CS Tg-CS

Figure A2 Flow cytometric analysis of protein binding to Raw 264.7 macrophages. Samples are indicated on the x-axis and the y-axis indicates the number of events emitted at 488 nm. The data shown is the averged number of events (n = 3) + SEM. Viable cells in this experiment were gated based on PI exclusion. The results shown are representative of one independent experiments.

To confirm that the fluorescent emissions given off by Raw 264.7 cells, when analysed by flow cytometry, was a result of proteins binding to viable cells, the samples (cells alone and cells incubated with FITC-labelled BSA, CS, Tg, and the Tg-CS complex) were fixed, wet-mounted, and examined using a Leica TCS confocal microscope (Figure A3).

230

Raw 264.7 BSA Tg CS Tg-CS

Figure A3 Confocal microscopy images of Raw 264.7 macrophages. Each column represents images for a single sample (labelled at the top). The top row depicts a grey-scale image of the cells, the middle row shows fluorescent emission against a black background, and the bottom row depicts an overlay of the fluorescence on the grey-scale image. It should be noted that the faint, green smears inside of cells are not representative of cell binding, but are a result of reflection off of the background. The data shown here is representative of one experiment.

231