Inhibiting Phosphorylation and Aggregation of Tau Protein Using R Domain Peptide Mimetics a Dissertation Presented to the Facul

Inhibiting Phosphorylation and Aggregation of Tau Protein Using R Domain Peptide

Mimetics

A dissertation presented to

the faculty of

the College of Arts and Sciences of Ohio University

In partial fulfillment

of the requirements for the degree

Doctor of Philosophy

Najah A. Alqaeisoom

August 2019

This dissertation titled

Inhibiting Phosphorylation and Aggregation of Tau Protein Using R Domain Peptide

Mimetics

NAJAH A. ALQAEISOOM

has been approved for

the Department of Chemistry and Biochemistry

and the College of Arts and Sciences by

Justin M. Holub

Assistant Professor of Biochemistry

Joseph Shields

Interim Dean, College of Arts and Sciences 3

ABSTRACT

ALQAEISOOM, NAJAH A., Ph.D., August 2019, Chemistry

Inhibiting Phosphorylation and Aggregation of Tau Protein Using R Domain Peptide

Mimetics

Director of Dissertation: Justin M. Holub

Tau protein plays a crucial role in stabilizing microtubules inside neuronal axons and maintaining the structural integrity of neurons. Binding of tau to microtubules at tau repeat domains (R) is regulated by phosphorylation. This phosphorylation is regulated by a family of enzymes called kinases. Under pathological conditions, tau is hyperphosphorylated by elevated activity of kinases such as the microtubule affinity- regulating kinase (MARK) proteins, leading to complete detachment of tau, microtubule collapse and ultimately, neuronal cell death. The free, hyper-phosphorylated tau proteins aggregate into insoluble prion-like oligomers which have been implicated in neurodegenerative diseases, including Alzheimer's disease (AD) and frontotemporal dementia. There is currently no treatment to prevent the progression of AD; all medications available today only reduce the symptoms of the disease. Moreover, using small molecule kinase inhibitors as treatment can cause serious negative side effects because of their lack of specificity. The research outlined in this work aims to develop a metabolically stable, selective peptide-based MARK kinase inhibitor that targets MARK proteins. This peptide-based inhibitor, designated tR1, was designed as a direct sequence memetic of the microtubule-binding R1 repeat domain of tau. Here, we show that tR1 4 peptides can inhibit MARK2 activity and reduce the level of tau phosphorylation in vitro and in cultured rat primary cortical neurons.

In the second segment of this project, we attempted to inhibit tau aggregation in vitro using peptide-based aggregation inhibitors. Here, we synthesized peptides designated (an-R3, PHF6, and PHF6*) which mimic nucleating sites in the microtubule binding repeat domain of full-length tau. We hypothesized that these peptides would associate with tau protein and block further tau aggregation. We assessed the ability of these three peptides to inhibit tau aggregation using in vitro heparin-induced tau aggregation assay. The aggregation products were analyzed by SDS-PAGE analysis and by circular dichroism (CD) spectropolarimetry. We provide evidence that the nucleation site located in R3 repeat domain of tau is more prone to aggregation than in R2 repeat domain. Moreover, we show that amphiphilic peptide sequences, in which polar or charged residues alternate with hydrophobic amino acids, are important for tau nucleation and aggregation.

DEDICATION

I dedicated this work to my parents, husband and my family

ACKNOWLEDGMENTS

First, I would like to acknowledge my advisor, Dr. Justin Holub for his support, patience, understanding, and his step by step guidance. I also would like to thank my dissertation committee members, Dr. Marcia Kieliszewski, Dr. Robert Colvin, Dr. Jana

Houser, and Dr. Jixin Chen for their time, efforts and their valuable comments to edit and improve my work. Special thanks to the Saudi Arabian Cultural Mission (SACM) for their financial support and covering all the cost of my study at Ohio University.

I also would like to sincerely thank my lab mates: Tang Tang, Ranju Pokhrel,

Chang Xu, Nahar Khairun, and Maya Sattler for their kindness, care, assistance, support and for creating a friendly and convenient research environment. Thanks to Dr. Michael

Held group and Dr. Jennifer Hines group who always help me whenever I ask them for help.

I acknowledge my collaborator Dr. Cheng Qian form Dr. Robert Colvin group for his cooperation with me on the work reported in Chapter 1. His significant contributions were of great help in getting the results of my first project. Special thanks to Danushka

Arachchige from Dr. Holub’s group for assisting in my research and helping me when I was not able to work in the lab.

Finally, I would like to thank my family, my husband, and my friends for their love, support and for wishing me the success. 7

TABLE OF CONTENTS

Page

Abstract ...... 3 Dedication ...... 5 Acknowledgments ...... 6 List of Tables...... 10 List of Figures ...... 11 List of Abbreviations ...... 13 Chapter 1: Introduction ...... 15 The Discovery of Tau Protein and its Role in Stabilizing Microtubules ...... 15 Biochemical Characterization of Tau ...... 15 Pathological Tau, Causes and Consequences ...... 17 Hyperphosphorylation of Tau Protein and the Formation of NFTs ...... 19 Pathological Tau Transmission ...... 23 Therapeutic Strategies for the Inhibition of Tauopathies...... 25 Inhibiting Tau Hyperphosphorylation ...... 26 Increasing of Microtubule Stability ...... 28 Increasing Tau Clearance ...... 30 Tau Immunotherapy ...... 33 Active Immunization ...... 33 Passive Immunization...... 34 Inhibition of Tau Aggregation ...... 35 Inhibiting Tau Misfolding ...... 37 Disrupting Tau Dimerization ...... 38 Accelerating Tau Aggregation ...... 39 β-sheet Breakers ...... 39 Peptide-Based Therapeutic for Tauopathies ...... 41 MARK Protein Activity, Inhibition by Small Molecules Versus Peptides ...... 43 Chapter 2: Inhibiting Tau Phosphorylation Using Human Tau Peptide-Based R Domain Mimetics ...... 46 Introduction ...... 46 Materials ...... 49 8

Methods ...... 51 Peptide Preparation ...... 51 Peptide Synthesis ...... 51 Labeling of Synthesized Peptides ...... 52 Capping of Synthesized Peptides ...... 52 Cleavage of the Peptides from the Resin ...... 53 Purification and Quantification of Peptides ...... 53 Characterization of Peptides by Analytical HPLC and Mass Spectrometry ... 54 Protein Preparation ...... 54 MARK2 Preparation ...... 54 hTau K18 Preparation ...... 56 Structural Analysis for MARK2 and hTau K18 by Circular Dichroism ...... 58 Evaluating the In Vitro Activity of Recombinant MARK2 ...... 59 In Vitro tR1 Stability Assessment ...... 60 Preparation of Rat Primary Cortical Neuron Culture...... 61 Cell Uptake of tR1 Peptide by Primary Cortical Neurons ...... 62 Evaluating the Toxicity of tR1 Peptide Using MTT Assay ...... 63 Immunofluorescence to Study the Ability of tR1 Peptide in Inhibiting MARK2 in Rat Primary Cortical Neurons ...... 64 Cell Imaging Using Fluorescence Microscopy ...... 65 Quantitative Image Analysis ...... 65 Western Blot to Study the Ability of tR1 Peptide in Inhibiting MARK2-Mediated Tau Phosphorylation in Rat Primary Cortical Neurons ...... 66 Western Blot Membrane Stripping and Re-blotting ...... 67 Densitometric Analysis of Western Blot Bands ...... 67 Data Analysis and Statics ...... 68 Results and Discussion ...... 68 Peptide Design ...... 68 tR1 Peptides Inhibit MARK2 Activity In Vitro and Reduce Tau Phosphorylation at Ser262 ...... 73 Stability of tR1 Peptide in Biological Media ...... 83 tR1 Toxicity and Internalization in Neurons ...... 85 tR1 Peptides Inhibit Phosphorylation of Tau at Ser262 in Rat Primary Cortical Neurons ...... 90 9

tR1 Peptides Inhibit Tau Phosphorylation at Ser262 and not at Thr231...... 95 Conclusion ...... 99 Chapter 3: Inhibiting Tau Aggregation Using Tau-Derived Peptide Mimetics ...... 100 Introduction ...... 100 Materials ...... 102 Methods ...... 103 Peptide Preparation ...... 103 Peptide Synthesis ...... 103 Capping of Synthesized Peptides ...... 104 Cleavage of the Peptides from the Resin ...... 104 Peptide Purification by HPLC ...... 105 Characterization of Peptides by Analytical HPLC and Mass Spectrometry . 105 Expression, Purification, and Characterization of Recombinant hTau K18 ...... 106 In Vitro hTau K18 Aggregation Assay ...... 107 SDS-PAGE Analysis ...... 108 Circular Dichroism (CD) Spectropolarimetry ...... 108 Results and Discussion ...... 108 Peptide Design ...... 108 Peptide Characterization ...... 113 Optimizing Conditions for the In Vitro hTau K18 Aggregation Assay ...... 115 Optimizing Incubation Time at 37 °C ...... 115 DTT Promotes hTau K18 Aggregation In Vitro ...... 119 Optimizing hTau K18 Concentration for CD Analysis ...... 122 Optimizing Inhibiting Peptide Concentration ...... 125 Effect of Tau-Derived Peptides on hTau K18 Aggregation In Vitro...... 129 Studying the Effect of Tau-Derived Peptides on hTau K18 Aggregation In Vitro Using Circular Dichroism (CD) Spectropolarimetry ...... 137 Conclusion ...... 140 Chapter 4: Summary and Future Work ...... 142 References ...... 145

LIST OF TABLES

Page

Table 1 Sequences of β-sheet breakers identified by mirror image phage display...... 41 Table 2 Calculated and observed masses for fluorescently labeled peptides synthesized in this work...... 71 Table 3 Calculated and observed masses for capped peptides synthesized in this work. 72 Table 4 Sequences, calculated and observed mass for peptides synthesized in this project...... 114

LIST OF FIGURES

Page

Figure 1: The six isoforms of tau encoded by the MAPT gene...... 17 Figure 2: Role of tau phosphorylation in regulating axonal transport...... 20 Figure 3: Schematic representation of tauopathy...... 22 Figure 4: Bar diagram of MARK2 showing the different domains and the location of these domains...... 47 Figure 5: (a) Schematic representation of tau protein bound to microtubules through its four binding sites (R1-R4)...... 70 Figure 6: Characterization of fluorescently labeled peptides by analytical HPLC...... 72 Figure 7: Characterization of capped peptides by analytical HPLC...... 73 Figure 8: Purification and characterization of recombinant hTau K18...... 74 Figure 9: Purification and characterization of recombinant MARK2...... 76 Figure 10: Antibody-based fluorescence polarization assay presenting the ability of recombinant MARK2 in phosphorylating tR1 peptide in vitro at Ser262...... 79 Figure 11: Recombinant MARK2 is active as it phosphorylates tR1 peptide and the recombinant hTau K18 in vitro at Ser262...... 80 Figure 12: tR1 peptides inhibit MARK2 in vitro...... 82 Figure 13: In vitro peptide stability tests suggest that tR1 peptide is stable in various reaction media...... 84 Figure 14: tR1 peptide is nontoxic in rat primary cortical neurons up to 300 μM...... 86 Figure 15: tR1 peptides penetrate neuronal cells...... 89 Figure 16: tR1 peptides inhibit tau phosphorylation at Ser262 in neuronal cells treated with endosomal disruptors...... 92 Figure 17: Western blot assay demonstrating the ability of tR1 peptide to inhibit tau phosphorylation at Ser262 in cultured primary neurons...... 95 Figure 18: tR1 peptide does not inhibit tau phosphorylation at Thr231 in cultured primary neurons...... 98 Figure 19: Structure of polyanions used to accelerate tau aggregation...... 110 Figure 20: Sequence of the an-R3 peptide synthesized for this project and its potential association with the tau R3 domain...... 111 Figure 21: Bar diagram representing the location and the sequence of the two nucleation sites PHF6*(red) and PHF6 (green) in the microtubule-binding domain of tau protein. 113 Figure 22: Characterization of peptides used in hTau K18 aggregation assay by analytical HPLC...... 114 12

Figure 23: Sequence of the peptides synthesized for use in aggregation assay...... 115 Figure 24: Optimizing incubation time at 37 °C for the in vitro hTau K18 aggregation assay...... 118 Figure 25: Aggregation kinetics of tau proteins...... 119 Figure 26: In vitro hTau K18 aggregation is enhanced by the presence of DTT...... 121 Figure 27: Dithiothreitol (DTT) reduces the disulfide linkages and keeps tau K18 monomer in the unfolded form...... 122 Figure 28: A standard far-UV CD spectra of recombinant hTau K18 monomers and aggregates...... 123 Figure 29: Far-UV CD spectra of recombinant hTau K18 aggregates in an aggregation buffer...... 125 Figure 30: Optimizing peptide concentration for in vitro hTau K18 aggregation assay. 127 Figure 31: Column graph showing the effect of a gradient concentration of an-R3 peptide on the concentration of hTau K18 aggregates...... 128 Figure 32: Percent inhibition of hTau K18 aggregation by gradient concentrations of an- R3 (0-500 µM) as quantified from SDS-PAGE experiment...... 129 Figure 33: Effect of peptides an-R3, PHF6 and PHF6* on in vitro hTau K18 aggregation...... 131 Figure 34: A graph representing the effect of an-R3, PHF6 and PHF6* on the concentration of hTau K18 monomers...... 132 Figure 35: Percent inhibition of hTau K18 aggregation by 40 µM of an-R3, PHF6 or PHF6* peptides as quantified from SDS-PAGE experiment...... 133 Figure 36: Far-UV CD spectra for isolated an-R3, PHF6 or PHF6* under standard aggregation conditions...... 135 Figure 37: Far-UV CD spectra showing the effect of an-R3, PHF6 or PHF6* on hTauK18 aggregation...... 139

LIST OF ABBREVIATIONS

Aβ ………..……………………………………………………………..amyloid β-peptide ACN…………………………………………………………………………….acetonitrile AD……………………………………………………………..……. .Alzheimer’s disease Ac-R1 or tR1………………………………………………….acylated R1 domain peptide ALP………………………………………….……………..autophagy-lysosomal pathway an-R3…………………………………………………………..anionic R3 domain peptide ATP………………………………………………………………...adenosine triphosphate AU……………………………………………………………………….Absorbance Units BBB…………………………………………………………………….blood-brain barrier BSA…………………………………………………………………bovine serum albumin β-sheet……………………………………………………………………………beta-sheet Baf……………………………………………………………………….…bafilomycin A1 5-CF………………………………………………………………….5-carboxyfluorescein CLQ……………………………………………………………………………chloroquine DTT………………………………………………………………………….. dithiothreitol ECL……………………………………………………..…..enhanced chemiluminescence EM………………………………………………………………..…...electron microscopy ESI……………………………………………………………….…electrospray ionization FP………………………………………………………………...fluorescence polarization FlutR1…………………………………………………..…fluorescently R1 domain peptide Fmoc-PAL-AM……………………………………………...…Fmoc-Pal-linker-AM resin GSK-3…………………………………………………………. glycogen synthase kinase3 HOBt……………………………………………………………..….hydroxybenzotriazole hTau K18………………………………….…repeat domain of human Tau (R1,2,3 and 4) kDa……………………………………………………………………………… kilodalton LB Broth…………………………………………………………….....Luria-Bertani broth MARK………………………………………..…… microtubule affinity-regulating kinase MAP……………………………………………………… microtubule-associated protein 14

MB…………………………………………………..……………………...methylene blue MS………………………………………………………………..…..…mass spectrometry MTBR…………………………………………………microtubule binding repeat domain mTOR ……………………………………………………mammalian target of rapamycin NFT…………………………………………………………..……..neurofibrillary tangles NMM…………………………………………………………………N-methylmorpholine NMP………………………………………………………………N-methyl-2-pyrrolidone NMR……………………………………………………….… nuclear magnetic resonance O.D…………………………………………………………………….……optical density PAO…………………………………………………………………….phenylarsine oxide PBS………………………………………………………..…….phosphate buffered saline PHF6……………………..……nucleation site in R3 of full-length tau (306VQIVYK311) PHF6*…………………..….…nucleation site in R2 of full-length tau (275 VQIINK 280) PHF………………………………………………………..………..paired helical filament PTM…………………………………………………...…...post translational modification PVDF………………………………………………..polyvinylidene difluoride membrane PyClock………………………………..6-chloro-benzotriazole-1-yl-oxy-tris-pyrrolidino- phosphoniumhexafluorophosphate (M.W. 554.85), C18H27N6OF6ClP2 R1, R2, R3 or R4……………………………………………………..repeat domain of tau RP-HPLC………………………reversed-phase high-performance liquid chromatography SDS-PAGE…………………..sodium dodecyl sulfate-polyacrylamide gel electrophoresis SPPS……………………………………………..………...... solid phase peptide synthesis TBS……………………………………………………………….……Tris-buffered saline TauO……………………………………………………………………...…..tau oligomers TCA……………………………………………………….……...... trichloroacetic acid TFA…………………………………………………………….……….trifluoroacetic acid UPP……………………………………………………...…ubiquitin-proteasome pathway V……………………………………………………………..……………………..voltage

CHAPTER 1: INTRODUCTION

The Discovery of Tau Protein and its Role in Stabilizing Microtubules

Tau protein is a member of the microtubule-associated protein (MAP) family. Tau is abundant in the brain, specifically in the axons of neurons (Binder, Frankfurter, &

Rebhun, 1985). Tau protein was discovered in 1975 when Marc Krischner and coworkers were searching for factors necessary for microtubule stabilization. They found that tau plays an important role in stabilizing microtubules, which are polymers of tubulin organized in a cylindrical shape inside the axons (Cleveland, Hwo, & Kirschner, 1977a;

Weingarten, Lockwood, Hwo, & Kirschner, 1975). Further studies showed that tau maintains the structural integrity of axons and facilitates the axonal transport of organelles and proteins (Ebneth et al., 1998; Maday, Twelvetrees, Moughamian, &

Holzbaur, 2014).

Biochemical Characterization of Tau

The first research performed after the discovery of tau was the determination of its biochemical properties (Cleveland et al., 1977a; Cleveland, Hwo, & Kirschner,

1977b). Tau is hydrophilic, dynamic, positively charged and natively unstructured under normal physiological conditions (Jeganathan, von Bergen, Mandelkow, & Mandelkow,

2008; Mietelska-Porowska, Wasik, Goras, Filipek, & Niewiadomska, 2014; Mukrasch et al., 2009; Schweers, Schönbrunn-Hanebeck, Marx, & Mandelkow, 1994). The structure of tau is divided into two primary domains based on its amino acid charges: a positively charged repeat domain at the C-terminus (G. Lee, Cowan, & Kirschner, 1988) and a projection domain consisting of an acidic insert of 29 amino acids at the N-terminus. This 16 dipolar property gives tau the ability to bind to the microtubules and the ability to aggregate. Tau proteins bind to microtubules through highly conserved repeat (R) domains, each consisting of 18 amino acids (Mietelska-Porowska et al., 2014).

In the central nervous system of adults, there are six isoforms of tau, each of which differs in the number of microtubule binding domains and in the presence or absence of the projection domain. These isoforms are encoded in the MAPT gene located on chromosome 17 in locus 17q21.3 (Álmos et al., 2008; Arendt, Stieler, & Holzer, 2016;

Neve, Harris, Kosik, Kurnit, & Donlon, 1986) and are produced by alternative splicing of exons 2, 3 and 10 of tau mRNA (Goedert, Spillantini, Jakes, Rutherford, & Crowther,

1989) (Figure 1).

Figure 1: The six isoforms of tau encoded by the MAPT gene. Gray color represents tau binding domains (R1-4), red color (N) represents the projection domain. The 4R/2N isoform is the longest isoforms with 4 repeat (R) domains and 2 inserts (N). The 3R/0N isoform is the shortest with only 3 R domains. R2 domain shown in green does not exist in all tau isoforms. Name of each isoform and number of amino acids that comprise the isoform is shown on the right. Image adapted from (Ballatore, Lee, & Trojanowski, 2007).

Pathological Tau, Causes and Consequences

Under normal physiological conditions, tau binds to tubulin inside the axons of neurons and stabilizes the structure of the microtubules. However, under pathological conditions, tau remains permanently detached from tubulin leading to microtubule collapse and neuronal cell death. The unattached tau proteins accumulate to form insoluble aggregates known as neurofibrillary tangles (NFTs). NFTs are classified as abnormal fibers that accumulate inside the cytoplasm of neuronal cells (Spillantini et al.,

1997). NFTs are the histopathological hallmark of a family of neurodegenerative diseases 18 collectively known as tauopathies (Iqbal, Liu, Gong, & Grundke-Iqbal, 2010).

Alzheimer’s disease (AD) (Iqbal et al., 2010), Down’s syndrome (D. P. Hanger et al.,

1991), frontotemporal dementia, Parkinsonism linked to chromosome 17 (Hutton et al.,

1998), and progressive supranuclear palsy (PSP) (Gozes et al., 2012) are examples of tauopathies.

The exact mechanism that leads to the formation of pathological tau is not known, but there are many factors inside the cell that facilitate tau misfolding and aggregation.

These factors include neutralizing the positive charge of tau monomers to overcome the electrostatic repulsions and thus facilitate dimerization of two tau monomers. This can be achieved either by extensive phosphorylation of tau or by binding of tau to polyanions such as fatty acids or nucleic acids (Kampers, Friedhoff, Biernat, Mandelkow, &

Mandelkow, 1996). Other factors that facilitate tau aggregation are functional impairment of specific metabolic pathways including: repairing system (chaperone) (Dickey et al.,

2007), clearance system (proteasome system) (Keller, Hanni, & Markesbery, 2000), energy source and antioxidant protection system (mitochondria and peroxisomes)

(Dumont et al., 2011), cleavage of tau by cellular proteases (Yin & Kuret, 2006), mutation in the MAPT gene (Arawaka et al., 1999) or cross-seeding by nucleators, which are fragments of tau aggregates that serve as a template-directed cleavage of full-length tau to produce microtubule-binding repeat domains (MTBRs). MTBRs were thought to be the core of paired helical filaments (PHFs) (C. M. Wischik, Edwards, Lai, Roth, &

Harrington, 1996). 19

Moreover, tau protein is highly modulated by many post-translational modifications (PTM) including phosphorylation (G. Lee, 2004), acetylation (T. J. Cohen et al., 2011), oxidation (Landino, Skreslet, & Alston, 2004), O-GlcNAcylation (Arnold et al., 1996), ubiquitination (Cripps et al., 2006), nitration (Reyes et al., 2008), isomerization (Miyasaka et al., 2005), glycation (Ledesma, Bonay, Colaço, & Avila,

1994), proteolytic cleavage (Yin & Kuret, 2006), polyamination (Wilhelmus et al., 2009), and sumoylation (Dorval & Fraser, 2006). Among all of these PTMs, hyperphosphorylation is suggested to be the primary reason for the occurrence of tau aggregates and tauopathies (Alonso, Zaidi, Novak, Grundke-Iqbal, & Iqbal, 2001; J.

Biernat, Gustke, Drewes, Mandelkow, & Mandelkow, 1993).

Hyperphosphorylation of Tau Protein and the Formation of NFTs

Although microtubule-bound tau is crucial for stabilizing microtubules, it is also thought to prevent axonal transport by acting as an obstacle to motor proteins and axonal transporters (Ebneth et al., 1998). Coordinated action of kinases and phosphatases allow axonal transport by causing a partial detachment of microtubule-bound tau (Cuchillo-

Ibanez et al., 2008; Eva-Maria Mandelkow, Thies, Trinczek, Biernat, & Mandelkow,

2004, p. 1; Mietelska-Porowska et al., 2014) (Figure 2).

Figure 2: Role of tau phosphorylation in regulating axonal transport. (a) Tau bound to microtubule forms a barrier for kinesin and other axonal transporters. (b) MARK2 and other kinases phosphorylate tau in its MTBR causing tau to disengage from tubulin and allowing axonal transporters to pass. Image adapted from (Eva-Maria Mandelkow et al., 2004). http://what-when-how.com/molecular-biology/microtubules- molecular-biology/

There have been 85 phosphorylation sites determined on tau protein sequence using mass spectrometry or antibodies specific for phospho-tau (Mietelska-Porowska et al., 2014). Two primary classes of kinases are involved in the phosphorylation of tau: proline-directed kinases and non-proline directed kinases. Under certain pathological conditions, some tau kinases are upregulated and hyperphosphorylate tau, causing it to dissociate from microtubules (Gerard Drewes, Ebneth, Preuss, Mandelkow, &

Mandelkow, 1997). If left untreated, the unbound tau can aggregate to form PHFs and

NFTs (Goedert, Spillantini, Cairns, & Crowther, 1992) (Figure 3). 22

Figure 3: Schematic representation of tauopathy. Tau bound to the microtubule maintains the structural integrity of the neuronal cell axon. Hyperphosphorylation of tau at R domains dissociates tau from microtubules, leading to microtubule collapse, axonal transport impairment, and synaptic loss. Hyperphosphorylated tau proteins self-assemble into dimers and higher order aggregates causing neuronal cell collapse and neurodegenerative diseases. Image adapted from (L. Martin, Latypova, & Terro, 2011).

The first PHFs were described with the aid of electron microscopy in 1963 as two filaments twisted around each other (Kidd, 1963; C. M. Wischik, 1985). Composition of

PHFs revealed that the core of PHF is composed of the MTBR of tau (Goedert, Wischik,

Crowther, Walker, & Klug, 1988) containing either the four repeat domains of tau or just three of the repeats (R1, R3 and R4) (Goedert, Spillantini, Potier, Ulrich, & Crowther,

1989; G. Lee et al., 1988). PHFs further accumulate to form NFTs.

Hyperphosphorylation of tau and the formation of PHFs and NFTs have been found in many tauopathies. Moreover, a greater amount of NFTs inside neuronal tissue of AD patient enhances the severity of the disease (Arriagada, Growdon, Hedley-Whyte, &

Hyman, 1992). Even though tauopathies all share the accumulation of misfolded tau, they present a wide variety of pathological and clinical symptoms ranging from movement disorders to cognitive defects. These differences in the clinical symptoms of tauopathies are a result of the variation in tau isoform composition of tau aggregates and also because of the diversity in PTM of tau protein (Murray et al., 2014).

The last stage of tauopathies is the formation of extracellular ghost tangles which are similar in composition to the intracellular NFTs, yet they accumulate outside dead cells in different areas of the brain (Endoh, Ogawara, Iwatsubo, Nakano, & Mori, 1993).

Pathological Tau Transmission

Tauopathies are progressive diseases (Gao & Hong, 2008) in which tau pathology starts to form in the entorhinal cortex, then transfer to the hippocampus and finally to all neocortex regions (Braak & Braak, 1991). However, recent evidence suggests that tau pathology is initiated in the locus coeruleus (Kelly et al., 2017). Internalization and 24 propagation of tau aggregates has been studied in cultured neurons (Guo & Lee, 2013;

Nonaka, Watanabe, Iwatsubo, & Hasegawa, 2010), non-neuronal cells (Frost, Jacks, &

Diamond, 2009; Guo & Lee, 2011) and in mice (Clavaguera et al., 2013, 2009; Iba et al.,

2013; Lasagna-Reeves et al., 2012) using either recombinant full-length tau (Rosenmann et al., 2006), synthetic tau aggregates prepared from the full length tau or tau K18 (a truncated isoform of tau-containing only the four R domains) (Frost et al., 2009; Guo &

Lee, 2011), cell lysate derived from transgenic mouse model of tauopathies (Clavaguera et al., 2009) or cell lysate extracted from human brain (Clavaguera et al., 2013; Lasagna-

Reeves et al., 2012). Remarkably, in all of these studies, the extracellular tau pathology penetrates the cells and recruits endogenous tau to form NFTs which spread to the other cells and develop neurodegenerative disease overtime (Clavaguera et al., 2013; Frost et al., 2009; Rosenmann et al., 2006). This result exemplifies the cell to cell transmission of tauopathy. The mechanism by which tau is released from the affected neurons and transduced to the neighboring neurons remains elusive, yet, several mechanisms have been proposed. One possible mechanism of tau transmission is through synaptic connections (L. Liu et al., 2012). However, a growing body of evidence suggest that tau aggregates enter the cell through endocytic pathways specifically by macropinocytosis

(Frost et al., 2009; J. W. Wu et al., 2013) which allows the extracellular cargo to enter the cytoplasm of the synaptic cells through vesicles formed by invagination of the cell membrane (Steinman, 1995; Swanson & Watts, 1995). Moreover, tau oligomers (tauO) have the potential to form pores in the membrane and thus permeate the cell (Flach et al.,

2012). Also, tunneling nanotubes (TNTs) in which a delicate membranous bridge is 25 established between neurons provide a direct connection and accessible pathway for the transmission of the pathogenic tau (Tardivel et al., 2016). Once inside the cell, pathogenic tau recruits the aggregation of native tau and then releases a small fragment of tau aggregates either in a free form or in an extracellular vesicle called an exosome (Asai et al., 2015; Y. Wang et al., 2017). These exosomes containing tau aggregates can fuse with the membrane of the recipient cell and deliver their cargo in order to continue disease propagation.

Therapeutic Strategies for the Inhibition of Tauopathies

AD is the most common age-related tauopathy that causes cognitive loss and dementia (Murray et al., 2014). This globally prevalent disease was discovered by Alois

Alzheimer in 1907 and now affect 5.5 million people in the United States; this number is expected to reach 13.8 million cases by 2050 (“2017 Alzheimer’s disease facts and figures,” 2017). The prevalence of AD is causing a steep economic burden. Indeed, in

2017, AD and the related costs to treating dementia have reached nearly $259 billion in the United States alone (“2017 Alzheimer’s disease facts and figures,” 2017). Moreover, no effective treatment has been found for AD.

The final stage in several neurodegenerative diseases including AD is thought to include the formation of extracellular aggregates called amyloid plaques consisting of truncated amyloid β (Aβ) protein and intracellular NFTs formed by the accumulation of pathological tau. Previously, amyloid protein, which is the monomer of Aβ, was proposed as the primary cause of AD. Therefore, intensive research had been employed to investigate a treatment to decrease or remove Aβ aggregates from the patient’s brain 26

(Panza et al., 2016). The efforts of searching for such treatments for Aβ aggregation did not fully succeed in which the current medications available commercially only reduce the symptoms of mild AD (Funke & Willbold, 2012). For example, passive anti-Aβ immunization was not efficient to cure AD; it was only helpful for some extent in relieving the symptoms of AD in mildly affected patients (Panza, Solfrizzi, Imbimbo, &

Logroscino, 2014). Hence, attention now is devoted toward targeting pathological tau

(Panza et al., 2016), not only because of the failure to obtain an effective cure for AD by targeting Aβ, but also because only NFTs are formed in some neurodegenerative diseases, including frontotemporal dementia. This finding suggests that tau is a primary cause of specific neurodegenerative diseases. Therapeutic strategies targeting tau include: inhibition of tau phosphorylation, tau immunotherapy, increasing of microtubule stability, increase tau clearance or inhibition of tau aggregation (Khanna, Kovalevich, Lee,

Trojanowski, & Brunden, 2016).

Inhibiting Tau Hyperphosphorylation

Protein phosphorylation by kinases is a fundamental intracellular process involving the transfer of a phosphate group from a donor such as a nucleotide adenosine triphosphate (ATP), to a polar side chain of a specific amino acid (Ser, Thr or Tyr) in the substrate protein. Phosphorylation by kinases regulates protein function and trafficking, signal transduction, cellular metabolism, transcription, apoptosis, and other cellular functions. There are approximately 518 known kinases in eukaryotic cells categorized into 50 superfamilies based on similarity in the catalytic domain and similarity in the biological activity (Manning, 2002). Because kinases are key regulators of nearly all 27 cellular processes, impairment in the kinase function result in many diseases. Indeed, the building block of PHFs found in many tauopathies is hyperphosphorylated tau (Bramblett et al., 1993). Several kinases including proline-directed kinases such as glycogen synthase kinase 3 (GSK-3) or non-proline directed kinases like microtubule-affinity regulating kinases (MARK1-4) are known to hyperphosphorylate tau and lead to neurodegenerative diseases (Gerard Drewes et al., 1997). The concentration of phosphorylated tau is often three to four times higher in AD patients compared to normal healthy human brain tissue (Ksiezak-Reding, Liu, & Yen, 1992).

Preventing tau hyperphosphorylation by inhibiting the kinases that are overexpressed or upregulated during disease pathogenesis is a potential therapeutic route to treat neurodegenerative diseases (Mietelska-Porowska et al., 2014). For this purpose, researchers have developed small molecules that competitively block the ATP-binding cavity of kinases implicated in neurodegenerative diseases. Even though some of the small molecules are effective in blocking the targeted kinase, they also inhibit other off- target kinases. For example, Le Corre and his group developed a library of the small molecule derived from Staurosporin (Yanagihara et al., 1991) to inhibit the kinase GSK-

3β. The strongest derivative, SRN-003556, was capable of inhibiting GSK-3β along with five other kinases (Le Corre et al., 2006). The lack of specificity of small molecule kinase inhibitors is one of the major issues that cause many negative side effects. The similarity between the ATP-binding cavities of kinases has led to difficulty in developing small molecule inhibitors that specifically targets only one kinase. Therefore, it is essential to 28 find an alternative therapeutic method that overcomes the disadvantages of the conventional small molecule method of inhibition.

Increasing of Microtubule Stability

Microtubules maintain neuronal morphology (Jacobs, 1986), establish and maintain neuronal polarity (Baas, Deitch, Black, & Banker, 1988) and facilitate axonal transport in which motor proteins walk over the outer surface of microtubules to deliver cargo to organelles (Conde & Cáceres, 2009). Microtubules are tube-like structures that are composed of alternating strands of α and β tubulin subunits, and they have two ends designated plus (+) and minus (-). In a healthy neuron, microtubules undergo a cycle of growing and shrinking known as “dynamic instability”. Microtubule stability is maintained for several hours whereas the dissociation of microtubules lasts only for minutes (Conde & Cáceres, 2009; Y. Li & Black, 1996). Furthermore, the plus (+) end of microtubules are more dynamic compared to the minus (-) end (R. A. Walker et al.,

1988). This dynamic instability of microtubules can be shifted in favor of more stable structures by microtubule-associated proteins (MAPs) such as tau in the axons and MAP2 in the dendrites (Bernhardt & Matus, 1984). Microtubule dissociation causes impairment in axonal transport (Hempen & Brion, 1996) which is common in neurodegenerative diseases.

Enhancing microtubule assembly has been suggested as a therapeutic route to treat tauopathies. Chemotherapeutics such as paclitaxel were used as microtubule stabilizing agent (Saloustros, Mavroudis, & Georgoulias, 2008), but their poor ability to penetrate the blood-brain barrier (BBB) limited their usage to treat tauopathies (Fellner et 29 al., 2002). Moreover, the strong anti-mitotic impact of such drugs resulted in dangerous side effects including neutropenia and peripheral neuropathies (Bedard, Di Leo, &

Piccart-Gebhart, 2010; Marupudi et al., 2007; Rowinsky, Eisenhauer, Chaudhry, Arbuck,

& Donehower, 1993). On the other hand, the small molecule Epothilone D showed good

BBB penetration and ability to improve microtubule stability with minimal side effects

(Brunden et al., 2010). Agents that are more effective than the small molecule Epothilone

D include the synthetic neuroprotective peptide Davunetide. This medication, also known as NAP, is an octapeptide (Asn-Ala-Pro-Val-Ser-Ile-Pro-Gln) mimetic of the activity- dependent neuroprotective protein (ADNP). ADNP is an essential protein for brain development normally expressed in glial cells (Bassan et al., 1999). Even though the exact mechanism of action of NAP is still unclear (Magen & Gozes, 2013), it is well established that NAP has a strong neuroprotective effect against neurodegenerative disorders such as dementia (Shiryaev et al., 2009) or AD (Matsuoka et al., 2008), oxidative stress (Busciglio et al., 2007), β amyloid peptide (Gozes, Divinski, & Piltzer,

2008), head injury (Gozes et al., 2005) and many other neuronal damaging factors (Sari,

2009; Wilkemeyer et al., 2003; Zemlyak, Manley, Sapolsky, & Gozes, 2007). Moreover,

NAP improves memory and learning ability (Bassan et al., 1999), reduces the level of phospho-tau (Matsuoka et al., 2008), stabilizes microtubules in neurons and enhances axonal transport possibly by binding to the β three tubulin (Divinski, Holtser-Cochav,

Vulih-Schultzman, Steingart, & Gozes, 2006). Recent evidence suggests that NAP enhances the tau: tubulin interaction, thus stabilizing microtubules through concomitant binding to tau and to the end-binding proteins (EBs) (Y Ivashko-Pachima, Sayas, 30

Malishkevich, & Gozes, 2017). The NAP peptide contains a Ser-Ile-Pro (SIP) motif that recognizes EB and interacts with it. This interaction mediates the binding of NAP to tau and microtubule, which increases microtubule stability (Yanina Ivashko-Pachima, Maor,

& Gozes, 2018). The neuroprotective ability of NAP against amyloid β peptide is attributed to its ability to prevent self-assembly of this peptide to form Aβ aggregates

(Ashur-Fabian et al., 2003). The NAP peptide also contains unique properties that make it a capable of inhibiting Aβ aggregation. NAP contains polar residues (Gln and Asn) at the end of the sequence to enhance peptide solubility. It also contains a hydrophobic core that allows it to bind to tau aggregates and a Pro residue to prevent β-sheet formation (Gozes,

Steingart, & Spier, 2004).

Increasing Tau Clearance

There are currently two cellular pathways to remove misfolded or damaged proteins from cells: the ubiquitin-proteasome pathway (UPP) and the autophagy- lysosomal pathway (ALP). In the ALP pathway, the unneeded proteins or organelles are encapsulated by a double membrane forming what is called an autophagosome. This autophagosome is then transferred and fuses with the lysosome, which leads to the degradation of the cargo by the enzymes in the lysosome (Dunn, 1990). The majority of undesired proteins are degraded by UPP (D. H. Lee & Goldberg, 1998). Degradation by this pathway is initiated when E3 ligase adds a polyubiquitin chain tag to the unwanted protein. Then, the tagged protein is recognized and degraded by the by the 26S proteasome. The 26 proteasome particle is a large cylindrical protein (2000 kDa) consisting mainly of the 20S catalytic domain and two 19S binding domains. Because 31 structured proteins cannot fit into the narrow opening of the 20S proteasome tunnel

(Thrower, 2000), the ubiquitinated protein is denatured in the 19S domain, and subsequently sent to the 20S particle where it is degraded (Hershko, 1988).

Misfolded or damaged proteins must be degraded and cleared to avoid cellular damage caused by binding of the misfolded protein with other cellular components or because of the self-assembly of the misfolded protein (Huang & Figueiredo-Pereira,

2010). It has been hypothesized that impairment in ALP (Menzies, Fleming, &

Rubinsztein, 2015) or UPP could be one of the reasons for the accumulation of the misfolded tau protein in neurodegenerative diseases (Alves-Rodrigues, Gregori, &

Figueiredo-Pereira, 1998; Keller et al., 2000). Hence, activation of UPP or ALP to either prevent tau aggregation or stimulate clearance of tau aggregates is a potential therapeutic route for neurodegenerative diseases.

The 19S binding domain in the proteasome compartment activates the 20S catalytic domain by deubiquitinating the protein and by hydrolyzing ATP. Activating the

19S subunit which in turn activates the UPP is a potential therapeutic target allowing the clearance of tau (Huang & Figueiredo-Pereira, 2010). Activating the proteasome can also be achieved by the consumption of oleuropein, a phenolic compound found in Olea europea leaf extract, olives and olive oil (Katsiki, Chondrogianni, Chinou, Rivett, &

Gonos, 2007). Some other natural products such as the antioxidant sulforaphane increase expression of proteasome and thus help to dispose misfolded tau (Kwak, Cho, Huang,

Shin, & Kensler, 2007). Moreover, PROTACS, an abbreviation for PROteolysis

TArgeting Chimera moleculeS, are novel synthetic molecules used to induce degradation 32 of a specific protein. The unique structure of PROTACS allows them to capture the target protein specifically from the bulk of the cytoplasm and bind it to the E3 ligase for ubiquitination; ultimately leading to degradation by the proteasome (Schneekloth,

Pucheault, Tae, & Crews, 2008). In 2018, Lu and his group synthesized a novel peptide

PROTACS that contained two motifs: one that binds selectively to tau protein and the other motif binds to the Keap1-Cullin 3 based E3 ligase complex (M. Lu et al., 2018).

Keap1, an abbreviation for Kelch-like ECH-associated protein 1, is an adaptor protein of

Cullin 3 based E3 ligase complex (A. Kobayashi et al., 2004). The most common substrate of the Keap1-Cullin 3 based E3 ligase is the nuclear factor erythroid 2 related factor 2 (Nrf2) which plays a vital role in protecting the cell from oxidative stress through regulating the expression of antioxidant enzymes and proteins (A. Kobayashi et al.,

2004). The activity of Nrf2 is inhibited by ubiquitination mediated by binding to Keap1

(McMahon, Itoh, Yamamoto, & Hayes, 2003). In certain diseases such as AD or

Parkinson’s disease, the activity of Nrf2 drops (Ramsey et al., 2007). Thus, modulating

Keap1-Nrf2 pathway is a potential therapeutic route to treat several neurodegenerative diseases (Deshmukh, Unni, Krishnappa, & Padmanabhan, 2017). Recently, Lu and coworkers succeeded in designing a peptide PROTAC that recruits intracellular tau proteins to the Keap1-Cullin 3 based E3 ligase complex for ubiquitination and degradation by the UPP (M. Lu et al., 2018).

Activating ALP helps to clear NFTs since these large aggregates cannot fit into the narrow opening of the proteasome. Several studies conducted in yeast, fly and mouse animal model of tauopathies have shown a decrease in the NFTs upon treatment with the 33 autophagy activator, rapamycin (Ozcelik et al., 2013) (Noda & Ohsumi, 1998).

Rapamycin induces autophagy by inhibiting the activity of the mammalian target of rapamycin (mTOR) (Ravikumar et al., 2004; L. Wu et al., 2013). mTOR is a Ser/Thr kinase involved in regulating a wide range of cellular process including cell viability and growth (Hay, 2004; Switon, Kotulska, Janusz-Kaminska, Zmorzynska, & Jaworski,

2017). Extra and intracellular stimuli regulate the activity of mTOR which in turn has a downstream effect on transcription, translation, autophagy and many other cellular processes (Switon et al., 2017). Hence, disruption of mTOR signal transduction by rapamycin suppresses the translation of tau mRNA (Morita & Sobue, 2009) and causes other negative side effects. The autophagy enhancer lithium chloride (LiCl) is a good alternative for rapamycin. LiCl induces autophagy enabling tau aggregates clearance. It also reduces tau hyperphosphorylation by inhibiting the enzyme GSK-3 (Shimada et al.,

2012).

Tau Immunotherapy

Initially, it was believed that immunotherapy is not suitable to treat tauopathies because antibodies are not cell permeable and thus cannot reach tau aggregates accumulating inside neuronal cells. Yet, cell to cell transmission of pathological tau

(Frost et al., 2009) allows extracellular antibodies to bind to the released misfolded tau and prevent it from infecting neighboring cells (Yanamandra et al., 2013).

Active Immunization

First attempts to develop tau immunotherapy using active immunization have shown that injecting a wild-type mouse with 50 μg of full-length recombinant tau caused 34 axonal impairment and formation of NFTs (Rosenmann et al., 2006). However, a reduction in tau aggregates was observed after injecting transgenic mouse model of tauopathy with phospho-tau 379–408 phosphorylated at Ser396 and Ser404 (Asuni,

Boutajangout, Quartermain, & Sigurdsson, 2007). The same result was obtained when injecting 100 μg of a combination of three short peptide sequences of phospho-tau

(tau195-213 phosphorylated at 202 and 205, tau 207-220 phosphorylated at 212 and 214 and tau 224-238 phosphorylated at 231) (Boimel et al., 2010). Even though it seems safe to use phospho-tau as a tau immunotherapy, injecting phospho-tau several times caused neuroinflammation (Rozenstein-Tsalkovich et al., 2013). Hence, extreme caution should be taken when designing and using active immunotherapy for tauopathies.

Passive Immunization

Passive immunization against tau pathology is an active area of research. Several monoclonal antibodies have been developed against specific epitopes in pathological tau.

MC1, for instance, is an antibody that recognizes an early conformational change of tau that produces misfolded tau (d’Abramo, Acker, Jimenez, & Davies, 2013; Jicha, Bowser,

Kazam, & Davies, 1997). Antibodies such as PHF1 recognize phospho-tau at Ser 396 or

Ser 404, phosphorylation sites exist in normal tau and in PHFs (d’Abramo et al., 2013; L.

Otvos et al., 1994). Other antibodies such as tau oligomer monoclonal antibody (TOMA) are developed against tauO (Castillo-Carranza et al., 2014). Results of administration of passive immunizations to mouse models of tauopathies are promising in which clearance of tau lesions and a reduction in the progression of tauopathies was achieved (d’Abramo et al., 2013; Schroeder, Joly-Amado, Gordon, & Morgan, 2016). 35

Inhibition of Tau Aggregation

Tau protein is classified as an amyloidogenic protein because of its ability to self- assemble into fibrils (Nizynski, Dzwolak, & Nieznanski, 2017). Indeed, pathological accumulation of tau is a common feature of neurodegenerative diseases. Although the exact mechanism and the key event triggering tau aggregation is still under debate, it is generally accepted that tau aggregation is a multistep process starting with misfolding of tau proteins (Chirita, Congdon, Yin, & Kuret, 2005), followed by dimerization of misfolded tau (Wille, 1992). These dimers accumulate to form prefibrillar tauO

(Friedhoff, von Bergen, Mandelkow, Davies, & Mandelkow, 1998) (Sahara et al., 2007) which further aggregate to produce mature, higher order tau aggregates (PHFs and

NFTs).

During the aggregation process, the intrinsically unfolded tau monomers undergo a transition from random coil to β-sheet structure, resulting in the highly organized secondary structure of PHFs (Martin von Bergen, Barghorn, Biernat, Mandelkow, &

Mandelkow, 2005). The core of PHFs is made up of the repeat domain of tau proteins

(Crowther & Wischik, 1985; C. M. Wischik, 1985). Two amphiphilic hexapeptide motifs consisting of hydrophobic amino acids followed by charged or polar amino acids within the repeat domain of tau are fundamental for filament formation and act as nucleating sites. These motifs are PHF6* (VQIINK) located at the N-terminus of the R2 repeat and

PHF6 (VQIVYK) at the N-terminus of R3 repeat domain (M. von Bergen et al., 2000;

Martin von Bergen et al., 2001). 36

The rate-limiting step of tau aggregation is the dimerization. This was determined by studying the in vitro thiazine red induced full-length tau aggregation as a function of time and tau concentration. Results of this study were examined using transmission electron microscopy. Following that the obtained data were fitted into a simple homogeneous nucleation model to estimate the equilibrium between tau monomers and tau aggregates and to study tau aggregation kinetics (Congdon et al., 2008).

The ability of tau to dimerize is attributed mainly to the two main interaction forces. The first one is the π-stacking between aromatic rings of the two nucleating sites of tau protein (Gazit, 2002). The second force is hydrophobic interaction between β-sheet strands (Zheng, Baghkhanian, & Nowick, 2013) promoted by the formation of intermolecular disulfide bonds (Schweers, Mandelkow, Biernat, & Mandelkow, 1995).

Tau contains Cys291 and Cys322 in R2 and R3 domains respectively (Schweers et al.,

1995), and the oxidation of the Cys brings the nucleating sites of tau into close proximity facilitating β-sheet formation (M. von Bergen et al., 2000).

In the past, NFTs were considered the toxic form of tau aggregates that cause synaptic lesion and neuronal death, but because synaptic and memory defects occur long before the formation of NFTs, it has been hypothesized that the prefibrillar aggregates, tauO, are the most toxic form of tau (Lasagna-Reeves et al., 2011). Several studies have shown that tauO disrupt the integrity of the membrane causing cell death, cognitive disorders and neurodegenerative disease, inflammation and neuronal damage, and mitochondrial and synaptic impairment (Flach et al., 2012; Gerson et al., 2016; Lasagna-

Reeves et al., 2011; Nilson et al., 2016). Thus, disrupting early steps that lead to the 37 formation of tau oligomers could represent a viable therapeutic strategy for curing tauopathies.

Inhibiting Tau Misfolding

Tau misfolding is a very early step in tau aggregation pathway so, inhibiting this process could theoretically prevent the entire aggregation process. In the cell, protein conformation is controlled by molecular chaperones which consist of conserved families of related proteins such as heat shock proteins (Hsp) (Dou et al., 2003). In vitro studies and a transgenic mouse model of tauopathy showed that molecular chaperones decrease tau aggregation. It was found that Hsp70 together with Hsp90 interact with tau monomer or tauO (Dou et al., 2003; Patterson, Ward, et al., 2011) to adjust the conformation of the misfolded tau monomer or to mediate the degradation of misfolded tauO.

Moreover, Lu and coworkers have shown that the mitotic regulator, prolyl cis/trans isomerase Pin1, catalyzes a conformational change of phosphorylated tau during

AD, allowing it to attach to microtubules, thus promoting microtubule assembly. The isomerase enzyme Pin1 is usually found in the nucleus (Ping Lu, Hanes, & Hunter, 1996) and catalyzes the prolyl isomerization of phospho-Ser or phospho-Thr preceding a proline reside in mitotic phosphoproteins (Yaffe et al., 1997). However, in the AD brain,

Pin1 translocates to the cytoplasm of the neuronal cells and there it recognizes only one site of phospho-Thr-Pro in the phosphorylated tau and catalyzes the cis-to-trans prolyl isomerization resulting in the twist of phospho-tau to the normal, healthy structure which restores the function of tau protein and prevents tau aggregation (Driver, Zhou, & Lu,

2015; P.-J. Lu, Wulf, Zhou, Davies, & Lu, 1999). Furthermore, it was found that Pin1 is 38 depleted (P.-J. Lu et al., 1999) or inactivated (Driver et al., 2015) during AD causing tau aggregation and NFT formation. Since Pin1 regulates the function of tau during neurodegenerative disease (P.-J. Lu et al., 1999), it is possible that Pin1 may serve as a target that corrects the structure of misfolded tau and restore its normal function.

Disrupting Tau Dimerization

Preventing tau homodimerization could be a viable therapeutic route to prevent the formation of the toxic tauO (V. M.-Y. Lee, Brunden, Hutton, & Trojanowski, 2011).

Methylene blue (MB) also known as methylthioninium chloride is a tricyclic aromatic phenothiazine used to treat malaria (Wainwright & Amaral, 2005) and methemoglobinemia. In 1996, Wischik and his group demonstrated the ability of MB in preventing tau dimerization and aggregation in vitro (C. M. Wischik et al., 1996). The inhibitory effect of MB was also observed in a transgenic mouse model (Hosokawa et al.,

2012; Claude M. Wischik, Bentham, Wischik, & Seng, 2008). In the absence of a reducing agent, MB oxidizes cysteine resides within tau repeat domain resulting in the formation of an intramolecular disulfide linkage and a folded tau monomer that is unable to dimerize with another tau (Crowe et al., 2013). Even though MB succeed in inhibiting tau aggregation, it failed phase III clinical trials in which LMTM, a reduced form of MB did not improve cognition in mild or moderate AD patients (Gauthier et al., 2016).

Furthermore, MB has been found to interact with cellular proteins other than tau causing adverse side effects (Gillman, 2011; Oz, Lorke, Hasan, & Petroianu, 2011).

Besides MB, there are many other small molecule inhibitors of tau aggregation such as rhodanines, phenylthiazolyl hydrazides, polyphenols, N-phenylamines, 39 anthraquinones, benzothiazoles, phenothiazines, porphyrins (Bulic et al., 2009). For most of these small molecule inhibitors, the exact mechanism of action and the specific cellular protein targets is poorly understood (T. Liu & Bitan, 2012). Importantly, such small molecule inhibitors are often nonspecific as they can bind to tau and other proteins and thus interfere with multiple cellular processes.

Accelerating Tau Aggregation

Since the accumulation of tau to NFTs seems to protect the cell from the toxic effect of the prefibrillar tauO, increasing the rate of NFT formation is proposed as a therapeutic treatment method for neurodegenerative diseases (T. Liu & Bitan, 2012). A recent study performed by Lo and coworkers revealed the ability of Azure C, which is a derivative for MB, as a potent enhancer for tau aggregation. It protects the cell from the toxic effect of tauO by accelerating tauO aggregation to nontoxic higher order aggregates

(Lo Cascio & Kayed, 2018) which can be cleared by ALP.

β-sheet Breakers

β-sheet breakers are a novel class of therapies which specifically target the amyloidogenic protein and block its nucleation site. The structure of the β-sheet breaker is similar to the nucleation site of the amyloidogenic protein and has the same level of hydrophobicity enabling it to bind the core of the amyloidogenic protein. On the other hand, the structure of β-sheet breakers are modified so that they prevent β-sheet formation (Soto, Kindy, Baumann, & Frangione, 1996), which in turn inhibits the formation of the toxic, oligomeric form of amyloid fibrils. Examples for modifications in the sequence of β-sheet breakers that prevent β-sheet formations are incorporation of : 1) 40 a proline amino acid which is known to have a strong ability in preventing β-sheet structure (Soto et al., 1996); 2) alternating N-methyl amino acids which creates two faces: an unmodified face capable of binding to β-sheet strands and a methylated face unable to form hydrogen bonds with the adjacent β-sheet strands (Amijee, Madine,

Middleton, & Doig, 2009; E. Hughes, Burke, & Doig, 2000); and 3) ester bonds that alternate with the amide bonds of the peptide backbone. This modification also creates two faces with one being unable to form hydrogen bond networks with a new tau monomer or nearby β-sheet strands (Gordon & Meredith, 2003).

The first β-sheet breaker for tau protein was generated by Sievers and his group in

2011 using Rosetta software. The all D-amino acid peptide TLKIVW was designed based on the atomic structure of aggregates of tau nucleation site, PHF6 (306 VQIVYK 311).

The ability of this peptide in binding to the tau nucleation site and preventing the aggregation process was proven in vitro using thioflavin S, nuclear magnetic resonance

(NMR) and electron microscopy (Sievers et al., 2011).

In 2016, Dimmers and coworkers applied mirror image phage display to identify a library of peptides that bind to PHF6 fibrils and prevent further aggregation (Dammers et al., 2016). Mirror image phage display is a technique used to screen a library of L- peptides (the amino group appears to the left of the isomeric α-carbon atoms) for their ability to bind to the unnatural D-target protein (the amino group appears to the right of the isomeric α-carbon atoms). Once bound peptides are identified, their D-form mirror images will be able to bind to the naturally occurring L-target protein (Schumacher et al.,

1996). 41

The sequence of tau aggregation inhibitory peptides is shown in Table 1. These peptides demonstrated capability in inhibiting tau aggregation and crossing the cell membrane (Dammers et al., 2016).

Table 1

Sequences of β-sheet breakers identified by mirror image phage display. Peptide name Sequence Peptide charge at pH 7.4 APT D-APTLLRLHSLGA-OH +1

KNT D-KNTPQHRKLRLS-OH +1

TL28 L-TTSLQMRLYYPP-OH +1

TD28 D-TTSLQMRLYYPP-OH +1

TD28rev D-PPYYLRMQLSTT-OH +1

Note: Adapted from (Dammers et al., 2016).

Peptide-Based Therapeutic for Tauopathies

In general, medications fall into two main categories: 1) small molecules treatments which are organic compounds with a low molecular weight (less than 500 Da); and 2) biologic treatments such as peptides, recombinant proteins or antibodies with a molecular weight higher than 5000 Da (Craik, Fairlie, Liras, & Price, 2013). Over the past 20 years, peptide therapeutics have emerged as powerful agents to treat a wide variety of diseases (Henninot, Collins, & Nuss, 2018). Peptides are short oligomers consisting of 2 to 50 amino acids. They occur naturally and play important roles by acting as hormones (Saugy, 2006), neurotransmitters (Giordano, MarchiÃ2, Timofeeva, & 42

Biagini, 2014), or anti-bacterial agents (Padhi, Sengupta, Sengupta, Roehm, &

Sonawane, 2014). Insulin extracted from a living organism was the first peptide treatment introduced to the market in 1923 (Banting, Best, Collip, Campbell, & Fletcher, 1922).

Since 2000, peptide treatments have flourished and in 2017, there were more than 50 clinically-approved peptide-based therapeutics commercially available in the pharmaceutical markets (Lau & Dunn, 2018).

In some cases, peptides have proven suitable alternatives for small molecule drugs

(Funke & Willbold, 2012). Peptides provide a large surface area for interaction with target proteins and can adopt the appropriate conformation to bind specifically to the desired target. High specificity may prevent interference with other biomolecules, minimize toxicity and unwanted side effects. Besides their high specificity and relative safety, peptides are more advantageous over small molecules because of their high efficiency, biological activity, structural diversity and their low accumulation inside the cell (Craik et al., 2013; Goyal, Shuaib, Mann, & Goyal, 2017). Despite all these advantages, using peptides as drugs still presents two major challenges. The first challenge for using peptides as therapeutics is problems with the drug delivery resulting from the poor ability of peptides to cross the cell membrane or BBB. The second challenge is the rapid degradation of the peptides in the plasma by serum proteases which dramatically shorten the half-life of the peptide. However, development in peptide design and peptide synthesis techniques have overcome many of these issues (Poduslo, Curran,

Kumar, Frangione, & Soto, 1999). Susceptibility of peptides for enzymatic degradation can be overcome by simple modifications in peptide structure such as switching from L 43 to D-enantiomeric amino acids (Robson, 1996; Tugyi et al., 2005) or by protecting the ends of the peptide by incorporating an amide bond at the C-terminus and acetyl group at the N-terminus (Sadowski et al., 2004). Moreover, covalent conjugation of polyamines to the peptide sequence enhance the ability of the peptide to cross the BBB (Poduslo &

Curran, 1996; Poduslo et al., 1999). Additional peptide modifications to improve pharmacokinetic properties are reviewed elsewhere (Avan, Hall, & Katritzky, 2014;

Henninot, Collins, & Nuss, 2018; Werner, Cabalteja, & Horne, 2016).

MARK Protein Activity, Inhibition by Small Molecules Versus Peptides

The MARK proteins are a family of Ser/Thr kinases that phosphorylate MAPs including tau at KXGS motifs of the MTBR (G. Drewes et al., 1995; Naz, Anjum, Islam,

Ahmad, & Hassan, 2013). In humans, the MARK family consist of four known isoforms

(MARK1-4) (G Drewes, 2004; Gerard Drewes et al., 1997; Trinczek, Brajenovic, Ebneth,

& Drewes, 2004). Several lines of evidence demonstrate that the balance of both the activity and the level of expression of MARK proteins is of vital importance in preventing tauopathies (Chen et al., 2006; Ebneth et al., 1998; Yu et al., 2012). Indeed, hyperactive MARK proteins phosphorylate tau at Ser262 which can disrupt the attachment of tau to microtubules (J. Biernat et al., 1993). Following detachment, the free tau can be hyperphosphorylated by cytosolic MAPs kinases including GSK-3β (Hong,

Chen, Klein, & Lee, 1997; Lucas et al., 2001; Takashima et al., 1998) and Cyclin- dependent kinase 5 (Cdk5) (Cruz, Tseng, Goldman, Shih, & Tsai, 2003). Thus, MARK hyperactivity is suggested to play a central role in initiating tau hyperphosphorylation and ultimately the occurrence of tauopathies (Ando et al., 2016; G Drewes, 2004). Reducing 44 the activity of MARK and other major kinases implicated in the pathogenesis has been considered as a promising therapeutic approach for tauopathies (Annadurai, Agrawal,

Džubák, Hajdúch, & Das, 2017; Caccamo, Oddo, Tran, & LaFerla, 2007; Levy et al.,

2013).

The most common strategy to inhibit kinase activity is to competitively target the

ATP-binding pocket of an inactive kinase using small molecules (Y. Liu & Gray, 2006).

Since the ATP-binding pocket is highly conserved in many different kinases, selective inhibition of a particular kinase cannot be easily achieved (Bain, McLauchlan, Elliott, &

Cohen, 2003; Fabian et al., 2005). Even though there are some small molecules inhibitors that have been specifically designed to target MARK, none of these inhibitors have reached clinical trials (Annadurai et al., 2017). Staurosporine, for instance, is a small molecule that shows an inhibitory effect on MARK activity. However, toxicity and low pharmaceutical properties (Gurley, Umbarger, Kim, Bradbury, & Lehnert, 1998) limited its usage as a MARK inhibitor (Annadurai et al., 2017). Other small-molecule inhibitors of MARK proteins lack specificity; for example, the compound 30019 inhibits all four

MARK isoforms with similar affinity (Timm et al., 2011).

An alternative strategy to avoid the toxicity of small molecules and to achieve selective inhibition of kinases is the development of peptide-based kinase inhibitors

(Eldar-Finkelman & Eisenstein, 2009). Such inhibitory peptides can be developed to mimic the binding site of the kinase substrate. Because each kinase binds to a specific motif and a specific sequence of amino acids in the substrate, selective inhibition of the kinase can, in theory, be achieved by designing a peptide that mimics the natural 45 substrate and contains the phosphorylation site surrounded by specific residues important for kinase recognition. Such peptides can compete with the natural substrate for the active site of the target kinase which results in selective inhibition of the impairment kinase

(Eldar-Finkelman & Eisenstein, 2009).

In the current study reported in this dissertation, we have successfully developed a novel peptide-based inhibitor for the MARK family isoform MARK2. Peptide-based inhibitors are preferred over small molecule inhibitors because of the high specificity, low toxicity, and the ease of design (Eldar-Finkelman & Eisenstein, 2009). Here we show that a peptide derived from the R1 domain of tau, (designated tR1) is capable of inhibiting the activity of MARK2 in vitro and in cultured rat primary cortical neurons.

The second study was performed to explore the mechanisms of tau aggregation and to develop a peptide-based inhibitor of tau aggregation. For this purpose, we synthesized three peptides: an-R3, PHF6 and PHF6*, which were generated on the basis of specific sequences in the MTBR of tau. We show that these synthetic peptides have a slight inhibitory effect on in vitro heparin-induced tau K18 aggregation. The modest ability of an-R3, PHF6 or PHF6* to inhibit tau aggregation maybe attributed to the self- assembly property of these peptides which prevent them from associating with tau and significantly inhibit tau aggregation. Our results also suggest that the nucleation site

PHF6 in the R3 domain of tau is more prone to aggregation than PHF6* in the tau R2 domain. 46

CHAPTER 2: INHIBITING TAU PHOSPHORYLATION USING HUMAN TAU

PEPTIDE-BASED R DOMAIN MIMETICS

The text of this chapter contains materials that are originally published as:

Alqaeisoom, N., Qian, C., Arachchige, D., Colvin, R. A., & Holub, J. M. (2018).

Inhibiting Phosphorylation of Tau (τ) Proteins at Ser262 Using Peptide-Based R1

Domain Mimetics. https://doi.org/10.1007/s10989-018-9689-6

Reprinted with permission from Springer Science Business Media, LLC, part of Springer

Introduction

The MARK proteins are a group of Ser/Thr kinases involved in maintaining cell polarity (D. Cohen, Brennwald, Rodriguez-Boulan, & Müsch, 2004; Timm et al., 2011) and facilitating neurite outgrowth (Jacek Biernat et al., 2002). In humans, the MARK family includes four isoforms (MARK1-4) (G Drewes, 2004; Gerard Drewes et al.,

1997; Trinczek et al., 2004). These isoforms have a similar conserved structure consisting of an N-terminal domain, catalytic domain, ubiquitin domain, spacer and C-terminal tail

(Gerard Drewes et al., 1997; Marx et al., 2006) (Figure 4).

Figure 4: Bar diagram of MARK2 showing the different domains and the location of these domains. The amino acid number is shown above the bar diagram. CD: the common docking domain, UBA: the ubiquitin associated domain. Image adapted from (Marx et al., 2006).

The activity of MARK is controlled by multiple signaling pathways. MARK2, for instance, is activated by phosphorylation of T208 in its catalytic domain by the kinase

MARKK/TAO-1 (Timm et al., 2003) or by Liver kinase B1 (LKB1) (Lizcano et al.,

2004, p. 1). Phosphorylation of MARK2 at T208 by the two indicated enzymes enable the substrate to access the catalytic domain of MARK2 (Huse & Kuriyan, 2002). On the other hand, phosphorylation of MARK2 at S212 in the catalytic domain by β-isoform of

GSK-3 inactivates it (Timm et al., 2003).

MARK isoforms phosphorylate MAPs specifically within the KXGS motifs located in each one of the MTBR of MAPs (G. Drewes et al., 1995; Gerard Drewes et al.,

1997; Ebneth, Drewes, Mandelkow, & Mandelkow, 1999). The respective phosphorylation of tau at Ser262, Ser293, Ser324 and Ser356 by MARK2 in the R1, R2,

R3, and R4 domains cause tau to dissociate from microtubules which in turn decreases microtubule stability (Annadurai et al., 2017; J. Biernat et al., 1993; G. Drewes et al.,

1995; Gerard Drewes et al., 1997). Even though microtubule instability is a normal process required for cell polarization and cell growth (Jacek Biernat et al., 2002), the sensitive equilibrium of MARK and phosphatase activity can be easily disrupted, 48 resulting in the disassembly of microtubule and neuronal degeneration. The upregulated activity of MARK causes the early stage of neurodegenerative disease (Augustinack,

Schneider, Mandelkow, & Hyman, 2002). Moreover, MARK is overexpressed during AD

(Gerard Drewes et al., 1997) and has been shown to colocalize with NFTs (Chin et al.,

2000).

A general strategy used to inhibit MARK proteins and other kinases is blocking the ATP-binding site of the kinase with small molecule inhibitors (Y. Liu & Gray, 2006).

Since small molecule kinase inhibitors are typically non-specific and result in adverse side effects (Eldar-Finkelman & Eisenstein, 2009), we searched for an alternative strategy that overcomes these disadvantages. Our approach is the use of a peptide-based

MARK2 inhibitor. To achieve our goal, we designed a library of peptides that mimic the repeat domains of tau which are known to be phosphorylated by MARK2 and other major enzymes such as GSK-3β and Cdk5 during AD (Cruz et al., 2003; G Drewes, 2004;

Takashima et al., 1998). We anticipated that these peptides would specifically inhibit

MARK2 activity, and could be developed as a peptide-based treatment for AD or as a chemical genetic agent to better understand the intricate nature of the MARK:tau interaction.

Our in vitro studies revealed that tR1 peptides reduce the phosphorylation level of tau at Ser262 in a dose-dependent manner. The inhibitory effect of tR1 peptides to

MARK2 activity was also demonstrated in primary cortical neurons treated with phenylarsine oxide (PAO) to elevate the activity of MARK2. Our studies also indicated that tR1 peptides are taken up by cultured rat primary cortical neurons through an energy- 49 dependent mechanism. Inside the cell, tR1 peptides may become trapped in endosomes and can be delivered into the cytosol when co-treated with an endosomal disruptor such as bafilomycin A1 or chloroquine. Inhibition of MARK2 is observed once tR1 peptides are released into the cytoplasm. Moreover, we found that the inhibitory effect of the tR1 peptide was selective for inhibiting tau phosphorylation at MARK2 phosphorylation site

(Ser262) and not other phosphorylation sites. For example, tau phosphorylation at

Thr231, which is modified by the kinase GKS-3β, was not inhibited by tR1. Taken together, these results established tR1 as a novel peptide-based kinase inhibitor for

MARK2 activity. We anticipate that tR1 peptides will facilitate the development of peptide-based therapeutics to treat AD and other neurodegenerative diseases in which

MARK proteins are implicated. Furthermore, tR1 may be developed as a chemical genetic agent that could be used as a biological tool to study the interaction between

MARK2 and tau proteins.

Materials

All Fmoc-protected amino acids, Fmoc-PAL-AM resin and coupling reagents were obtained from Novabiochem (Billerica, MA). N,N-diisopropylethylamine (DIEA),

N-methyl-2-pyrrolidone (NMP), piperidine, isopropyl β-D-1-thiogalactopyranoside

(IPTG), ammonium persulfate, sodium azide, Tween-20, benzamidine HCl, sodium phosphate (monobasic) monohydrate, 3-[(3-cholamidopropyl)-dimethylammonio]-1- propanesulfonate hydrate (CHAPS), bovine serum albumin, adenosine-5'-triphosphate disodium salt hydrate (ATP), 2-mercaptoethanol, phenylmethanesulfonyl fluoride

(PMSF), triisopropylsilane (TIPS), N-methylmorpholine (NMM), phenol red, 50 bafilomycin A1 (Baf), chloroquine (CLQ), acetic anhydride, trichloroacetic acid, Triton

X-100, ethylene glycol-bis (2-aminoethylether)-N,N,N′,N′-tetraacetic acid (EGTA), trypsin, and rabbit anti-τ (tau) polyclonal antibody (#T6402) were purchased from

Sigma-Aldrich (St. Louis, MO). Glycine and chloramphenicol were obtained from EMD

Millipore (Billerica, MA). LB Medium, n-Butanol, Coomassie brilliant blue R-250 were obtained from MP Biomedicals (Santa Ana, CA). Tris (base), bisacrylamide, sodium dodecyl sulfate (SDS), tetramethylethylenediamine (TEMED), methanol, phenol, imidazole, LB agar, sodium chloride, ampicillin, RPMI medium 1640, bacterial protein extraction reagent (B-PER) and protease inhibitors were purchased from Thermo Fisher

Scientific (Waltham, MA). Human AB serum (35-060-CI) was purchased from Corning

Inc. (Corning, NY). Potassium chloride, potassium phosphate, calcium chloride, 2- propanol, dichloromethane (DCM) and magnesium chloride were obtained from Fisher

Scientific (Fair Lawn, NJ). Neurobasal (NB) medium and B27 supplement were purchased from Invitrogen (Carlsbad, CA). L-glutamine and amphotericin B were purchased from Calbiochem (Darmstadt, Germany). Protein marker was obtained from

New England Biolabs (Ipswitch, MA). Trifluoroacetic acid (TFA) and phenylarsine oxide

(PAO) were purchased from Acros Organics (Morris Plains, NJ). 5-carboxyfluorescein

(5-CF) and dimethyl sulfoxide (DMSO) was obtained from Santa Cruz Biotechnology

(Dallas, TX). Acetonitrile (ACN) was purchased from NeoBits (Sunnyvale, CA). Ni-

NTA agarose resin was purchased from Molecular Cloning Laboratories (San Francisco,

CA). Bromophenol blue was obtained from Eastman (Kingsport, TN). Amersham ECL

Prime western blotting detection reagents were obtained from GE Health Care 51

(Pittsburgh, PA). Protein markers, Coomassie brilliant blue G-250 and PVDF membranes were purchased from Bio-Rad (Hercules, CA). Mouse monoclonal anti-β-actin antibody

(#A5316) was purchased from Sigma-Aldrich. Rabbit polyclonal anti-Tau (pS262) antibody (#ab4856) and goat anti-rabbit HRP-linked IgG (#ab6721) were purchased from

Abcam (Cambridge, MA). Mouse monoclonal anti-Tau (pT231) antibody (#MN1040),

Alexa-488 goat -rabbit secondary antibody (#A11034) and Alexa-546 goat α-mouse secondary antibody (#A11030) were purchased from ThermoFisher. All other reagents were obtained from commercial sources and used without further purification unless stated otherwise.

Methods

Peptide Preparation

Peptide Synthesis

All peptides used in this research were synthesized by Fmoc-based solid phase peptide synthesis (SPPS) using Fmoc-PAL-AM resin (25 µmol scale). The cycle starts by removing Fmoc protecting group before the addition of a new amino acid. The removal of Fmoc protecting group was achieved by treating the resin with 25% piperidine (v/v) in

NMP solution containing 0.1 M HOBt which decreases the formation of aspartimide

(Palasek, Cox, & Collins, 2007) and placing the reaction in a microwave accelerated reaction system (CEM, Matthews, NC) operated by a software program written in house.

After deprotection and washing the resin 5 × with NMP, the coupling reaction performed inside the microwave accelerator by the addition of 5 equivalents of amino acid, 5 equivalents of PyClock and 10 equivalents of DIEA in NMP to the resin. All equivalents 52 were based on PAL-AM resin loading level. Once the coupling of the amino acid was achieved, the resin was washed 5 × with NMP. The cycle of coupling and deprotection was repeated until the entire sequence of the desired peptide was obtained.

Labeling of Synthesized Peptides

One half of the synthesized peptide-resin was kept unlabeled, and the N-terminus of the unlabeled peptide was protected with acetyl protection group (vide infra). The other half of the synthesized peptide-resin was labeled on the N- terminus with 5- carboxyfluorescein (5-CF) using the following method. First, the terminal Fmoc protecting group was removed by treating the resin with 25% piperidine (v/v) in NMP containing 0.1 M HOBt and allowing the deprotection reaction to proceed inside the microwave. After deprotection, a mixture consisting of 3 equivalents of 5-CF, 3 equivalents of HCTU and 7.5 equivalents DIEA in NMP was added to the reaction vessel. All equivalents were based on the loading level of the resin. Labeling was allowed to stir in the dark at room temperature for 24 hours. Following the reaction, the resin- bound peptide was washed with NMP and dichloromethane and was dried under vacuum for 4 to 6 hours to remove residual solvents.

Capping of Synthesized Peptides

The synthesized peptides were capped at the N-terminus with an acetyl protecting group. Peptide acetylation was performed by allowing deprotected peptides to react with a solution of 6% (v/v) acetic anhydride and 6% (v/v) NMM in NMP for 15 minutes at room temperature with shaking. Following that, the resin-bound peptide was washed with

NMP and dichloromethane and dried under vacuum for 4 to 6 hours before cleavage. 53

Cleavage of the Peptides from the Resin

After completing the synthesis, the peptide was globally deprotected and cleaved from the resin by adding a mixture composed of 88% TFA, 5% phenol, 2% triisopropylsilane and 5% H2O (v/v) to the resin-bound peptide. Then, the reaction vessel containing the peptide was incubated in the microwave at 38 ºC for 30 min. Following that, the peptide was precipitated in cold diethyl ether, pelleted by centrifugation and resuspended in 2-3mL of 15% ACN (v/v) in water. This suspension was then frozen under -

80 ºC and lyophilized to dryness.

Purification and Quantification of Peptides

Following lyophilization, the crude peptide powder was re-suspended in milliQ water or 5% ACN (v/v) in water depending on the solubility of the peptide. After that, the peptide was purified across a reversed-phase semi-preparatory C18 column (Grace, 10

µM, 250 x 10 mm) using a ProStar HPLC system (Agilent). The method used was a linear gradient of 15-50% solvent B (0.1% TFA in ACN) over solvent A (0.1% TFA in water) at a flow rate of 4 mL/min. All major peaks were collected and were analyzed by mass spectrometry. The purified peaks that contained the peptide were combined, frozen and lyophilized twice. The peptide was then resuspended in water and stored under 4 °C

(if the peptides contained Cys, they were stored at -20 °C as a powder form to avoid oxidation). The concentration of the pure peptide was determined either by dry weight

(capped) or by using an extinction coefficient for 5-CF of 38,000 M-1cm-1 at 450 nm in water. 54

Characterization of Peptides by Analytical HPLC and Mass Spectrometry

The purity of the peptides was examined by analytical HPLC. Briefly, peptides were dissolved in water at a final concentration of 2.5 M. The dissolved peptides were then loaded onto a reversed-phase analytical C18 column (Grace, 5 m, 50 x 2.1 mm) and eluted within 20 minutes using the method 5-95% solvent B (0.1% TFA in ACN) over solvent A (0.1% TFA in water). Peptide purity was measured by calculating the percentage of the peak area of the peptide in proportion to other peaks in the spectrum eluted between 10 to 40 minutes. All peptide products were purified to > 95% (except

FlutR2 in which its purity level was > 85%) as determined by product peak integration of analytical HPLC chromatograms.

The identity of the peptides was confirmed by directly injecting around 200 µL peptide dissolved in water or in 10% ACN into an electrospray ionization mass spectrometry (ESI-MS) source at a flow rate 10 L/min and observed peaks in the range of 500-2000 m/z. Mass data were visualized using Xcalibur v3.0 (Thermo) and deconvoluted using MagTran v1.0 (Amgen, Thousand Oaks, CA).

Protein Preparation

MARK2 Preparation

pMCSG7 plasmids coding for full-length human MARK2 with an N-terminal

His6 tag were supplied in DH5-α bacterial cells. Once obtained by our lab, the plasmids were extracted from DH5-α cells using a plasmid extraction kit and then transformed into competent Escherichia coli BL21(DE3) cells. Following transformation, cells containing the plasmids were stored as glycerol stocks at -80 °C. 55

For expression, the cells were streaked on LB agar plates supplemented with 100

µg/mL ampicillin and incubated for 18 hours at 37 °C. The next day, a single colony was isolated and inoculated in 5 mL selective LB broth at 37 °C with vigorous shaking at 225 rpm for overnight. Following that, the 5 mL culture was inoculated into 1L selective LB broth and grown at 37 °C with shaking at 225 rpm. Bacterial growth was monitored by measuring the O.D. at 600 nm on a Cary 50 UV/Vis spectrophotometer until reaching an

O.D.600 of 0.6. Then, isopropyl-1-thio-β-D galactopyranoside (IPTG) was added at a final concentration of 1 mM to induce expression of MARK2. The induction was performed at

16 °C with shaking for 18 hours. Following induction, the cultures were centrifuged at

3500 rpm for 20 minutes at 4 °C. The pellet was collected and stored at -80 °C until further use.

In order to purify MARK2, the bacterial pellet obtained from 1 L cultures was gently thawed on ice and dissolved in 10 mL buffer A (50 mM Tris-HCl, 200 mM NaCl,

50 mM imidazole, 5 mM 3-[(3-cholamidopropyl) dimethylammonio] propanesulfonate

(CHAPS), 2 mM benzamidine, 1 mM β-mercaptoethanol (βME), 1 mM phenylmethylsulfonyl fluoride (PMSF), pH 7.5) (Jacek Biernat et al., 2002). Then, sonication was performed on ice 3 × for 10 seconds (10 seconds on, 1 minute off).

Following sonication, centrifugation was carried out 2 × at 17,000 rpm for 30 minutes each. The cleared lysate was then collected, loaded onto a nickel-nitrilotriacetic acid- agarose (Ni-NTA column) and incubated for 1 hour at 4 °C with shaking. Then, the column containing cell lysate was washed 3 × with 10 mL buffer A (50 mM Tris-HCl,

200 mM NaCl, 50 mM imidazole, 5 mM CHAPS, 2 mM benzamidine, 1 mM βME, 1 56 mM PMSF, pH 7.5). MARK2 protein was eluted 5 × with 3 mL buffer B (50 mM Tris-

HCl, 200 mM NaCl, 500 mM imidazole, 5 mM CHAPS, 2 mM benzamidine, 1 mM

βME, 1 mM PMSF, pH 7.5) (Jacek Biernat et al., 2002). A small sample from the washed and eluted fractions was analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS–PAGE). The remaining volumes of the eluted fractions containing the protein were combined and dialyzed 2 × in buffer C (50 mM Tris-HCl, 5 mM MgCl2,

2 mM ethylene glycol-bis (2-aminoethylether)-N,N,N′,N′-tetraacetic acid (EGTA), 0.5 mM dithiothreitol (DTT), 0.5 mM benzamidine; 0.5 mM PMSF, pH 8.0) (Timm et al.,

2011). Purified MARK2 was concentrated, aliquoted, flash frozen and stored under -80

°C until further usage. hTau K18 Preparation

The plasmid pProEx-HTa-Myc-K18 coding for human tau K18 (hTau K18) with an N-terminal His6 tag was a generous gift from Professor Kevin G. Moffat, University of

Warwick, (Coventry, UK). Once received, pProEx-HTa-Myc-K18 plasmids were transformed into Rosetta2 DE3 chemically competent cells and stored as glycerol stocks at -80 °C.

For expression, the cells were streaked on selective LB agar plates supplemented with 100 µg/mL ampicillin and 35 µg/mL chloramphenicol and incubated for 18 hours at

37 °C. Following overnight growth, a single colony was isolated and cultured at 37 °C in

5 mL selective LB broth with shaking at 225 rpm for 18 hours. Then, the 5 mL culture was inoculated into 1L selective LB broth and grown at 37 °C with shaking at 180 rpm.

Bacterial growth was monitored by measuring O.D.600 on a Cary 50 UV 57 spectrophotometer until reaching an O.D.600 of 0.6. Expression of hTau K18 was induced by adding IPTG at a final concentration of 0.5 mM. The induction was performed at 37

°C for 1 hour with shaking (200 rpm). Following induction, the cultures were centrifuged

2 × at 4 °C at 3500 rpm for 20 minutes. The supernatant was removed, and the pellet was immediately resuspended in 10 ml of 50 mM NaH2PO4.H2O buffer (pH 7.5). The resuspended solution was stored at -80 °C until further use.

Prior to purification, the lysate was gently heated at 42 ºC for 10 minutes to thaw.

Then, a protease inhibitor cocktail (Pierce mini tablet EDTA-Free, 1 tablet/ 50 ml lysate

+ 5 ml of 1 x B-PER bacterial protein extraction reagent + 0.25 mg DNAse I) was added to the lysate. The mixture was allowed to incubate at room temperature for 1 hour.

Following incubation, the suspension was sonicated at 70% power for 1 minute and centrifuged at 4 ºC for 45 minutes at 17,000 rpm. The supernatant was filtered through a cold 0.22 µm sterile filter tube on ice (Millipore, Darmstadt, Germany). Following that, purification was performed by loading the supernatant (clear lysate) onto a Ni-NTA column that was previously washed with milliQ water and with buffer A (50 mM

NaH2PO4.H2O, 500 mM NaCl, 10 mM imidazole, pH 7.0). The clear lysate was allowed to incubate on the column for 1 hour at 4 °C with shaking. Next, the crude extract was collected by gravity filtration. The column was re-washed with buffer B (50 mM

NaH2PO4.H2O, 500 mM NaCl, 25 mM imidazole, pH 7.0) and hTau K18 was eluted with buffer C (50 mM NaH2PO4.H2O, 500 mM NaCl, 500 mM imidazole, pH 7.0). Then, the protein was dialyzed two times against the dialysis buffer (50 mM Tris-HCl, 100 mM

NaCl, pH 7.5). Washing and elution fractions collected during hTau K18 purification 58 were separated using 15% SDS-PAGE (Karikari et al., 2017). Purified His6-hTau K18 proteins were concentrated, aliquoted, flash frozen and stored under -80 °C until further usage.

Structural Analysis for MARK2 and hTau K18 by Circular Dichroism

The solution-phase structure of His6-MARK2 was analyzed by wavelength- dependent CD spectropolarimetry (Jasco J-715 CD spectropolarimeter). To perform this analysis, His6-MARK2 was diluted to a final concentration of 5 µM in buffer C (50 mM

Tris-HCl, 5 mM MgCl2, 2 mM EGTA, 0.5 mM DTT, 0.5 mM benzamidine, 0.5 mM

PMSF, pH 8.0). CD spectrum was recorded from 245 nm to 195 nm at 25 °C. Four scans were averaged and corrected by subtracting the background scan (buffer only). In order to assess the thermal stability of MARK2 in aqueous solution, His6-MARK2 was diluted to a final concentration of 5 µM in buffer C and the temperature-dependent CD signal was monitored at 220 nm from 5 to 95 °C. Melting temperature (Tm) was calculated by taking the first derivative of the melting curve.

The solution-phase structure of recombinant hTau K18 was also analyzed by wavelength-dependent circular dichroism (CD) spectropolarimetry (Jasco J-715 CD spectropolarimeter). To perform this analysis, hTau K18 was diluted to a final concentration of 10 µM in a phosphate buffer (10 mM NaH2PO4.H2O, pH 7.4). CD spectra were recorded from 190 nm to 280 nm at 25 °C (Karikari et al., 2017). Four scans were averaged and corrected by subtracting the background (buffer only). 59

Evaluating the In Vitro Activity of Recombinant MARK2

The in vitro activity of the purified MARK2 was examined by antibody-based fluorescence polarization assays and by western blot. To test the activity of the recombinant MARK2 by antibody-based fluorescence polarization assays, 60-µl reaction mixture containing 100 µM ATP, 10 µM of FluR1 and 10 nM MARK2 were suspended in phosphorylation buffer (50 mM Tris-HCl, 5 mM MgCl2, 2 mM EGTA, 0.5 mM PMSF,

0.5 mM DTT, 0.5 mM benzamidine, pH 8.0) and incubated at 30 °C for 1 hour.

Following incubation, 3 µg of the primary antibody α-Tau pS262 was added to the reaction to bind phosphorylated R1 peptide. The reaction of the antibody with the phosphorylated R1was allowed to proceed for 1 hour at room temperature. Then, the fluorescence polarization of each solution was measured using a SpectraMax M5e multi- mode plate reader (Molecular Devices, Sunnyvale, CA). The excitation wavelength used was 498 nm, and the emission wavelength was 525 nm.

For western blotting, an in vitro phosphorylation assay was performed in a 30 µl reaction mixture that contained 50 µM ATP, 5 µM of hTau K18, 10 nM MARK2 and varying concentrations of tR1(0-50 µM) in phosphorylation buffer (50 mM Tris, 5 mM

MgCl2, 2 mM EGTA, 0.5 mM PMSF, 0.5 mM DTT, 0.5 mM benzamidine, pH 8.0). The reaction mixture was allowed to incubate for 1 hour at 32 °C. Immediately after incubation, phosphorylation reaction was terminated by the addition of SDS sample buffer containing (84 mM Tris-HCl, 20% glycerol, 4.6% SDS, 10% 2-mercaptoethanol, and 0.004% bromophenol blue, pH 6.8) and then boiling for 3 minutes at 95 °C. The reaction products were then separated by SDS-PAGE on a 15% polyacrylamide gel at 60

200 V. Following separation; the products were transferred from the gel to PVDF membrane for 2 hours at room temperature at 200 mA. After the transfer, the membrane was blocked overnight at 4 °C with 5% (w/v) nonfat dry milk in TBST (Tris-buffered saline + 0.1% Tween 20). The membrane was then washed 3 × for 5 minutes each with

TBST. Following blocking and washing, the membrane was incubated for 2 hours at room temperature with the primary antibody α-Tau pS262 diluted in the ratio (1:1000) in a solution made of 5% bovine serum albumin (w/v), 0.2% NaN3 (w/v) and phenol red in

TBST buffer (pH 7.4). Unbound antibodies were washed 3 × 5 minutes each with TBST.

Then, the membrane was incubated with the secondary antibody [1:5000 dilution Goat polyclonal α-rabbit IgG H&L (HRP)] suspended in a solution made of 5% (w/v) non-fat dry milk in TBST for 2 hours at room temperature. The membrane was washed 3 × for 5 minutes each with TBST before signal development using enhanced chemiluminescence

(ECL). Images were acquired by Bio-Rad Gel Doc EZ imager (Bio-Rad), and the bands were analyzed using Image Lab Software v5.2.1 (Bio-Rad).

In Vitro tR1 Stability Assessment

The stability of tR1 peptide was tested in human serum or neuronal basal media

(NB) using a method described previously (Laszlo Otvos, 2008). Briefly, 2 µL of tR1 peptide stock solution (5 mg/mL in DMSO) was added to 198 µL of preheated NB or

RPMI supplemented with 25% (v/v) heat-inactivated human serum. Each degradation reaction was allowed to incubate at 37 °C for either 0, 1, 4 or 24 hours. Following incubation, the degradation was terminated by the addition of 400 µL 15% (w/v) trichloroacetic acid (TCA). The samples were cooled at 4 ºC and then centrifuged 3 × at 61

14,000 rpm. The supernatants were collected and analyzed across an analytical reversed- phase C18 column (Grace, 5 µm, 50 x 2.1 mm) using the following method: 30-minute linear gradient of 0-50% solvent B (0.1% TFA in ACN) over solvent A (0.1 TFA in water), at 1 mL/min flow rate. To quantify the fraction of undigested tR1 peptide using

HPLC, we subtracted background signals produced by reaction media. Then, we integrated the area of the product peak (peak of tR1 peptide subjected to the digestion but still undegraded) and normalized it to the undigested control (tR1 peptide that was not subjected to degradation).

Trypsin digestion of tR1 peptide was used as a positive control to show how the

HPLC spectra of digested tR1 peptide resolved. To get the spectrum for this control, 1.0

g/L tR1 peptides were added to 100 L digestion buffer containing 10 ng/L trypsin,

100 mM Tris-HCl and 1 mM CaCl2 (pH 7.8). The reaction was incubated for 15 minutes at 37 °C before the addition of 100 L of (50%, v/v) aqueous trifluoroacetic acid (TFA) to terminate the digestion. After terminating the digestion using TFA, digestion products were treated as described above. Autolytic fragments of trypsin were ruled out by running an HPLC for trypsin alone in the buffer and we observed no autolytic peaks.

Preparation of Rat Primary Cortical Neuron Culture

The process of dissecting brain of embryonic (E18) Sprague-Dawley rats

(Envigo) is described elsewhere (Colvin, Lai, Holmes, & Lee, 2015). Cultivation of dissociated neurons was achieved by plating the neurons on coverslips coated with 0.05% polyethylenimine (poly-E) (50% solution) in borate buffer (pH 8.2) for immunofluorescence or on poly-E coated plates for MTT assay or western blot. At 37 °C 62 and under a 5% CO2 humidified atmosphere, dissociated neurons were allowed to attached to 0.5 mL minimum essential medium (MEM, Corning Mediatech) containing

26 mM sodium bicarbonate, 55 mM glucose, 50 μg/mL gentamycin (Amresco), 20 mM

KCl, 1 mM pyruvic acid (MP Biomedicals), 2 mM L-glutamine and 10% (v/v) heat- inactivated fetal bovine serum (Atlas Biologicals). Then, the culture medium was replaced with neurobasal (NB) media supplemented with 0.5 mM L-glutamine, 2.8

μg/mL amphotericin B and 2% (v/v) B27.

Cell Uptake of tR1 Peptide by Primary Cortical Neurons

Primary cortical neurons from fetal rats were cultured as described previously.

Then, cultured neurons were grown for 5 to 7 days in vitro (DIV) to ensure maturation of neuronal processes. Before starting cell uptake study, the culture medium was replaced with NB media not supplemented with B27, and neuronal cells were incubated in this fresh NB media overnight. These neurons were then incubated for 4 hours at 37 °C under

Flu a 5% CO2 humidified atmosphere with 10 µM tR1 peptides (final concentration). To investigate the cellular uptake mechanism of FlutR1, 10 mM of the metabolic inhibitor sodium azide (NaN3) (H. Kobayashi, Maeda, & Anraku, 1977; Noumi, Maeda, & Futai,

1987) was added to the neurons and allowed to incubate for 30 minutes. Next, FlutR1 at a final concentration of 10 µM was added and incubated in the pretreated neurons for 4

Flu hours at 37 °C under a 5% CO2 humidified atmosphere. To facilitate uptake, tR1 peptides were allowed to associate with the cell membrane by incubating 10 µM FlutR1 in cultured neurons for 5 minutes. Then, the neurons were incubated with 50 nM bafilomycin A1 (Baf) or 100 µM chloroquine (CLQ) for 4 hours at 37 °C under a 5% 63

CO2 humidified atmosphere to disrupt endosomes. Following incubation, treated neurons were rinsed with warm PBS buffer and then fixed by treatment with 4% (w/v) para- formaldehyde in PBS for 8 minutes at 25 °C. To image the cells, fixed neurons were rinsed with PBS and mounted on glass slides using Anti-Fade/DAPI reagent.

Evaluating the Toxicity of tR1 Peptide Using MTT Assay

Cytotoxicity of tR1 peptide was determined in high-density rat primary cortical neuronal cells grown for 5 to 7 DIV on poly-E coated 24-well plates. Before starting the assay, cell culture media was switched from neurobasal supplemented with B27 to neurobasal media without B27 to remove growth factor from the media which prevents maturation during the assay. Immediately after switching the media, various concentrations of tR1 peptide ranging from 0 to 300 µM were added to the cells. After 72 hours incubation, the media and the peptides were replaced with 50 µg/mL of the tetrazolium MTT reagent and allowed to incubate at 37 °C for one hour. The solution of

MTT reagent was carefully aspirated and replaced with 0.5 mL DMSO to extract formazan. Absorbance was then measured at 550 nm and 650 nm using a UV spectrophotometer (Cary UV 50, Varian). The background absorbance at 650 nm was subtracted from the absorbance of formazan at 550 nm. Following that, corrected absorbances at 550 nm were normalized to the control (0 µM peptide) and plotted as a function of peptide concentration. 64

Immunofluorescence to Study the Ability of tR1 Peptide in Inhibiting MARK2 in Rat

Primary Cortical Neurons

Rat primary cortical neurons were cultured on poly-E coated coverslips in 24-well culture plates for 5 to 7 DIV to ensure maturation of neuronal processes. Prior for phosphorylation assay, culture media was switched 2 × within 30 minutes to NB without

B27 supplement to ensure that growth factors in the culture medium are completely consumed. tR1 peptides at final concentrations of 0, 10 or 30 M were incubated in the cells for 5 minutes before the addition of 50 nM Baf or 100 M CLQ. Baf and CLQ were added to the neurons and allowed to incubate for 4 hours. Following that, 5 M PAO was incubated with the neurons for 45 minutes. Following treatment, the cells were rinsed with warm PBS buffer and then fixed by treatment with 4% (w/v) para-formaldehyde in

PBS for 8 minutes at 25 °C. Cultured neurons were washed 3 × in PBS before incubation with 0.02% Triton-X100 for 1 minute at 25 °C to permeabilize the cells. Next, neurons were incubated for two hours at 25 °C with the primary antibody α-Tau pSer262 diluted

1:1000 or α-pTauT231 diluted 1:1000 in 5% (w/v) non-fat milk in TBS supplemented with protease and phosphatase inhibitors. Neurons were then incubated in the dark for 2 hours at 25 °C with the secondary antibody Alexa-488 goat -rabbit secondary antibody diluted 1:2000 or Alexa-546 goat -mouse secondary antibody diluted 1:2000 in 5% non- fat milk in TBS supplemented with protease and phosphatase inhibitors. Before imaging the cells, coverslips were rinsed with PBS and mounted on glass slides using Anti-

Fade/DAPI reagent. 65

Cell Imaging Using Fluorescence Microscopy

Fluorescence imaging was performed using an inverted epifluorescence microscope (Nikon, Diaphot 300) equipped with a Nikon Plan Apo 60x,1.40 oil- immersion objective lens for immunofluorescence, and a Nikon Plan Apo 100x,1.40 oil- immersion objective lens for cell uptake assays. FITC-HYQ filter sets were used for visualizing Alexa 488-tagged antibodies and FlutR1 peptide; TRITC Dil filter sets were used for visualizing Alexa 546-tagged antibodies; UV-2E/DAPI filter sets were used for

DAPI staining of nuclei. Images were captured using a CCD camera (SPOT imaging) assisted with SPOT software v5.1. For cell selection, the coverslip was first viewed with the UV-2E/DAPI filter to visualize DAPI-staining as a blind sampling method to identify nuclei that appeared healthy.

Quantitative Image Analysis

Fluorescence intensity of fluorescent images was quantified using ImageJ

(Schneider, Rasband, & Eliceiri, 2012). To quantify the fluorescence intensity of the cell body, the entire area of the neuronal cell including cell body, dendrites and axon were highlighted. Then, the mean fluorescent intensity in the cell body area was measured.

Fluorescent intensities for the cell body was quantified with background subtraction. To quantify FlutR1 puncta inside the endosomes using ImageJ particle analysis, the threshold was manually adjusted to highlight the clear puncta. Also, particle size was controlled to

3-infinity pixel3 to prevent interference with fluorescence particles in the background.

The total number of fluorescence particles from each fluorescence image determined by

ImageJ was plotted against each treatment condition. 66

Western Blot to Study the Ability of tR1 Peptide in Inhibiting MARK2-Mediated Tau

Phosphorylation in Rat Primary Cortical Neurons

Rat primary cortical neurons were cultured on poly-E coated coverslips in six- well culture plates for 5 to 7 DIV. Before performing phosphorylation assay, culture media was changed 2 × within 30 minutes to NB that was not supplemented with B27 to ensure that growth factors in the culture medium are completely consumed. tR1 peptides with final concentrations of 0, 10 or 30 M were incubated with the cells for 5 minutes.

Next, cultured neurons were incubated with 50 nM Baf or 100 M CLQ for 4 hours.

After incubation, 5 M PAO was incubated in the neurons for 45 minutes. Then, neurons were rinsed with warm PBS buffer and fixed using 4% (w/v) para-formaldehyde in PBS for 8 minutes at 25 °C. Following fixation, cell lysis buffer containing (TBS, 1% Triton-

X100 (v/v), protease/phosphatase inhibitors and 5 mM EDTA) was used to break down the cell membrane. The cell lysate was treated with the broad-spectrum kinase inhibitor staurosporine (J. N. Hughes, Wong, Lau, Rathjen, & Rathjen, 2014) at a final concentration of 50 nM to prevent any residual kinase activity. The cell lysate was then centrifuged at 10,000 rpm for 10 minutes at 4 ºC, and the clear supernatant was collected.

Total protein concentration in the supernatant was calculated using a Lowry protein assay

(Lowry, Rosebrough, Farr, & Randall, 1951). Protein concentrations were then diluted to

1.2 mg/mL in lysis buffer containing (TBS, 1% Triton-X100 (v/v), protease/phosphatase inhibitors and 5 mM EDTA). Before running SDS-PAGE, 4 x Laemmli sample loading buffer (Bio-Rad) supplemented with 10% 2-mercaptoethanol (v/v) was added to the proteins. These proteins were then denatured by boiling at 95 ºC for 10 minutes. 18 µg 67 proteins were loaded to each lane and the proteins were separated on 12% polyacrylamide gel. Following separation, the proteins were transferred to a nitrocellulose membrane under 200 mA current for 2 hours. These membranes were blotted and developed as described for the in vitro phosphorylation assays (vide supra).

Western Blot Membrane Stripping and Re-blotting

The removal of the primary antibody -Tau pS262 from western blot membranes that were blotted and developed previously was achieved by incubating the membrane in an excess volume of a mild stripping buffer (200 mM glycine, 3.5 mM SDS and 1% (v/v)

Tween-20, pH 2.2) for 10 minutes at 25 °C. Next, the buffer was discarded, and the membrane was washed 2 × with PBS (pH 7.4) buffer for 10 minutes. Then, the membrane was washed 2 × with TBST (pH 7.4) for 5 minutes. Following washing, the stripped membrane was blocked using 5% non-fat milk in TBST overnight at 4 °C. Once the membrane was blocked, it was washed briefly with TBST (pH 7.4) and then blotted with either -Tau (total tau) primary antibody diluted 1:2000 or with anti--actin primary antibody diluted 1:2000. After incubating with the primary antibody, the membrane was washed 3 × with TBST before incubating with the secondary antibody -rabbit HRP- linked IgG diluted 1:5000 or -mouse HRP-linked IgG diluted 1:5000. Next, the membrane was developed using ECL reagent. Images were acquired using a Bio-Rad Gel

Doc EZ imager and the bands were analyzed using ImageJ.

Densitometric Analysis of Western Blot Bands

To quantify the intensity of the bands in western blot membranes using ImageJ, the integrated density of each band is measured. The integrated density of phosphorylated 68 tau band in each lane was normalized to the corresponding density of total tau band.

Percent inhibition was calculated by dividing the intensity of phosphorylated tau bands in each lane from the untreated sample (0 µM tR1). Then, the ratio between the intensities of phospho-tau bands and the intensity of the control was multiplied by 100.

Data Analysis and Statics

The plots and the graphs shown in this work were produced by GraphPad Prism v4.0 graphing software (GraphPad Software) or KaleidaGraph v4.5 (Synergy Software).

Student’s t-test or one-way ANOVA with Tukey’s multiple comparison test was used to analyze data mean. If the probability values were less than 0.05, we conclude that there is a significant difference in the means.

Results and Discussion

Peptide Design

The attempt to produce a peptide-based inhibitor for MARK2 was initiated by designing synthetic peptides that mimic the MTBR of tau. Tau binds to the microtubule through the conserved KXGS motif located in each of the repeat domains of tau (Figure 5 a). MARK proteins phosphorylate tau within the conserved motif KXGS (Figure 5 b).

Phosphorylation of tau at Ser262, Ser293, Ser324 and Ser356 in the respective R1, R2,

R3, and R4 domains disrupts binding of tau to the microtubule which decreases microtubule stability (G. Drewes et al., 1995; Gerard Drewes et al., 1997). Moreover, phosphorylation of Ser262 alone by MARK2 significantly reduces the affinity of tau for microtubules (J. Biernat et al., 1993). 69

All peptides synthesized for this project contain the conserved KXGS motif that is phosphorylated by MARK2 surrounded by 4 to 5 amino acids essential for MARK2 recognition (Figure 5 b and c, Tables 2 and 3). We anticipated that these peptide mimetics would compete with full-length tau for the binding site of MARK2 and would specifically inhibit tau phosphorylation at Ser262, Ser293, Ser324, and Ser356. These peptides were generated by SPPS and characterized by analytical HPLC and mass spectrometry (see Methods section). The purity of all the peptides was above 95% except for FlutR2 (purity level was > 85%) as determined by analytical HPLC chromatograms

(Figures 6 and 7). Masses were determined by ESI-MS and are outlined in Tables 2 and

Figure 5: (a) Schematic representation of tau protein bound to microtubules through its four binding sites (R1-R4). The microtubules on the left are stabilized; however, phosphorylation of tau within the four repeat domains cause tau to detach, which destabilizes the microtubules. (b) Figure showing the four binding sites (R1-R4) of human tau, repeat sequence are underlined, and phosphorylation sites are shown in red. (c) Sequences of R domain peptide mimetics designed to inhibit MARK2. Phospho-Serine residues are shown in red. KXGS motifs are underlined.

Table 2

Calculated and observed masses for fluorescently labeled peptides synthesized in this work. Flu: fluorescently labeled with 5-CF. Peptide Sequence Calculated Mass Observed Mass (+m/z) (+m/z)

Flu tR1 Flu-NVKSKIGSTENLK-NH2 1774.109 1774.88

Flu tR2 Flu-NVQSKCGSKDNIK-NH2 1777.029 1776.85

Flu tR3 Flu-KVTSKCGSLGNIH-NH2 1700.018 1699.76

Flu tR4 Flu-GRVQSKIGSLDNIT-NH2 1844.140 1844.90

Figure 6: Characterization of fluorescently labeled peptides by analytical HPLC. All spectra were acquired at 214 nm, AU: normalized absorbance units.

Table 3

Calculated and observed masses for capped peptides synthesized in this work. Ac: acetylated peptide. Peptide Sequence Calculated Mass Observed Mass (+m/z) (+m/z)

tR1 Ac-NVKSKIGSTENLK-NH2 1459.65 1459.82

tR4 Ac-RVQSKIGSLDNIT-NH2 1472.67 1471.82

Figure 7: Characterization of capped peptides by analytical HPLC. All spectra were acquired at 214 nm, AU: normalized absorbance units.

tR1 Peptides Inhibit MARK2 Activity In Vitro and Reduce Tau Phosphorylation at Ser262

The capability of tau R domain peptide mimetics to inhibit MARK2-mediated tau phosphorylation was initially tested in vitro by using recombinant MARK2 and hTau

K18 (a truncated tau isoform that contains the full repeat domains of tau). We preferred to use hTau K18 instead of full-length tau because of the ease of hTau K18 purification, high yield and enhanced solubility of hTau K18 (Figure 8 a) (Karikari et al., 2017). Both hTau K18 and MARK2, each containing an N-terminal His6 tag, were expressed and purified from bacterial cells.

Following expression and purification, the SDS gel for the recombinant hTau K18 shows a single band at 22 kDa which is the approximate molecular weight of hTauK18

(Figure b). Wavelength-dependent CD spectrum in the far ultraviolet area was recorded for hTau K18 at 25 °C in 10 mM NaH2PO4.H2O (pH 7.4) buffer. The CD spectrum displayed a classical signature of a random coil (unfolded protein) (Barghorn et al., 2000) with a defined minimum at around 200 nm and a shoulder around 220 nm (Figure 8 c). 74

Tau protein is natively unstructured (Schweers et al., 1994), and the CD scan confirmed that the recombinant hTau K18 we prepared is unstructured in aqueous solution.

Figure 8: Purification and characterization of recombinant hTau K18. (a) Schematic representation for hTau K18 used for the in vitro study. Top bar diagram represents the full-length tau. The lower diagram represents hTau K18. (b) SDS-PAGE gel stained with Coomassie blue. An arrow on the right of the gel points to the recombinant tau K18. The mass of hTau K18 is 22 kDa. Protein size in kilodalton (kD) is shown to the left of the gel. M: size marker (protein standards), FT: flow through, W1-3: column washes, E1-5 column elutions. (c) Wavelength-dependent circular dichroism (CD) spectrum of 10 µM hTau K18 in 10 mM NaH2PO4.H2O (pH 7.4) at 25 °C.

Curiously, the SDS-PAGE gel for MARK2 showed more than one band in each lane of the elutions (Figure 9 a). MARK2 is a relatively large protein (molecular weight is 87.9 kDa). Its structure consists of N-terminal domain, catalytic domain, ubiquitin domain, spacer and C-terminal tail (Timm, Marx, Panneerselvam, Mandelkow, & 75

Mandelkow, 2008) (Figure 4). Hence, we speculated that MARK2 may break down by the effect of the reducing condition (βME) and boiling at 95 °C prior to performing SDS-

PAGE. The masses of MARK2 fragments were calculated using an online peptide calculator (BACHEM website). It was found that the masses of the bands correlated well with the calculated masses of different domains of MARK2. To study the solution phase structure of MARK2, wavelength-dependent CD scan in the far-UV region was recorded from 195 nm to 245 nm at 25 °C in the phosphorylation buffer (see Methods section)

(Figure 9 b). CD spectra for MARK2 showed a characteristic of predominantly alpha- helical protein with two minima at λ =220 nm and at 205 nm (Figure 9 b). The study of thermal stability of MARK2 revealed that the recombinant His6-MARK2 was stable in aqueous solution at a temperature up to 61 °C and undergoes a cooperative melting transition around 61 °C (Figure 9 c). Taken together, these data suggest that His6-

MARK2 is folded and has a well-defined secondary structure in aqueous solution.

Figure 9: Purification and characterization of recombinant MARK2. (a) 10% SDS-PAGE gel of separated MARK2 protein fraction stained with Coomassie blue. The mass of MARK2 is 87.9 kDa. Protein size (kD) is shown on the left of the gel. M: size marker, FT: flow through, W1-3: column washes; E1-5 column elutions. Arrows on the right of the gel point to the full-length and truncated structures of MARK2. Bar diagram on the right of the gel represents the structure of full-length MARK2 and truncated structures of MARK2; the size of each truncated structure in (kD) is shown on the top of each bar diagram. The masses of MARK2 fragments were calculated by an online peptide calculator (www.bachem.com). (b) Wavelength-dependent CD spectra of 5 µM MARK2 in phosphorylation buffer (50 mM Tris-HCl; 5 mM MgCl2, 2 mM EGTA, 0.5 mM DTT, 0.5 mM benzamidine, 0.5 mM PMSF, pH 8.0). (c) Thermal stability CD of MARK2 (5 µM in phosphorylation buffer) at 220 nm; melting temperature Tm = 61.1 ºC.

Analyses of the activity of recombinant MARK2 was tested in vitro using antibody-based fluorescence polarization direct binding assay and western blot.

Antibody-based fluorescence polarization assays were carried out by incubating 100 µM

ATP, 10 µM FluR1 and 10 nM MARK2 in a phosphorylation buffer at 30 °C for 1 hour 77 followed by incubation with 3 µg of the primary antibody α-Tau pS262 for 1 hour at 25

°C. Then, fluorescence polarization (FP) was measured by using a SpectraMax M5e multi-mode plate reader. The excitation wavelength used was 498 nm, and the emission wavelength was 525 nm. The principle of fluorescence polarization assay is that when a fluorophore covalently attached to a protein is excited by a plane polarized light at a wavelength suitable for the excitation of the fluorophore, the fluorophore emits light with a degree of polarization inversely proportional to the speed of molecular rotation of the fluorescently labeled protein (Moerke, 2009). This emitted light passes through two polarizing filters: a parallel polarizing filter and perpendicular polarizing filter with respect to the plane of polarized excitation light. Both the parallel and perpendicular lights pass through a final filter that selects the emission wavelength before reaching the detector. The fluorescence polarization (FP) is then calculated by the following equation

(Moerke, 2009):

FP = I|| − I⊥ / I|| + I⊥

Where I|| is the intensity of the emitted light parallel to the excitation plane, and I⊥ is the intensity of the emitted light perpendicular to the excitation plane.

Since small peptides tumble rapidly in solution, the orientation of the fluorophore changes, reducing the strength of the FP signal. However, binding of the peptide to a large biomolecule such as an antibody slows the movement of the peptide resulting in high FP signal. Hence, high FP would indicate that FlutR1 serves as a substrate for

MARK2. Low FP signal would indicate that MARK2 is not active or FlutR1 is not a suitable substrate for this enzyme (Figure 10 a). As shown in Figure 10 b, in the presence 78 of ATP, MARK2 catalyzed the transfer of γ-phosphate from ATP to FlutR1 peptide. This phosphorylation is strongly suggested by the high FP obtained from the binding of the primary antibody α-Tau pS262 to FlutR1. However, low FP was obtained in the absence of MARK2 or in the absence of the active MARK2 or when there is no antibody. We speculated that slightly elevated FP in the absence of antibody may result from binding of the active MARK2 to FlutR1.

To confirm MARK2 activity by western blot analysis, 5 µM of hTau K18, 50 µM

ATP, 10 nM MARK2 were mixed in phosphorylation buffer and incubated for 1 hour at

32 °C. Next, the phosphorylation reaction was terminated by the addition of SDS sample buffer and boiling the sample for 3 minutes at 95 °C. The proteins were then separated by

SDS-PAGE and then transferred to a PVDF membrane. Western blot was performed as described in the methods section and the membrane was blotted with the primary antibody α-Tau pS262. After signal development using ECL reagents, imaging using Bio-

Rad Gel Doc EZ imager and analyzing band intensity using ImageJ, the membrane was stripped and reblotted with an antibody against total tau to confirm equal loading of hTau

K18. The reblotted membrane was then developed, and the band intensity was measured by ImageJ software. Western blot results shown in Figure 11 a and b revealed that ATP is necessary for MARK2 activity (compare Figure 11 b, lanes 1 and 2). Also, phosphorylation of hTau K18 occurs only by MARK2 and not by any other factor such as autophosphorylation by the reaction media (Figure 11 b, lanes 3 and 4). Moreover, tR1 peptide serves as a substrate for MARK2 (Figure 11 a). Collectively, the results obtained from antibody-based fluorescence polarization assay and western blot analysis confirmed 79 that the recombinant MARK2 was active and capable of phosphorylating hTau K18 and tR1 peptides at Ser262 in vitro (Figures 10 and 11).

Figure 10: Antibody-based fluorescence polarization assay presenting the ability of recombinant MARK2 in phosphorylating tR1 peptide in vitro at Ser262. (a) Schematic representation for antibody-based fluorescence polarization assay. The scheme shows peptides attached to a green fluorophore, MARK2 is the enzyme, ATP: adenosine triphosphate, ADP: adenosine diphosphate; phospho group is shown in yellow; the antibody is shown in blue. FlutR1: fluorescently-labeled tR1 peptide. The FlutR1 peptide in solution does not give high FP signal. Once FlutR1 peptides are phosphorylated by MARK2 at Ser262, they bind -Tau pSer262 antibody which leads to an increase in FP signal. (b) Graph showing a comparison of the activity of MARK2 in the presence or absence of MARK2, ATP, 1° Ab, or FluR1. Results were obtained from antibody-based fluorescence polarization assay. Error bars represent SD. Each reaction was performed in triplicate n=3. 80

Figure 11: Recombinant MARK2 is active as it phosphorylates tR1 peptide and the recombinant hTau K18 in vitro at Ser262. (a) Western blot membrane demonstrating that hTau K18 is a substrate for MARK2. Lanes 1 and 2 show that ATP is required for MARK2-mediated phosphorylation of hTau K18, lanes 3 and 4 show that MARK2 is required for phosphorylation of hTau K18. Top membrane is blotted using α-Tau pSer262 antibody; lower membrane was blotted using α-Tau (total-Tau) antibody. Size marker (kD) is shown to the left of the image. (b) Western blot membrane showing that tR1 is a substrate for MARK2. The tR1 peptide used in western blot is acylated on the N-terminus. Gradient concentration of tR1 (0-100 µM) in phosphorylation buffer were incubated with MARK2 in the presence of ATP for 1 h at 32 °C and then separated by SDS-PAGE. The membrane was blotted with the antibody -Tau pSer262.

We next moved forward to examine the ability of R domain peptide mimetics to inhibit MARK2 and reduce the level of hTau K18 phosphorylation in vitro. The in vitro phosphorylation assay was performed by mixing 5 µM of hTau K18, 10 nM MARK2, 50

µM ATP, and 0 to 50 µM tR1 in a phosphorylation buffer and incubating at 32 °C for 1 hour. The reaction was terminated by adding SDS sample buffer and boiling for 3 minutes at 95 °C. Next, the proteins were separated by SDS-PAGE and then western blot 81 was performed. The level of phospho-hTau K18 at Ser262 was quantified by blotting the membrane with a primary antibody specific for phospho-tau Ser262, followed by imaging and densitometric analysis. Equal loading of hTau K18 in each well was confirmed by stripping the membrane and reblotting it with an antibody specific for total tau. To calculate percent inhibition, we compared the phosphorylation level under each treatment condition to the phosphorylation level of the untreated control. Results obtained from the in vitro experiment demonstrated that tR1 peptide is effective in inhibiting hTau K18 phosphorylation at Ser262 in a concentration-dependent manner. The inhibition was around 4% when using 0.05 μM tR1, and it raises up to 66% at 50 μM tR1 treatment concentration (Figure 12 a and b). Moreover, 1 μM tR1 was capable of inhibiting hTauK18 phosphorylation by around 42%. This result is significant because it counters the assumption that the inhibition we observed in hTau K18 phosphorylation is only a consequence of depleting ATP when MARK2 phosphorylates tR1 peptides. Indeed, we still observe inhibition in hTau K18 phosphorylation even though ATP is readily available for MARK2 to phosphorylate hTau K18 as we only use a low concentration of tR1 (1 μM) (Figure 12 a and b). Therefore, the in vitro activity of MARK2-mediated hTau K18 phosphorylation is inhibited by tR1 peptides and not by the lack of ATP in the reaction media.

Figure 12: tR1 peptides inhibit MARK2 in vitro. (a) Western blot assay demonstrating the inhibitory effect of tR1 peptide on MARK2- mediated hTau K18 phosphorylation. The gradient concentration of tR1 used in the assay is written on the top of each lane. The top blot shows phospho-hTau K18 at Ser262 as it was blotted with -Tau pSer262 antibody, lower blot displays total hTau K18 loaded in each lane. This membrane was blotted using -Tau (total-tau) antibody. Protein size (kD) is shown on the left of the membranes. (b) Quantitative analysis of the western blot membranes. The graph presents percent inhibition of MARK2 mediated hTau K18 phosphorylation at Ser262 by tR1 peptide in vitro. Error bars represent standard deviation. Each treatment was performed in three separate experiments n=3. Statistical differences are compared with untreated control (0 µM). Statistical differences for samples above 1 µM calculated by Student’s t-test are significant *, P < 0.05.

Stability of tR1 Peptide in Biological Media

Peptide instability is one of the major issues affecting the potency of peptide- based therapeutics (Mathur et al., 2016). Most short peptides are degraded within 2 to 30 minutes because of the proteolytic activity of enzymes existing in the biological media

(Penchala et al., 2015; Pollaro & Heinis, 2010). To examine the stability of the tR1 peptide under various conditions, we incubated tR1 for up to 24 h at 37 °C in neurobasal

(NB) media or in RPMI media supplemented with 25% (v/v) human serum. Tryptic degradation of tR1 was used as a positive control. After incubation, the digestion reaction was terminated, and digestion products were analyzed using analytical HPLC. The tR1 peptide was degraded as expected within 15 minutes incubation with 10 ng/µL trypsin in the digestion buffer (Zhu et al., 2002) (Figure 13, top spectrum). Trypsin cleaves the carboxyl side of lysine, and since the sequence of tR1 contains three lysines, it was expected to degrade tR1 in the presence of trypsin. However, we observed no degradation for the tR1 peptide in RPMI media supplemented with human serum or in NB media even at incubation times up to 24 hours (Figure 13 b and c). Usually, the peptide will be degraded very fast in human serum (Böttger, Hoffmann, & Knappe, 2017), but tR1 was extraordinarily stable even after one-day incubation. Figure 13 d shows that tR1 peptides incubated for up to 24 hours in NB media or RPMI media supplemented with human serum remain intact by more than 90%. 84

Figure 13: In vitro peptide stability tests suggest that tR1 peptide is stable in various reaction media. (a) Analytical RP-HPLC spectra showing the effect of 10 ng/µL trypsin in a digestion buffer on tR1 peptide. Bottom spectrum displays tR1 peptides at 0 minutes incubation with trypsin. The top spectrum shows tR1 peptide after incubating with trypsin for15 minutes. (b) Analytical RP-HPLC spectra showing the effect of RPMI media supplemented with 25% (v/v) human serum on tR1 peptides. (c) Analytical RP-HPLC spectra showing the effect of neurobasal (NB) media on tR1 peptide. (d) Quantitative analysis of tR1 peptide stability measured from spectra shown in (b) and (c). Circles represent tR1 stability in RPMI media supplemented with human serum; squares in NB media. AU: absorbance units at 214 nm. Incubation time is written to the right of each HPLC spectrum.

tR1 Toxicity and Internalization in Neurons

Rat primary cortical neurons isolated from day 18 (E18) embryo of Sprague-

Dawley rats were maturated for 5 to 7 DIV in culture media. In vitro maturated rat primary cortical neurons were used to perform the in-cell studies due to the high similarity of MARK2 and tau isoforms expressed in rat and in human. MARK2 expressed in human and in rat present 100% similarity in the amino acids 49 to 363 that covers the catalytic domain, CD and UBA domain (Ahrari & Mogharrab, 2014, p. 208). MTBR domain (tau K18) containing interaction sites of MARK2 is located at 244-372 in the sequence of the full-length tau (Mukrasch et al., 2005). This domain is highly conserved in human and in rat with only one single amino acid difference at position 257 (Lys 257 in human tau, Arg 246 in rat tau) (Petry et al., 2014). This single amino acid difference may not prevent tR1 from inhibiting MARK2 in rat primary neurons since this difference is not included in the KXGS recognition motif of tau.

The toxicity of tR1 peptide was assessed by incubating neuronal cells with varying concentrations of tR1 (0–300 μM) for 72 hours, then quantifying cell viability by a colorimetric MTT assay. The result shows no significant toxic effect for tR1 peptide even at high concentrations (300 μM) compared to the untreated control (Figure 14).

Figure 14: tR1 peptide is nontoxic in rat primary cortical neurons up to 300 μM. MTT assay presenting the effect of the tR1 peptide on the vitality of cultured rat primary cortical neurons. Various concentrations of tR1 peptides were incubated in cultured neurons for 72 h. The viability of the cells was then quantified using a colorimetric MTT assay. Cell viability percentage was calculated relative to untreated neurons.

Cellular uptake of the tR1 peptide was examined by incubating neuronal cells cultured for 5 to 7 DIV with 10 μM FlutR1 peptide for 4 hours. After washing, fixation and staining the neurons with DAPI, the intracellular distribution of FlutR1 was visualized using fluorescence microscopy (Figure 15 a). Cell uptake images show no fluorescence when neurons are not incubated with FlutR1 (Figure 15 a, left image). Once these neurons are incubated with FlutR1, distinct fluorescent puncta were observed distributed throughout the cell soma and neurites (Figure 15 a, middle image). Neurons co-treated

Flu with tR1 and the metabolic inhibitor NaN3 (Sato, Nagai, Mitsui, Ryoko Yumoto, &

Takano, 2009) show a considerable reduction in the fluorescent puncta compare to neurons only incubated with FlutR1 (Figure 15 a, right image). These results indicate that

FlutR1 peptides penetrate the cell through an energy-dependent mechanism (perhaps endocytosis) and not by passive diffusion. To evaluate the mechanism of uptake, the 87 endosomal disruptor Baf or CLQ were added to neurons pre-incubated with 10 μM

FlutR1. Baf is a macrolide antibiotic medicine that acts by preventing lysosome acidification (Tolstikov, Cole, Fang, & Pincus, 1997; Yoshimori, Yamamoto, Moriyama,

Futai, & Tashiro, 1991). Upon treatment with 50 nM Baf and 10 μM FlutR1, it was observed that fluorescent puncta disappeared and are replaced with more diffuse fluorescence (Figure 15 b, left image) indicating that the puncta seen where indeed endosomes.

CLQ is a 4-aminoquinoline-based medication used to treat malaria (Sáenz, Mutka,

Udenze, Oduola, & Kyle, 2012; Wellems & Plowe, 2001). It is believed that the antimalarial activity of CLQ is attributed to the ability of this medication to inhibit the polymerization of the free heme molecules produced by degradation of hemoglobin by the malaria parasite (Sullivan, Gluzman, Russell, & Goldberg, 1996).

CLQ passively diffuses through the cell membrane and endosomes because it is neutral at physiological pH. However, the acidic nature of the endosome protonates CLQ which prevents the passive diffusion of CLQ to the outside of the endosome. The accumulation of CLQ inside the endosome causes endosomal rupture (Caron,

Quenneville, & Tremblay, 2004; Madani et al., 2013). Neuronal cells pre-incubated with

10 μM FlutR1 and then with 100 μM CLQ displayed less fluorescent puncta and showed more diffuse, larger vesicular structures as compared to the control sample treated with

FlutR1 alone (Figure 15 b, right image).

The degree of FlutR1 penetration was quantified from fluorescent images of treated neurons using ImageJ software. For each treatment condition, the fluorescence 88 was quantified by displaying three to five fluorescent images of single neurons acquired from different neuronal cultures and counting the number of fluorescent particles

(puncta) inside the neurons. Then, the number of fluorescent puncta were plotted against the corresponding treatment condition (Figure 15 c). This analysis showed a significant reduction in the number of fluorescent puncta for neurons co-incubated with the endosomal disruptor Baf or CLQ compared to neurons treated with FlutR1 alone. We speculate that these fluorescence puncta are vesicles containing FlutR1 peptides trapped inside and cotreating the cells with Baf or CLQ disrupts these vesicles. Once the vesicles containing the FlutR1 peptides are disrupted, the peptides escape and distribute into a larger area. Individual, distributed FlutR1 peptides display low incidence of fluorescence compared to a concentrated group of FlutR1 peptides trapped inside a small vesicle.

Collectively, these results suggest that FlutR1 peptides are internalized through an endocytic pathway. FlutR1 peptides might be trapped inside endosomes and only released into the cytoplasm when co-treated with an endosomal disruptor.

Figure 15: tR1 peptides penetrate neuronal cells. (a) Fluorescently labeled tR1 peptides (FlutR1) are internalized by cultured rat primary cortical neurons. The image on the left displays the fluorescence of neuronal cells not treated with FlutR1 peptide. Cell nuclei are stained with DAPI (blue). Image in the middle displays the fluorescence in cells incubated with 10 μM FlutR1 peptides, arrows point to fluorescent particles accumulated inside the cells; image on the right is a fluorescent image of cells pre-incubated for 30 min with the metabolic inhibitor sodium azide (NaN3) followed by incubation with 10 μM FlutR1. (b) FlutR1 peptides are released into the cytosol of neuronal cells with the aid of endosomal disruptors. The image on the left displays the fluorescence in neuronal cells co-treated with 10 μM FlutR1 and 50 nM bafilomycin A1 (Baf), on the left is an image displaying the fluorescence of cells co- treated with 10 μM FlutR1 peptides and 100 μM chloroquine (CLQ), arrows point to large vesicular structures inside the cell. Scale bar is 10 μm. (c) Quantitative analysis of the mean number of fluorescent particles (puncta) present in each fluorescent image in panels (a) and (b). Each bar represents the mean ± SEM of fluorescent puncta found in 3-5 images of neuronal cells for each treatment condition. Differences shown in graph (c) are compared with neurons treated with FlutR1 alone and calculated using one-way ANOVA and Tukey’s multiple comparison test. The differences are significant *, P < 0.05.

90 tR1 Peptides Inhibit Phosphorylation of Tau at Ser262 in Rat Primary Cortical Neurons

The ability of tR1 peptides to inhibit endogenous tau phosphorylation at Ser262 was tested in cultured rat primary cortical neurons using immunofluorescence assay and western blot. To perform immunofluorescence assay, dissociated neurons were cultured for 5–7 DIV, then maintained in NB media with no B27. Following growth and maturation, the cells were incubated with 0 or 30 μM tR1 for 4 hours. The neurons were then incubated with 5 μM PAO for 45 minutes to activate intracellular kinases.

Specifically, the thiol-reactive reagent PAO increases tau phosphorylation at Ser262 and

Ser356 in R1 and R4 of MTBR respectively (Jenkins & Johnson, 1999). PAO stimulates signaling cascades that lead to the activation of MARK, which in turn phosphorylates endogenous tau at Ser262 and Ser356 (Jenkins & Johnson, 2000). Following incubation with PAO, the neurons were washed with PBS, fixed and the level of phosphorylated

Ser262 was visualized using immunofluorescence microscopy (Figure 16 a, top row).

Quantification of the level of tau phosphorylation at Ser262 was performed by measuring the mean fluorescence intensity from each image using ImageJ software and plotting the mean fluorescence intensity against each corresponding treatment condition (Figure 16 b). The results revealed that neurons treated with only PAO showed high fluorescence intensity inside the neuronal cell which indicates the presence of a high level of phosphorylated tau at Ser262 (Figure 16 a, top row). As it can be seen in Figure 16 a, top row, neurons treated with only 30 µM tR1 peptide did not show inhibition of tau phosphorylation at Ser262 (no reduction in fluorescence intensity compared to neurons treated with PAO alone). Cellular uptake studies (Figure 15) performed in this project 91 suggest that tR1 peptides may become trapped inside endosomes upon internalization.

We, therefore, hypothesized that there is a need to add an endosomal disruptor to release tR1 peptide inside the cytosol and thus inhibit tau phosphorylation at Ser262. To release tR1 into the cytoplasm, we co-incubated neurons with either 0, 10 or 30 μM tR1 and 50 nM Baf (Figure 16 a, middle row) or 100 μM CLQ (Figure 16 a, bottom row). We also incubated neurons with only 50 nM Baf or 100 μM CLQ without tR1 peptide to see if the addition of the endosomal disruptors affects phosphorylation of tau at Ser262. Here, we observed that Baf or CLQ alone did not affect endogenous tau phosphorylation at Ser262

(Figure 16 a and b). Notably, a clear reduction in tau phosphorylation at Ser262 was observed in neurons treated with tR1 and with Baf or CLQ indicating that the tR1 peptide was capable of inhibiting endogenous tau phosphorylation with the aid of an endosomal disruptor (Figure 16 a and b). The level of inhibition depends on tR1 peptide concentration as we observed a more pronounced reduction in tau phosphorylation when the tR1 concentration is increased from 10 to 30 μM. Here, we observed a nearly 60% reduction in phospho-tau when using 30 μM tR1.

For each treatment condition, typical results (Figure 16 a) are shown in the large images. The inset images represent the weakest responding neurons within each population. The broad range of intensities in each treatment condition shown in Figure 16 b may result from including fluorescence intensities of the weakest responding neurons quantified from the small images. Indeed, the slight variation in tR1 inhibitory effect may be attributed to the variation in the uptake of tR1 peptide by cultured neurons and also because of some variation in the ability of tR1 peptide to escape from the endosomes. 92

Figure 16: tR1 peptides inhibit tau phosphorylation at Ser262 in neuronal cells treated with endosomal disruptors. (a) Rat primary cortical neurons were treated with PAO to induce tau phosphorylation at Ser262, followed by treatment with 10 or 30 µM tR1 peptide and the endosomal disruptor Baf or CLQ. Cells were fixed, and immunofluorescence was used to visualize phospho- tau at Ser262. The three top images represent non-treated neurons (image on the left), neurons treated with 5 μM phenylarsine oxide (PAO) alone (middle image), or neurons co-treated with PAO and 30 µM tR1 peptide (image on the right). The three immunofluorescence images in the middle row represent neurons co-treated with 5 µM PAO, 50 nM bafilomycin A1 (Baf) and 0 μM tR1 peptide (image on the left), 10 μM tR1(middle image), or 30 μM tR1(image on the right). The three images on the bottom represent immunofluorescence images for neurons co-treated with 5 µM PAO, 100 µM chloroquine (CLQ) and 0 μM tR1 peptide (left image), 10 μM tR1(middle image), or 30 μM tR1 (right image). Insets show the weakest inhibition of tau phosphorylation at Ser262 by tR1 peptide observed in some rat primary cortical neurons. Scale bar is 10 μm. (b) Quantitative analysis of mean fluorescence intensity of fluorescent images displayed in panel (a). Error bar represents the mean ± SEM of three separate immunofluorescence experiments for each treatment condition. Differences shown in graph (b) are compared with neurons treated with PAO alone and calculated using one-way ANOVA and Tukey’s multiple comparison test; *, P < 0.05; ns: not significant.

The inhibitory effect of tR1 peptide was also investigated in neurons using western blot (Figure 17 a). In order to obtain an appropriate amount of protein to perform western blot, rat primary cortical neurons were cultured at a higher density as compared to cultures used for immunofluorescence. Following initial attachment, culture media was switched to NB media that is not supplemented with B27. Next, 0 or 30 μM tR1 peptide was incubated with the cultured neurons for 5 minutes followed by 4 hours incubation with 100 μM CLQ. After that, 5 μM PAO was added to the neurons and allowed to proceed for an additional 30 minutes. Treated neurons were lysed and the cellular proteins were separated by SDS-PAGE and transferred to a nitrocellulose membrane.

Phospho-tau at Ser262 was detected by blotting the membrane with primary antibody specific for phospho-tau Ser262. Bands detected with a primary antibody specific for total tau or β-actin were used as loading controls (Figure 17 a). Band intensity of phospho-tau Ser262 from at least three separate experiments was quantified and normalized to the intensity of total tau (loading control) located in the same lane. Then, the ratio between integrated band intensity of phospho-tau and total tau was plotted against each treatment condition (Figure 17 b).

Western blot results revealed that untreated cells had a low content of endogenous phospho-tau Ser262 (Figure 17 a, lane 1). On the other hand, cells treated with only PAO contained a high level of phospho-tau Ser262 (Figure 17 a, lane 2). Upon treatment with tR1 and CLQ, we observed a reduction in the level of phospho-tau Ser262 as compared to the level of phospho-tau in cells treated with PAO alone (Figure 17 a, compare lanes 2 and 3). These results are in good agreement with the results obtained previously using 94 immunofluorescence assay (Figures 16 and 17). These results suggest that PAO was effective in elevating the level of phospho-tau Ser262 inside the cells. Treatment with tR1 alone or CLQ alone has no effect on endogenous tau phosphorylation at Ser262 (Figure

17 a, lanes 4 and 5). Only when treating neurons with tR1 peptide and CLQ did we see a reduction in the level of phospho-tau. This suggests that tR1 peptides are trapped in endosomes inside the cells and an endosomal disruptor is necessary to release the peptides into the cytoplasm, which in turn inhibits MARK2 from phosphorylating tau at

Ser262.

Figure 17: Western blot assay demonstrating the ability of tR1 peptide to inhibit tau phosphorylation at Ser262 in cultured primary neurons. (a) Top western blot membrane displays the level of phospho-tau at Ser262 resulting from each treatment condition shown underneath each lane. This membrane was blotted with the primary antibody α-Tau pSer262. The middle and the bottom membranes show loading control blotted with antibody either specific for total tau (middle membrane) or antibody specific for β-actin (below membrane). Size marker is displayed in kDa to the left of the membranes. (b) Quantitative analysis of phospho-tau Ser262 resulted from each treatment condition. Each band of phospho-tau Ser262 in the top membrane was quantified and normalized to the band of total tau in the same lane. Error bars represent the mean ± SEM of three separate experiments for each treatment condition. Differences shown in graph (b) are compared with neurons treated with PAO alone and calculated using one-way ANOVA and Tukey’s multiple comparison test. *, P < 0.05; ns: differences are not significant.

tR1 Peptides Inhibit Tau Phosphorylation at Ser262 and not at Thr231

Next, we wanted to evaluate the specificity of tR1 inhibitory effect by testing the ability for tR1 to inhibit tau phosphorylation at another phosphorylation site: Thr231. tR1 peptide could be considered selective in inhibiting tau phosphorylation at Ser262 only if it does not reduce the level of phosphorylation at other sites within the tau protein.

Thr231 is not a phosphorylation site for MARK2, but many kinases such as protein 96 kinase A (PKA) (J.-Z. Wang, Grundke-Iqbal, & Iqbal, 2007), adenosine monophosphate- activated kinase (AMPK) (Vingtdeux, Davies, Dickson, & Marambaud, 2011), CDK5 and GSK-3β (F. Liu, Iqbal, Grundke-Iqbal, & Gong, 2002) are known to phosphorylate tau at Thr231.

The selectivity of the tR1 peptide was tested in rat primary cortical neurons that were maturated for 5–7 DIV. Cell culture media was then replaced with NB media not supplemented with B27. Neurons were incubated with 30 μM tR1 and 100 μM CLQ as described in the Methods section. After 4 hours of incubation, treated neurons were washed with PBS buffer and fixed. Following that, an immunofluorescence assay was performed to show the level of tau Thr231 phosphorylation (Figure 18 a). To observe staining, cultured neurons were washed with PBS and permeabilized by incubating with

0.02% Triton-X100 for 1 minute at 25 °C. Then, neurons were allowed to incubate for two hours at 25 °C with the primary antibody α-Tau pT231 diluted 1:1000 in 5% (w/v) non-fat milk in TBS supplemented with protease and phosphatase inhibitors. This primary antibody binds to pThr231 located in the proline-rich region of tau (Avila et al.,

2016; Reynolds et al., 2008). Following incubation, neurons were incubated in the dark for 2 hours at 25 °C with the secondary antibody Alexa-546 goat -mouse secondary antibody diluted 1:2000 in 5% non-fat milk in TBS supplemented with protease and phosphatase inhibitors. pTau Thr231 was quantified by measuring the mean fluorescence intensity of the fluorescent images using ImageJ. Then, the mean fluorescence intensity for each image was plotted against the treatment conditions (Figure 18 b). The result of immunofluorescence assay revealed that neurons treated with only tR1 or with only CLQ 97 did not show any difference in phospho-Tau Thr231 compared to the untreated neurons indicating that neither tR1 alone nor CLQ alone affect endogenous tau phosphorylation at

Thr231 (data not shown). Notably, the level of phospho-tau at Thr231 was also not affected by co-treatment with tR1 and CLQ indicating that tR1 does not significantly inhibit tau phosphorylation at Thr231(Figure 18 a and b).

To further validate this result, western blotting was performed using rat primary cultured neurons (Figure 18 c). After maturating dissociated neurons as described, the media was switched to NB media without B27. Then, 30 μM tR1 was added to the media and allowed to incubate for 5 minutes before the addition of 100 μM CLQ. Following 4 hours incubation, neuronal cells were lysed, and the extracted proteins were separated using SDS-PAGE. Western blot analysis was then performed and phospho-tau Thr231 was detected by blotting the membrane with an antibody specific for phospho-tau Thr231 or total tau as a loading control (Figure 18 c). Western blot results shown in Figure 18 c displayed the level of endogenous phospho-tau at Thr231 in the untreated neurons

(Figure 18 c, lane 1). It can be seen from Figure 18 that incubating neurons with tR1 and

CLQ did not influence the level of endogenous phospho-tau at Thr231 compared to untreated neurons (Figure 18 c, lane 2). Collectively, data from immunofluorescence and western blot demonstrated that tR1 peptides do not reduce tau phosphorylation at Thr231, implying that tR1 peptides are at least moderately selective in inhibiting kinases that phosphorylate endogenous tau at Ser262.

Figure 18: tR1 peptide does not inhibit tau phosphorylation at Thr231 in cultured primary neurons. (a) Immunofluorescent images show the level of phospho-tau at Thr231. The two images on the left are non-treated neurons (top image) or neurons co-treated with 30 μM tR1 and 100 μM CLQ (bottom image). (b) Quantitative analysis of mean fluorescence intensity of fluorescent images displayed in panel (a). (c) Western blot assay showing that tR1 peptide has no inhibitory effect on tau phosphorylation at Thr231in cultured neurons. Top western blot membrane displays the level of phospho-tau at Thr231. This membrane was blotted with the primary antibody α-Tau pThr231. The lower image shows loading control blotted with an antibody specific for total tau. Protein size is displayed in kDa to the left of the membranes. (d) Quantitative analysis of phospho-tau Thr231. Each band of phospho-tau Thr231 in the top membrane was quantified and normalized to the band of total tau in the same lane. Error bars in the graphs (b) and (d) represent the mean ± SEM of three separate experiments for each treatment condition. Difference shown in the graphs are compared to untreated neurons and calculated using one-way ANOVA and Tukey’s multiple comparison test; ns: differences are not significant. 99

Conclusion

Throughout this project, we succeeded in synthesizing a stable, cell-permeable peptide-based kinase inhibitor of MARK2. This novel peptide named tR1 is a direct sequence mimetic of the amino acids 255-267 in R1 repeat domain of tau protein. The peptide tR1 was effective in inhibiting phosphorylation of tau at a primary MARK2 phosphorylation site, (Ser 262), within its R1 domain. The ability for tR1to inhibit

MARK2 was observed in vitro and in cultured rat primary cortical neurons. The development of a peptide-based kinase inhibitor tR1 might be beneficial in understanding the complex nature of MARK:tau interaction. For instance, we may be able to use tR1 peptides to study the interaction of MARK with tau and explore various binding sites on

MARK proteins. Furthermore, by using a substrate-based inhibition approach for targeting MARK2, we can in theory, knock out MARK2 activity and study cell signaling cascades of this enzyme in the cell. Moreover, we are interested in using tR1 as a tool to map the specific binding site of tau in MARK proteins which may facilitate understanding the role of particular phosphorylation sites in the MTBR of tau proteins.

Perhaps most importantly, the development of tR1 as a peptide-based inhibitor for

MARK2 may serve as a template to design next-generation medications for the treatment of AD and other neurodegenerative diseases.

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CHAPTER 3: INHIBITING TAU AGGREGATION USING TAU-DERIVED PEPTIDE

MIMETICS

Introduction

Under normal physiological conditions, tau protein is soluble and unstructured.

However, under pathological conditions, tau can misfold and multimerize to form higher- order insoluble aggregates (Y. P. Wang, Biernat, Pickhardt, Mandelkow, & Mandelkow,

2007). The path from functional tau bound to the microtubule to large deposits of NFTs is thought to be a multistep process starting with the formation of dimers, then oligomers, followed by PHFs, and finally NFTs (Alonso et al., 2001; Ballatore et al., 2007).

For many years, it was speculated that intracellular NFTs were the toxic form of tau responsible for neuronal cell death and synaptic damage. However, recent evidence suggests that tauO that consist of 6 to 8 tau monomers (Ganguly et al., 2015; Sahara et al., 2007) and pre-fibrillar intermediates are the most toxic forms (Caughey & Lansbury,

2003; Klein, Krafft, & Finch, 2001). Therefore, efforts have shifted toward targeting tauO for therapeutic effect (Lasagna-Reeves et al., 2012; Yoshiyama, Lee, &

Trojanowski, 2013). There are currently several therapeutic strategies for the treatment of tau aggregation. Increasing the clearance of misfolded tau by activating ubiquitin- proteasome pathway (UPP) or autophagy-lysosomal pathway (ALP) is one of these therapeutic methods. Moreover, passive immunization using antibodies against tauO is being developed as a method to clear extracellular tauO (Asuni et al., 2007). Antibodies such as TOCI and TOMA have been shown to be effective in clearing tauO from outside neuronal cells and indirectly assist clearing of tauO accumulating inside the neurons 101

(Castillo-Carranza et al., 2014; Patterson, Remmers, et al., 2011). Small molecules have been also used to inhibit tau aggregation. Studies have shown that methylene blue

(methylthioninium chloride) is capable of inhibiting tau aggregation by reducing cysteine thiols in tau (Akoury et al., 2013). Moreover, some small molecules, such as curcumin, inhibit tau aggregation by inhibiting π stacking (Gazit, 2002). Other therapies used in the treatment of tau aggregation are inhibiting tau misfolding, disrupting tau dimerization (V.

M.-Y. Lee et al., 2011), or accelerating tau aggregation (T. Liu & Bitan, 2012).

In this study, we focused on developing a novel strategy that involves the use of peptide-based aggregation inhibitors. This approach is advantageous over the other therapeutic approaches because peptides exhibit low cellular toxicity, high specificity, biological potency, and structural diversity. Moreover, peptides can be engineered to cross the cell membrane or BBB (Goyal et al., 2017). Throughout this research, we targeted an essential, early step in tau aggregation: tau dimerization. Indeed, inhibiting tau dimerization could be of vital importance to prevent the formation of the toxic tauO and entirely disrupt the tau aggregation pathway (V. M.-Y. Lee et al., 2011).

To initiate this study, we synthesized three peptides that mimic short segments in the MTBR of full-length tau. Tau dimerization can occur through the formation of disulfide bonds between Cys291 or Cys322 in the R2 and R3 MTBRs (Schweers et al.,

1995). Such tau dimers can aggregate to form a β-sheet structure, which can act as a fundamental nucleation site for the formation of higher order aggregates. Based on this, we designed an anionic peptide mimetic of the R3 domain of tau (designated an-R3) that contains Cys322. Moreover, the sequence of tau protein contains two respective 102 hexapeptide motifs: PHF6* and PHF6 in the R2 and R3 MTBRs respectively that act as nucleating sites for tau aggregation (E.-M. Mandelkow & Mandelkow, 2012). Therefore, we synthesized peptides that mimic these two nucleation sites. We hypothesized that these peptides would be capable of associating with tau and block the nucleation sites, thus preventing further aggregation.

We assessed the ability of the synthetic peptides an-R3, PHF6 and PHF6* to inhibit tau aggregation by using an in vitro heparin-induced tau aggregation assay. The resulting tau aggregates were then quantified by SDS-PAGE analysis and by CD spectropolarimetry. Here, we provide evidence that the isolated PHF6 sequence, located in the R3 repeat domain of tau, are more prone to aggregation than the isolated PHF6* in the R2 domain. Also, we found that binding of PHF6* with PHF6 partially inhibits tau aggregation. In addition, we found that amphiphilic peptide sequences such as our an-R3 peptide or VYK (Goux et al., 2004), in which polar or charged residues alternate with hydrophobic amino acids, are important for tau nucleation and aggregation. These findings are in good agreement with previously published results (Akoury et al., 2013;

Ganguly et al., 2015; Goux et al., 2004).

Materials

All Fmoc-protected amino acids, Fmoc-PAL-AM resin, and coupling reagents were obtained from Novabiochem (Billerica, MA). DNAse I (DN25), N,N- diisopropylethylamine (DIEA), N-methyl-2-pyrrolidone (NMP), piperidine, isopropyl β-

D-1-thiogalactopyranoside (IPTG), ammonium persulfate, benzamidine HCl, bovine serum albumin (BSA), 2-mercaptoethanol, triisopropylsilane (TIPS), N- 103 methylmorpholine (NMM), acetic anhydride were purchased from Sigma-Aldrich (St.

Louis, MO). Chloramphenicol was obtained from EMD Millipore (Billerica, MA).

Heparin sodium salt (porcine average molecular weight of 3000 g/mol), LB Medium and n-Butanol and Coomassie brilliant blue R-250 were obtained from MP Biomedicals

(Santa Ana, CA). Tris (base), sodium acetate, bisacrylamide, sodium dodecyl sulfate

(SDS), tetramethylethylenediamine (TEMED), methanol, phenol, imidazole, LB agar, sodium chloride, ampicillin, bacterial protein extraction reagent (B-PER) and protease inhibitors were purchased from Thermo Fisher Scientific (Waltham, MA).

Dichloromethane (DCM) was obtained from Fisher Scientific (Fair Lawn, NJ). Protein marker was obtained from New England Biolabs (Ipswitch, MA). Trifluoroacetic acid

(TFA) was purchased from Acros Organics (Morris Plains, NJ). Acetonitrile (ACN) was purchased from NeoBits (Sunnyvale, CA). Ni-NTA agarose resin was purchased from

Molecular Cloning Laboratories (San Francisco, CA). Bromophenol blue was obtained from Eastman (Kingsport, TN). 0.22 µm sterile syringe filter was ordered from Millipore

(Darmstadt, Germany).

Methods

Peptide Preparation

Peptide Synthesis

The three peptides: an-R3, PHF6 and PHF6*, were synthesized by SPPS using

Fmoc-PAL-AM resin (50 µmol scale). For the full procedure of SPPS see Chapter 2

Methods section. Briefly, the amino acids were coupled to the Fmoc-PAL-AM resin sequentially until the desired peptide sequence was obtained. Before the addition of a 104 new amino acid, the Fmoc protecting group was removed by treating the resin with 25% piperidine (v/v) in NMP solution containing 0.1 M HOBt to prevent aspartimide formation (Palasek et al., 2007). Deprotection reactions were performed inside a microwave accelerated reaction system (CEM). Following deprotection, the peptide attached to the resin was washed 5 × with NMP and 5 equivalents of the amino acid, 5 equivalents of PyClock and 10 equivalents of DIEA in NMP was added to the resin. All amino acid couplings were achieved inside the microwave. Equivalents were based on

PAL-AM resin loading level. After coupling the amino acid, the resin was washed 5 × with NMP. Each amino acid in the sequence was added following similar cycles of deprotection and coupling until the desired sequence was obtained.

Capping of Synthesized Peptides

The synthesized peptides were capped at the N-terminus with an acetyl protecting group. Peptide acetylation was achieved by allowing Fmoc-deprotected peptides to react with a solution of 6% (v/v) acetic anhydride and 6% (v/v) NMM in NMP for 15 minutes at room temperature with shaking. After that, the resin-bound peptide was washed with

NMP and dichloromethane and dried under vacuum for 4 to 6 hours.

Cleavage of the Peptides from the Resin

After completing the synthesis, the peptides were globally deprotected (only the uncapped peptides) and cleaved from the resin by adding a cleavage cocktail composed of 88% TFA, 5% phenol, 2% triisopropylsilane and 5% H2O (v/v). Then, the reaction vessel containing the resin-bound peptide and the cleavage cocktail was incubated in the microwave at 38 ºC for 30 min. Following that, the peptide was precipitated in cold 105 diethyl ether, pelleted by centrifugation and re-suspended in 2-3 mL of 15% ACN (v/v) in water. This suspension was then frozen at -80ºC and lyophilized to dryness.

Peptide Purification by HPLC

Following synthesis and cleavage, crude peptides were resuspended in water and injected across a reversed-phase semi-preparative C18 HPLC column. The an-R3 peptide was separated using a linear gradient of 15-50% solvent B (0.1% TFA in acetonitrile) over solvent A (0.1% TFA in water) in 35 minutes (1% per minute) at a flow rate 4 mL/min. PHF6 and PHF6* peptides were separated using a linear gradient of 15-45% solvent B (0.1% TFA in ACN) over solvent A (0.1% TFA in water) in 23 minutes (1.3% per minute) at a flow rate 4 mL/min.

Characterization of Peptides by Analytical HPLC and Mass Spectrometry

All major peaks were collected and analyzed by direct-inject ESI mass spectrometry to identify the peptides. The peaks containing the purified peptide were combined, frozen and lyophilized twice. The purity of the peptides was evaluated by analytical reversed-phase HPLC. The peptides were eluted in 20 minutes using a linear gradient of 5-95% solvent B (0.1% TFA in ACN) over solvent A (0.1% TFA in water).

All spectra were monitored at 214 nm. To measure the purity of the peptide, the percentage of the peak area of the peptide was calculated in proportion to other peaks in the spectrum eluted between 10 to 40 minutes. The purity of all the peptides was above

95%. 106

Expression, Purification, and Characterization of Recombinant hTau K18

The plasmid pProEx-HTa-Myc, encoding hTau K18 with an N-terminal His6 tag was a generous gift from Professor Kevin G. Moffat, University of Warwick, (Coventry,

UK). Upon arrival in our lab, the plasmids were immediately transformed into Rosetta2

DE3 competent cells (Novagen, Darmstadt, Germany). The transformants were stored under -80 ºC as glycerol stocks.

To express hTau K18, cells containing the plasmid pProEx-HTa-Myc were streaked on selective LB agar plates supplemented with ampicillin (100 µg/mL) and chloramphenicol (35 µg/mL) and incubated for overnight at 37 °C. Following overnight growth, a single colony was isolated and inoculated into 5 mL selective LB broth at 37 °C with shaking at 225 rpm for 18 hours. The 5 mL culture was then inoculated into 1L selective LB broth and grown at 37 °C with shaking at 180 rpm. Bacterial growth was monitored until reaching an O.D600 of 0.6. Then, IPTG was added at a final concentration of 0.5 mM to induce expression of hTau K18. Following induction, the cultures were centrifuged at 4 °C at 3500 rpm for 20 minutes. Then, the pellet was collected and resuspended in phosphate buffer (50 mM NaH2PO4.H2O pH 7.5). Following suspension, the pellet was stored at -80 °C until further use.

Prior to purification, the lysate was gently thawed at 42 ºC for 10 min. Then, one pellet of protease inhibitor cocktail, 5 ml of 1 x B-PER bacterial protein extraction reagent and 0.25 mg DNAse I was added to the lysate. The mixture was left at room temperature for 1 hour, sonicated at 70% power for 1 minute and centrifuged for 45 minutes at 17,000 rpm 4 ºC. The supernatant was then filtered through a cold 0.22 µm 107 sterile filter tube (Millipore, Darmstadt, Germany). The cleared lysate containing hTau

K18 was added to a Ni-NTA column that was previously washed with milliQ water and with buffer A (50 mM NaH2PO4.H2O, 500 mM NaCl, 10 mM imidazole, pH 7.0). The clear lysate was allowed to incubate in the column for 1 hour at 4 °C with shaking.

Following incubation, the crude extract was collected and the column was re-washed with buffer B (50 mM NaH2PO4.H2O, 500 mM NaCl, 25 mM imidazole, pH 7.0). hTau

K18 protein was eluted with buffer C (50 mM NaH2PO4.H2O, 500 mM NaCl, 500 mM imidazole, pH 7.0) and the protein was dialyzed overnight against the dialysis buffer (50 mM Tris HCl, 100 mM NaCl, pH 7.5) (Karikari et al., 2017). Following dialysis, the protein was concentrated, aliquoted, flash frozen and stored under -80 °C until further use. A complete protocol for hTau K18 extraction, expression, purification, and characterization is described in more detail in Chapter 2 (Methods section).

In Vitro hTau K18 Aggregation Assay

Depending on the detection method, 10-40 µM hTau K18 proteins were incubated at 37 °C for 3 days in an aggregation buffer containing heparin (added at a molar ratio of

1:1 heparin to hTau K18), 100 mM sodium acetate buffer (pH 7.0), and 2 mM dithiothreitol (DTT) (T. J. Cohen et al., 2011). The volume of the aggregation reaction mixture was dependent upon the assay that was being performed. The aggregation reaction was supplemented with 1 mM DTT every 24 hours to prevent the formation of intramolecular disulfide bond, which is shown to inhibit tau aggregation (Akoury et al.,

2013). The effect of the peptides (an-R3, PHF6 or PHF6*) on hTau K18 aggregation was evaluated by adding 40 µM peptide to the aggregation reaction immediately upon mixing. 108

The aggregation of hTau K18 proteins was monitored using either SDS-PAGE or CD spectropolarimetry.

SDS-PAGE Analysis

To detect hTau K18 aggregates by SDS-PAGE, the aggregation assay as described above was performed in a total volume of 30 µL using 10 µM of hTau K18 and the reaction was allowed to incubate for 3 days. Following incubation, the aggregation reaction was separated on 15% SDS-PAGE gel at 200 V for 30 minutes. The gel was then stained with Coomassie brilliant blue R-250 and imaged by using a Gel Doc E-Z imager

(Bio-Rad). Densitometric analysis of each band was acquired using Image Lab Software v5.2.1 (Bio-Rad).

Circular Dichroism (CD) Spectropolarimetry

The solution-phase structure of hTau K18 aggregates was analyzed by wavelength-dependent CD spectropolarimetry using a Jasco J-715 CD spectropolarimeter. To perform this analysis, 12-40 µM hTau K18 in 200 μL aggregation buffer containing equal molar ratio of heparin, 100 mM sodium acetate buffer (pH 7.0), and 2 mM DTT was incubated at 37 °C for 3 days. CD spectra were then recorded from

190 nm to 280 nm at 25 °C. Four scans were averaged and corrected by subtracting the background (sodium acetate buffer only).

Results and Discussion

Peptide Design

At physiological pH, the individual repeat domains of tau proteins are positively charged (+9) (Jeganathan et al., 2008) which is thought to facilitate binding to negatively 109 charged microtubules (Shammas et al., 2015). Tau is a natively unstructured protein. This is primarily because of the high percentage of hydrophilic amino acids in tau sequence which makes it soluble and natively unfolded in aqueous environments (E.-M.

Mandelkow & Mandelkow, 2012; Shammas et al., 2015). However, the unstructured tau proteins can dimerize, and form organized higher order β-sheet-like structures when the positive charges on tau are neutralized. Moreover, the sequence of tau protein contains two nucleation sites known as hexapeptide motifs located in the MTBRs of tau. These two nucleation sites make tau susceptible to aggregation (E.-M. Mandelkow &

Mandelkow, 2012).

It is widely accepted that the initial driving force for tau aggregation is neutralizing the positive charge on tau proteins. The electrostatic repulsion between two tau monomers prevents them from binding to each other. However, anionic cofactors such as heparin or ionized fatty acids, are believed to accelerate tau aggregation by neutralizing the positive charge on tau and eliminating the electrostatic repulsion between tau-tau proteins (Figure 19) (Lippens et al., 2007). In addition, the R2 and R3 domains of tau contain Cys291 and Cys322 respectively that make them capable of forming a disulfide bridge with another tau protein (Schweers et al., 1995). These dimers of tau can then align to form β-sheet-like structures which act as a platform for further aggregation

(Friedhoff et al., 1998; Xu, Brunden, Trojanowski, & Lee, 2010).

110

Figure 19: Structure of polyanions used to accelerate tau aggregation. Top: structure of heparin (sulfated glycosaminoglycans). Below: structure of arachidonic acid.

To prevent tau aggregation in the presence of anionic cofactors such as heparin, we began by designing a peptide that mimics the R3 domain

(317 KVTSKCGSLGNIHHK 331) of full-length hTau. The only difference between the wild type R3 domain (wt R3) and our synthetic peptide is that all positively charged amino acid side chains in R3 repeat domain of tau (K 317, K321, H329, H330, and K331) were replaced with negatively charged amino acids (Figure 20). We hypothesized that anionic R3 (an-R3) with its negative charges would bind and neutralize the positive charges on R3 repeat domain of tau, and potentially form a disulfide bond between 111

Cys322 in the wt R3 and Cys10 of the synthetic an-R3 (Figure 20). We reasoned that binding of an-R3 (peptide sequence: DEEINGLSGCDSTVD-NH2) to the R3 repeat domain of tau would disrupt tau-tau dimerization and thus prevent further tau aggregation

(Nizynski et al., 2017).

Figure 20: Sequence of the an-R3 peptide synthesized for this project and its potential association with the tau R3 domain. The figure represents the electrostatic interaction of tau protein (wtR3, top) with the anionic R3 mimetic (an-R3, bottom) and the formation of a disulfide bridge. Cationic residues in tau protein are colored orange. Anionic resides in an-R3 are blue. Cysteines are colored purple. Amino acid number from N to C terminus are shown on both sides of peptide sequences. a charge at pH 7.0

The second driving force for tau aggregation is the existence of two hexapeptide motifs acting as nucleation sites that facilitate tau aggregation (E.-M. Mandelkow &

Mandelkow, 2012). The full-length tau protein contains two nucleation sites for aggregation. The first site, known as PHF6* (275 VQIINK 280) is located at the N- terminus of the R2 domain. The second nucleation site is PHF6 (306 VQIVYK 311) which is located at the N-terminus of the R3 domain. While the two nucleation sites,

PHF6 and PHF6*, play a crucial role in initiating tau aggregation through the formation of β-sheet structure (Martin von Bergen et al., 2001), only PHF6 is essential for tau 112 aggregation (W. Li & Lee, 2006). Moreover, it was found that the core of PHF6 is the amphiphilic motif VYK, and it is suggested that the interaction between short amphiphilic motifs of tau proteins can induce tau nucleation and aggregation (Goux et al.,

2004). In 2015, Ganguly et al. studied the homodimerization ability between R3 peptides and the heterodimerization between R3-R2 experimentally and by computer simulation.

They found that the binding of R2 to R3 slows down R3 aggregation in the presence of heparin and inhibits R3 aggregation in the absence of heparin (Ganguly et al., 2015).

Notably, Ganguly and co-workers only used short peptide segments containing PHF6* and PHF6 to represent R2 and R3 repeat domain of the full-length tau in their aggregation assay. However, they did not conduct their study in the presence of tau protein. To expand on their studies, we included tau proteins in our experiments and tested the ability of PHF6 and PHF6* to inhibit hTau K18 aggregation. We reasoned that incorporating hTau K18 in this assay will make the in vitro study more physiologically relevant. To perform our study, we synthesized two peptide mimetics for the nucleation sites PHF6 and PHF6* and used them as tools to block the nucleation sites on hTau K18 and prevent the aggregation of hTau K18 (Figure 21).

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Figure 21: Bar diagram representing the location and the sequence of the two nucleation sites PHF6*(red) and PHF6 (green) in the microtubule-binding domain of tau protein. The upper diagram represents the longest isoform of human Tau (hTau) with the projection domain at the N-terminus and the repeat domain at the C-terminus. The middle diagram shows the repeat domains (hTau K18) and the nucleation sites. Bottom sequences show PHF6* and PHF6 and their locations within the tau R domains. Amino acid numbers are shown below each diagram.

Peptide Characterization

Our attempts to prevent hTau K18 aggregation were initiated by designing a synthetic peptide mimetic of the R3 domain, and mimetics of the two nucleation sites of full-length tau. The two peptides that mimic the nucleating sites, PHF6 and PHF6* were acylated at the N-terminus (Table 4). However, an-R3 was left unacylated because we wanted to test the effect of N-terminal modification of the peptide on enhancing or inhibiting hTau K18 aggregation. We also prepared a standard unacylated tau aggregator peptide (Table 4) (Sievers et al., 2011). The peptides were prepared by SPPS and characterized by analytical RP-HPLC and by mass spectrometry (see Methods section).

The purity of all the peptides was found to be greater than 95% (Figure 22). 114

Table 4

Sequences, calculated and observed mass for peptides synthesized in this project. Ac: an acetylated peptide. Peptide Sequence Calculated Mass Observed Mass (+m/z) (+m/z) an-R3 DEEINGLSGCDSTVD-NH2 1551.63 1551.60

Ac-PHF6* Ac-VQIINK-NH2 754.50 755.45

Ac-PHF6 Ac-VQIVYK-NH2 789.46 790.48

Tau aggregator TLKIVW-NH2 757.47 758.60

Figure 22: Characterization of peptides used in hTau K18 aggregation assay by analytical HPLC. All spectra were acquired at 214 nm, AU: normalized absorbance units. 115

Figure 23: Sequence of the peptides synthesized for use in aggregation assay. The single letter designation of the amino acid is written above each corresponding structure in red. Structures generated by ChemSketch. ACD/ChemSketch Freeware, version 2018.12, Advanced Chemistry Development, Inc., Toronto, ON, Canada, www.acdlabs.com, 2019.

Optimizing Conditions for the In Vitro hTau K18 Aggregation Assay

Optimizing Incubation Time at 37 °C

hTau K18 has been used previously in aggregation assays to monitor tau aggregation (T. J. Cohen et al., 2011; Shammas et al., 2015). Using tau K18 which only contains the MTBR domains of tau offers several advantages over the full-length tau because of the ease of purification, high expression level and high aggregation ability 116

(Karikari et al., 2017; Shammas et al., 2015). Incubating tau K18 in an aggregation buffer under 37 °C is a common strategy used to aggregate tau proteins. However, the exact time it takes to form aggregates remains to be determined. To optimize the incubation time, we incubated hTau K18 in the aggregation buffer for different times (0, 3, 7 and 10 days) and monitored the extent of aggregation using SDS-PAGE. SDS-PAGE has been used to indicate the formation of insoluble tau aggregates in the aggregation reaction mixture (Flach et al., 2012). The gradual disappearance of the 22 kDa hTau K18 monomers by increasing incubation time is a qualitative indication of the formation of tau aggregates (Flach et al., 2012). The underline principle of this assay is that the gel only displays hTau K18 monomers and not hTau K18 aggregates as the latter does not enter the gel during electrophoresis and thus cannot be observed following staining (Flach et al., 2012). Even though tau aggregates are not shown in the gel, we still can estimate the amount of tau aggregates based on the amount of protein added to each well. The more hTau K18 monomers present in the gel, the less hTau K18 aggregates are formed and the less hTau K18 monomers present in the gel, the more hTau K18 aggregates are formed.

Our results indicate that 3 days of incubation are sufficient to form a detectable amount of hTau K18 aggregates (Figure 24). During this relatively long period of incubation time, tau monomers pass through three stages (Figure 25). In the first stage known as the lag phase, the monomers of tau start to dimerize and from nuclei essential for protein aggregation. The lag phase appears only in the unseeded aggregation reaction

(Nizynski et al., 2017). However, in the seeded aggregation reaction in which mature tau aggregates are added to recruit the aggregation of tau monomers the lag phase is not 117 observed (Nizynski et al., 2017; Weismiller et al., 2018). The second stage named as the elongation phase, tau dimers start to accumulate to form oligomers and larger protein aggregates. Some of these large aggregates break down to smaller fragments that act as seeds for further aggregation. In the last phase, the aggregation process stops and no further aggregation occur (Figure 25) (Knowles, Vendruscolo, & Dobson, 2014). In a study conducted by Flach et al to monitor the kinetics of heparin-induced tau aggregation, it was found that the amount of tau oligomers (tauO) increased after 6 hours incubation with heparin and reached the highest level at 48 hours. No more increase in tauO was observed after 48 hours incubation most probably because of the depletion of tauO to produce higher order tau aggregates (Flach et al., 2012).

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Figure 24: Optimizing incubation time at 37 °C for the in vitro hTau K18 aggregation assay. (a) 15% SDS-PAGE gel showing the concentration of hTau K18 monomers. 10 µM hTau K18 proteins were incubated at 37 °C for 0, 3, 7 and 10 days with heparin added at a molar ratio of 1:1 heparin: hTau K18 in 100 mM sodium acetate buffer (pH 7.0) containing 2 mM dithiothreitol (DTT). The aggregation reactions were supplemented with 1 mM DTT every 24 hours. The control (0 hour) contains only hTau K18 in sodium acetate buffer with no heparin, DTT or incubation. After the incubation time, the aggregation reaction was separated on 15% SDS-PAGE gel at 200 V. The gel was stained with Coomassie brilliant blue R-250, destained and imaged by Gel Doc E-Z imager (Bio- Rad). (b) Column graph represents the effect of incubation time at 37 °C on the concentration of hTau K18 monomers. Densitometric analysis of each band of hTau K18 monomers was acquired using Image Lab Software v5.2.1 (Bio-Rad).

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Figure 25: Aggregation kinetics of tau proteins. The three stages of tau aggregation are indicated on the sigmoidal curve. Figure adapted from (Nizynski et al., 2017).

DTT Promotes hTau K18 Aggregation In Vitro

To assess the influence of reducing agents on hTau K18 aggregation, we incubated 10 µM hTau K18 in an aggregation reaction at 37 °C for 3 days and supplemented only one aggregation reaction with 1 mM DTT every 24 hours (Figure 26 a, sample 4). Following incubation, we separated the proteins by SDS-PAGE to assess the extent of aggregation in the reactions. Interestingly, hTau K18 aggregation was enhanced by the addition of DTT as compared to the aggregation reaction that was not supplemented with DTT every 24 hours (Figure 26 a and b, the samples 3 and 4). Sugino and co-workers observed the same result when they studied the effect of DTT on tau K18 aggregation. They observed that tau K18 aggregation is significantly enhanced by the presence of DTT (Sugino et al., 2009). DTT is a reducing agent capable of cleaving 120 disulfide bonds in proteins (Figure 27). Because tau K18 contains two cysteine amino acids in R2 and R3 repeat domains, intramolecular disulfide linkages can, therefore, be formed (S. Walker, Ullman, & Stultz, 2012). These results indicate that the oxidized tau

K18 is not the preferred structure for tau K18 aggregation and that the most suitable structure for tau K18 aggregation is the linear, intrinsically disordered structure that contains reduced disulfide bonds.

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Figure 26: In vitro hTau K18 aggregation is enhanced by the presence of DTT. (a) 15% SDS-PAGE gel showing the concentration of hTau K18 monomers. 10 µM hTau K18 proteins were incubated at 37 °C for 3 days with heparin at a molar ratio of 1:1 heparin to hTau K18 in 100 mM sodium acetate buffer (pH 7.0) containing 2 mM dithiothreitol (DTT). Only sample 4 was supplemented with 1 mM DTT every 24 hours. After the incubation, the proteins were separated by SDS-PAGE and stained with Coomassie blue. (b) Column graph showing the effect of DTT on the concentration of hTau K18 monomers. Densitometric analysis of each band of hTau K18 monomer was acquired using Image Lab Software v5.2.1 (Bio-Rad). Each column represents five separate experiments (n=5). Error bars represent standard deviation. p-values are calculated by Student’s t-test. ns not significant. 122

Figure 27: Dithiothreitol (DTT) reduces the disulfide linkages and keeps tau K18 monomer in the unfolded form. Bond dissociation energy of a disulfide bond is 251 kJ/mol (Cremlyn, 1996), DTT concentration is 2 mM (T. J. Cohen et al., 2011). Image adapted from (X. Li, Gao, & Serpe, 2015).

Optimizing hTau K18 Concentration for CD Analysis

The structure of the tau monomer is generally understood to be a random coil in solution whereas the structure of tau aggregates is thought to be primarily β-sheet

(Karikari et al., 2017). The CD spectrum for a random coil peptide (tau monomers) has a minimum in between 195 and 200 nm, and CD for β-sheet structure has a minimum at around 217 nm (Goux et al., 2004). The transition of tau monomers to tau aggregates causes a shift in CD minimum to approximately 217 nm, which is indicative for the formation of β-sheet structure (Figure 28) (Karikari et al., 2017).

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Figure 28: A standard far-UV CD spectra of recombinant hTau K18 monomers and aggregates. These CD spectra are used as standards to show the position of the minimum of hTau K18 monomers and the shift toward 220 nm caused by the formation of hTau K18 aggregates. To obtain the CD spectrum, 12 μM hTau K18 was allowed to aggregate at 37 °C for 7 days in the presence of 6 μM heparin, 10 mM Na2PO4 pH 7.4, 50 mM ammonium acetate and 1 mM DTT. Following incubation, aggregation mixture was centrifuged at 100,000 × g for 1 hour at 4 °C. The supernatant was discarded, and the pellet was resuspended in 200 μl of 10 mM Na2PO4 pH 7.4. Reprinted with permission from Elsevier, 130, Karikari, T. K., Turner, A., Stass, R., Lee, L. C. Y., Wilson, B., Nagel, D. A., Hill., E.j., Moffat, K. G, Expression and purification of tau protein and its frontotemporal dementia variants using a cleavable histidine tag, 44–54, Copyright © 2019.

10 µM Tau K18 was sufficient to study hTau K18 aggregation using SDS-PAGE.

However, the attempt to aggregate 10 µM hTau K18 and detect the aggregates by CD gave a spectrum with low signal to noise ratio (data not shown). This noisy spectrum does not clearly show the transition of hTau K18 monomers to tau aggregates probably because the concentration of hTau K18 aggregates was too low for CD analysis. Indeed, accurate protein concentration is critical when analyzing the solution phase structure of a protein using far-UV circular dichroism (S. R. Martin & Bayley, 2002). Therefore, we tested a series of hTau K18 concentrations ranging from 12 to 40 µM. The result shown in Figure 29 suggests that concentrations higher than 10 µM hTau K18 monomers form 124 sufficient amount of hTau K18 aggregates indicated by the clear shift in minimum from around 195 nm to 210 nm. The shift of the minimum in our spectra did not reach 217 nm

(Goux et al., 2004), possibly because tau aggregates in our samples are mixed with traces of tau monomers. Ultra-centrifugation may allow the separation of tau aggregates from the remaining tau monomers in solution and hence will produce a typical CD spectrum for β-sheet structures of tau aggregates with a minimum at 217 nm. We did not perform the ultra-centrifugation step; therefore, our CD spectra for tau aggregates may be influenced by the presence of tau monomers (Figure 29).

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Figure 29: Far-UV CD spectra of recombinant hTau K18 aggregates in an aggregation buffer. 12-40 µM hTau K18 were aggregated in 100 mM sodium acetate buffer (pH 7.0) containing heparin added at a molar ratio of 1:1 heparin to hTau K18 and 2 mM dithiothreitol (DTT). The mixtures were incubated at 37 °C for 3 days. 1 mM DTT was added to the aggregation reactions every 24 hours. hTau K18 monomers (black curve) contains only hTau K18 in sodium acetate buffer with no heparin, no DTT or incubation. The spectrum for hTau K18 monomers shows a minimum at around 195 nm. This minimum is shifted towards 210 nm when hTau K18 is aggregated. All spectra were collected at 25 °C.

Optimizing Inhibiting Peptide Concentration

To determine the optimal peptide concentration that has the greatest influence on hTau K18 aggregation, we mixed 10 µM hTau K18 in the aggregation buffer with varying concentrations of an-R3 peptide ranging from 0 to 500 µM. Reaction products were analyzed using SDS-PAGE gel and Coomassie blue staining. As it can be seen in

Figure 30, only concentrations of 40 µM an-R3 and higher caused an increase in hTau 126

K18 monomers compared to the untreated sample. The results shown in Figure 30 and

Figure 31 indicate that an-R3 is capable of modesty inhibiting hTau K18 aggregation.

This inhibition was concentration-dependent, ranging from 6% inhibition at treatment concentration of 40 µM an-R3 to 16% at 500 µM an-R3 (Figure 32). The peptides an-R3 contains Glu amino acids that might form salt bridges with Lys of hTau K18 (Zhang,

Holmes, Lockshin, & Rich, 1993). The salt bridge, along with intermolecular disulfide bond, may allow the binding of an-R3 to hTau K18 which block the nucleation site and prevent hTau K18 aggregation. Nevertheless, it seems like an-R3 has propensity to self- assemble ability because it has an amphiphilic sequence (Figure 23 and Table 4). The self-assembly of an-R3 would presumably deplete the free an-R3 peptides and therefore, negate the inhibitory effects of these peptides on hTau K18 aggregation. However, when using excess peptide concentrations (e.g., 40 µM), some of these peptides will be free and available to dimerize with hTau K18 monomers and inhibit hTau K18 aggregation.

Therefore, the concentration 40 µM peptide was used to study the effect of the synthesized peptides, an-R3, PHF6 and PHF6*, on the in vitro hTau K18 aggregation.

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Figure 30: Optimizing peptide concentration for in vitro hTau K18 aggregation assay. The image displays a 15% SDS-PAGE gel showing the presence of hTau K18 monomers. 10 µM of recombinant hTau K18 proteins were mixed with heparin added at a molar ratio of 1:1 heparin to hTau K18 in 100 mM sodium acetate buffer (pH 7.0) containing 2 mM dithiothreitol (DTT). Gradient concentrations of an-R3 peptide (0-500 µM) were added to the aggregation reactions. The mixtures were incubated at 37 °C for 3 days. 1 mM DTT was added to the aggregation reactions every 24 hours. Lane 1 contains only hTau K18 in sodium acetate buffer, no heparin, DTT or peptide were added. Lane 1 was not incubated under 37 °C to prevent the formation of tau aggregates. Lane 4 was treated with 100 µM tau aggregator peptide (Sievers et al., 2011). After the incubation time, the aggregation reaction was separated by SDS-PAGE. The gel was stained with Coomassie brilliant blue R-250, destained and imaged by Gel Doc E-Z imager (Bio-Rad).

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Figure 31: Column graph showing the effect of a gradient concentration of an-R3 peptide on the concentration of hTau K18 aggregates. The content of each sample is indicated underneath each column. Column 1 contains only hTau K18 in sodium acetate buffer with no incubation, no heparin, DTT or peptide were added. Column 4 was treated with 100 µM tau aggregator peptides (Sievers et al., 2011). Densitometric analysis of each band of tau monomers in the above SDS-PAGE gel was acquired using Image Lab Software v5.2.1 (Bio-Rad). Each column represents three separate experiments (n=3). Error bars represent standard deviation. p-values are calculated by student’s t-test. ns not significant.

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Figure 32: Percent inhibition of hTau K18 aggregation by gradient concentrations of an- R3 (0-500 µM) as quantified from SDS-PAGE experiment. Each column represents the average of three separate experiments (n=3). To calculate % inhibition, the band intensity of samples treated with an-R3 was subtracted from the band intensity of the untreated sample (0 µM). The result was divided by the intensity of sample number 1 (see Figure 30 for sample content). Then, the ratio was multiplied by 100 to get percent inhibition.

Effect of Tau-Derived Peptides on hTau K18 Aggregation In Vitro

To evaluate the efficiency of the synthesized peptides an-R3, PHF6 and PHF6* on influencing hTau K18 aggregation, hTau K18 was incubated with 40 µM of an-R3,

PHF6 or PHF6* separately in an aggregation buffer. The resultant aggregation products were then evaluated using SDS-PAGE or CD spectropolarimetry.

The result obtained from SDS-PAGE gel indicated that all three peptides, an-R3,

PHF6 and PHF6* moderately inhibit hTau K18 aggregation as compared to the untreated 130 sample (0 µM) (Figure 33 and 34). As it can be seen in Figure 35, PHF6* inhibited hTau

K18 aggregation the most (18%), followed by PHF6 that cause 11% inhibition and finally an-R3 that caused the least inhibition (4%) (Figure 35).

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Figure 33: Effect of peptides an-R3, PHF6 and PHF6* on in vitro hTau K18 aggregation. 15% SDS-PAGE gel showing the concentration of hTau K18 monomers. 10 µM of recombinant hTau K18 proteins were mixed with heparin at a molar ratio of 1:1 heparin to hTau K18 in 100 mM sodium acetate buffer (pH 7.0) containing 2 mM DTT. 40 µM of an-R3, PHF6 or PHF6* were added to the aggregation reactions. The mixtures were incubated at 37 °C for 3 days. 1 mM DTT was added to the aggregation reactions every 24 hours. Lane 1 contains only hTau K18 in sodium acetate buffer, no incubation, heparin, DTT or peptide was added. Lane 4 contains 100 µM of the peptide tau aggregator (Sievers et al., 2011). After incubation, the aggregation reactions were separated by SDS-PAGE. The gel was stained with Coomassie brilliant blue R-250, destained and imaged by Gel Doc E-Z imager (Bio-Rad).

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Figure 34: A graph representing the effect of an-R3, PHF6 and PHF6* on the concentration of hTau K18 monomers. The content of each sample is indicated underneath each column. Column 1 contains only hTau K18 in sodium acetate buffer with no incubation, no heparin, DTT or peptide added. Column 4 was treated with 100 µM tau aggregator peptide (Sievers et al., 2011). Densitometric analysis of each band of hTau monomer in the SDS-gel shown in Figure 33 was acquired using Image Lab Software v5.2.1 (Bio-Rad). Each column represents three separate experiments (n=3). Error bars represent standard deviation. p-values are calculated by Student’s t-test. ns not significant.

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Figure 35: Percent inhibition of hTau K18 aggregation by 40 µM of an-R3, PHF6 or PHF6* peptides as quantified from SDS-PAGE experiment. Each column represents the average of three separate experiments (n=3). To calculate % inhibition, band intensity of samples treated with an-R3, PHF6 or PHF6* was subtracted from the band intensity of the untreated sample (0 µM). The result was divided by the intensity of sample in lane 1 (see Figure 33 for sample content). Then, the ratio was multiplied by 100 to get percent inhibition.

These results indicate that PHF6, PHF6* and an-R3 bind to hTau K18 monomers and disrupt tau aggregation. This is illustrated by the increase in hTau K18 monomers when incubated with the peptides as compared to the untreated sample (0 µM) (Figures

33, 34 and 35). Yet, the effect of an-R3, PHF6 and PHF6* in inhibiting hTauK18 aggregation might be restricted by the high aggregation tendency of the peptides themselves. In order to evaluate this, CD analysis was carried out to test the self- assembly of isolated an-R3, PHF6 and PHF6* peptides under aggregation conditions and 134 without the presence of hTau K18. CD spectra for 100 µM of isolated PHF6 and isolated

PHF6* show a minimum at around 217 nm which may reveal the ability of these peptides to self-assemble into a β-sheet structure (Figure 36, lower image on the left). However, the CD spectrum of 100 µM isolated an-R3 peptides reveals a random coil structure with a minimum at around 200 nm (Figure 36, the lower image on the left). Although isolated an-R3 may not form β-sheet structures on their own, they still aggregate. β-sheet structure often facilitates protein aggregation, but it is not required because some peptides that form β-sheet structures do not aggregate (Lopez de la Paz et al., 2002). Moreover, some proteins form disordered (amorphous) aggregates without forming β-sheets (Fink, 1998).

Therefore, it might be possible that an-R3 peptides aggregate without the formation of β- sheet structure. The peptide an-R3 is partially soluble in water even though it contains polar residues (Figure 23). Moreover, analytical HPLC spectra for a freshly prepared an-

R3 solution shows a clearly defined peak, but this peak disappeared when incubating an-

R3 under aggregation conditions or even when left under 4 °C for few days in water alone (data not shown). The disappearance of the peak may indicate the formation of larger particles of an-R3 peptides that cannot pass through the small pores of the analytical guard column.

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Figure 36: Far-UV CD spectra for isolated an-R3, PHF6 or PHF6* under standard aggregation conditions. The top image is far-UV CD spectra for the controls hTau K18 monomers, hTau K18 aggregates and also a CD for 100 µM heparin. hTau K18 monomer (black spectrum) contains 15 µM hTau K18 in sodium acetate buffer with no heparin, DTT or incubation. The lower images are far-UV CD for the three peptides. 40 or 100 µM of an-R3, PHF6 or PHF6* were mixed with heparin at a molar ratio of 1:1 heparin to the peptide in 100 mM sodium acetate buffer (pH 7.0) containing 2 mM DTT. The mixtures were incubated at 37 °C for 3 days. 1 mM DTT was added to the aggregation reactions every 24 hours. All spectra were collected at 25 °C.

136

According to previous studies, PHF6 can homodimerize with itself or heterodimerize with PHF6* (Ganguly et al., 2015). Heterodimerization of PHF6* with

PHF6 inhibits tau aggregation in the absence of heparin and slows down tau aggregation in the presence of heparin. The ability of PHF6-PHF6* heterodimerization to only slow down and not inhibit tau aggregation in the presence of heparin is mainly because PHF6 prefers to interact with heparin (Ganguly et al., 2015).

Our result shown in Figure 33 supports Ganguly’s et al. finding in which we observed that binding of PHF6* peptides with PHF6 reduces tau aggregation. Similarly, binding of PHF6 peptide with PHF6* decreases tau aggregation, but with lower efficiency compared to PHF6* alone (Figures 33, 34 and 35, see PHF6). We speculate that the inhibitory effect of PHF6 peptide is less than PHF6* because homodimerization between PHF6 peptides is much stronger and more preferred than homodimerization between PHF6* peptides, (Figures 33, 34 and 35, see PHF6 and PHF6*) and (Figure 36).

As a result, less PHF6 monomers are available to dimerize with hTauK18 to inhibit hTau

K18 aggregation compared to PHF6*. The greater tendency of PHF6 to homodimerize compared PHF6* is attributed to the higher stability of PHF6 dimers (Ganguly et al.,

2015). PHF6 dimers are stabilized by several hydrogen bonds mainly from Q307, V309 and K311 amino acids side chains. As a result of the formation of hydrogen bonds between two PHF6 peptides, the hydrophobic face of each peptide is buried in the middle. PHF6 homodimers are also stabilized by an attractive force between negative and positive charges positioned in the middle of the dimer. Moreover, PHF6 peptides are capable of forming parallel orientation β-sheet structures which accelerate the 137 aggregation of PHF6 peptides. Unlike anti-parallel orientation, parallel β-sheets facilitate the alignment of the β strands to build a steric zipper, an oligomer essential for protein aggregation (Ganguly et al., 2015). On the other hand, PHF6* peptides organize as antiparallel β-sheets, which slows down the aggregation process. Moreover, homodimerization of PHF6* brings the two positively charged amino acids K280 and

K281 into close proximity, and the repulsive forces disrupt the PHF6* dimerization

(Ganguly et al., 2015). For all of these reasons, the self-assembly tendency of PHF6 peptides is stronger and faster compared to PHF6* in the presence of heparin (Figure 36).

As self-assembly of PHF6 is higher than self-assembly of PHF6*, less free PHF6 peptides are available to bind with hTauK18 and inhibit hTau K18 aggregation compared to PHF6*. Therefore, PHF6 peptides are less able to inhibit hTau K18 aggregation compared to PHF6* peptides.

Studying the Effect of Tau-Derived Peptides on hTau K18 Aggregation In Vitro Using

Circular Dichroism (CD) Spectropolarimetry

Far-UV CD spectra for hTau K18 incubated with the three peptides, an-R3, PHF6 or PHF6*, under aggregation conditions for three days displayed a shift in minimum from

195 nm as compared the control hTau K18 monomers. This shift in minimum from 195 nm towards 210 nm may reveal the transformation of hTau K18 monomers to hTau K18 aggregates (Figure 37 upper and lower images). Figure 37 lower image shows that PHF6 cause the greatest increase in hTau K18 aggregation followed by PHF6* and finally an-

R3 as judged by the negative intensity of the minimum (Figure 37 lower image). As mentioned above, both the peptides (an-R3, PHF6 or PHF6*) and hTau K18 aggregate to 138 form β-sheet structures (Figure 36, upper and lower image), and CD analysis displays the combination (overall) structure of the two aggregates without distinguishing between tau or the peptide (Figure 37 lower image). Variation in the ability of PHF6, PHF6* and an-

R3 in inhibiting hTau K18 aggregation observed in the CD result can be explained by the difference in the amount of β-sheet structure produced by each peptide. This variation has made an-R3 peptide the strongest hTau K18 aggregation inhibitor among the other two peptides since an-R3 does not form β-sheet (Figure 37 lower image). Therefore, CD analysis may not be an optimal quantification method to test tau aggregation inhibitory effect of peptides capable of forming β-sheet structure.

139

Figure 37: Far-UV CD spectra showing the effect of an-R3, PHF6 or PHF6* on hTauK18 aggregation. The top image is far-UV CD spectra of the controls: hTau K18 monomer, hTau K18 aggregates and hTau K18 treated with 100 µM tau aggregator peptides. hTau K18 monomer (black spectrum) contains 15 µM hTau K18 in sodium acetate buffer with no heparin, DTT or incubation. Lower image is far-UV CD spectra for 15 µM hTau K18 mixed with heparin at a molar ratio of 1:1 heparin to hTau K18 in 100 mM sodium acetate buffer (pH 7.0) containing 2 mM DTT. 40 µM of an-R3, PHF6 or PHF6* were added to the aggregation mixtures. The mixtures were incubated at 37 °C for 3 days. 1 mM DTT was added to the aggregation reactions every 24 hours. All spectra were collected at 25 °C. 140

Conclusion

The inhibitory effect of tau aggregation of the peptides an-R3, PHF6 and PHF6* was more obvious in SDS-PAGE analysis than by CD. In the SDS-PAGE gel analysis, the disappearance of hTau K18 monomers enabled us to determine the relevant extent of tau aggregation semi-quantitatively. SDS-PAGE analysis revealed that PHF6* caused the greatest inhibition of hTau K18 aggregation followed by PHF6 and finally an-R3

(Figures 33, 34 and 35). However, CD analysis displayed an-R3 as the strongest peptide in inhibiting hTau K18 β-sheet formation then PHF6* and finally PHF6 (Figure 37). The high ability of PHF6 peptide to self-assemble into a β-sheet structure (Figure 36 lower image) (Ganguly et al., 2015) creates aggregates that may influence the CD signal for hTau K18. PHF6* also aggregate to form β-sheet structure, but less than PHF6 and therefore has a lower impact on CD spectrum of hTau K18 (Figures 36 and 37, lower image). CD analysis shows the totality of β-sheet structures without distinguishing whether these β-sheets are from hTau K18 proteins or from the peptide PHF6. Therefore,

CD results show the greatest increase in hTau K18 aggregation in the presence of PHF6 peptide as compared to an-R3 that may not form any β-sheets structure (Figure 37).

Taken together, the results from SDS-PAGE analysis and CD suggest that the three peptides an-R3, PHF6 or PHF6* do not show a significant impact on inhibiting hTau K18 aggregation (Figures 33 and 37). This is likely because of the high self- assembly property of an-R3, PHF6 and PHF6* peptides (Figure 36). However, these peptides may be developed as tau aggregators for clearance by antibodies. Furthermore, using such synthetic peptides as biological tools to study the early event of tau 141 aggregation could illuminate important details about the molecular mechanisms of tau aggregation. Moreover, this research provides decent lead compounds for the discovery of potential therapeutic for neurodegenerative disease. 142

CHAPTER 4: SUMMARY AND FUTURE WORK

This dissertation focused on developing peptide-based inhibitors of tau phosphorylation and aggregation. We succeeded in designing a peptide-based inhibitor for tau phosphorylation at Ser262. Furthermore, our search for a peptide-based inhibitor for tau aggregation enriched our knowledge about the in vitro aggregation of hTau K18.

Future work for inhibiting tau phosphorylation using tR1 peptide will require testing the ability of tR1 to cross BBB in an animal model. If tR1 failed in crossing BBB, then we can overcome this problem by designing a library of very short peptides (maybe two amino acids to make it small and thus capable of penetrating BBB) derived from tR1 peptide sequence and test the ability of these short derivatives in inhibiting tau phosphorylation in vitro, cultured neurons and in vivo. In addition, other proteins could be used to “shuttle” the tR1 peptides across the BBB (Kumagai, Eisenberg, & Pardridge,

1987). Furthermore, mass spectrometry (MS) is a powerful and sensitive tool that could be employed to further investigate the specificity of the tR1 peptide in inhibiting tau phosphorylation (Diane P. Hanger et al., 2007). tR1 specificity was tested in our research using antibodies. Each one of these antibodies can only bind to one specific phospho- epitope which makes it difficult to detect all the inhibited phosphorylation sites.

However, MS sequencing will allow the detection of phosphorylated amino acids on hTau K18 phosphorylated by MARK2 in vitro, and may also show all the inhibited phosphorylation sites caused by the tR1 peptide. We can also try to inhibit newly detected tau phosphorylation sites by designing new inhibitory peptides based on these regions of tau. 143

For the second project focusing on inhibiting hTau K18 aggregation using tau R domain peptide mimetics, future testing the toxicity of an-R3, PHF6 and PHF6* to inhibit tau aggregation in cultured neurons will be necessary. It will be also important to involve a standard control that is known to inhibit tau aggregation when testing the ability of our synthetic peptides in inhibiting tau aggregation. We can, for example, use the all D-amino acid peptide TLKIVW as a standard inhibitor for tau aggregation (Sievers et al., 2011).

Moreover, it will be interesting to examine the ability of an-R3, PHF6 and PHF6* to aggregate tau in the absence of heparin. We can, for example, use an aggregation buffer containing HEPES and KCl to induce tau aggregation without the need for heparin

(Patterson, Remmers, et al., 2011). Also, since hTau K18 aggregation is easier than aggregating the full-length tau, we can try to aggregate hTau K18 by incubating it under

37 °C for long periods (>10 days) in a buffer continuing 100 mM sodium acetate buffer

(pH 7.0) and 2 mM DTT and supplementing the aggregation reaction mixture with 1 mM

DTT every 24 hours. It should be noted that incubation time required to produce a detectable amount of hTau K18 aggregates will vary between different patches of recombinant hTau K18. This is mainly because each patch of hTau K18 contains different amount of hTau aggregates formed during expression and purification process.

It will also be interesting to compare the effect of N-terminal acylated an-R3, PHF6 and

PHF6* on the in vitro tau aggregation with the non-acylated an-R3, PHF6 and PHF6*. In order to improve the detection method of tau aggregation, we can use Fourier transform infrared spectroscopy (FTIR), NMR, MS, transmission electron microscopy or filter trap assays. 144

One of the most important future experiments will be to evaluate the effect of phosphorylation on the rate of tau aggregation. Moreover, when designing a new peptide- based tau aggregation inhibitors, it will be beneficial to modify the structure of the inhibitory peptide to create two faces: 1) unmodified face capable of binding to β-sheet strands and 2) a modified face unable to form hydrogen bonds with the adjacent β-sheet strands so that further aggregation is blocked by the inhibitory peptide. Incorporating N- methylated amino acid in the sequence of the peptide-based tau aggregation inhibitor is an example of such a modification. Also, it will be important to review Zheng’s article which studies the importance of the position of hydrophobic residues when designing a strong inhibitor for tau aggregation (Zheng et al., 2013). Zheng and coworkers mapped the macrocyclic β-sheet peptide 1, an inhibitor for Ac-PHF6 aggregation, for amino acids important for the inhibitory effect. They found that the hydrophobic face of macrocycle 1 is essential for the inhibition of amyloidogenic protein aggregation and that the position of these hydrophobic amino acids is also critical (Zheng et al., 2013).

145

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