Role of the Slingshot-Cofilin and Ranbp9 Pathways in Alzheimer's Disease Pathogenesis Jung a Woo University of South Florida, Vinny7351@Gmail.Com

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10-12-2015 Role of the Slingshot-Cofilin and RanBP9 pathways in Alzheimer's Disease Pathogenesis Jung A Woo University of South Florida, [email protected]

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Role of the Slingshot-Cofilin and RanBP9 Pathways

in Alzheimer’s Disease Pathogenesis

JungA Alexa Woo

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy with a concentration in Neuroscience Department of Molecular Medicine Morsani College of Medicine University of South Florida

Co-Major Professor: David E. Kang, Ph.D. Co-Major Professor: Dave Morgan, Ph.D. Chad A. Dickey, Ph.D. Yu Chen, Ph.D. Kevin Nash, Ph.D.

Date of Approval: Oct 6, 2015

Keywords: Amyloid β, Alzheimer’s Disease, Cofilin, Mitochondrial Dysfunction, Synaptic dysfunction, Slingshot, RanBP9

DEDICATION

박사를 마치면서 석사 연구를 시작하기도 전에 썼던 연구 계획서를 읽어보았다. 그곳에는 포부와

열정에만 차있는 아직은 미성숙한 내가 있었다. 학부과정에 배운 지식이 전부이지만 모든 연구를 수행함이

있어서 가장 중요한것은 끈기와 겸손, 그리고 열정이라고. 그리고 궁극적으로는 사람을 살리는, 사람을

이롭게 하는 학문을 하고 싶다고 말했던 25 살의 나. 앞으로도 더 나은 연구를 통해 사람을 살리는 학문을 할

수 있는 과학자가 될 수 있기를 소망해본다.

이 길을 지나오기까지 무조건적인 지지와 사랑을 보내준 우리 부모님, 사랑하는 가족들, 긴

석.박사기간동안 과학자로써의 역량을 키울 수 있게 도와주신 지도교수님, 마지막으로 지혜와 능력을

허락해주신 하나님께 이 박사학위를 바칩니다.

This dissertation is dedicated to God for giving me strength and wisdom to continue on my research journey. I also dedicated to this thesis to my precious and beloved parents, Un-ha

Woo and Hyang-yeon Jung for their endless love, support, and encouragement. Without your support, love and prayer, I would not have come this far. I love you Mom & Dad!

ACKNOWLEDGMENTS

지금까지 건강과 지혜와 능력을 허락해주신 하나님께 감사드립니다.

박사학위를 마치기까지 감사해야할 사람들이 참 많이 있습니다. 지도교수님, Dr. David Kang

교수님이 없었다면 지금의 연구자로써의 저는 없을 것입니다. 처음 파이펫 한번 제대로 잡아보지 못했던

제가 5 년만에 제 연구에 자부심을 가지고 치열하게 연구하는 과학자로 성장하게 해 주신 것 감사드립니다.

처음에 교수님을 뵜을때 나는 언제쯤 내 스스로 실험가설을 세우고 연구를 진행하고 논문을 쓰고 할 수

있을까 까마득하게만 느껴졌는데 돌이켜보니 어느새 저도 한 사람의 과학자가 되어 있네요. 감사하다는

말로는 표현이 안될 것 같습니다. 앞으로도 배운 지식으로 더 나은 과학자가 되겠습니다. 제 공동 지도교수가

되주신 Dr. Dave Morgan, 학위 심사 위원분들, Dr. Yu Chen, Dr. Chad Dickey, Dr. Kevin Nash, 그리고

마지막으로 Emory 에서 제 박사학위 심사를 위해 먼 걸음 해주신 Dr. Allan Levey 께 모두 감사의 말씀

드립니다.

우리 실험실 가족들, 실험실 동료라고 정의하기엔 너무나 과분한 사람들. 모두 감사합니다. 우리

실험실 가족들이 없었다면 지금의 저, 저의 연구또한 없을 겁니다. Courtney Uhlar, Taylor Boggess, Richard

Witas, Emilio De Narváez, Shazia Majid, Parth Patel, Anusha Bukhari, Kamal Makati, Catherine Fang, Tian Liu,

Xingyu Zhao, Patrick LePochat, Drew Maslar, Josh Vega, 그리고 Evy Panero -모두 제가 동료 이상의

사람들입니다. 다들 너무나 고맙습니다. Courtney Penn (Soon to be Trotter), Hirah Khan- 너희와 같이 연구할

수 있었던 지난 시간들 감사하고 행복한 시간들이었어. 내게 가족만큼 소중한 존재가 되어줘서 감사해.

앞으로 우리 앞에 펼쳐질 과학자로써의 미래가 나는 더욱 기대돼. Justin Trotter, 그리고 Amirthaa

Suntharalingam, Joe Grieco 여기와서 얻게된 소중한 친구들, 앞으로도 좋은 연구로 함께 세상을 더 이롭게

하는 긍정적인 역할을 함께 감당하자! 한국에서 항상 지지와 격려를 아끼지 않고 학위 심사 발표 응원하러

미국까지 날아와 준 내 친구 유리 고마워,

마지막으로 사랑하는 우리 엄마아빠, 엄마아빠의 기도와 지지가 없었다면 여기까지 오지

못했을꺼예요. 멀리 있느라 제대로 효도도 못하는 딸이지만 앞으로도 엄마아빠가 자랑스러워 할 수 있는

딸이 될게요. 엄마아빠가 기도하시는데로 늘 초심을 잃지 않고 인류에 도움이 되고, 세상을 이롭게 하는

과학자가 되겠습니다. 성교 & 지은이 사랑한다. 앞으로도 행복하게 인생의 새 시작을 잘 감당하길 바란다.

사랑하는 우리 외할머니, 건강히 오래오래 사세요! 외숙모 외삼촌, 끊임없는 기도와 격려에 감사하고

사랑합니다.

제가 사랑하는 모든 이들에게 이 논문을 바칩니다.

I have so many people to thank for supporting me during my Ph.D. years. Foremost, I would like to show my sincere gratitude to my advisor Dr. David Kang who has taught me how to be a great scientist. Without your support, patience and immense knowledge, there is absolutely no way I would have been where I am now. Having you as my mentor was the best luck I’ve ever had. You offered me continuous advice and encouragement throughout all these years. It has been such a great journey working with you. I remember I just had some ambiguous cynical ideas about biological research. I was totally unexperienced without knowing what I want to do in my life. With your guidance and support, now I know what I want, where I want to go. (I still remember the day when I got surface LRP western blot data and how much you were

excited about!) Based on knowledge that I was given by you, I will always try to be a better scientist in my career path. Truth to be told, there is NO way I can thank you enough for everything Dr. Kang.

I acknowledge my gratitude to my co-Professor and committee members Dr. Dave

Morgan, Dr. Yu Chen, Dr. Chad Dickey, and Dr. Kevin Nash for the several years of support. I also would like to express my sincere gratitude to Dr. Allan Levey at Emory University for serving as my Ph.D. thesis outside chair.

In my daily life, I have been amazingly blessed with the best group of people with whom

I’ve had to pleasure to work with in my lab. Without these people, this work simply would

NEVER have happened. Courtney Uhlar, Taylor Boggess, Richard Witas, Emilio De Narváez,

Shazia Majid, Parth Patel, Anusha Bukhari, Kamal Makati, Catherine Fang, Tian Liu, Xingyu

Zhao, Patrick LePochat, Drew Maslar, Josh Vega, and Evy Panero-You guys are not just my students or lab folks, but my family and my best friends. It has been such an honor and pleasure to have all of you in my life. I really appreciate all your hard work and I will never forget all those days and nights that we spent together in the lab. Special thanks go to Hirah Khan and

Courtney Penn (Soon to be Courtney Trotter). Thank you for being part of my life. You guys have been the best friends and colleagues anyone could ask for. I loved our times together in lab.

Many thanks and appreciation to my dear friends, Justin Trotter in Dr. Sudhof’s lab: Thank you for being an amazing friend Justin, You inspire me and you give me motivation to become a better scientist. I appreciate our friendship! I miss you, Amirthaa Suntharalingam in Dr. Dickey

Lab: You are like my big little sister. I love you, and Joe Grieco in Dr. Weeber Lab: Thank you for being a great friend and colleague.

My best friend who was willing to come from 6823miles away (From Korea) just for my defense, Yuri Seo, I love you! Thank you for putting up with me for past more than 10 years.

Finally, I thank my beloved family for supporting me throughout all of my studies. Mom and Dad, you showed me unconditional love. You taught me to always be grateful for life. You have always respected and supported my endeavors with an open mind - even when I wanted to go to military school to become an army official in Korea. SeongKyo, my beloved brother, I love you and always miss you. I pray for your next chapter of your life with Jieun Kwon (My lovely future sister-in-law). I am also thankful to my best aunt (Soon Kang) and uncle (Byungdae Jung), thank you for all your prayers and support. I love you. My beloved grandma, I always miss you.

To all of you, thanks for always being there for me. God bless you!

TABLE OF CONTENTS

List of Tables ...... iv

List of Figures ...... v

Abstract ...... vii

Chapter 1 Introduction ...... 1 Alzheimer’s disease (AD) overview ...... 1 Amyloid plaques and neurofibrillary tangles ...... 2 Genetic basis of Alzheimer’s disease (AD) ...... 3 Early Onset AD ...... 3 Late Onset AD ...... 4 Molecular basis of Alzheimer’s disease (AD) ...... 5 Soluble Aβ oligomer hypothesis and hyper phosphorylation of tau ...... 5 Amyloid β and its receptors ...... 7 Neuroinflammation in AD ...... 10 Mitochondrial dysfunction in AD ...... 10 Cytoskeletal aberrations in AD ...... 14 Role of Cofilin in F-Actin dynamics and mitochondrial dysfunction ...... 15 RanBP9 in APP processing, mitochondria-mediated neurotoxicity, and integrin signaling ...... 17

Chapter 2 Slingshot-Cofilin activation medicates Mitochondrial and synaptic dysfunction via Amyloid β ligation to β1 integrin conformers ...... 26 Background ...... 26 Materials and Methods ...... 28 Cells, cDNA constructs, siRNA sequences, and transfection ...... 28 Antibodies and reagents ...... 29 Cell/tissue lysis and Immunoblotting ...... 30 Cell surface biotinylation assay ...... 30 Annexin V/PI apoptosis assay ...... 31 FITC-Aβ oligomer binding assay ...... 31 Purification of β1-integrin-Fc and binding to Aβ oligomers ...... 31 Mice and human samples ...... 32 Immunofluorescence ...... 33 JC-1, Mitosox-Red, and H2DCF imaging ...... 34 Stereotaxic surgery ...... 34 Electrophysiology in mouse brain slices ...... 35 Fear conditioning, open field, and rotarod behavior ...... 36 Statistical analysis & graph ...... 37 Results ...... 37 i

Aβ oligomers induces loss of cell surface β1-integrin but not TfR or PrPc and reduces β1-integrin activation ...... 37 β1-integrin allosteric modulation or activation reduces Aβ oligomers binding to neurons, HT22 cells, and Purified β1-integrin integrin ...... 38 Essential role of β1-integrin in Aβ oligomer-induced cofilin activation and reciprocal requirement of cofilin in Aβ oligomers induced depletion of surface β1-integrin and FCs ...... 39 Essential roles of cofilin and β1-integrin in Aβ oligomer-induced depletion of F-actin and F-actin-associated synaptic proteins in primary hippocampal neurons ...... 41 Requirement of SSH1 in Aβ oligomer-induced cofilin translocation to mitochondria and accumulation of mitochondrial cofilin in AD brains ...... 42

Aβ oligomer-induced ROS production, mitochondrial dysfunction, and apoptosis via the β1-integrin–SSH1–Cofilin activation pathway ...... 43 Cofilin reduction rescues APP/Aβ42 oligomers-induced gliosis and loss of synaptic protein, as well as LTP and contextual memory deficits in APP/PS1 mice ...... 44 Discussion ...... 46 Conclusion ...... 49

Chapter 3 RanBP9 at the intersection between Cofilin and Amyloid β pathologies ...... 66 Background ...... 66 Methods ...... 68 Cells, cDNA constructs, siRNA sequences, and transfection ...... 68 Antibodies, reagents, and Aβ oligomer preparation ...... 69 Cell/tissue lysis and Immunoblotting ...... 70 Mice ...... 70 Immunofluorescence ...... 70 Fear conditioning, open field, and rotarod behavior ...... 71 Electrophysiology in mouse brain slices ...... 72 Results ...... 73 Endogenous RanBP9 mediates Aβ42 oligomer-induced translocation of cofilin to mitochondria and promotes cofilin activation via positively regulating SSH1 ...... 73 Endogenous RanBP9 and SSH1 promote cofilin–actin rod formation and Aβ oligomer-induced collapse of growth cones in primary hippocampal neurons ...... 75 Endogenous RanBP9 mediates the depletion of postsynaptic proteins induced by Aβ oligomers in primary hippocampal neurons ...... 76 Significant increase in endogenous RanBP9 in APP/PS1 mice brains ...... 77 Rescue of neuroinflammation, synaptic damage, and Aβ accumulation by RanBP9 reduction in APP/PS1 mice ...... 77 Accumulation of cofilin–actin rods in APP/PS1 mice and significant attenuation of cofilin–actin rod formation by RanBP9 reduction ...... 78 RanBP9 reduction rescues deficits in synaptic plasticity and contextual memory associated with APP/PS1 mice ...... 79 Discussion ...... 80 Conclusion ...... 84

Chapter 4 Discussion ...... 95 Summary ...... 95 Conclusion ...... 96

References ...... 97

Appendix A IACUC Approval for Animal Research ...... 123

Appendix B Copyright Permissions ...... 124

Appendix C Copyright Permissions ...... 125

iii

LIST OF TABLES

Table 1-1 Genes associated with Alzheimer’s disease ...... 20

TABLE OF FIGURES

Figure 1-1. Amyloid β oligomer receptor ...... 21

Figure 1-2. The actin-integrin linkage ...... 22

Figure 1-3. Mitochondrial damage in Alzheimer’s disease ...... 23

Figure 1-4. Cofilin regulates actin remodeling ...... 24

Figure 1-5. Schematic model of the RanBP9-cofilin pathway in AD pathogenesis ...... 25

Figure 2-1. Aβ42 oligomers induces loss of cell surface β1-integrin but not TfR or PrPc and reduces β1-integrin activation ...... 51

Figure 2-4. Essential roles of Cofilin and β1-integrin in Aβ42 oligomer-induced depletion of F-actin and associated synaptic proteins in primary hippocampal neurons ...... 54

Figure 2-5. Requirement of Slingshot-1 (SSH1) in Aβ42 oligomer-induced Cofilin translocation to mitochondria and accumulation of mitochondrial Cofilin in AD brains...... 55

Figure 2-6. Aβ42 oligomer-induced ROS production, mitochondrial dysfunction, and apoptosis via the β1-integrin–SSH1–Cofilin activation pathway ...... 56

Figure 2-7. Cofilin reduction rescues APP/Aβ-induced gliosis and loss of synaptic proteins as well as LTP and contextual memory deficits in APP/PS1 mice ...... 57

Supplementary Figure 2-1. Aβ42o Induces Loss of Cell Surface β1-Integrin ...... 58

Supplementary Figure 2-2. Aβ42 Oligomers but not Monomers or Fibrils Bind to Purified β1-integrin-Fc ...... 59

Supplementary Figure 2-3. Aβ42o Disrupt Integrin-dependent Focal Complexes in HT22 Cells, Primary Neurons, and APP/PS1 Mouse Brains ...... 60

Supplementary Figure 2-4. Cofilin Mediates Aβ42o-induced Depletion of Focal Vinculin in HT22 Cells ...... 61

Supplementary Figure 2-5. AD and Control Subject Information ...... 62

Supplementary Figure 2-6. Allosteric Modulation of β1-Integrin Rescues Aβ42o-induced Apoptosis in Primary Neurons and HT22 Cells ...... 63

Supplementary Figure 2-7. Aβ42o Reduces 14-3-3/SSH1 Complex and Increases SSH1/Cofilin Complex in HT22 Cells ...... 64

Supplementary Figure 2-8. Reduction in Cofilin Protein in the Hippocampus and Cortex of Cofilin+/- Mice with No Apparent Abnormarmalities in Gliosis or Synaptic Proteins...... 65

Figure 3-1. RanBP9 mediates Aβ oligomer-induced translocation of cofilin to mitochondria and lowers cofilin activation via SSH1...... 85

Figure 3-2. RanBP9 mediates cofilin–actin rod formation and Aβ oligomer-induced collapse of growth cones in primary hippocampal neurons...... 86

Figure 3-3. RanBP9 mediates the depletion of postsynaptic proteins and F-actin induced by Aβ oligomers in mature primary hippocampal neurons...... 87

Figure 3-4. Increased endogenous RanBP9 levels in APP/PS1 mice ...... 88

Figure 3-5. RanBP9 reduction rescues neuroinflammation and synaptic protein depletion in APP/PS1 transgenic mice...... 89

Figure 3-6. Mitigation of Aβ accumulation by RanBP9 reduction in APP/PS1 mice...... 90

Figure 3-7. Accumulation of cofilin–actin rod/aggregate structures in APP/PS1 mice...... 91

Figure 3-8. Rescues of synaptic plasticity and contextual memory impairments in APP/PS1 mice by RanBP9 reduction...... 92

Supplementary Figure 3-1. Short-term Aβ Monomer Exposure Does Not Alter Cofilin Translocation to Mitochondria or Drebrin/F-Actin Levels ...... 93

ABSTRACT

Alzheimer’s disease (AD) is a neurodegenerative disorder characterized by two major pathological hallmarks, amyloid plaques and neurofibrillary tangles. The accumulation of amyloid-β protein (Aβ) is an early event associated with synaptic and mitochondrial damage in

AD. Therefore, molecular pathways underlying the neurotoxicity and generation of Aβ represent promising therapeutic targets for AD. Recent studies have shown that actin severing protein,

Cofilin plays an important role in synaptic remodeling, mitochondrial dysfunction, and AD pathogenesis. However, whether Cofilin is an essential component of AD pathogenesis and how

Aβ induced neurotoxicity impinges its signals to Cofilin are unclear.

In my dissertation studies, we found Aβ oligomers bind with intermediate activation conformers of β1-integrin to induce the loss of surface β1-integrin and activation of Cofilin via

Slingshot homology-1 (SSH1) activation. Specifically, conditional loss of β1-integrin prevented

Aβ induced Cofilin activation, and allosteric modulation or activation of β1-integrin significantly reduced Aβ binding to neurons and mitigated Aβ42-induced reactive oxygen species (ROS) generation, mitochondrial dysfunction, synaptic proteins depletion, and apoptosis. Furthermore, we found that SSH1 reduction, which mitigated Cofilin activation, prevented Aβ-induced mitochondrial Cofilin translocation and apoptosis, while AD brain mitochondria contained significantly increased activated/oxidized Cofilin. In mechanistic support in vivo, we demonstrated that APP transgenic mice brains contain decreased SSH/Cofilin and SSH1/14-3-3 complexes which indicates that SSH-Cofilin activation occurred by releasing of SSH from 14-3-

vii

3. We also showed that genetic reduction in Cofilin rescues APP/Aβ-induced synaptic protein loss and gliosis, as well as impairments in synaptic plasticity and contextual memory in vivo.

Our lab previously found that overexpression of the scaffolding protein RanBP9 increases Aβ production in cell lines and in transgenic mice, while promoting Cofilin activation and mitochondrial dysfunction. However, how endogenous RanBP9 activates cofilin and whether endogenous RanBP9 accelerates Aβ-induced deficits in synaptic plasticity, cofilin- dependent pathology, and cognitive impairments were unknown. In my dissertation studies, we found that endogenous RanBP9 positively regulates SSH1 levels and mediates A-induced translocation of Cofilin to mitochondria. Moreover, we demonstrated that endogenous RanBP9 mediates A-induced formation of Cofilin-actin rods in primary neurons. Endogenous level of

RanBP9 was also required for Aβ-induced collapse of growth cones in immature neurons and depletion of synaptic proteins in mature neurons. In vivo, we also found APP transgenic mice exhibit significantly increased endogenous RanBP9 levels and that genetic reduction in RanBP9 rescued APP/Aβ-induced synaptic protein loss, gliosis, synaptic plasticity impairments, and contextual memory deficits. These findings indicated that endogenous RanBP9 not only promotes Aβ production but also meditate Aβ induced neurotoxicity via positively regulating

SSH1. Taken together, these novel findings implicate essential involvement of β1-integrin–

SSH1/RanBP9–Cofilin pathway in mitochondrial and synaptic dysfunction in AD pathogenesis.

viii

CHAPTER 1

INTRODUCTION

Alzheimer’s disease (AD) overview

Alzheimer’s disease (AD) is a neurodegenerative disorder that is the most common cause of dementia. It is a progressive disease that leads to memory loss, confusion, disorientation, and difficulty having a conversation. Prevalence rates of AD range from ~10% of individuals >65 and up to 50% of individuals greater than 85 years of age. The clinical course of AD ranges from

5 to 15, depending on the age of onset. Historically, AD was first identified in 1901 by a

German psychiatrist Alois Alzheimer, whose patient named Auguste Deter presented with AD- like symptoms, and upon autopsy found ‘milliary foci’ and ‘neurofibrils’ in brain, which were later biochemically identified as amyloid plaques and neurofibrillary tangles, respectively.

Alzheimer’s disease (AD) currently afflicts 5.4 million individuals in the USA. Due to the aging population, this number is estimated to rise to 13.2 million by 2050 if no effective treatment is found. In 2012, the cost of caring for AD and related dementia patients in the US stood at $200 billion per year, and this figure is projected to rise to $1.1 trillion per yearr by 2050 (2012 dollars). Therefore, AD is a tremendous cost to patients and families in both human and financial terms.

Amyloid plaques and neurofibrillary tangles

The major pathological hallmarks of Alzheimer’s disease are the Amyloid plaques and neurofibrillary tangles. Amyloid plaques are the accumulations of amyloid β (Aβ) protein, largely in extracellular space, while neurofibrillary tangles are the intraneuronal inclusion of hyperphosphorylated tau. Aβ is a neurotoxic peptide derived from amyloid precursor protein which is cleaved by β-, and γ-secretases (APP PROCESSING FIGURE). The vast majority of

APP is normally cleaved in the middle of the A sequence by -secretase

(ADAM10/TACE/ADAM17) in the non-amyloidogenic pathway, thereby preventing the generation of an intact A peptide. Alternatively, a small proportion of APP is cleaved in the amyloidogenic pathway, leading to the secretion of A peptides (37 to 42 amino acids) via two proteolytic enzymes, - and -secretase, known as BACE1 and presenilin, respectively (De

Strooper & Annaert, 2000a). The proteolytic processing of APP to generate A requires endocytosis from the cell surface and localization to cholesterol-rich membrane rafts where both

BACE1 and the presenilin complex are enriched (Koo & Squazzo, 1994; Perez et al., 1999;

Wahrle et al., 2002).

Neurofibrillary tangles are composed of the microtubule associated protein tau.

Hyperphosphorylated tau assembles into twisted filaments seen by electron microscopy as paired helical filaments or PHFs, which are immunopositive with AD-specific diagnostic antibodies, such as AT100 or AT8 (Avila, Lucas, Perez, & Hernandez, 2004). Although tau stabilizes/associates with microtubules within somatoaxonal compartments under normal physiological conditions, tau becomes hyperphosphorylated during early stages of AD, which weakens its association to microtubules and often mislocalizes to somatodendritic compartments.

Thus, hyperphosphorylated tau is unable support microtubule stabilization, mislocalizes to

dendritic compartments, and becomes increasing prone to self-aggregation (Gendron &

Petrucelli, 2009).

Genetic basis of Alzheimer’s disease (AD)

Early onset AD

Early onset AD occurs before the age 65. Of this group, familial early-onset AD (FAD) patients are those whose genetic causes have been identified. In 1991, the amyloid hypothesis was postulated that the accumulation of A is the fundamental cause of AD rather than simply pathological hallmarks of the disease (Hardy & Allsop, 1991). This hypothesis was initially supported by the identification of mutations in APP that co-segregated with FAD patient families. Of important note, all of the FAD mutations identified thus far are concentrated near the -secretase or -secretase cleavage sites in APP such that either total A or the more pathogenic A42 peptide is increased. While presenilin-dependent -secretase cleavage of APP at A residue 42 (A42) is a minor species (~10%) compared to A40 (>80%), many studies have shown that A42 is more neurotoxic and is required to efficiently seed the aggregation of

A40 (Glabe, 2008). Recently, a new Icelandic mutation in near the BACE1 cleavage site was identified, which protects against the risk of AD and reduces A secretion by ~40% (Jonsson et al., 2012). Furthermore, individuals with trisomy 21 (Down’s syndrome) with an extra copy of the APP gene on chromosome 21 invariably exhibit AD pathology by the 4th decade of life.

Closer examination of Down’s syndrome brains showed that A42 containing senile plaques develop much earlier than neurofibrillary tangles, further supporting the amyloid hypothesis

(Hyman, 1992; Hyman, West, Rebeck, Lai, & Mann, 1995). In 1995, mutations in two homologous genes, Presenilin 1 (PS1) and Presenilin 2 (PS2) were discovered that co-segregated with the most aggressive forms of autosomal dominant early-onset FAD. Thus far, more than 3

100 different FAD mutations in PS1 and PS2 have been identified (St George-Hyslop & Petit,

2005). Further adding support to the amyloid hypothesis, all of the presenilin mutations studied were found to increase the more pathogenic and faster aggregating A42 peptide (Table 1-

1)(Figure1-2)(Herz & Beffert, 2000).

Late onset AD

The vast majority of AD (>95%) is the late-onset sporadic form and it is not inherited in an autosomal dominant pattern. Various different genes have been identified as late onset genetic factors, and the most common and strongest genetic risk factor to date is the 4 allele of the apolipoprotein E (APOE) gene. APOE ε4 allele increases the risk for the disease by ~3-fold among hemizygotes and by ~15-fold among homozygotes (Holtzman, 2001). ApoE4 also promotes the accumulation of Amyloid beta far more efficiently than apoE3 (Strittmatter et al.,

1993). Therefore, all known and confirmed genetic component of AD either increase total A, the more pathogenic A42 peptide, or aggregation states of A. Recently, genomic-wide association studies (GWAS) have allowed the identification of variation in over 20 loci that contribute to disease risk such as INPP5D, MEF2C, ZCWPW1, CLU, PTK2B, PICALM,

SORL1, CELF1, CR1, BIN1, MS4A4/MS4A6E, SLC24A4/RIN3, FERMT2, CD33, ABCA7,

CASS4, TREM2, CD2AP, HLA-DRB1/HLA-DRB5, EPHA1, NME8. Furthermore, rare variants associated with Late-onset AD have been identified in TREM2 and PLD3 gene (Figure1-2).

Even though various different genes have been found, many of their roles in disease pathogenesis still have to be explored.

Molecular basis of Alzheimer’s disease (AD)

Soluble A oligomer hypothesis and hyperphosphorylation of tau

Biochemical and cellular studies on A-induced neurotoxicity have shown that distinct states or species of A have different biological properties. First, A42, while at much lower concentration than A40, aggregates faster and seeds the aggregation of A40 (Glabe, 2008).

Transgenic mice engineered to produce only A40 or A42 cleaved from the familial British and

Danish Dementia-related BRI protein shows that A40 cannot form aggregates into amyloid plaques even by 18 months of age, while a lesser concentration of A42 induces robust amyloid plaque formation even at 12 months of age. Furthermore, the BRI-A42 mice crossed with APP

Tg2576 mice bearing the “Swedish” mutation exponentially worsens amyloid plaque formation

(McGowan et al., 2005). Second, A can exist as soluble monomers, dimers, trimers, and higher order oligomers (also known as A-derived diffusible ligands; ADDLs) prior to forming protofibrils and insoluble amyloid fibrils. These pathological soluble A oligomers cause synaptic dysfunction to neurons even at picomolar concentrations. Studies have shown that SDS- stable dimers and trimers derived from the 7PA2 cell line expressing an FAD APP mutation impair long term potentiation (LTP), the physiological correlate of learning and memory, at subnanomolar concentrations (D. M. Walsh et al., 2002). Furthermore, soluble SDS-resistant A dimers derived from AD brains lead to hyperphosphorylation of tau and neuritic degeneration in primary hippocampal neurons at picomolar concentrations (Jin et al., 2011). Recently, a new mutation in APP was identified in a Japanese family with AD-type dementia. This mutation occurred as a deletion of residue 22 (glutamic acid) of the A peptide (E22). Biochemically, the E22 mutation is more resistant to degradation, unable to form fibrils, but far more prone to self-association as A oligomers in the form of SDS-stable dimers and trimers (Tomiyama et al., 5

2008). Expression of this APP mutation in transgenic mice leads to learning and memory deficits associated with impaired LTP, enhanced neuroinflammation, and tau hyperphosphorylation in the absence of any thioflavin S positive amyloid plaques. This indicated that A oligomers are sufficient and fibrillar amyloid deposition is not necessary for A-induced neurotoxicity and learning and memory deficits (Nishitsuji et al., 2009). Interestingly, loss of one copy of IGF-1 receptor, which lengthens lifespan, was shown to be protective by promoting faster aggregation of A and increasing the density of amyloid plaques, thereby sequestering toxic soluble A oligomer species into amyloid fibrils (Cohen et al., 2009). While it appears that amyloid fibrils per se might be less neurotoxic than soluble A oligomers, it is important to note that amyloid fibrils can disaggregate into soluble A oligomers, and thus, may serve as stable sources of these neurotoxic species.

Many studies have shown that A promotes the hyperphosphorylation of tau in vitro and in vivo (Avila et al., 2004; Gendron & Petrucelli, 2009). A also increases the number of neurofibrillary tangles in transgenic mice engineered to express a Frontotemporal Dementia

(FTDP-17) tau mutation (Lewis et al., 2001). Moreover, depletion of A by injection of an antibody directed against A also leads to reduced tau pathology in the APP/tau/presenilin-1 mutant triple transgenic mice (Oddo, Billings, Kesslak, Cribbs, & LaFerla, 2004), further supporting the model that A promotes tau pathology in vivo. In addition, recent studies have demonstrated that many of the toxic effects of A require tau expression, indicating that tau is indeed downstream of A-induced neurotoxic signals. Neurite retraction and progressive neuronal atrophy are seen when neurons are treated with A but not in neurons derived from tau knockout mice (Jin et al., 2011; Rapoport, Dawson, Binder, Vitek, & Ferreira, 2002).

Furthermore, learning and memory impairment as well as high sensitivity to excitotoxin 6

treatment are present in mutant APP transgenic mice but not in the same transgenic mice on a tau homozygous tau knockout background, even though A deposition is not affected by tau

(Roberson et al., 2007). A also induces impairments in LTP and axonal transport of mitochondria; however, such impairments are not seen in tau knockout neurons (Shipton et al.,

2011; Vossel et al., 2010), indicating that tau is required for multiple facets of A-induced neurotoxicity. Indeed, A oligomers rapidly lead to disassembly of microtubules but only in cells expressing tau (King et al., 2006). This is particularly important in neurons, as the microtubule network is critical for processes as axonal transport of key proteins and organelles to the synapse.

Therefore, A-induced hyperphosphorylation of tau appears to partially explain microtubule disassembly and impaired excitatory synaptic transmission. It is important to note that like soluble A oligomers, soluble hyperphosphorylated tau rather than PHF tau may represent the toxic species, since turning off FTD mutant tau expression in an inducible transgenic model does not remove insoluble PHF-1 positive tangle-like structures over several months but improves learning and memory (Santacruz et al., 2005).

Amyloid β and its surface receptors

Although it is clear that Aβ oligomers are neurotoxic at least in part by transmitting neurotoxic signals, what types of neuronal surface receptors and intracellular signals are needed to transmit these Aβ-induced neurotoxic signaling has remained to be fully elucidated. It has been reported that Aβ binds to various different neuronal receptors such as N-methyl-D-aspartate

(NMDA), nicotinic (α7nAChR), p75NTR receptors. Aβ binding to NMDA receptors has been reported to increase calcium influx and potentiate NMDAR activation, but other conflicting reports have shown that Aβ oligomers depress LTP and cause progressive loss of dendritic spines associated with reduced excitatory synaptic transmission and removal of glutamate 7

receptors from spines (Hsieh et al., 2006b; Pena, Gutierrez-Lerma, Quiroz-Baez, & Arias, 2006;

G. M. Shankar et al., 2007). Conflicting reports have also shown that Aβ can act as both agonist and antagonist of α7nAChR (Dineley et al., 2001; Pettit, Shao, & Yakel, 2001). The binding of

Aβ to p75NTR, especially in cholinergic neurons of the basal forebrain where this receptor is enriched, is associated with death of these neurons. In p75NTR knockout mice, such Aβ-induced degeneration of the basal forebrain is not observed (Yaar et al., 1997). Therefore, targeting the

Aβ/p75NTR interaction appears to represent a promising therapeutic strategy, especially for the cholinergic system. Recently, 2 other neuronal surface receptors for A oligomers have been proposed, namely PrPc/mGluR5(Lauren, Gimbel, Nygaard, Gilbert, & Strittmatter, 2009; Um et al., 2013) and Pir-B(T. Kim et al., 2013). These receptors bind specifically to A oligomers but not fibrils and are required for A-induced deficits in LTP in acute hippocampal slices (Figure 1-

4)(Overk & Masliah, 2014).

In addition to the aforementioned receptors, integrin adhesion receptors have also been shown to mediate A-induced neurotoxicity and deficits in LTP. The integrin family mediates both cell-cell and cell-substratum adhesion by binding their extracellular ligands such as laminin and fibronectin. Integrins consist of an α and a β subunit, and each subunit has a large extracellular portion, a single transmembrane segment, and a short cytoplasmic tail (Hynes,

1992). These cytoplasmic tails bind to intracellular ligands to mediate dynamic cellular responses, such as cell adhesion, migration, neurogenesis, apoptosis, and synaptic stability.

Hence, integrins provide a transmembrane link for the bidirectional transmission of signals across the plasma membrane by binding both extracellular and intracellular ligands (Zamir &

Geiger, 2001). The ability to connect to the actin cytoskeleton is an important part of the adhesive function of the integrin family. Within focal adhesions, structural proteins such as

vinculin and talin anchor -integrins to the actin cytoskeleton, while signaling proteins such as focal adhesion kinase (FAK), Pyk2, Paxillin, and Src mediate downstream signaling events in a transient and controlled manner (Cabodi et al., 2010) (Figure 1-2) (Vicente-Manzanares, Choi, &

Horwitz, 2009). Integrin signaling is also associated with changes in synaptic plasticity and neuronal excitability. Glutamate receptors (NMDA and AMPA receptors) are the target of integrin-dependent regulation. Interfering with integrin-dependent adhesion by antagonists or conditional deletion of 3, 5, and/or 1 integrins leads to defective LTP (Becchetti, Pillozzi,

Morini, Nesti, & Arcangeli, 2010). Therefore, integrins represent important adhesion receptors that can regulate multiple facets of neuronal function and viability. Integrin blocking antibodies to α1/β1 or echistatin (a highly selective and potent integrin inhibitor) blocks fibrillar Aβ- induced MAPK activation and Aβ-induced neurotoxicity in primary neuronal cultures (Anderson

& Ferreira, 2004). Moreover, antibodies against αv-integrins block the inhibition in LTP induced by Aβ. A small molecule nonpeptide antagonist of αv-containing integrins (SM256) and a potent disintegrin echistatin also block Aβ-induced inhibition of LTP. Accordingly, a potent ligand of

αv/ β1 integrin, superfibronectin, also efficiently inhibits Aβ-induced inhibition of LTP (Q.

Wang et al., 2008a). As glutamate receptors are targets of integrin-mediated regulation, it is likely that integrin-mediated alterations in glutamate receptors underlie Aβ-induced inhibition of

LTP. Addition of soluble Aβ together with fibrillar Aβ dramatically enhances the meshwork of

Aβ deposition on cortical neurons and induces neurotoxicity in an α2/β1 and αv/β1 integrin dependent manner (Wright et al., 2007a). Also, Antibodies against β1, α2, and αv but not α1, α3,

α4, α5, α6, α9, αv/β 3, αv/β5, or β3 integrins inhibit Aβ-induced neurotoxicity in human primary neurons (Wright et al., 2007a).

Neuroinflammation in Alzheimer’s disease

Multiple studies suggest Alzheimer’s disease pathogenesis is not restricted to neuronal compartments in the brain. In AD, it has been shown that there is a high expression of inflammatory mediators around Aβ deposition and neurofibrilarry tangles, which are associated with neurodegeneration (Akiyama et al., 2000). Moreover, it has been also established a link between chronic use of non-steroidal anti inflammatories (NSAIDs) and reduced risk of AD

(Vlad, Miller, Kowall, & Felson, 2008). More Recently, Genome-wide association studies identified immure CD33 and TREM2 as AD risk factors indicates the link between immune alterations and AD (Guerreiro & Hardy, 2013) . TREM2 (Triggering Receptor Expressed On

Myeloid Cells 2) is a microglial activator and act via DAP12 to activate syk-mediated tyrosine phosphorylation to promote microglial phagocytosis. CD33 encodes a cell-surface protein of the sialic acid-binding Ig-like lectin (SIGLEC) family and provides another example of a myeloid- cell- or microglial-cell-expressed gene in which variants have been associated with an increased risk of developing AD. CD33 expression was upregulated on microglia in human AD. Recently,

CD33 SNP rs3865444A, which is associated with decreased AD risk has been found (Malik et al., 2013; Raj et al., 2014), which is associated with reduced CD33 expression.

Mitochondrial dysfunction in Alzheimer’s disease (AD)

Mitochondria, called powerhouse of the cell provide cellular energy, and also involved in various different cellular signaling. As the human brain utilizes 20% of total body oxygen consumption, mitochondria serve an important capacity in neuronal homeostasis and mechanisms of neurodegeneration. Mitochondria have various different biological functions such as ATP production, control the energy efficiency, calcium homeostasis, reactive oxygen species

(ROS) production, and regulation of apoptotic signaling, all of which play integral roles in neurodegeneration seen in AD.

Given that mitochondria are the most metabolically active organelles, mitochondrial dysfunction leads to brain metabolism failure. In AD, aberrant activity of tricarboxylic acid cycle

(TCA) complexes, pyruvate and isocitrate dehydrogenases, have been reported. Several studies have also shown Aβ deposition is associated with local levels of brain metabolism and neuronal activity. Interestingly, Aβ can be directly transported into mitochondria, which perturbs oxygen consumption and activity of enzymes in complex III and IV of the respiratory chain. Moreover, it has been reported that Aβ in mitochondria is associated with an increase in hydrogen peroxide and a decrease in COX activity.

Mitochondria play essential roles in regulating Ca2+ homeostasis which is pathologically linked to neurotoxicity. When Ca2+ levels increase in the cytosol, mitochondria potently take up

Ca2+ through the mitochondrial Ca2+ uniporter (MCU), resulting in Ca2+ buffering and increased metabolism and production of ATP (Bezprozvanny & Mattson, 2008; Matter, Zhang, Nordstedt,

& Ruoslahti, 1998). However if excess amounts of Ca2+ enter the mitochondria (calcium overload), it leads to decreased mitochondrial membrane potential (MMP) and oxidative stress.

This, in turn, results in formation of the mitochondrial permeability-transition pore (mtPTP) and failure to exert Ca2+ buffering capacity, leading to secondary excitotoxicity (Hengartner, 2000).

Multiple studies have demonstrated that A can dysregulate Ca2+ homeostasis via several Ca2+ signaling components (Bezprozvanny, 2009): calcineurin (Kuchibhotla et al., 2008), NMDARs

(De Felice et al., 2007), AMPARs (Hsieh et al., 2006a, 2006b), and P/Q-type VGCCs (Nimmrich et al., 2008). Furthermore, FAD mutations in presenilins (PS), which potentiate A42 generation, induce excessive Ca2+ release from the ER through IP3 and ryanodine receptors (Cheung et al.,

2008; Rybalchenko, Hwang, Rybalchenko, & Koulen, 2008). In AD brains, intracellular Ca2+ levels are positively correlated with neurofibrillary tangle formation (McKee, Kosik, Kennedy,

& Kowall, 1990; Nixon, 2003). We and others have also shown that Aβ induces aberrant changes in mitochondrial membrane potential in cultured neurons and astrocytes, which can lead to the disruption of mitochondrial Ca2+ homeostasis. In addition, many studies have shown that Aβ can overactivate NMDA and AMPA receptors that can ultimately lead to mitochondrial Ca2+ overload and mitochondrial damage. This, in turn, results in mitochondrial dysregulation and apoptosis.

Mitochondria serve as prominent sources of reactive oxygen species (ROS) production, reactive molecules containing oxygen that regulate cellular signaling and cell death. ROS, at low to moderate concentrations, is an important mediator of defense mechanisms against infectious agents, various cellular signaling events, and synaptic function. However, if ROS levels exceed the amount that can be neutralized by anti-oxidant mechanisms, it eventually results in mitochondrial dysfunction and neuronal damage (Andreyev, Kushnareva, & Starkov, 2005), the molecular targets of which are DNA, lipids, and protein machineries within the cell and mitochondria (Knight, 1997). The role of mitochondrial ROS as inducers of Ca2+ dysregulation is well-established. On the other hand, a major cause of ROS production is also Ca2+ dysregulation. Thus, oxidative stress and Ca2+ regulation are intricately linked and can cooperatively contribute to AD pathogenesis. It has been shown that Aβ oligomers can form

Ca2+-permeable channels in the plasma membranes (Arispe, Rojas, & Pollard, 1993) and that the association of Aβ to membranes is enhanced in the presence of excess phosphatidylserine

(PtdS) (Lee, Pollard, & Arispe, 2002). Other studies have demonstrated that Aβ treatment promotes caspase 3, 8, and 9 activation (Fossati et al., 2010; Z. F. Wang et al., 2010)

Furthermore, Aβ induces abnormal mitochondrial dynamics. Specifically, Aβ peptide impairs axonal transport of mitochondria, attenuates mitochondrial motility, and alters mitochondrial distribution in mouse hippocampal neurons, resulting in synaptic degeneration (Calkins &

Reddy, 2011). Aβ induces Ca2+ dysregulation by membrane-associated oxidative stress and activates calcineurin, an important mediator of synaptic plasticity (Celsi et al., 2007). In addition,

Aβ induces synaptic dystrophy and dysregulation at least in part due to decreased expression of

NMDA and EphB2 receptors at the synapse, an event that can be corrected by NMDAR agonists

(Lacor et al., 2007). Though the exact mechanisms of ROS-induced synaptic degeneration remain to be further elucidated, ROS is well known to play important roles in synaptic plasticity.

Indeed, oxidation of cysteine residues on RynR increases Ca2+ release from the ER and oxidation (Hidalgo, 2005) of the same residues on NMDAR decreases the receptor activity

(Lipton et al., 2002). Therefore, it appears that Aβ induces synaptic deficits via both increasing oxidative damage and dysregulating Ca2+ dynamics at the synapse and mitochondria, both of which cooperatively contribute to neurodegeneration in AD (Figure 1-3)(Luque-Contreras,

Carvajal, Toral-Rios, Franco-Bocanegra, & Campos-Pena, 2014).

Tau pathology is also linked to mitochondrial dysfunction. Transgenic mice overexpressing the P301L mutant demonstrate a significant reduction in complex V and NADH- ubiquinone oxidoreductase activity together with impairments in mitochondrial respiration and

ATP synthesis (David et al., 2005). Tau can be cleaved by caspases 3, 7, and 8 after Asp-421, yielding a 421 residue tau fragment that assembles into PHF-like fibrils more rapidly than full length tau. Overexpression of this cleaved tau leads to a significant decrease in mitochondrial membrane potential and loss of mitochondrial membrane integrity (Quintanilla, Matthews-

Roberson, Dolan, & Johnson, 2009). Moreover, introduction of tau in mature hippocampal

neurons results in degeneration of synapses by perturbing mitochondria transport and ATP levels at the synapse (Thies & Mandelkow, 2007). N-terminal tau fragments have toxic effects on mitochondria and lead to the mitochondrial dysfunction associated with impairments in oxidative phosphorylation by distorting the enzyme structure of complex V and the level of adenine nucleotide translocator, which ultimately perturbs ATP synthesis in mitochondria (Atlante et al.,

2008). Interestingly, these N-terminal tau fragments are localized in AD synaptic mitochondria

(Amadoro et al., 2010). Taken together, both A and tau exert deleterious effects on mitochondria, events that involve ROS generation and Ca2+ dysregulation, which negatively impact synaptic function and neuronal viability.

Cytoskeletal aberrations in AD

The cytoskeleton is a dynamic structure mainly composed of microfilaments, intermediate filaments and microtubules. Cytoskeleton has multiple biological functions, including provision of mechanical resistance, anchorage to the extracellular matrix, and functioning as scaffolds for transport of need proteins and organelles (i.e. microtubules). These functions regulate various cellar pathways, including cell adhesion, migration, division, and survival. Actin is a globular protein that can forms microfilaments, and it is major components of the cytoskeleton that can mediate cell motility. Actin has two distinct states in the cells which are a monomeric G-actin and filaments F-actin. F-actin preferentially polymerizes at one end of the filament at steady state, and the difference in polymerization rates between the two ends result in a net turnover of the filaments. Multiple studies have shown actin dynamics play important roles in dendritic spine morphogenesis, membrane protein trafficking, and synaptic plasticity.

Role of Cofilin in F-Actin dynamics and mitochondrial dysfunction

ADF/Cofilin, a family of actin-binding protein is a major regulator of actin dynamics.

Cofilin modulates actin dynamics depending on the ratio of cofilin to actin. At low cofilin to actin ratios, cofilin acts to sever actin filaments, whereas at high cofilin to actin ratios, cofilin nucleates actin assembly and stabilizes F-actin (B.W. Bernstein & J.R. Bamburg, 2010). The activity of cofilin is tightly regulated by phosphorylation and dephosphorylation on serine-3 of cofilin by LIMK and Slingshot-1L (SSH), respectively (Eiseler et al., 2009b). Phosphorylation of cofilin by LIMK1, a downstream component of Rac-PAK signaling upon focal adhesion activation, serves to inactivate cofilin, whereas dephosphorylation of cofilin by Slingshot-1L

(SSH) activates cofilin (Figure 1-4)(Chiu, Patel, Shaw, Bamburg, & Klip, 2010). It has also been shown that reelin, via a pathway involving Dab1, Src, and PI3K, promotes the phosphorylation of cofilin on serine-3, thereby inactivating cofilin (Chai, Forster, Zhao, Bock, & Frotscher,

2009). Therefore, proper relay of focal adhesion signals downstream of integrin and/or reelin activation leads to the phosphorylation and inactivation of cofilin by LIMK1, Src, and potentially other kinases, leading to reduced actin dynamics. Multiple studies have shown that cofilin is involved in oxidative stress induced cell death via critical mitochondrial mechanism. Upon oxidative stress, activated cofilin becomes oxidized on several cysteine residues. This allows cofilin to lose its affinity for actin and to translocate to mitochondria, where it induces swelling, drop in mitochondrial membrane potential, and cytochrome c release by promoting the opening of the permeability transition pore. Furthermore, knockdown of endogenous cofilin by siRNA also inhibits both oxidant and staurosporin induced apoptosis, indicating that cofilin is critical for mitochondria-mediated apoptosis. Cellular stimuli that generate ROS not only oxidize cofilin to promote mitochondrial dysfunction but also activate SSH via oxidation of 14-3-3, a chaperone

protein that normally functions to inhibit SSH in the nonoxidized form (J.S. Kim, T.Y. Huang, &

G.M. Bokoch, 2009). Therefore, enhanced ROS generation by A, tau, and/or other oxidative stress can dephosphorylate and activate cofilin. Another critical pathway of cofilin activation is via calcium. Calcineurin, a calcium activated phosphatase, dephosphorylates SSH, thereby promoting cofilin dephosphorylation and activation (Y. Wang, Shibasaki, & Mizuno, 2005a).

Given that A increases intracellular Ca2+, it has been reported that A also activates calcineurin and dephosphorylates cofilin (Homma, Niino, Hotta, & Oka, 2008). Furthermore, A-induced dendritic spine loss is also mediated by a pathway involving calcineurin and dephosphorylation of cofilin (G.M. Shankar et al., 2007). Indeed, inhibition of calcineurin by FK506 is neuroprotective in transgenic animal models of AD (Hong et al., 2010; Rozkalne, Hyman, &

Spires-Jones, 2011). Finally, it has been shown that active PAK, a downstream component of focal adhesion and Rac signaling, is severely reduced in AD brains. A oligomers induce defects in PAK signaling, leading to the activation of cofilin and synaptic pathology. Overexpression of active PAK reverses these effects of A oligomers (Ma et al., 2008; Zhao et al., 2006).

Therefore, A-induced alterations in Ca2+, ROS, and focal adhesion signaling can also induce the activation of cofilin, which in turn, dysregulates cytoskeletal dynamics, mitochondria, and tau .

Activated and oxidized cofilin can also form cofilin-actin rods in the form of rod shaped bundles of filaments. Cofilin-actin rods form in different cultures cells including primary neurons in response to heat shock, osmotic stress, and ATP depletion, and Aβ. Cofilin-actin rods have been found in both axons and dendrites of mouse hippocampal and cortical neurons and in organotypic hippocampal slices. These cofilin-actin rods can block axonal transports as well as transport within neurites. Moreover, a subset of cofilin-actin rods contains hyperphosphorylated

tau, indicating that cofilin-actin cytoskeletal dynamics also alter tau pathology (Whiteman et al.,

2009). Indeed, it has been demonstrated that transgenic mice and flies overexpressing FTDP-17 mutant tau promote F- actin assembly, and hyperphosphorylated tau coprecipitates in rods together with cofilin and actin (Fulga et al., 2007). These rod structures resemble striated neuropil threads. Cofilin also plays a key role in the organization of actin cytoskeleton in dendritic spines as well as Aβ induced spine loss (G. M. Shankar et al., 2007). Thus, cofilin and tau may cooperatively control such activity and eventually precipitate in rods that contain cofilin, actin, and tau when neurons become further stressed by Aβ-induced neurotoxic signals.

RanBP9 in APP processing, mitochondria-mediated neurotoxicity, and integrin signaling

RanBP9, also known as RanBPM, acts as a multi-modular scaffolding protein, bridging interactions between the cytoplasmic domains of a variety of membrane receptors and intracellular signaling targets. These include Axl and Sky (Hafizi, Gustafsson, Stenhoff, &

Dahlback, 2005), MET receptor protein tyrosine kinase (D. Wang, Li, Messing, & Wu, 2002), and 2-integrin LFA-1 (Denti et al., 2004). Similarly, RanBP9 interacts with Plexin-A receptors to strongly inhibit axonal outgrowth (H. Togashi, E.F. Schmidt, & S.M. Strittmatter, 2006) and functions to regulate cell morphology and adhesion (Dansereau & Lasko, 2008; Valiyaveettil et al., 2008b). We previously demonstrated that the C-terminal 37 amino acids of the low density receptor-related protein (LRP) robustly promoted A generation independent of FE65 and specifically interacted with Ran Binding Protein 9 (RanBP9). Our initial studies found that

RanBP9 strongly increases BACE1 cleavage of APP and A generation. In cells expressing wild type APP, RanBP9 reduced cell surface APP and accelerated APP internalization, consistent with enhanced -secretase processing in the endocytic pathway. The N-terminal half of RanBP9 containing SPRY-LisH domains not only interacted with LRP but also with APP and BACE1.

Overexpression of RanBP9 resulted in the enhancement of APP interactions with LRP and

BACE1 and increased lipid raft association of APP. Importantly, knockdown of endogenous

RanBP9 significantly reduced A secretion in Chinese Hamster Ovary cells and in primary neurons, demonstrating its physiological role in BACE1 cleavage of APP. We also found that

RanBP9 overexpression in transgenic mice (APP/PS1;RanBP9-TG) significantly increased Aβ load and thioflavin-S positive amyloid in brain compared to single APP/PS1 transgenic mice.

Moreover, we found that an N-terminal bioactive fragment of RanBP9 protein was significantly increased in brains of AD patients (M. K. Lakshmana et al., 2012).

In addition to its effects on APP processing, our studies have shown a significant role of

RanBP9 in neurotoxicity and apoptosis. RanBP9 single transgenic mice demonstrated significantly increased synapse loss, neurodegeneration, gliosis, and spatial memory deficits.

Consistent with these in vivo observations, RanBP9 overexpression promoted apoptosis and potentiated A-induced neurotoxicity independent of its capacity to promote A generation.

Conversely, RanBP9 reduction by siRNA or gene dosage mitigated A-induced neurotoxicity in

HT22 neuroblastoma cells. Importantly, RanBP9 overexpression led to the activation of cofilin, a key regulator of actin dynamics and mitochondria-mediated apoptosis, and siRNA knockdown of cofilin abolished both A and RanBP9-induced apoptosis (Figure 1-5)(J. A. Woo et al., 2012).

We later went on to show in primary hippocampal neurons that RanBP9 potentiates Aβ-induced reactive oxygen species (ROS) overproduction, apoptosis, and calcium deregulation. Analyses of calcium handling measures demonstrated that RanBP9 selectively delays the clearance of cytosolic Ca2+ mediated by the mitochondrial calcium uniporter (MCU) through a process involving the translocation of cofilin to mitochondria and oxidative mechanisms (Roh et al.,

2013b). Further, RanBP9 retarded the anterograde axonal transport of mitochondria in primary

neurons and decreased synaptic mitochondrial activity in brain (Roh et al., 2013b). These data indicated that RanBP9, cofilin, and A mimic and potentiate each other to produce mitochondrial dysfunction, ROS overproduction, and calcium deregulation, leading to neurodegenerative changes reminiscent of those seen in AD.

Because APP, LRP, and RanBP9 all physically interact with -integrins, we also investigated whether RanBP9 alters integrin-dependent cell adhesion and focal adhesion signaling. We found that RanBP9 overexpression dramatically disrupts integrin-dependent cell attachment and spreading in NIH3T3 and hippocampus-derived HT22 cells, concomitant with strongly decreased Pyk2/Paxillin signaling and Talin/Vinculin localization in focal adhesion complexes (J. A. Woo et al., 2012). Conversely, RanBP9 knockdown robustly promoted cell attachment and spreading, as well as, focal adhesion signaling and assembly (J. A. Woo et al.,

2012). Cell surface biotinylation and endocytosis assays revealed that RanBP9 overexpression and RanBP9 siRNA potently reduces and increases surface 1-integrin and LRP by accelerating and inhibiting their endocytosis, respectively. Primary hippocampal neurons derived from

RanBP9 transgenic mice also demonstrated severely reduced levels of surface 1-integrin, LRP, and APP as well as neurite arborization (J. A. Woo et al., 2012). Therefore, these data indicated that RanBP9 simultaneously inhibits cell adhesive processes and enhances A generation by accelerating APP, LRP, and 1-integrin endocytosis.

Table 1-1 Genes associated with Alzheimer’s disease

Dominant mutations that cause early-onset AD have been identified in APP, PS1 and PS2, and cause a relative increase in APP processing to AThe ApoE4 allele has been identified to late-onset AD in a large number of studies. LRP, a multifunctional receptor that binds ApoE and numerous other ligands has been found to be weakly linked to late-onset AD by several studies.

Figure 1-1 Amyloid β oligomer receptors

Putative Aβ oligomer receptors, signaling pathways, and therapeutic targets. A number of potential cell surface molecules (insulin receptor, NGF receptor, PrPc, mGluR5, EphA4, EphB2, NMDA, and Flizzed) that mediate the synaptotoxic effects of Aβ oligomers have been identified.

Figure 1-2 The actin-integrin linkage

The linkage between the extracellular matrix (ECM, red strand on top) and the actin cytoskeleton (represented by yellow beaded coils) is depicted. Integrins (represented by the α- and β-transmembrane subunits in light blue and pink) can bind directly to the talin head domain (red sphere). Through its tail domain (red rod), talin can bind directly to actin as well as to other components of the linkage, such as vinculin (shown in purple). Vinculin can also bind to actin directly, as well as to the actin cross-linker α- actinin (shown as a dimer, in green). Both vinculin and α-actinin are anchored to the membrane, and their activity is modulated by interactions with phosphatidylinositol (4,5)-bisphosphate (PIP2). Finally, vinculin and FAK (shown in blue) can bind to the actin nucleator Arp2/3 (shown as a heptamer in grey).

Figure 1-3 Mitochondrial damage in Alzheimer’s disease

Aβ overproduction damages mitochondria causing dysfunction of mitochondrial complexes I and IV, which result in reactive oxygen species (ROS) overproduction and adenosine triphosphate (ATP) depletion. In neurons, ATP depletion may lead to neurotransmission dysfunction and altered axonal transport, thus provoking mitochondrial dynamics abnormalities. ATP depletion also causes dysfunction of the ATP-dependent ion channels, leading to altered ion balance in the cytosol. ROS increase in turn leads to mitochondrial permeability transition pore (MPTP) aperture, which increases mitochondrial damage by allowing calcium entrance into the mitochondrial matrix, worsening the electron transport chain and oxidative phosphorylation disruption.

Figure 1-4 Cofilin regulates actin remodeling

The accumulation of polymerized F-actin poses a stimulatory factor in the phosphatase activity of SSH, which leads to net dephosphorylation and activation of cofilin. Hereon, the actin-severing function of cofilin maintains the flexibility of remodeled actin and enables regeneration of free monomeric actin for further polymerization.

Figure 1-5 Schematic model of the RanBP9-cofilin pathway in AD pathogenesis

As the binding of Aβ to various integrins is critical to transmit its neurotoxic signals, Aβ oligomers are proposed to activate/dephosphorylate cofilin via the recruitment of RanBP9 to integrin/APP/LRP complexes. The binding of RanBP9 to integrin/APP/LRP complexes accelerates their endocytosis, thereby promoting Aβ generation, disrupting focal adhesions, and activating cofilin. Under conditions of oxidative stress such as that induced by Aβ oligomers, oxidized, and dephosphorylated cofilin translocates into the mitochondria to open up the mitochondrial permeability transition pore, which induces mitochondrial dysfunction and promotes apoptotic processes. Activated cofilin can lead to the formation of cofilin-actin rods, which attract hyperphosphorylated tau and promote tau pathology.

CHAPTER 2

SLINGSHOT-COFILIN ACTIVATION MEDIATES MITOCHONDRIAL AND

SYNAPTIC DYSFUNCTION VIA AMYLOID BETA LIGATION TO BETA1-

INTEGRIN CONFORMERS

Permissions Statement

The information in chapter 2, “Slingshot-Cofilin Activation Mediates Mitochondrial and

Synaptic Dysfunction” has been legally reproduced under the Creative Commons Attribution

(CC-BY) license. This means the publication is accessible online without any restrictions and can be re-used in any way subject only to proper citation.

Woo, J. A., Zhao, X., Khan, H., Penn, C., Wang, K., Joly-Amado, A., Weeber, E., Morgan, D.,

Kang, D. E (2015). “Slingshot-Cofilin activation mediates mitochondrial and synaptic

dysfunction via Aβ ligation to β1-integrin conformers.” Cell death and differentiation

(2015) 22, 921-934

Background

The defining pathological hallmark of Alzheimer’s disease (AD) is the accumulation of

A in brain associated with tau pathology, synapse loss, cytoskeletal aberrations, mitochondrial dysfunction, and cognitive decline. Soluble oligomeric forms of A are thought to be the most 26

toxic species, resulting in synaptic loss and downstream neurotoxicity (D. M. Walsh et al., 2002).

An early and consistent impairment secondary to A oligomer treatment in primary neurons is the shrinkage of dendritic spines (Selkoe, 2008) involving the rearrangement of F-actin cytoskeleton in spines and loss of spine-associated proteins such as PSD95 and Drebrin(G.M.

Shankar et al., 2007; Zhao et al., 2006), as well as impaired mitochondrial function(Du et al.,

2010; Moreira, Santos, Moreno, & Oliveira, 2001). Studies have implicated an involvement of the F-Actin-severing protein Cofilin in A-induced dendritic spine changes(G.M. Shankar et al.,

2007; Zhao et al., 2006), accumulation of Cofilin-Actin aggregates / rods in AD brains

(Minamide, Striegl, Boyle, Meberg, & Bamburg, 2000), and increased Cofilin activity in brains of AD patients (T. Kim et al., 2013). Cofilin normally functions as a key regulator of Actin dynamics that destabilizes F-Actin. Cofilin is inactivated by phosphorylation on Ser3 by LIMK1, whereas its dephosphorylation by SSH1 activates Cofilin(B. W. Bernstein & J. R. Bamburg,

2010; Kurita, Watanabe, Gunji, Ohashi, & Mizuno, 2008; Niwa, Nagata-Ohashi, Takeichi,

Mizuno, & Uemura, 2002). Upon oxidative stress and/or Ca2+ elevation(B. W. Bernstein & J. R.

Bamburg, 2010; J. S. Kim, T. Y. Huang, & G. M. Bokoch, 2009; Roh et al., 2013a), SSH1 is activated and active Cofilin becomes oxidized on cysteine residues, resulting in rapid mitochondrial translocation to promote mitochondria-mediated apoptosis and induction of

Cofilin-Actin pathology(B. W. Bernstein, Shaw, Minamide, Pak, & Bamburg, 2012; Klamt et al.,

2009). Despite the circumstantial evidence for the involvement of Cofilin in AD pathogenesis, no direct evidence thus far has been presented.

Heterodimeric Integrins ( &  subunits) comprise major adhesion receptors that regulate multiple facets of cellular function, including adhesion, motility, survival, and synaptic plasticity(Becchetti et al., 2010). A primary function of Integrins is to link the extracellular

matrix to the F-actin cytoskeleton via structural scaffolding proteins such as Talin and Vinculin

(C. Kim, Ye, & Ginsberg, 2011; Zamir & Geiger, 2001). Among several proposed surface A oligomer receptors such as PrPc/mGluR5(Lauren et al., 2009; Um et al., 2013) and Pir-B(T. Kim et al., 2013), it has been shown that 2/1 and v/1-integrins are also required to mediate A- induced apoptosis and impairment in LTP(Q. Wang et al., 2008b; Wright et al., 2007b).

However, whether A binds directly to integrins and how A engagement alters downstream integrin function are unknown. In this study, we explored the mechanistic relationships among

A42O, 1-integrin, and Cofilin activities in vitro, HT22 cells, primary neurons, and genetically modified mice. Here we show that A42O exhibits direct high-affinity binding 1-integrin, inducing its conformational alteration, loss of surface 1-integrin, and disruption of integrin- associated focal complexes as well as mitochondrial and synaptic dysfunction via a pathway involving ROS-dependent activation of SSH1 and Cofilin.

Materials and Methods

Cells, cDNA Constructs, siRNA Sequences, and Transfections

Hippocampus derived HT22 and CHO 7WD10 cells were maintained in DMEM containing 10% FBS. HT22 and CHO7WD10 cell lines were obtained from Prof. David

Schubert (Salk Institute, San Diego) and Prof. Edward Koo (UC, San Diego). GFP-Vinculin construct was obtained from Professor Ballestrem (University of Manchester, UK)(Humphries et al., 2007). The 1-Fc Integrin construct was obtained from Dr. Mould at the University of

Manchester (UK). The 1-integrin-Fc construct encodes residues 1-708 (1 extracellular region) fused in-frame to the hinge regions and CH2 and CH3 domains of human IgG1(Valdramidou,

Humphries, & Mould, 2008). RNAi sequences targeting Cofilin (5’-

GGAGGACCUGGUGUUCAUC-3’) and SSH1 (5'-GAGGAGCUGUCCCGAUGAC-3') were 28

obtained from Dharmacon. All cells were transfected for 48 hours using lipofectamine 2000

(Invitrogen) according to the manufacturer’s instructions with either cDNA expression constructs, empty vector controls, siRNA duplexes, or control single-stranded denatured control siRNAs prior to performing biochemical and/or immunocytochemical assays. All siRNAs were transfected at a final concentration of 100nM. Cortical and hippocampal primary neurons were derived from postnatal day 0 (P0) pups and grown on Poly-D-Lysine coated coverslips or plates as previously described (J. A. Woo et al., 2012).

Antibodies and Reagents

Antibodies to APP (6E10, Covance), sAPP (IBL-America), sAPP (IBL-America), 1- integrin (M-106, Santa Cruz), activated 1-integrin (HUTS-4, Millipore), allosteric 1-integrin neutralizing (MAB1965, Millipore), 6D11 (Santa Cruz), Talin (H-300, Santa Cruz), Vinculin

(hVIN-1, Sigma-Aldrich), Cofilin (D3F9, Cell Signaling), phospho-Cofilin (77G2, Cell

Signaling), SSH1 (SP1711, ECM Biosciences), pan-14-3-3- (Santa Cruz), Actin (AC-74, Sigma

Aldrich), Tubulin (TU-02, Santa Cruz), A (D5XD2, Cell Signaling), biotin (B7653, Sigma-

Aldrich), GFAP (Invitrogen), Synapsin I (Invitrogen), PSD95 (Abcam), Drebrin (Abcam),

MAP2 (Millipore), , HRP-linked secondary antibodies (Jackson Immunoresearch), and fluorescently labeled secondary antibodies (Invitrogen) were obtained from the indicated sources. The mouse monoclonal anti-RanBP9 antibody was a generous gift from Prof. Elizabetta

Bianchi (Pasteur Institute, France). Trolox (Cayman), JC-1 (Invitrogen), Mitosox-Red

(Invitrogen), H2DCF (Invitrogen), NOX inhibitor III (Millipore) were purchased from the indicated sources. Synthetic A1-42 and FITC-A1-42, and biotin-A1-42 peptides were purchased from American Peptide (Sunnyvale, CA). A1-42 oligomers and fibrils were prepared precisely as previously characterized (Dahlgren et al., 2002). Briefly, A1-42 powder was

dissolved in HFIP at 1mM for 30min at room temperature, aliquoted to eppendorf tubes, allowed to evaporate overnight in fume hood, and subjected to speed vacuum for 1h to remove traces of

HFIP or moisture. To prepare A oligomers, A1-42 film was then dissolved in DMSO (5mM), and F-12 cell culture medium (without phenol) was added to a final concentration of 100M

A1-42 and incubated at 40C for 24h.

Cell/Tissue Lysis and Immunoblotting

Cultured cells or brain homogenates were lysed with lysis buffer (50mM Tris-Cl, 150mM

NaCl, 2mM EDTA and 1% TritonX-100) unless stated otherwise. For separation of cytosolic versus focal adhesion-enriched (F) fractions, cytosolic proteins were first released with saponin buffer (0.1% saponin, 25mM Hepes, 75mM potassium acetate) for 10min on ice followed by extraction of remaining proteins by 2X SDS sample buffer and sonication (Kang et al., 2002;

Soriano et al., 2001). For isolation of mitochondria, mitochondrial isolation kit (Thermo

Scientific, IL, USA) was used according to the manufacturer’s instructions for cultured cells.

Protein quantification was performed by a colorimetric detection reagent (BCA Protein Assay,

Pierce). Equal amounts of protein were subjected to SDS-PAGE and transferred to nitrocellulose membranes for immunoblotting. After probing with the primary antibody, the corresponding peroxidase-conjugated secondary antibody was detected by ECL western blot reagents (Pierce).

ECL images were captured by the Fuji LAS-4000 imager and quantified using the Image J software.

Cell Surface Biotinylation Assay

Surface biotinylation assays were performed as we previously documented(Lakshmana et al., 2009). Briefly, confluent cells in 6 well plates were washed three times in PBS and treated with 2.0 mg/ml sulfo-NHS-LC-biotin in Phosphate buffered saline (PBS), pH 8.0 under gentle 30

shaking for 1 h on ice. The cells were then washed three times in PBS and lysed in 1% Triton-X-

100 lysis buffer. Biotinylated proteins were isolated by pull down with anti-biotin antibody together with anti-mouse agarose beads (American Qualex). APP (6E10, Covance) and 1- integrin (M-106, Santa Cruz Biotechnology) within anti-biotin immune complexes were detected by immunoblotting with the indicated antibodies.

Annexin V / PI Apoptosis Assay

To examine the stage of apoptotic cells, measurements were carried out using FACS

Calibur (BD Bioscience, San Jose, CA) or fluorescence microscopy. Cells were either trypsinized/collected stained in solution or directly stained on glass coverslips with Annexin V- fluorescein and PI for 20 min at 25℃ in the dark state using the FITC Annexin V apoptosis detection kit I (BD, San Diego, CA). After washing with ice-cold PBS, cells were measured by flow cytometry or by fluorescence microscopy as previously described (J. A. Woo et al., 2012).

FITC-A1-42 Oligomer Binding to Primary Neurons and HT22 Cells

DIV21 hippocampal neurons or HT22 cells were incubated with FITC-A42O (100nM) for 1-2h. Cells were then washed extensively with PBS, fixed with 4% paraformaldehyde, stained with DAPI, and fluorescent images were acquired using the Olympus FV10i confocal microscope. All comparison images were acquired with identical laser intensity, exposure time, and filter. Adjustments to the brightness/contrast were applied equally to all comparison images.

Purification of 1-integrin-Fc and Binding to A42O

CHO cells in 10cm plates were transfected with 12g 1-integrin-Fc constructs for 48h.

Conditioned medium was collected for 24h thereafter and spun for 5min at 1000g. Proteins in expressed in conditioned medium were captured by the anti-Fc affinity column and eluted by

several rounds. 96-Well plates were coated with goat anti-human-Fc at a concentration of 5g/ml in DPBS (50 l/well) for 16h. Wells were washed with DPBS (100 l/well) and then blocked for

1 h with 100l of 5% (w/v) bovine serum albumin (BSA), 150 mM NaCl, 0.05% (w/v) NaN3, 25 mM Tris-Cl, pH 7.4 (blocking buffer). The blocking solution was removed, and wells were then washed three times with 100l of 150mM NaCl, 25mM Tris-Cl, pH 7.4, and containing 1 mg/ml

BSA (washing buffer). Purified 1-Fc was added to the plate incubated at room temperature for

1 h. Wells were washed three times in washing buffer, and biotin labeled A1-42 (50l, monomer, oligomer or fibrils at the indicated concentrations were added for 1h at 37°C. After washing wells 3 times in washing buffer, streptavidin-HRP (1:2000 dilution in washing buffer,

Cell Signaling) was added (50l/well) for 30 min at room temperature. Wells were washed 4x in washing buffer, and color was developed using the TMB substrate (Cell Signaling) and read at

450nM wavelength. Background A binding to conditioned medium from mock-transfected

CHO cells was subtracted from all measurements (specific binding).

Mice and Human Samples

APP/PS1, wild type (WT), Cofilin+/-, and 1-integrin fl/fl mice were all bred in the

C57BL6 background for at least 3 generations prior to interbreeding with each other. APP/PS1 mice were obtained from Jackson laboratory (Jankowsky et al., 2004). We generated Cofilin+/- knockout mice from an embryonic stem (ES) cell gene trap clone originally made by Lexicon

Genetics and deposited into the MMRRC repository (UC, Davis). Cofilin+/- mouse brains exhibited ~50% reduction in endogenous Cofilin protein in both hippocampus and cortex compared to littermate wild type mice (Supplemental Fig. S8). While Cofilin+/- mice were viable and fertile with no salient abnormalities, Cofilin+/-X Cofilin+/- crosses yielded no viable

Cofilin-/- pups (embryonic lethal). Detailed genotypic and phenotypic information about these 32

mice are available (http://www.informatics.jax.org/external/ko/lexicon/2682.html). Information on 1-integrin fl/fl mice are also available (http://jaxmice.jax.org/strain/004605.html). Post- mortem brain samples were obtained from the UC, Irvine, ADRC. Ages, gender, and PMI are indicated in Supplemental Fig. S5.

Immunofluorescence

Immunohistochemistry and immunocytochemistry were performed as previously described (J. A. Woo et al., 2012). Briefly, animals were perfused with 4% paraformaldehyde in

PBS, and the brains were post-fixed in the same fixative for 24 hr. The brains were then cryoprotected in 30% sucrose and sectioned (30 µm) on a cryostat or microtome. For primary neurons and HT22 cells, cells were fixed in 4% paraformaldehyde for 15min at room temperature. For cell surface protein detection only, Triton-X-100 was omitted. After blocking with normal goat serum, primary antibodies were applied overnight at 40C, and secondary antibodies were applied for 45min at room temperature, followed by counterstaining with

Hoechst33342 or DAPI. Immunoreactivities were quantitated from every 12th serial section through an entire hippocampus or anterior cortex. In brain, focal Vinculin was separated from diffuse cytosolic Vinculin by using the ‘thresholding’ function in Image J software, where the threshold was adjusted to convert the highest intensity signal from green to red while not altering the diffuse cytosolic Vinculin signal. The average area and size/puncta were quantitated with the

Image J software. In primary neurons, vinculin immunoreactivity was quantified in dendrites and dendritic spines by subtracting diffuse cytosolic intensity in dendrites, which proves an internally controlled measure of focal vinculin vs. diffuse cytosolic vinculin. All images were acquired with the Olympus FV10i confocal microscope and quantified using the Olympus Fluoview

Software. All comparison images were acquired with identical laser intensity, exposure time, and filter. Adjustments to the brightness/contrast were applied equally to all comparison images.

JC-1, Mitosox-Red, and H2DCF Imaging

HT22 cells seeded in 24-well plates were treated with/without HUTS-4 (1:250),

MAB1965 (1:250), or Nox inhibitor III (40nM) for 45min, then treated with or without A42O

(1M) for 2h. For Mitosox-Red imaging, cells were then washed with PBS and Mitosox-Red added to a final concentration 5M in PBS for 15min at 370C, after which cells were washed with PBS, fixed, and imaged under the Olympus FV10i confocal microscope. For H2DCF imaging, after washing cells in PBS, H2DCF was added at a final concentration of 50nM in

HBSS for 15min at 370C, followed by live cell imaging at 370C with the Nikon Eclipse Ti fluorescence microscope. For JC-1 imaging, after washing cells in PBS, JC-1 was added at a final concentration of 0.5M in PBS and incubated for 15min at 370C, followed by live cell imaging at 370C with the Nikon Eclipse Ti fluorescence microscope. All comparison images were acquired with identical laser or mercury lamp intensity, exposure time, and filter.

Adjustments to the brightness/contrast were applied equally to all comparison images.

Stereotaxic Surgery

Eight-month old APP/PS1 mice were stereotaxically injected with 2l of either control

IgG (control ascites diluted 1:10 in PBS) into the left hemisphere or 1-integrin neutralizing IgG

(MAB1965 ascites diluted 1:10 in PBS) into the right hemisphere of the hippocampus (AP 2.7,

Lat -2.7, DV -3.0) using the convection enhanced delivery method described previously (Carty et al., 2010). Forty-eight hours later, mice were subjected to transcardial perfusion with 4% paraformaldehyde followed by tissue sectioning and immunohistochemistry.

Electrophysiology in Mouse Brain Slices

Hippocampus slices were prepared from 3-month-old wild type (WT), APP/PS1, and

APP/PS1;Cofilin+/- mice and subjected to input/output curved, paired pulse facilitation, and

LTP as previously described (Weeber et al., 2002). Briefly, animals were sacrificed, brains were harvested and sectioned horizontally (400 m) in ice-cold cutting solution (110 mM sucrose, 6 mM NaCl, 3 mM KCl, 26mM NaHCO3, 1.25mM NaH2PO4, 7mM MgCl2, 0.5mM CaCl2, 10g/L

Glucose, pH 7.3–7.4). The hippocampus was dissected and acclimated in 50:50 solution

(cutting:artificial cerebrospinal Fluid (ACSF)) for 10 min at room temperature. Then the slices were transferred to ACSF (125mM NaCl, 2.5mM KCl, 1.25mM NaH2PO4, 0.26mM NaHCO3,

1.2mM MgCl2, 2.0mM CaCl2, and 10g/L Glucose, pH 7.3–7.4, saturated with 95% O2 and 5%

CO2). Slices were recovered in ACSF at room temperature at least 40 min, followed by a final incubation in ACSF for 1 hour at 30°C.

Extracellular field potential recording, LTP: The recording chamber was held at 30 ±

0.5°C with ACSF flow rate of 1 ml/min. The stimulating electrode was placed in the Schaffer collaterals of the hippocampus. The recording glass electrode loaded with ACSF was positioned at the CA1 stratum radiatum below the pyramidal cell layer. Stimulating pulses were generated by the Digidata 1322A interface (Molecular Devices, Sunnyvale, CA) and a stimulus isolator

(model 2200; A-M Systems, Sequim, WA) under control of Clampex 10.0 software (Molecular

Devices). Field excitatory postsynaptic potentials (fEPSP) were amplified using a differential amplifier (model 1800; A-M Systems), filtered at 1 kHz, and digitized at 10 kHz.

Input-output curve was generated by stepping stimulation amplitude from 1 to 15mV.

Stimulation amplitude that elicited half-maximal fEPSP was determined by the Input-output curve and stimulation rate of 0.05Hz was used through the whole experiment. Paired-pulse

facilitation (PPF), which is short-term plasticity, was evoked by two pulses with interpulse intervals from 20ms to 300ms. Percentage of the facilitation was calculated by dividing field excitatory postsynaptic potential (fEPSP) slope elicited by the second pulse with the fEPSP slope elicited by the first pulse. Long-term potentiation (LTP) was induced by theta burst stimulation

(TBS) (5 trains of four pulses at 200Hz separated by 200ms, repeated 6 times with an inter-train interval of 10 sec). LTP was sampled 60 min after the induction, and calculated by dividing the slope of 60 min post-induction responses with the average slope of 20 min baseline responses.

Fear Conditioning, Open Field, and Rotarod Behavior

Fear conditioning (contextual & cued), rotarod, and open field tasks were performed as previously described (Bolognani et al., 2007). For fear conditioning, an aversive stimulus (in this case a mild foot shock, 0.5mA) was paired with an auditory conditioned stimulus (white noise) within a novel environment. Training consisted of two mild shocks paired with two conditioned stimuli with a 3 min interval between each shock. Freezing on the training day in response to the foot shock was used as an estimate of learning during the acquisition trial. To test conditioning to the context, animals were re-introduced to the same training chamber for 6 min and freezing behavior was recording by tracking software (any-maze) every second. To test conditioning to the tone, animals were introduced to a novel context, consisting of a chamber with different shape, floor and olfactory cues from the training chamber. Mice were scored for 3 min, before and after the tone in the same manner described above. Learning was assessed by measuring freezing behavior (i.e. motionless position) every second. The open field was used as a standard test of general activity. Briefly, animals were monitored for 15 minutes in a 40 cm square open field with a video tracking software (Any-maze). General activity levels were evaluated by measurements of total distance traveled and total time immobile. Motor performance was

evaluated by an accelerating rotarod apparatus with a 3cm diameter rod starting at an initial rotation of 4 RPM slowly accelerating to 40 RPM over 5 minutes. The time spent on the rod during each of four trials per day for two consecutive days was measured previously described in

(Brownlow, Benner, D'Agostino, Gordon, & Morgan, 2013)

Statistical analysis & graphs

Statistical data were analyzed by the GraphPad Prizm 6.0 software (GraphPad Software,

San Diego, CA) using Student’s t-test, 1-way or 2-way ANOVA. One-way or 2-way ANOVA was followed by Tukey posthoc test. All quantitative graphs were expressed as mean + SEM.

Differences were deemed significant when P<0.05.

Results

c Aβ42O Induces Loss of Cell Surface β1-Integrin but Not TfR or PrP and Reduces β1- Integrin Activation

We prepared A1-42 oligomers (A42O) as previously characterized (Dahlgren et al.,

2002). These A42O preparations contained SDS-resistant dimers, trimers, and tetramers

(Supplemental Fig. S2-1A). The indicated A42O concentrations heretofore are based on the monomer concentration. To determine whether A42O alters surface levels or activation status of

1-integrin, we plated the hippocampus-derived HT22 mouse cell line in the presence and absence of fibronectin (FN) coating, treated them with A42O for 2 hours and performed cell surface biotinylation experiments. Cell surface 1-integrin normalized to total was significantly reduced by ~60% regardless of FN coating after A42O treatment without altering total 1- integrin, although FN coating also significantly increased surface 1-integrin (Fig. 2-1A,B;

Supplemental Fig. S2-1B,C). While FN coating modestly but significantly increased surface/total transferrin receptor (TfR), A42O (1M) had no significant effect on surface TfR (Fig. 2-1A,C). 37

c Surface PrP was unaffected by either FN coating or A42O treatment (Fig. 2-1A,D). In FN- coated coverslips, A42O (2h) also significantly reduced total surface 1-integrin levels by ~45% and further reduced activated surface 1-integrin by ~75% as detected by the HUTS-4 antibody

(Luque et al., 1996) in non-permeabilized cells (Fig. 2-1E,F). In DIV14 fixed and permeabilized neurons, A42O treatment per se significantly reduced the FITC-HUTS-4-positive activated 1- integrin by >50%, which was significantly prevented by the 1-integrin allosteric modulating antibody (MAB1965)(Aljamal-Naylor et al., 2012) and the PrPc function blocking antibody

(6D11)(Lauren et al., 2009)(Fig. 2-1G,H).

β1-Integrin Allosteric Modulation or Activation Reduces Aβ42O Binding to Neurons, HT22 Cells, and Purified β1-Integrin

We next assessed A42O binding to mature DIV21 hippocampal neurons by adding

0 100nM FITC-A42O for 1h at 37 C with prior incubation (1h) with control IgG, MAB1965, or

HUTS-4 monoclonal IgGs. As expected, FITC-A42O associated with dendrites and dendritic spines, and such binding was significantly reduced by MAB1965 IgG by ~50% (Fig. 2-2A,B).

The HUTS-4 IgG, which has been shown to enhance ligand binding to 1-integrin(Valdramidou et al., 2008), also significantly reduced FITC-A42O binding to neurons by >60% (Fig. 2-2A,B).

As previously shown(Lauren et al., 2009), 6D11 IgG significantly reduced FITC-A42O binding to neurons by ~50% (Fig. 2-2A,B). In 1-integrin fl/fl neurons (conditional knockout, cKO)(Raghavan, Bauer, Mundschau, Li, & Fuchs, 2000) transduced with high-titer lentiviruses expressing Cre-mRFP, FITC-A42O binding to neurons was significantly reduced by >60% (Fig.

2-2C,D) while reducing 1-integrin expression by ~80-90% (Fig. 2-2E). We next determined whether A binds directly to 1-integrin-Fc (extracellular domain fused with human IgG-

Fc).(Valdramidou et al., 2008) Biotin-A42O bound to purified 1-Integrin-Fc (Fig. 2F) coated 38

on ELISA plates with high affinity in the presence of 5mM MgCl2, which induces an intermediate open integrin conformation (Fig. 2-2G, Kd=11.3nM based on monomer calculation).

In the absence of cation, minimal binding of A42O to 1-integrin-Fc was detected and did not appreciably increase with increasing biotin-A42O (Fig. 2-2G). Surprisingly, MnCl2 (0.5mM), which induces the maximal open activation state of integrins(Valdramidou et al., 2008), failed to support A42O binding to 1-integrin-Fc (Fig. 2-2G), indicating that A42O prefers an intermediate activation state of 1-integrin. Accordingly, FITC-A42O binding to HT22 cells was also significantly reduced in the presence of 0.5mM MnCl2 or MAB1965 but not MgCl2

(Fig. 2-2H,I). Unlike A42O, A42 monomers and fibrils bound poorly to 1-integrin-Fc

(Supplemental Fig. S2-2).

Essential Role of β1-Integrin in Aβ42O-induced Cofilin Activation and Reciprocal Requirment of Cofilin in Aβ42O-induced Depletion of Surface β1-Integrin and Focal Complexes

A42O (2h, 1M) treatment to primary cortical neurons significantly induced Cofilin dephosphorylation/activation (Fig. 2-3A,B). However, such Cofilin dephosphorylation was absent in 1-integrin cKO neurons transduced with Lenti-Cre-mRFP (Fig. 2-3A,B). We next determined whether Cofilin per se is also required for the observed A42O-induced depletion of cell surface 1-integrin in HT22 cells. A42O reduced surface 1-integrin, and Cofilin siRNA reduced Cofilin levels (Fig. 2-3C). However, Cofilin siRNA increased surface 1-integrin levels and prevented the A42O-induced reduction in surface 1-integrin (Fig. 2-3C). Quantification of surface 1-integrin and PrPc levels secondary to Cofilin siRNA demonstrated that Cofilin knockdown significantly increases the surface/total 1-integrin ratio without affecting

c surface/total PrP ratio (Fig. 2-3D,E), similar to that observed with A42O treatment.

A direct consequence of 1-integrin internalization is the disruption of Integrin and F-

Actin-associated focal complexes, structurally organized by Talin and Vinculin. We treated

DIV14 cortical neurons or HT22 cells with or without A42O (1M) and subjected them to sequential extraction of the cytosolic fraction on ice with the surfactant saponin, followed by extraction of membrane and cytoskeletal elements with SDS sample buffer (FC enriched fraction). A42O saliently reduced FC-enriched but not cytosolic (cyto) Vinculin and Talin in primary neurons (Supplemental Fig. S2-3A) and HT22 cells (Supplemental Fig. S2-3B,C). To observe focal Vinculin in a different way, we cultured primary hippocampal neurons to DIV21 and treated them with or without A42O for 2 hours followed by staining for Vinculin and F-

Actin. Regardless of A42O treatment, Vinculin nearly completely co-localized with F-Actin, and discrete puncta positive for both Vinculin and F-Actin were seen in dendritic spines (Fig. 2-

3F). However, A42O exposure significantly reduced focal Vinculin immunoreactivity in spine- containing dendrites after subtracting the diffuse cytosolic stain in dendrites (Fig. 2-3F,G). In 7- month wild type mice, Vinculin immunoreactivity in the CA3 pyramidal neurons demonstrated punctate immunoreactivity throughout neuronal cell bodies, indicative of focal complexes, with little diffuse staining. However, in 7-month old APP/PS1 transgenic littermates(Jankowsky et al.,

2004), Vinculin immunoreactivity appeared markedly more diffuse and cytosolic in the CA3 pyramidal neurons (Supplemental Fig. S2-3D)with significant 75% and ~50% reductions in area and size of focal Vinculin, respectively, in APP/PS1 mice compared to wild type littermates

(Supplemental Fig. S2-3E).

A42O treatment (1M) to wild type neurons depleted focal Vinculin and Talin (Fig. 2-

3H). However, the same treatment in littermate Cofilin+/- neurons did not significantly deplete

Vinculin and Talin in the FC enriched fraction compared to wild type (WT) neurons (Fig. 2-

3H,I). Accordingly, Cofilin knockdown also prevented A42O-induced depletion of Focal

Vinculin in HT22 cells (Supplemental Fig. S2-4A,B). Next, we stereotaxically injected 8-month old APP/PS1 mice hippocampi with 2l 1-integrin allosteric modulating / neutralizing antibody

(MAB1965 prediluted 1:10) into the right hemisphere and control IgG (control ascites fluid prediluted 1:10) into the left hemisphere. Forty-eight hours post-injection, MAB1965 IgG significantly increased focal Vinculin area and size in CA3 pyramidal neurons compared to control IgG in the contralateral hemisphere (Fig. 2-3J,K), indicating that endogenous 1-integrin mediates APP/A-induced depletion of focal Vinculin in brain.

Essential Roles of Cofilin and β1-Integrin in Aβ42O-Induced Depletion of F-Actin and F- Actin-Associated Synaptic Proteins in Primary Hippocampal Neurons

The primary function of Cofilin is to sever F-Actin, which promotes synaptic remodeling via accelerating Actin dynamics. (Shirao & Gonzalez-Billault, 2013). As A42O induced Cofilin activation, we cultured hippocampal neurons derived from Cofilin+/- and wild type (WT) littermate mice and cultured them to DIV21. A42O (1M, 2h) significantly depleted F-Actin,

Drebrin, and PSD95 in dendritic spines and spine-containing neurites of WT neurons (Fig. 2-

4A,B). However, the same A42O treatment had no significant effects in Cofilin+/- neurons (Fig.

2-4A,B), indicating that a critical threshold of endogenous Cofilin is required for A42O-induced depletion of F-Actin, Drebrin, and PSD95. Likewise, allosteric modulation of 1-integrin with

MAB1965 IgG completely prevented the depletion of F-Actin and PSD95 induced by A42O

(Fig. 2-4C,D), indicating that the 1-integrin-Cofilin pathway mediates A42O-induced changes in F-Actin and F-Actin-associated postsynaptic proteins.

Requriement of Slingshot-1 (SSH1) in Aβ42O-Induced Cofilin Translocation to Mitochondria and Accumulation of Mitochondrial Cofilin in AD brains

Activated and oxidized Cofilin is known to translocate to mitochondria, where it induces mitochondrial dysfunction (Klamt et al., 2009; Roh et al., 2013a). We treated HT22 cells with or without A42O (1M) for 0, 2, and 4 hours and fractionated intact mitochondria from cytosol.

A42O rapidly increased the translocation of Cofilin to mitochondria, while decreasing phospho-

Cofilin in the cytosol (Fig. 2-5A, B). No phospho-Cofilin was detected in the mitochondrial fraction (Fig. 2-5A), consistent with its activation requirement for translocation. RNAi-mediated knockdown of SSH1, the phosphatase that activates Cofilin(Niwa et al., 2002), significantly prevented A42O-induced translocation of Cofilin to mitochondria together with increased

Cofilin phosphorylation (Fig. 2-5C), indicating that A42O activates and promotes Cofilin translocation to mitochondria via SSH1. We next isolated mitochondria from the frontal cortex of age-matched cognitively normal control (NC) and AD patients (Supplemental Fig S2-5). In the absence of DTT in sample buffer (non-reducing), we detected cysteine oxidized Cofilin monomers, dimers, trimers, and higher-order oligomers, which essentially all collapsed to the monomer form in the presence of DTT (reducing) (Fig. 2-5D). This banding pattern is similar to that observed from Cofilin-Actin rods, which also require activation and oxidation-induced disulfide bonding to form(B. W. Bernstein et al., 2012). Total Cofilin (all 4 bands in non- reducing condition), oxidized Cofilin oligomers (dimers or larger), and Cofilin monomers were significantly increased in mitochondria of AD (Fig. 2-4D,E), results that fulfill the prediction from cultured cells and suggest that activated and oxidized mitochondrial Cofilin plays a pathogenic role in AD.

Aβ42O-Induced ROS productions, Mitochondrial Dysfunction, and Apoptosis via the 1- Integrin-SSH1-Cofilin Activation Pathway 

HT22 cells were treated with A42O (1M) with or without vehicle, NOX inhibitor,

HUTS-4 IgG, or MAB1965 IgG and then subjected to live cell imaging for JC-1, Mitosox-Red, and H2DCF. As expected, A42O treatment (1M, 2h) significantly decreased mitochondrial membrane potential (increased JC-1 green/red ratio), increased mitochondrial superoxide

(Mitosox-Red), and increased ROS levels (H2DCF) (Fig. 2-6A,B; Supplemental Fig. S2-6A).

However, co-incubation with HUTS-4, MAB1965, or NOX inhibitor significantly prevented such changes induced by A42O, although the JC-1 membrane potential measure was not fully restored by these treatments (Fig. 2-6A,B; Supplemental Fig. 2-S6A). Accordingly, MAB1965 significantly prevented A42O-induced apoptosis in primary neurons and HT22 cells as assessed by Annexin V/PI staining (Supplemental Fig. S2-6B-D). Similarly, Annexin V/PI staining also demonstrated significant induction of early (Annexin V) and late (PI) apoptosis after 24h A42O

(1M) treatment, which was nearly completely blocked by SSH1 siRNA knockdown (Fig. 2-

6C,D), indicating that SSH1-dependent Cofilin activation is required for A42O-induced apoptosis.

A key mechanism of Cofilin regulation is via the binding of SSH1 to 14-3-3, which renders SSH1 sequestered in an inactive state. However, the oxidation of 14-3-3 releases SSH1, thereby leading to SSH1 activation (Eiseler et al., 2009a; J. S. Kim et al., 2009). In 7-month old

APP/PS1 mice, SSH1/Cofilin complexes and 14-3-3/SSH1 complexes were significantly increased and decreased, respectively, compared to wild type littermate mice (Fig. 2-6E,F), indicating the release of SSH1 from 14-3-3 and association of SSH1 with Cofilin in APP/PS1 mice. Accordingly, phospho-Cofilin was significantly decreased in APP/PS1 mice, despite no apparent changes in total 14-3-3, SSH1, or Cofilin in TX-100 solubilized extracts (Fig. 2-6E,F). 43

A42O treatment also reduced 14-3-3/SSH1 complexes in HT22 cells, which was largely restored with prior treatment with MAB1965 or the potent anti-oxidant Trolox (Supplemental

Fig. S2-7B). Likewise, A42O treatment significantly increased SSH1/Cofilin complexes in

HT22 cells (Supplemental Fig. S2-7C,D). These results therefore demonstrate that APP/A promotes Cofilin activation at least in part via the 1-integrin-ROS-14-3-3/SSH1 pathway.

Cofilin Reduction Rescues APP/Aβ-Induced Gliosis and Loss of Synaptic Proteins as well as LTP and Contextual Memory Deficits in APP/PS1 mice

We examined brains of 7-month old APP/PS1, APP/PS1;Cofilin+/-, and WT littermate mice for neurodegenerative phenotypes, including GFAP for astrogliosis as well as PSD95

(postsynaptic) and Synapsin I (presynaptic) for synaptic integrity. As expected, APP/PS1 mouse hippocampi demonstrated significantly increased GFAP immunoreactivity throughout the hippocampus as well as diminished Synapsin I and PSD95 immunoreactivity within the stratum lucidium (SL: synaptic terminating zone) of CA3 (Fig. 2-7A,B). In contrast, APP/PS1;Cofilin+/- mice showed significantly reduced GFAP immunoreactivity and increased Synapsin I and

PSD95 immunoreactivities compared to APP/PS1 littermate mice, essentially indistinguishable from wild type littermates (Fig. 2-7A,B).

We next tested short-term and long-term synaptic plasticity from acute hippocampal slices prepared from 3-month old wild type (WT), APP/PS1, and APP/PS1;Cofilin+/- mice. The stimulating electrode was placed in the Schaffer collaterals of the hippocampus, and the recording electrode was positioned at the CA1 stratum radiatum below the pyramidal cell layer.

As shown in Fig. 2-7C, the input-output curves did not markedly differ among WT, APP/PS1, and APP/PS1;Cofilin+/- slices. In paired pulse facilitation (PPF) experiments, we observed significant differences in fEPSP slope across genotypes among all interstimulus intervals (Fig. 2-

7D). However, correction for multiple comparisons showed that only APP/PS1;Cofilin+/- and

WT slices differed significantly at the 40ms interstimulus interval (Fig. 2-7D). For long-term potentiation measurements, we detected no differences in fEPSP slope among WT, APP/PS1, and APP/PS1;Cofilin+/- slices at baseline (Fig. 2-7E). However after theta burst stimulation, we observed significant differences in fEPSP slope across genotypes for all time points (Fig. 7E).

Correction for multiple comparisons showed that APP/PS1 slices were significantly impaired in fEPSP slope compared to WT and APP/PS1;Cofliln+/- slices at every time point up to 1 hour, indicating that Cofilin reduction rescues the deficits in LTP in APP/PS1 mice.

We carried out fear conditioning tests (contextual & cued), in which 7-month old WT,

APP/PS1, and APP/PS1;Cofilin+/- mice were trained on day 1 for both. On day 2, we tested mice for contextual and cued conditioning memory. We observed no significant differences between the genotypes in percent time spent freezing during the training phase (Fig. 2-7F).

However, we observed significant differences across genotypes in the percentage of time spent freezing during the hippocampus-dependent contextual fear conditioning test on day 2.

Correction for multiple comparisons showed that wild type and APP/PS1;Cofilin+/- mice significantly spend more time freezing than APP/PS1 mice (Fig. 2-7G). On the hippocampus- independent cued fear conditioning test, we did not observe significant differences in the percentage of time spent freezing times across genotypes (Fig. 2-7H). We also did not find significant differences among the 3 genotypes in the rotarod or open field (total distance traveled

& total time spent immobile) tests (Fig. 2-7I-K), indicating the absence of salient perturbations in generalized motor activity or coordination. Taken together, these results demonstrate that endogenous Cofilin is required to mediate the APP/PS1-induced deficits in synaptic plasticity as well as hippocampus-dependent contextual memory but not cued memory.

Discussion

Molecular pathways that govern the production and neurotoxicity of Aβ are attractive therapeutic targets for AD. While previous studies have shown Cofilin activation in AD brains and potential involvement of 1-integrin in A-induced apoptosis (T. Kim et al., 2013;

Minamide et al., 2000; Q. Wang et al., 2008b; Wright et al., 2007b), it was unknown whether endogenous Cofilin per se and different 1-integrin conformers are required for molecular events leading to A42O-induced neurodegeneration and how the 1-integrin-Cofilin signaling pathway mediates AD-relevant pathogenic processes.

In this study, we demonstrated for the first time that A42O induces the depletion of surface 1-integrin via its direct binding to low-intermediate activation conformers of 1- integrin, as detected by competition of A42O binding to neurons and HT22 cells with the 1- integrin activating antibody (HUTS-4) or MnCl2, both of which induce high conformational activation of 1-integrin (Luque et al., 1996; Valdramidou et al., 2008). Moreover, MnCl2 essentially eliminated Aβ42O binding to purified β1-integrin-Fc. However, A42O did not bind to

1-integrin-Fc in the absence of Mg2+, suggesting that an intermediate / partially open conformation of integrin is required for binding. While physiological ligands bind with high affinity to the highly active/open conformation of 1-integrin (Valdramidou et al., 2008), the human Ecovirus binds to the relatively inactive/closed integrin conformer (Jokinen et al., 2010), similar to what we observed with A42O. This finding raises the intriguing possibility of blocking A42O-induced neurotoxic actions by conformational 1-integrin activation without compromising integrin function. In addition to preferentially binding to low-intermediate activation 1-integrin conformers, A42O also increased the propensity of 1-integrin to assume a lower-activation conformation, probably by preventing bound 1-integrin from assuming a 46

high-activation state. As A42O reduced surface 1-integrin levels regardless of FN plating,

A42O likely does not compete with most physiological ligands (i.e. FN) for integrin binding, as seen by its preference for binding to lower-activation conformation of 1-integrin. Although we focused on 1-integrin in this study, it is well known that 1-integrin functions in concert with various -integrin subunits, such as 2 and αv, which have been shown to be involved in A neurotoxicity(Wright et al., 2007b), as well as α6, which has been implicated in microglial phagocytosis of fibrillar A together with β1(Koenigsknecht & Landreth, 2004).

c Our observation that the PrP blocking antibody (6D11) significantly prevented Aβ42O- induced reduction in β1-integrin activation was somewhat surprising. However, it has been

c reported that PrP and Pir-B, both of which bind Aβ42O, negatively regulate β1-integrin activation and signaling(Loubet et al., 2012; Pereira, Zhang, Takai, & Lowell, 2004), raising the possibility that loss of these receptors can also interfere with Aβ42O binding to β1-integrin in neurons. These receptors, including β1-integrin are typically clustered in pre- and postsynaptic membranes, and their activities are regulated by crosstalk at multiple points of convergence by other transmembrane receptors. For example, PrPc-dependent activation of Fyn requires Integrin engagement (Pantera et al., 2009), and glutamate signaling via NMDARs and AMPARs depends on various Integrin heterodimers (Becchetti et al., 2010). Indeed, β1-Integrin is also enriched in dendritic spines and is required for dendrite arborization, maintenance of synapses, and

LTP(Babayan et al., 2012; Bernard-Trifilo et al., 2005; Chan, Weeber, Kurup, Sweatt, & Davis,

2003; Warren et al., 2012). In view of these observations, it is plausible that Integrins operate physically and/or functionally in concert with other Aβ42O-binding receptors to mediate Aβ42O binding and neurotoxicity.

Deletion of β1-integrin resulted in the failure of Aβ42O to activate Cofilin. At the same time, Cofilin was also required for removal of surface β1-integrin, indicating that Cofilin normally plays a role in the trafficking and/or internalization of β1-integrin. As such, we focused on β1-integrin and F-Actin-associated structural focal components (Talin/Vinculin) and postsynaptic proteins known to function in concert with F-Actin and synaptic remodeling, because the direct consequence of β1-integrin internalization is the disruption of the structural components of focal complexes (Talin & Vinculin), which anchor the cytoplasmic tail of β1- integrin to F-Actin. Thus, it is likely that the F-Actin severing activity of Cofilin is responsible for the removal β1-integrin from the cell surface and the depletion of focal Talin/Vinculin by destabilizing or reducing the F-Actin linkage to β1-integrin, changes which were directly analogous to F-Action per se and F-Actin-regulated postsynaptic proteins (Drebrin & PSD95).

Cofilin activity is associated with dendritic spine growth and shrinkage during LTP and

LTD, respectively (Chen, Rex, Casale, Gall, & Lynch, 2007; Rex et al., 2009; Zhou, Homma, &

Poo, 2004), and Cofilin regulates the trafficking of AMPA receptors in dendritic spines(Gu et al.,

2010). Therefore, Aβ42O-induced Cofilin activation and resultant F-Actin severing/depolymerization also likely underlies the reduction in F-Actin and synaptic proteins

(PSD95 & Drebrin), both of which could be antagonized by Cofilin reduction or β1-integrin allosteric modulation. β-Arrestin 2 recruits Cofilin to dendritic spines upon NMDA receptor activation to control the remodeling of spines, and β-Arrestin 2 knockout neurons are resistant to

Aβ-induced dendritic spine loss through spatial control of Cofilin activation(Pontrello et al.,

2012). Likewise, Cofilin+/- neurons were resistant to Aβ-induced depletion of postsynaptic proteins. In view of these observations, it is plausible that Cofilin reduction prevents the

sustained over-activation of Cofilin by Aβ oligomers to mitigate the loss of postsynaptic proteins and the deficits in LTP.

Our findings indicated that Aβ42O-induced and β1-dependent Cofilin activation, mitochondrial dysfunction, and apoptosis require endogenous SSH1 at least in part via ROS and

14-3-3. This is consistent with the induction of ROS production by NADPH oxidases (NOX) via

Rac1-mediated signaling downstream of integrin(Werner & Werb, 2002), Aβ42O–induced

Cofilin-Actin pathology via a NOX/PrPc-dependent manner(K. P. Walsh et al., 2014), activation of SSH1 by oxidation of 14-3-3(J. S. Kim et al., 2009), Aβ-induced oxidation of 14-3-3(Boyd-

Kimball et al., 2005; Sultana et al., 2006), and Cofilin-dependent mitochondrial dysfunction downstream of oxidative stress(Roh et al., 2013a). Although a recent study reported that fibrillar

Aβ42 can activate LIMK1 via PAK1, they also observed increased Cofilin dephosphorylation; therefore, Aβ oligomers and fibrils may impact the Cofilin activation pathway via similar but also divergent mechanisms(Mendoza-Naranjo et al., 2012). In addition to ROS-dependent activation of SSH1, it is likely that Aβ42O can also activate Cofilin via a mechanism involving an increase in cytosolic Ca2+, which in part may be mediated by the PrPc-mGluR5 complex(Um et al., 2013), as Calcineurin dephosphorylates and activates SSH1(Y. Wang et al., 2005a). We have also shown that RanBP9, which activates Cofilin, also impairs mitochondrial Ca2+ buffering and mitochondrial homeostasis in a Cofilin-dependent manner (Liu, Roh, Woo, Ryu, & Kang,

2013; Roh et al., 2013a; J. A. Woo et al., 2012).

Conclusion

Our findings indicate that a molecular pathway involving SSH1-Cofilin activation via direct engagement of lower-activation conformers of 1-integrin is essential for both A42O- induced mitochondrial and synaptic dysfunction in AD. Therefore, promoting 1-integrin

activation and/or inhibiting excessive Cofilin activation (i.e. SSH1 inhibition) represent potentially viable and novel therapeutic strategies to combat AD pathogenesis.

Figure 2-1. Aβ42 oligomers induces loss of cell surface β1-integrin but not TfR or PrPc and reduces β1- integrin activation a-d) HT22 cells plated with/without Fibronectin with/without Aβ42 oligomers (1µM, 2hours), subjected to surface biotinylation (surface), and direct immunoblotting (total) or immunoprecipitation for biotin (surface) and immunoblotting for the indicated protein. b-d) Quantification of normalized surface/total β1, TfR, and PrPc (n>3 replicates, ANOVA, post hoc Tukey, *p<0.05, #p<0.005). Error bar represent S.E.M. e & f) HT22 cells treated with/without Aβ42 oligomers and immunostained for total surface β1-integrin or activated surface β1-integrin without membrane permeabilization (n>4 replicates, T-test, **P<0.005, #P<0.005). g & h) Direct staining for activated β1-integrin (FITC-HUTS-4 antibody) in fixed and permeabilized DIV14 primary hippocampal neurons after treatment with/without Aβ42 oligomers (1µM, 2 hours) and/or control ascites IgG (1:250), MAB1965 (1:250), or 6D11 (1:50). H) Quantification of FITC-HUTS-4 immunoreactivity (n=4 replicates, ANOVA, post hocTukey,*P<0.05, **P<0.005). 51

Figure 2-2. β1-Integrin allosteric modulation or activation reduces Aβ42 oligomers binding to neurons, HT22 cells, and purified β1-integrin as effectively as the loss of β1-integrin a & b) FITC-Aβ42O (100 nM, 1 hour) binding to DIV21 primary hippocampal neurons with/without prior treatment (1 h) of control IgG, MAB1965 (β1-integrin allosteric modulator, 1:250), HUTS-4 (β1-integrin allosteric activator, 1:250), and/or 6D11 (PrPc blocking, 1:50) monoclonal antibodies. Bottom panel magnified from regions of white rectangles. b) Quantification of FITC-Aβ42 oligomers binding (n>5 replicates, ANOVA, post hoc Tukey, #P<0.0005 compared with control IgG). Error bars represent S.E.M. c-e) FITC-Aβ42 oligomers (100 nM, 1 hour) binding to β1-integrin fl/fl neurons transduced with Lenti-mRFP (cont) or Lenti-Cre-mRFP (β1 cKO). d) Quantification of FITC-Aβ42O binding (n= 5 replicates,T-test, #P<0.0005). e) Representative blots of β1-integrin from cKO (Lenti-Cre-mRFP) versus cont (Lenti-mRFP) neurons (all neurons including nontransduced). f) Conditioned medium from CHO cells transfected with β1-integrin-Fc affinity purified with anti-Fc affinity column, sequentially eluted, and immunoblotted with anti-Fc. g) Purified β1-integrin-Fc from fractions 1 and 2 captured by 96- well plates coated with anti-Fc IgG, subjected to biotin-Aβ42 oligomers binding and/or incubated with different cations (MgCl2, 5mM and MnCl2, 0.5mM). h) Kd= 10.3 nM with MgCl2. Two-way ANOVA, post hoc Bonferroni, n=4 replicates, #P<0.0005. h & i) HT22 cells treated with FITC-Aβ42 oligomers (0.1μM, 2 hours) and/or β1-integrin blocking IgG (MAB1965, 1:500), MnCl2 (0.5mM), or MgCl2 (5mM), and subjected fluorescence microscopy. (h) Representative images of FITC-Aβ42 oligomers binding/uptake show inhibition by MAB1965 antibody and MnCl2. i) Quantification of mean intensities of FITC-Aβ42 oligomers binding/uptake (n= 4 replicates, ANOVA, post hoc Tukey, **P<0.005 compared with FITC-Aβ42 oligomers alone).

Figure 2-3. Essential role of β1-integrin in Aβ42 oligomer-induced Cofilin activation as well as reciprocal requirement of Cofilin in Aβ42 oligomer-induced depletion of surface β1-integrin and F-actin-associated FCs (Vinculin and Talin) a & b) DIV14 cortical WT or β1-integrin cKO neurons (transduced with Lenti-Cre) treated with/without Aβ42 oligomers (1μM, 2 hours) and subjected to immunoblotting. Note the decrease in p-Cofilin by Aβ42 oligomers in WT but not in β1 cKO neurons (n=4 replicates, T-test, *P<0.05). Error bars represent S.E.M. c) HT22 cells transfected with control or Cofilin siRNA, treated with Aβ42 oligomers (1 μM, 2 hours) and subjected cell surface biotinylation followed by direct immunoblotting or IP with anti- biotin and immunoblotting. Note that Cofilin knockdown prevents Aβ42 oligomers-induced depletion of surface β1-integrin. d & e) HT22 cells transfected with control or Cofilin siRNA and subjected to cell surface biotinylation followed by direct immunoblotting or IP with anti-biotin and immunoblotting. d) A representative experiment showing a dramatic increase in surfaceβ1-integrin but no change in surface PrPc after Cofilin knockdown. e) Quantification of β1-integrin normalized to total β1- integrin and surface PrPc normalized to total PrPc (n=4 replicates, T-test, #P<0.0005). f&g) DIV21 hippocampal primary neurons treated with/without Aβ42 oligomers (1μM, 2 hours) and subjected to staining for Vinculin, F-actin (rhodamine-phalloidin), and DAPI. g) Quantification of focal Vinculin mean intensity (FC Vinculin) in dendrites and dendritic spines after subtracting the mean diffuse cytosolic intensity in dendrites (n=5 replicates,T-test, #P<0.0005). Error bars represent S.E.M. h & i) DIV14 cortical +/− WT and Cofilin neurons treated with/without Aβ42 oligomers (1μM), subjected to separation of cytosol and FC-enriched +/− fractions followed by immunoblotting. h) Aβ42 oligomers-induced depletion FC Vinculin and Talin in WT but not in Cofilin neurons. i) Quantification of FC Vinculin and Talin after 2 hours Aβ42 oligomers treatment normalized to respective vehicle treated controls at 100%. (n=3 replicates, T-test,*P<0.05,**P<0.01). j&k) Eight-month-old APP/PS1 transgenic mice and their nontransgenic littermates (WT) stereotaxically injected with 2 μl β1-integrin blocking antibody (MAB1965, diluted 1 : 10) into the left hippocampus and 2 μl control IgG (control ascites fluid, diluted 1 : 10) into the right hippocampus and subjected to immunohistochemistry for Vinculin 48 hours post injection. j) Representative images of Vinculin FCs (red) in the CA3 of the hemisphere injected with β1-integrin blocking antibody compared with the control IgG injected contralateral hemisphere.. k) Quantification of focal Vinculin area and size normalized to control IgG (n=5 mice, 2 M and 3 F,T-test, **P<0.005).

Figure 2-4. Essential roles of Cofilin and β1-integrin in Aβ42 oligomer-induced depletion of F-actin and associated synaptic proteins in primary hippocampal neurons a & b) WT and Cofilin+/− DIV21 hippocampal neurons treated with/without Aβ42 oligomers (1 μM, 2 hours) and stained for F- actin (rhodamine-phalloidin), Drebrin, or PSD95. b) Quantification of mean intensities of Drebrin, PSD95, and F-actin in spine- containing neurites (n =4–6 replicates, ANOVA, post hoc Tukey, *P<0.05, **P<0.005, #P<0.0005). Error bars represent S.E.M. c & d) DIV21 hippocampal neurons treated with/without Aβ42 oligomers (1 μM, 2 h) and/or control IgG or β1-integrin IgG (MAB1965) and stained for PSD95 and F-actin (rhodamine-phalloidin). c) Representative images showing depletion of PSD95 and F-actin after Aβ42 oligomers treatment, which is prevented by MAB1965. d) Quantification of mean PSD95 and F-actin intensities in spine-containing neurites (n>6 replicates, ANOVA, post hoc Tukey, *P<0.05, **P<0.005)

Figure 2-5. Requirement of Slingshot-1 (SSH1) in Aβ42 oligomer-induced Cofilin translocation to mitochondria and accumulation of mitochondrial Cofilin in AD brains. a & b) HT22 cells treated with/without Aβ42 oligomers (1 μM) for the indicated times and subjected to separation of mitochondria and cytosol fractions followed by immunoblotting for the indicated proteins. a) Representative blots showing Cofilin dephosphorylation (cytosol) and translocation to mitochondria (mito) after Aβ42 oligomers treatment. b) Quantification of cytosolic and mitochondrial Cofilin (n=4 replicates, T-test, **P<0.005, #P<0.0005). c) HT22 cells transfected with control or SSH1 siRNA, treated with/without Aβ42 oligomers (1 μM, 4 hours), and subjected to separation of cytosol versus mitochondria followed by immunoblotting for the indicated proteins. Note that SSH1 siRNA prevents Aβ42 oligomers-induced Cofilin dephosphorylation and translocation to mitochondria (n=4 replicates, ANOVA, post hoc Tukey, #P<0.0005). d & e) Mitochondria isolated from mid-frontal cortex of AD (n=5) and cognitively NC (n=4) and immunoblotted for Cofilin and Timm50 with or without DTT in sample buffer. d) Representative blots showing abundance of Cofilin monomers (CofM), dimers (CofDi), trimers (CofTri), and high molecular weight species (CofH) in AD (blot on left run without DTT). Samples with asterisk (*) run with DTT and boiling show reduction of oxidized Cofilin oligomers to collapsed monomers (blot on right). e) Quantification of Cofilin total (all four Cofilin bands), oligomers (CofDi, CofTri, and CofH), and monomers (CofM).

Figure 2-6 Aβ42 oligomer-induced ROS production, mitochondrial dysfunction, and apoptosis via the β1- integrin–SSH1–Cofilin activation pathway a & b) HT22 cells treated with vehicle or Aβ42 oligomers (1 μM) for 2 hours with or without 1 hour prior treatment with Nox inhibitor (VAS2870, 40 nM), HUTS-4 (β1-integrin allosteric activating IgG, 1:250), or MAB1965 (β1-integrin allosteric modulating IgG, 1:250) and subjected to live-cell imaging for JC-1 (mitochondrial membrane potential, green/red ratio), Mitosox-Red (mitochondrial superoxide indicator), or H2DCF (general ROS indicator). Note that β1-integrin conformational modulation or inhibition of Nox significantly reduces Aβ42 oligomers-induced mitochondrial dysfunction and overall ROS increase. b) Quantification of JC-1 green/red ratio, Mitosox-Red, and H2DCF intensities (n=6, 1-way ANOVA, post hoc Tukey, *P<0.05, **P<0.01, or #P<0.005 compared with Aβ42 oligomers. c & d) HT22 cells transiently transfected with control or SSH1 siRNA, treated with / without Aβ42 oligomers (1 μM, 18 hours) in 1% FBS medium, and subjected to FITC Annexin V/propidium iodide (PI) staining and fluorescence confocal microscopy. c) Representative images showing reduction in Annexin V (early apoptosis) and PI (late apoptosis) staining in cells transfected with SSH1 siRNA (with Aβ42 oligomers). d) Quantification of Annexin V/PI staining (n=4 replicates, ANOVA, post hoc Tukey, *P<0.05, **P<0.005, #P<0.0005). e & f) Seven-month-old WT and APP/PS1 littermate mouse frontal cortex homogenates (1% Triton X-100) immunoprecipitated for pan-14-3-3 or SSH1 and immunoblotted for SSH1 or Cofilin. Same brain samples also directly immunoblotted for the indicated proteins. e) Representative blots showing decreased 14-3-3/SSH1 complex and increased SSH1/Cofilin complex in APP/PS1 mice. f) Quantification of SSH1/14-3-3 complex, Cofilin/SSH1 complex, p-Cofilin, and total Cofilin (n=4 mice/genotype, T-test, *P<0.02, **P<0.005)

Figure 2-7 Cofilin reduction rescues APP/Aβ-induced gliosis and loss of synaptic proteins as well as LTP and contextual memory deficits in APP/PS1 mice a & b) Seven-month-old WT, APP/PS1, and APP/PS1;Cofilin+/− mice immunostained for GFAP, Synapsin I, and PSD95. a) Representative images showing that Cofilin reduction ameliorates astrogliosis and synaptic damage associated with APP/PS1 mice. b-d) Quantification of mean PSD95 and Synapsin I intensities in the stratum lucidum (SL) (n=4 mice/genotype, ANOVA, post hoc Tukey, *P<0.05, #P<0.0005). c-e) Stimulating electrode placed in the Schaffer collaterals of the hippocampus and recording glass electrode positioned at the CA1 stratum radiatum below the pyramidal cell layer. c) Input/output analysis generated by stepping up stimulation amplitude from 1 to 15mV in WT, APP/PS1, and APP/PS1;Cofilin+/− acute slices. No significant differences observed (n= 24 slices from four mice, APP/PS1: 19 slices from four mice, APP/PS1;Cofilin+/−: n=20 slices from three mice). d) PPF showing no significant differences across genotypes and interstimulus interval except between APP/PS1;Cofilin+/− and WT slices at the 40-ms interstimulus interval (two-way ANOVA, post hoc Bonferroni, *P<0.05; WT: n=32 slices from four mice, APP/PS1: n= 31 slices from four mice, APP/PS1;Cofilin+/−:n=25 slices from three mice). e) LTP induced by the TBS showing significant differences in fEPSP slope in APP/PS1 compared with WT and APP/PS1;Cofilin+/− slices (two-way ANOVA, post hoc Bonferroni, P<0.0001 at all-time points). (WT: n= 28 slices from four mice, APP/PS1: n =33 slices from four mice, APP/PS1;Cofilin+/−: n=20 slices from three mice). Error bars represent S.E.M. f) Percentage of time spent freezing during training period on day 1 (no significant differences observed by one-way ANOVA or Kruskal–Wallis statistic; WT n=12, APP/PS1 n= 8, APP/PS1;Cofilin+/− n=6; equal distribution of gender). Error bars represent S.E.M. (2-7G) Percentage of time spend freezing during contextual fear conditioning (FC) on day 2 (Kruskal–Wallis statistic=9.66, P= 0.008, genotypes= 3, values=26; post hoc Dunn’s,*P<0.05; WT n=12, APP/PS1 n=8, APP/PS1;Cofilin+/ − n=6; equalized distribution of gender). h) Percentage of time spent freezing during cued fear conditioning (FC) freezing on day 2 (no significant differences observed by one-way ANOVA or Kruskal–Wallis statistic; WT n=12, APP/PS1 n =8, APP/PS1;Cofilin+/− n=6; equalized distribution of gender). i) Total time spent on rotarod test (no significant differences observed by one-way ANOVA or Kruskal–Wallis statistic; WT n= 11, APP/PS1 n=7, APP/PS1;Cofilin+/− n= 5; equalized distribution of gender). j) Total distance traveled during open- field test (no significant differences observed by one-way ANOVA or Kruskal–Wallis statistic; WT n=11, APP/PS1 n=7, APP/PS1;Cofilin+/− n= 5; equalized distribution of gender). k) Total time spent immobile during open-field test (no significant differences observed by 1-way ANOVA or Kruskal–Wallis statistic; WT n=11, APP/PS1 n= 7, APP/PS1;Cofilin+/− n=5; equalized distribution of gender)

Supplementary Figure 2-1 A42O Induces Loss of Cell Surface 1-Integrin

A) Immunoblotting of unconjugated and FITC-conjugated A42M (freshly solubilized monomer) and A42O (oligomer) preparations. Note that A42O run as monomers, dimers, trimers, and tetramers on SDS-PAGE. B&C) HT22 cells (without FN coating) treated with or without or without A42O (1M) for 2h and subjected to cell surface biotinylation and detection of surface 1-integrin (upper panel). Total 1-integrin without immunoprecipitation (lower panel). B) Quantitation of surface 1-integrin normalized to total 1-integrin (n=5 replicates, ANOVA, posthoc Tukey, **P<0.005). Error bars represent SEM.

Supplementary Figure 2-2 A42 Oligomers but not Monomers or Fibrils Bind to Purified 1-integrin-Fc

Purified 1-integrin-Fc from fractions 1 & 2 (Fig. 2F) captured by 96-well plates coated with anti-Fc IgG, subjected to binding to biotin-A42O (oligomer), biotin-A42M (monomer), or biotin-A42F (fibril) in the presence of 5mM MgCl2. 2-way ANOVA posthoc Bonferroni, n=4 replicates, #P<0.0005.

Supplementary Figure 2-3 A42o Disrupt Integrin-dependent Focal Complexes in HT22 Cells, Primary Neurons, and APP/PS1 Mouse Brains

A) DIV14 primary cortical neurons treated with or without 1M A42o for 2h, subjected to sequential extractions with saponin (cytosol) and 2x SDS sample buffer (FC), followed by immunoblotting for the indicated proteins. B,C) HT22 cells treated with A42o (0, 0.25, & 2M) for 0, 2, or 4h, and subjected to sequential extraction with saponin (cytosol) and lysis with 2x SDS sample buffer (FC), followed by immunoblotting for the indicated proteins. C) Quantification of FC Vinculin (n=3 replicates, ANOVA, posthoc Tukey, *P<0.05, **P<0.005 compared to control). F,G) Brains from 7-month old APP/PS1 mice and their wild type littermates subjected to immunohistochemistry for Vinculin. F) Representative images from the CA3 region of the hippocampus. Note that Vinculin appears as focal complexes (red) with little cytoplasmic staining in nontransgenic (WT) mice, whereas Vinculin focal complexes and diffuse cytoplasmic staining are decreased and increased, respectively, in APP/PS1 mice. Focal Vinculin signal is threshold adjusted to convert highest intensity signals from green to red puncta without altering diffuse cytosolic signal. G) Quantification of area covered by focal Vinculin and average size of focal Vinculin in the CA3 of APP/PS1 and WT mice (n=4 mice/genotype, 2F & 2M, T-test, *P<0.05).

Supplementary Figure 2-4 Cofilin Mediates A42O-induced Depletion of Focal Vinculin in HT22 Cells

A,B) HT22 cells transfected with control or Cofilin siRNA, treated with A42o (1M, 2h) and subjected to sequential extractions with saponin (cytosol) and 2x SDS sample buffer (FC), followed by immunoblotting for the indicated proteins. B) Quantification showing Cofilin knockdown abrogates A42o-induced depletion of FA Vinculin (n=4 replicates, ANOVA, posthoc Tukey, **P<0.005 compared to control).

Supplementary Figure 2-5 AD and Control Subject Information

Information on gender, ages, and post-mortem intervals (PMI) of cognitively normal controls (NC) and AD brain samples (mid- frontal gyrus). All NC and AD cases are of Caucasian ethnicity.

Supplementary Figure 2-6 Allosteric Modulation of 1-Integrin Rescues A42O-induced Apoptosis in Primary Neurons and HT22 Cells

A) HT22 cells HT22 cells treated with vehicle or A42O (1M) for 2h with or without 1h prior treatment with Nox inhibitor (VAS2870, 40nM), HUTS-4 (1-integrin allosteric activating IgG, 1:250), or MAB1965 (1-integrin allosteric modulating IgG, 1:250) and subjected to live cell imaging for H2DCF (general ROS indicator). Representative images show that 1-integrin conformational modulation or inhibition of Nox reduces A42O-induced overall ROS increase. Quantifications shown in main manuscript Fig. 6B. B,C) DIV14 hippocampal primary neurons treated with or without 1M A42O and 1-integrin allosteric modulating and neutralizing IgG (MAB1965; 1:500) for 18 hours under regular culture condition, followed by Annexin V/PI staining on glass coverslips. B) Quantification of Annexin V-positive (early apoptotic) and Annexin V/PI-positive (late apoptotic) neurons (n=4 replicates, ANOVA, posthoc Tukey, *P<0.05). D) HT22 cells serum starved (1% FBS) for 24h and treated with or without A42O (1M) and/or allosteric 1-integrin modulating / neutralizing IgG (MAB1965, 1:500) for 24h. Cells were then subjected to propidium iodide (PI) and Annexin V staining followed by FACS analysis. Lower right (Annexin V+) and upper right quadrant (Annexin V/PI+) indicate early apoptotic and late apoptotic cells, respectively. A representative experiment from 3 independent experiments is shown.

Supplementary Figure 2-7 A42O Reduces 14-3-3/SSH1 Complex and Increases SSH1/Cofilin Complex in HT22 Cells

A) HT22 cell lysates immunoprecipitated with anti-mouse IgG agarose beads alone or together with pan-14-3-3 monoclonal antibody and subjected to immunoblotting for SSH1. Note that SSH1 co-immunoprecipitates with 14-3-3 but not with anti-mouse IgG beads alone. B) HT22 cells treated with/without A42 (1M, 2h) and with/without prior treatment (30min) with vehicle O (DMSO), MAB1965 (1:250), or the antioxidant Trolox (200M). Lysates subjected to direct immunoblotting or IP with pan-14- 3-3 antibody followed by immunoblotting. Note that A42O reduces the 14-3-3/SSH1 complex, which is prevented by MAB1965 IgG and Trolox. C&D) HT22 cells treated with/without A42 (1M, 2h) and lysates subjected to IP with anti-rabbit agarose O beads alone or together with anti-SSH1 antibody. Note that Cofilin co-IPs with SSH1 specifically, and the Cofilin/SSH1 complex is increased with A42O treatment. D) Quantification of SSH1/Cofilin complex (n=3 replicates, T-test, **P<0.005).

Supplementary Figure 2-8 Reduction in Cofilin Protein in the Hippocampus and Cortex of Cofilin+/- Mice with No Apparent Abnormarmalities in Gliosis or Synaptic Proteins.

A) Brains from 6-month old Cofilin+/- and littermate mice were dissected and subjected to immunoblotting for Cofilin and Actin from hippocampal (HIPPO) and cortical (CTX) brain extracts. B) Seven-month old WT and Cofilin+/- littermate mice subjected to immunohistochemistry for GFAP, PSD95, and Synapsin I. Representative images show no apparent abnormalities in gliosis or synaptic integrity.

CHAPTER 3

RANBP9 AT THE INTERSECTION BETWEEN COFILIN AND AMYLOID BETA

PATHOLOGIES

Permissions Statement

The information in chapter 3, “RanBP9 at the intersection between Cofilin and Amyloid

β pathologies” has been legally reproduced under the Creative Commons Attribution (CC-BY) license. This means the publication is accessible online without any restrictions and can be re- used in any way subject only to proper citation.

Woo, J. A., Boggess, T., Uhlar, C., Wang, X., Cappos, G., Joly-Amado, A., Majid, S., Minamide,

L. S., Bamburg, J. R., Morgan, D., Kang, D. E (2015). “RanBP9 at the intersection

between cofilin and Aβ pathologies: rescue of neurodegenerative changes by RanBP9

reduction.” Cell death and disease (2015) 6, 1676; dio:10.1038

Background

The defining pathological hallmark of Alzheimer’s disease (AD) is the accumulation of

A in brain associated with tau pathology, synapse loss, cytoskeletal aberrations, mitochondrial dysfunction, and cognitive decline. The generation of A occurs via sequential- and -secretase processing of the Amyloid Precursor Protein (APP) by BACE1 and the Presenilin complex,

respectively(De Strooper & Annaert, 2000b). Soluble oligomeric forms of A are thought to be the most toxic species, resulting in synaptic loss and downstream neurotoxicity (D. M. Walsh et al., 2002). Despite the requirement for Tau in multiple aspects of A-induced neurotoxicity

(Morris, Maeda, Vossel, & Mucke, 2011), a large knowledge gap exists as to how the A oligomer-induced neurotoxic signals are transduced intracellularly to impair synaptic plasticity, eventually leading to neurodegeneration. Both Aand Tau promote Cofilin-Actin pathology (B.

W. Bernstein & J. R. Bamburg, 2010; Fulga et al., 2007), Cofilin-Actin pathology is widespread in AD brains(Minamide et al., 2000), and Cofilin activity is also increased in AD brains(T. Kim et al., 2013). Cofilin normally functions as a key regulator of Actin dynamics that destabilizes F-

Actin. Cofilin is inactivated by phosphorylation on Ser3 by LIMK1, whereas its dephosphorylation by SSH1 activates Cofilin(B. W. Bernstein & J. R. Bamburg, 2010). Upon oxidative stress and/or Ca2+ elevation(B. W. Bernstein & J. R. Bamburg, 2010; J. S. Kim et al.,

2009; Roh et al., 2013a), SSH1 is activated and active Cofilin becomes oxidized on cysteine residues, resulting in rapid mitochondrial translocation to promote apoptosis and induction of

Cofilin-Actin pathology(B. W. Bernstein et al., 2012; Klamt et al., 2009). An early and consistent impairment secondary to A oligomer treatment in primary neurons is the shrinkage of dendritic spines(Selkoe, 2008) involving the rearrangement of F-actin cytoskeleton in spines and loss of spine-associated proteins such as PSD95 and Drebrin(G.M. Shankar et al., 2007;

Zhao et al., 2006), as well as impaired mitochondrial function(Du et al., 2010; Moreira et al.,

2001).

We recently found that overexpression of the scaffolding protein RanBP9 increases A production in cell lines and in transgenic mice(M.K. Lakshmana et al., 2012; Lakshmana et al.,

2009). Moreover, RanBP9 is significantly increased in brains of AD patients and the J20 APP

transgenic model(M.K. Lakshmana et al., 2012; J. A. Woo et al., 2012). In studying the trafficking of APP, we also found that RanBP9 overexpression not only promotes the endocytosis of APP but also those of LRP and 1-integrin, the latter resulting in disassembly of

Integrin-associated focal complexes (Talin & Vinculin)(Woo, Roh, Lakshmana, & Kang, 2012).

In addition, RanBP9 overexpression promotes Cofilin activation and the translocation of Cofilin to mitochondria, resulting in overall mitochondrial dysfunction(Roh et al., 2013a; J. A. Woo et al., 2012). However, how RanBP9 activates Cofilin is unknown, and it is not clear whether reduction in endogenous RanBP9 protects against A oligomer-induced deficits in synaptic plasticity, Cofilin-dependent pathology, A accumulation, and memory impairment. Here we report that siRNA or genetic reduction in RanBP9 significantly reduces SSH1 levels and mitigates A-induced translocation of Cofilin to mitochondria, Cofilin-Actin rod/aggregate formation, and depletion of synaptic proteins, deficits in synaptic plasticity, A accumulation, and contextual memory deficits in vivo.

Methods

Cells, cDNA constructs, siRNA sequences, and transfection

Hippocampus-derived HT22 cells were maintained in Dulbecco’s modified Eagle's medium containing 10% FBS. HT22 cells were obtained from Professor David Schubert (Salk

Institute, San Diego, CA, USA). The RNAi targeting RanBP9 (5 ′ -

UCUUAUCAACAAUACCUGC-3′) and SSH1 (5′-GAGGAGCUGUCCCGAUGAC-3′) were obtained from GE Dharmacon (Lafayette, CO, USA). All cells were transfected for 48 h using lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions with siRNA duplexes or control single-stranded denatured control siRNAs before

performing biochemical and/or immunocytochemical assays. All siRNAs were transfected at a final concentration of 100 nM. Cortical and hippocampal primary neurons were derived from postnatal day 0 (P0) pups and grown on poly-D-lysine-coated coverslips or plates as previously described.19 Quantitation of digitized immunoblotting data was performed using the Image J software (NIH Image J, Bethesda, MD, USA).

Antibodies, reagents, and Aβ oligomer preparation

Antibodies to APP (6E10, Covance, Madison, WI, USA), cofilin (D3F9, Cell Signaling,

Danvers, MA, USA), phospho-cofilin (77G2, Cell Signaling), actin (AC-74, Sigma Aldrich, St.

Louis, MO, USA), tubulin (TU-02, Santa Cruz Biotechnology, Dallas, TX, USA), Aβ (D54D2,

Cell Signaling), GFAP (Invitrogen), synapsin I (Invitrogen), PSD95 (Abcam, Cambridge, MA,

USA), Drebrin (Abcam), microtubule-associated protein 2 (Millipore, Billerica, MA, USA),

HRP-linked secondary antibodies (Jackson Immunochemicals, West Grove, PA, USA), and fluorescently labeled secondary antibodies (Invitrogen) were obtained from the indicated sources. The mouse monoclonal anti-RanBP9 antibody was a generous gift from Professor

Elizabetta Bianchi (Pasteur Institute, France). Synthetic Aβ1-42 peptide was purchased from

American Peptide (Sunnyvale, CA, USA). Aβ1-42 oligomers were prepared as previously characterized. Briefly, Aβ1-42 powder wasdissolved in hexafluoro-2-propanol (HFIP) at 1 mM for 30 min at room temperature, aliquoted to eppendorf tubes, allowed to evaporate overnight in fume hood, and subjected to speed vacuum for 1 hour to remove traces of HFIP or moisture. To prepare Aβ oligomers, Aβ1-42 film was then dissolved in dimethyl sulfoxide (5 mM), and F-12 cell culture medium (without phenol) was added to a final concentration of 100 μM Aβ1-42 and incubated at 4 °C for 24 hours.

Cell/tissue lysis and immunoblotting

Cultured cells or brain homogenates were lysed with lysis buffer (50 mM Tris-Cl, 150 mM NaCl, 2 mM EDTA, and 1% Triton X-100) unless stated otherwise. For separation of cytosolic versus focal adhesion-enriched (F) fractions, cytosolic proteins were first released with saponin buffer (0.1% saponin, 25 mM HEPES, and 75 mM potassium acetate) for 10 min on ice followed by extraction of remaining proteins by 2 × SDS sample buffer and sonication. For isolation of mitochondria, mitochondrial isolation kit (Thermo Scientific, Rockford, IL, USA) was used according to the manufacturer’s instructions for cultured cells. Protein quantification was performed by a colorimetric detection reagent (BCA protein assay, Pierce, Rockford, IL,

USA). Equal amounts of protein were subjected to SDS-PAGE and transferred to nitrocellulose membranes for immunoblotting. After probing with the primary antibody, the corresponding peroxidase-conjugated secondary antibody was detected by ECL western blot reagents (Pierce).

ECL images were captured by the Fuji LAS-4000 imager (LAS-4000, Pittsburgh, PA, USA) and quantified using the ImageJ software (NIH ImageJ, Bethesda, MD, USA).

Mice

APP/PS1, WT, and RanBP9+/− mice19 were all bred in the C57BL6 background for at least three generations before interbreeding with each other. APP/PS1 mice, which express

‘Swedish’ APP and PS1 ΔE9 mutations, were obtained from Jackson Immunoresearch (West

Grove, PA, USA).

Immunofluorescence

Immunohistochemistry and immunocytochemistry were performed as previously described. Briefly, animals were perfused with 4% paraformaldehyde in PBS, and brains were post-fixed in the same fixative for 24 hours. The brains were then cryoprotected in 30% sucrose

and sectioned (30 μm) on a cryostat or microtome. For primary neurons and HT22 cells, cells were fixed in 4% paraformaldehyde for 15 min at room temperature. After blocking with normal goat serum, primary antibodies were applied overnight at 4 °C, and secondary antibodies were applied for 45 min at room temperature, followed by counterstaining with Hoechst33342 or

DAPI and mounting. For detection of cofilin–actin rods in brain, 70% ethanol was applied to sections for 5 min after washing the secondary antibody, followed by 0.1% Sudan Black B in

70% ethanol for 12 min, two washes with 70% ethanol, several washes in Tris-buffered saline

(TBS) before mounting. All immunoreactivities were quantitated from every 12th serial section through an entire hippocampus or anterior cortex. All images were acquired with the Olympus

FV10i confocal microscope (Tokyo, Japan) and quantitated using the Image J software.

Comparison images were acquired with identical laser intensity, exposure time, and filter.

Adjustments to the brightness/contrast were applied equally to all comparison images.

Fear conditioning, open field, and rotarod behavior

Fear conditioning (contextual and cued), rotarod, and open field tasks were performed as previously described. For fear conditioning, an aversive stimulus (in this case a mild foot shock,

0.5 mA) was paired with an auditory conditioned stimulus (white noise) within a novel environment. Training consisted of two mild shocks paired with two conditioned stimuli with a

3-min interval between each shock. Freezing on the training day in response to the foot shock was used as an estimate of learning during the acquisition trial. To test conditioning to the context, animals were re-introduced to the same training chamber for 6 min and freezing behavior was recording by tracking software (Any-maze, Wood Dale, IL, USA) every second.

To test conditioning to the tone, animals were introduced to a novel context, consisting of a chamber with different shape, floor and olfactory cues from the training chamber. Mice were

scored for 3 min, before and after the tone in the same manner described above. Learning was assessed by measuring freezing behavior (i.e., motionless position) every second. The open field was used as a standard test of general activity. Briefly, animals were monitored for 15 min in a

40 cm square open field with a video tracking software (Any-maze). General activity levels were evaluated by measurements of total distance traveled. Motor performance was evaluated by an accelerating rotarod apparatus with a 3 cm diameter rod starting at an initial rotation of 4 r.p.m. slowly accelerating to 40 r.p.m. over 5 min. The time spent on the rod during each of four trials per day for 2 consecutive days was measured previously described in.

Electrophysiology in mouse brain slices

Hippocampus slices were prepared from 3-month-old WT, APP/PS1, and

APP/PS1;RanBP9+/− mice and subjected to input/output curved, paired pulse facilitation, and

LTP as previously described. Briefly, animals were killed, brains were harvested, and sectioned horizontally (400 μm) in ice-cold cutting solution (110 mM sucrose, 6 mM NaCl, 3 mM KCl, 26 mM NaHCO3, 1.25 mM, NaH2PO4, 7 mM MgCl2, 0.5 mM CaCl2, 10 g/l glucose, pH 7.3–7.4).

The hippocampus was dissected and acclimated in 50:50 solution (cutting:artificial cerebrospinal fluid (ACSF)) for 10 min at room temperature. Then the slices were transferred to ACSF (125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 0.26 mM NaHCO3, 1.2 mM MgCl2, 2.0 mM

CaCl2, and 10 g/l glucose, pH 7.3–7.4, saturated with 95% O2 and 5% CO2). Slices were recovered in ACSF at room temperature at least 40 min, followed by a final incubation in ACSF for 1 hour at 30 °C. Extracellular field potential recording, LTP: the recording chamber was held at 30± 0.5°C with ACSF flow rate of 1 ml/min. The stimulating electrode was placed in the

Schaffer collaterals of the hippocampus. The recording glass electrode loaded with ACSF was positioned at the CA1 stratum radiatum below the pyramidal cell layer. Stimulating pulses were

generated by the Digidata 1322A interface (Molecular Devices, Sunnyvale, CA, USA) and a stimulus isolator (model 2200; A-M Systems, Sequim, WA, USA) under control of Clampex

10.0 software (Molecular Devices). Field excitatory postsynaptic potentials (fEPSP) were amplified using a differential amplifier (model 1800; A-M Systems), filtered at 1 kHz, and digitized at 10 kHz. Input–output analysis was performed by stepping stimulation amplitude from 1 to 15mV. Stimulation amplitude that elicited half-maximal fEPSP was determined by the input–output curve and stimulation rate of 0.05 Hz was used through the whole experiment. PPF, which is short-term plasticity, was evoked by two pulses with interpulse intervals from 20 to 300 ms. Percentage of the facilitation was calculated by dividing fEPSP slope elicited by the second pulse with the fEPSP slope elicited by the first pulse. LTP was induced by theta burst stimulation

(TBS) (five trains of four pulses at 200 Hz separated by 200 ms, repeated six times with an inter- train interval of 10 s). LTP was sampled 60 min after the induction, and calculated by dividing the slope of 60 min post-induction responses with the average slope of 20-min baseline responses.

Results

Endogenous RanBP9 mediates Aβ42 oligomer-induced translocation of cofilin to mitochondria and promotes cofilin activation via positively regulating SSH1

We assessed whether Aβ oligomers alter Cofilin translocation to mitochondria and whether siRNA knockdown of RanBP9 might influence this phenotype. We prepared Aβ1-42 oligomers precisely as previously characterized(Dahlgren et al., 2002), which contained SDS- resistant dimers, trimers, and tetramers (Aβ42O) whereas freshly dissolved Aβ1-42 only contained monomers (Aβ 42M) (Fig. 3-1A). All stated Aβ42O concentrations heretofore are based on the monomer concentration. Hippocampus derived HT22 cells were transiently transfected with control or RanBP9 siRNA for 48 hours and subjected to treatment with or without Aβ42O 73

(1µM) for 2 hours, followed by separation of intact mitochondria and cytosol fractions. Aβ42O treatment increased both Cofilin and RanBP9 translocation to mitochondria (Fig. 1B). However, siRNA knockdown of endogenous RanBP9 mitigated AβO-induced Cofilin mitochondrial translocation (Fig. 3-1B,C). Aβ42 monomer exposure (1µM, 2h) neither promoted Cofilin translocation to mitochondria nor altered Cofilin activation (Supplemental Fig. S3-1A). In vivo,

RanBP9 levels were greatly decreased in the hippocampus of 3-month old APP/PS1;RanBP9+/- mice in both cytosol and mitochondrial fractions compared to APP/PS1 mice(Jankowsky et al.,

2004) (Fig. 3-1D). While Cofilin levels were unchanged in the cytosol fraction between the genotypes, Cofilin was significantly reduced in the hippocampal mitochondrial fraction of

APP/PS1;RanBP9+/- mice compared to APP/PS1 mice (Fig. 3-1D,E), indicating that endogenous RanBP9 facilitates translocation of Cofilin to mitochondria in vivo. As expected, 3- month old RanBP9+/- mice exhibited reduced RanBP9 protein together with significantly increased inactive phospho-Cofilin without altering total Cofilin (Fig. 3-1F,G), which was accompanied by significantly decreased SSH1 protein in RanBP9+/- mice (Fig. 1F-H), indicating that RanBP9 positively regulates SSH1 levels. Indeed, SSH1 but not LIMK1 levels were significantly increased secondary to RanBP9 overexpression in HT22 cells (Fig. 3-1I,J).

Consistent with these observations, RanBP9 not only markedly increased SSH1 but also co- immunoprecipitated with SSH1 in brain (Fig 3-1K). Accordingly, siRNA knockdown of RanBP9 in HT22 cells strongly reduced the enhancement of Cofilin-SSH1 complex induced by Aβ42O treatment (Fig. 3-1L) while significantly decreasing SSH1 levels (Fig. 3-1L,M). In DIV21 primary cortical neurons, Aβ42O (2h) marked increased RanBP9-SSH1 and Cofilin-SSH1 complexes (Fig. 3-1N), suggesting that RanBP9 not only increases SSH1 levels but may also serve to facilitate Cofilin-SSH1 interaction upon Aβ42O exposure. To determine whether the

RanBP9-SSH1 complex alters SSH1 protein stability, we performed cycloheximide turnover experiments. Indeed, RanBP9 siRNA and overexpression marked accelerated and delayed the turnover of SSH1, respectively (Fig. 3-1O), confirming that RanBP9 positively regulates SSH1 protein stability.

Endogenous RanBP9 and SSH1 promote cofilin–actin rod formation and Aβ oligomer- induced collapse of growth cones in primary hippocampal neurons.

Hyperactive Cofilin can form Cofilin-saturated Actin filament bundles (or rods) when challenged by oxidative stress, excitotoxic insult, or Aβ oligomers in primary neurons. These

Cofilin-Actin rods accumulate in neurites, which blocks neuritic transport of key cargo to distal locations(Cichon et al., 2012; Mi et al., 2013). To assess the role of RanBP9 in Cofilin-Actin rod formation, we transiently transfected GFP-Cofilin in wild type and RanBP9+/- hippocampal neurons. On DIV8, we treated neurons with a low concentration of hydrogen peroxide (1µM) for

30 minutes, which is known to induce Cofilin-Actin rod formation. Wild type neurons formed significantly higher numbers of GFP-Cofilin rods per neuron and per neuronal cytoplasmic area

(NCA) compared to RanBP9+/- neurons (Fig. 3-2A,B). We next treated DIV8 wild type and

RanBP9+/- neurons with or without Aβ42O (1µM) for 24 hours, which has been shown to induce

Cofilin-Actin rod formation in 10-20% of neurons(Maloney, Minamide, Kinley, Boyle, &

Bamburg, 2005). We observed fewer percentage of neurons with Cofilin rod/aggregate-like structures in RanBP9+/- neurons (16.3% of WT neurons & 6.9% of RanBP9+/- neurons)(Fig. 3-

2C) as well as significantly reduced numbers of Cofilin rods/aggregates per neuron and per neuronal cytoplasmic area (NCA) compared to WT neurons upon Aβ42O treatment (Fig. 3-

2C,D). In addition, we observed the presence of F-Actin-positive growth cones in WT vehicle treated neurons, which largely collapsed in wild type neurons treated with Aβ42O (Fig. 3-2C,E).

However, F-Actin-positive growth cones were largely unaffected by Aβ42O in

RanBP9+/- neurons (Fig. 3-2C,E), indicating that RanBP9 reduction protects against Cofilin-

Actin rod formation as well as Aβ42O-induced collapse of growth cones. Such changes in

RanBP9+/- neurons were not due to gross alterations in the numbers or length of neurites (Fig.

3-2F,G). However, Scholl analysis demonstrated that RanBP9+/- neurons exhibited subtle but significant increases in numbers of neurite intersections at certain distances from the soma at basal state on DIV9 (Fig. 3-2F,G). Similar to RanBP9 reduction, SSH1 siRNA also significantly prevented Cofilin rod/aggregate structures and collapse of growth cones induced by Aβ42O

(1µM) exposure for 24 hours (Fig. 3-2H-J), indicating that RanBP9 reduction mitigates Aβ42O- induced effects at least in part via negative regulation of SSH1.

Endogenous RanBP9 mediates the depletion of postsynaptic proteins induced by Aβ oligomers in primary hippocampal neurons.

Dendritic spines are structures critical for excitatory synaptic transmission and highly enriched in postsynaptic proteins such as Drebrin, PSD95, and F-Actin(Shirao & Gonzalez-

Billault, 2013). To assess synaptic perturbations induced by Aβ42O, we cultured P0 hippocampal neurons derived from RanBP9+/- and wild type littermate mice. We cultured neurons to DIV21 to visualize mature dendritic spines and treated them with or without Aβ42O for 2 hours. As expected, immunocytochemical analysis demonstrated that Aβ42O significantly depleted

Drebrin, PSD95, and F-Actin in dendritic spines and spine-containing neurites in wild type neurons (Fig. 3-3A-D). However, the same Aβ42O treatment had no significant effects in

RanBP9+/- neurons (Fig. 3-3A-D), indicating that endogenous level of RanBP9 is required for

Aβ1-42O-induced depletion of postsynaptic proteins and F-Actin in mature neurons. Aβ42 monomer treatment (1µM, 2h) did not appreciably alter Drebrin or F-Actin levels (Supplemental

Fig. S3-1B). In DIV21 mature neurons, Scholl analysis demonstrated significantly increased 76

complexity and length of neurites beyond 50µm from the soma in RanBP9+/- vs. wild type neurons (Fig. 3-3E,F), supportive of a role for endogenous RanBP9 in the inhibition of neurite outgrowth and maturation.

Significant increase in endogenous RanBP9 in APP/PS1 mice brains.

We examined endogenous RanBP9 expression in the APP/PS1 mice expressing the

‘Swedish’ and PS1ΔE9 mutation by immunoblotting and immunohistochemistry. Endogenous

RanBP9 protein levels were significantly increased by 3.5-fold in the hippocampus of 8-month old APP/PS1 mice by immunoblotting (Fig. 3-4A,B), consistent with our previous observations in 12-month old J20 mice harboring the ‘Swedish’ mutation and in AD brains (M.K. Lakshmana et al., 2012; J. A. Woo et al., 2012). Further, immunohistochemical analysis of APP/PS1 mouse hippocampus demonstrated greatly enhanced RanBP9 levels in the dentate gyrus (DG) and CA3 regions compared to wild type littermates (Fig. 3-4C).

Rescue of neuroinflammation, synaptic damage, and Aβ ccumulation by RanBP9 reduction in APP/PS1 mice.

Given the robust upregulation of RanBP9 levels in APP/PS1 mice, we next determined whether genetic reduction in RanBP9 can prevent neuroinflammation and synapse loss associated with APP/PS1 transgenic mice. We performed immunohistochemistry for GFAP and

Iba1 to detect activated astrocytes and microglia as well as Synapsin I and PSD95 to detect pre- and post-synaptic proteins from 8-month old APP/PS1, APP/PS1;RanBP9+/-, and wild type

(WT) littermates. As expected, APP/PS1 mice demonstrated significantly increased GFAP and

Iba1 immunoreactivities throughout the hippocampus and areas of the anterior cortex associated with Aβ accumulation as well as diminished PSD95 and Synapsin I immunoreactivities within the stratum lucidium (SL: synaptic terminating zone) of CA3 (Fig. 3-5A,B). In contrast,

APP/PS1;RanBP9+/- mice showed significantly reduced GFAP and Iba1 immunoreactivities in the hippocampus and anterior cortex as well as significantly rescued Synapsin I and PSD95 immunoreactivities within the SL of CA3 compared to APP/PS1 littermate mice, essentially indistinguishable from wild type mice (Fig 3-5A,B). Because RanBP9 overexpression promotes

Aβ production in cultured cells and in vivo (M.K. Lakshmana et al., 2012; Lakshmana et al.,

2009), we next examined whether endogenous RanBP9 reduction alters Aβ burden in the hippocampus and anterior cortex of 8-month old APP/PS1 mice. Indeed, APP/PS1;RanBP9+/- mice demonstrated significantly decreased both total Aβ deposits and thioflavin S-positive fibrillar Aβ deposits within the hippocampus and anterior cortex compared to littermate

APP/PS1 mice (Fig. 3-6A-C). In addition, soluble Aβ levels were also significantly reduced in the hippocampus of APP/PS1;RanBP9+/- mice compared to littermate APP/PS1 mice (Fig. 3-

6D,E). These results demonstrate that RanBP9 reduction protects against Aβ accumulation as well as neuroinflammation and synaptotoxicity in vivo.

Accumulation of Cofilin-Actin Rods in APP/PS1 mice and Significant Attenuation of Cofilin-Actin Rod Formation by RanBP9 Reduction.

Given the positive regulation of SSH1 and Cofilin as well as Cofilin-Actin rods in primary neurons by RanBP9, we assessed Cofilin-Actin rods / aggregates in wild type, APP/PS1, and APP/PS1;RanBP9+/- brains. For this, we used Sudan Black B to quench background and the more diffuse non-rod staining. Therefore, only intense signals, such as rods, are visualized while removing normal cytoplasmic immunoreactivity. Using this method, we found that Cofilin rods / aggregates were readily detectable in the hippocampus and anterior cortex of 9-month old

APP/PS1 mice, while very few were found in wild type (WT) mice (Fig. 3-7A). Such Cofilin rod/aggregate structures, which colocalized with Actin, were significantly increased by ~10-fold in APP/PS1 vs. WT mice (Fig. 3-7A-C) and resembled those seen in AD brains (Fig. 3-7D). 78

RanBP9 reduction significantly attenuated Cofilin-Actin rod / aggregate formation, although not completely to the level of WT mice (Fig. 3-7A-C). In the AD entorhinal cortex, Cofilin rod / aggregate structures were numerous and overwhelmingly did not colocalize with 12E8-positive phospho-Tau immunoreactivity, albeit within the same region and in close proximity (Fig. 3-

7D). These findings in vivo confirm our findings in primary neurons that RanBP9 reduction ameliorates Cofiin-Actin pathology in an in vivo mouse model of Aβ accumulation.

RanBP9 reduction rescues deficits in synaptic plasticity and contextual memory associated with APP/PS1 mice.

We tested short-term and long-term synaptic plasticity from acute hippocampal slices prepared from 3-month-old WT, APP/PS1, RanBP9+/−, and APP/PS1;RanBP9+/− mice, an age when little to no Aβ plaques are detected. The stimulating electrode was placed in the Schaffer collaterals of the hippocampus, and the recording electrode was positioned at the CA1 stratum radiatum below the pyramidal cell layer. As shown in Figure 3-8A, input–output analysis did not markedly differ among WT, APP/PS1, and APP/PS1;RanBP9+/−, and RanBP9+/−slices.

However, the APP/PS1;RanBP9+/− slices showed a nonsignificant decrease in basal synaptic transmission compared with WT, RanBP9+/−, and APP/PS1 slices (Figure 3-8a). In paired pulse facilitation (PPF) experiments, significant differences were observed in fEPSP slope across genotypes among all interstimulus intervals (Figure 3-8b). Correction for multiple comparisons surprisingly showed that APP/PS1;RanBP9+/− slices exhibited significantly higher PPF compared with both WT, RanBP9+/−, and APP/PS1 slices across most interstimulus intervals

(Figure 3-8b), indicating a synergistic effect of RanBP9 and APP/PS1 genotypes on cooperative presynaptic efficacy. For long-term potentiation (LTP) measurements, we detected no differences in fEPSP slope among WT, APP/PS1, RanBP9+/−, and APP/PS1;RanBP9+/− slices at baseline (Figure 3-8c). However, after theta burst stimulation, we observed significant 79

differences in fEPSP slope across genotypes for all time points (Figure 3-8c). Correction for multiple comparisons showed that APP/PS1 slices were significantly impaired in fEPSP slope compared with WT, RanBP9+/−, and APP/PS1;RanBP9+/− slices at every time point up to

1 hour (Figure 3-8c). APP/PS1;RanBP9+/− slices exhibited significantly stronger fEPSP slope across all time points compared with WT and APP/PS1 slices (Figure 3-8c), indicating that

RanBP9 reduction greatly enhances both the induction and maintenance of LTP. Similarly,

RanBP9+/− slices exhibited significantly stronger LTP than WT slices up to 30 min and did not significantly differ from APP/PS1;RanBP9+/− slices (Figure 3-8c). Therefore, these results indicate that RanBP9 reduction not only rescues the deficits in synaptic plasticity associated with the APP/PS1 mice but also further potentiates synaptic plasticity.

Discussion

Molecular pathways that govern the neurotoxicity and production of Aβ are attractive therapeutic targets for AD. In this study, we utilized HT22 cells, primary neurons, acute brain slices, and genetically modified mice to gain insights into the role of endogenous RanBP9 in Aβ- induced synaptic degeneration, Cofilin activation, Cofilin-Actin rod formation, and Aβ accumulation in a mouse model of AD. We made a number of novel observations demonstrating that endogenous RanBP9 positively regulates SSH1 levels and is critical for Aβ-induced translocation of Cofilin to mitochondria, Cofilin-Actin rod/aggregate formation, collapse of growth cones, loss of synaptic proteins, and impairments in synaptic plasticity and learning/memory. Our findings underscore the critical importance of the RanBP9-SSH1-Cofilin pathway in AD pathogenesis.

A prerequisite for Cofilin translocation to mitochondria and formation of Cofilin-Actin rods is its activation via dephosphorylation and oxidation on several cysteine residues (B. W.

Bernstein et al., 2012; Klamt et al., 2009). Therefore, the observation that Aβ oligomers induced

Cofilin translocation to mitochondria and formation of Cofilin-Actin rods infers its rapid activation and oxidation by Aβ oligomers. Interestingly, AβO also increased RanBP9 translocation to mitochondria, which may in part be explained by its association with the pro- apoptotic mitochondria-associated protein P73 (Liu et al., 2013). Genetic reduction and siRNA knockdown of RanBP9 significantly mitigated both of these measures and reduced Cofilin activation. As Cofilin is activated by SSH1-dependent dephosphorylation and inactivated by

LIMK-dependent phosphorylation, it is plausible that RanBP9 alters one or both of these pathways. Indeed, we found that RanBP9 positively regulates SSH1 but not LIMK1 levels in

HT22 cells and in brain, which accounts for increased Cofilin activation by RanBP9. As RanBP9 levels are robustly increased in APP/PS1 mouse brains, it is likely that AβO also activates Cofilin via a mechanism involving RanBP9 and SSH1. Aβ is known to increase cytosolic Ca2+ and activate Calcineurin as well as enhance production of reactive oxygen species (ROS)(Butterfield,

Swomley, & Sultana, 2013; Reese & Taglialatela, 2011), both of which are known activators of

SSH1. Calcineurin-dependent dephosphorylation of SSH1 serves to activate SSH1 (Y. Wang,

Shibasaki, & Mizuno, 2005b), and cysteine oxidation of 14-3-3 removes the inhibition of SSH1 by 14-3-3(J. S. Kim et al., 2009). In addition, cysteine oxidation of Cofilin itself is required for both mitochondrial translocation and Cofilin-Actin rod formation (B. W. Bernstein et al., 2012;

Klamt et al., 2009). We recently showed that AβO promotes Cofilin-Actin rod formation via a pathway requiring PrPc and NADPH oxidase (NOX), the latter which is involved in the generation of ROS (K. P. Walsh et al., 2014). We have also shown that AβO and RanBP9 impair mitochondrial Ca2+ buffering and promote mitochondrial superoxide production in a Cofilin- dependent manner (Roh et al., 2013a; J. A. Woo et al., 2012). Therefore, the engagement of AβO

to its neuronal receptors likely involves downstream Ca2+ and ROS from multiple sources.

Activated/oxidized Cofilin likely contributes to sustaining this cycle by promoting mitochondrial superoxide production.

We observed that RanBP9+/- neurons are resistant to Aβ oligomer-induced collapse of F-

Actin containing growth cones in immature DIV9 neurons and depletion of postsynaptic proteins

(Drebrin, PSD95, and F-Actin) in mature DIV21 neurons. Such findings are likely explained, at least in part, by the reduction in Cofilin activation status in RanBP9+/- neurons. Specifically, phosphorylation and dephosphorylation of Cofilin are associated with dendritic spine growth and shrinkage during LTP and LTD, respectively (Chen et al., 2007; Rex et al., 2009; Zhou et al.,

2004), and Cofilin regulates the trafficking of AMPA receptors in dendritic spines (Gu et al.,

2010). Further, β-Arrestin 2 recruits Cofilin to dendritic spines upon NMDA receptor activation to control the remodeling of spines, and β-Arrestin 2 knockout neurons are resistant to Aβ- induced dendritic spine loss through spatial control of Cofilin activation (Pontrello et al., 2012).

Despite the crucial involvement of Cofilin in synaptic plasticity, the aforementioned mechanism alone is unlikely to explain the full extent of the rescue and enhancement of LTP and PPF associated with APP/PS1;RanBP9+/- mice, since both LTP and PPF in were strongly enhanced significantly beyond those of wild type slices. However, in the absence of the APP/PS1 genotype, RanBP9+/- mice exhibited normal PPF but significantly enhanced LTP. The exaggerated PPF in APP/PS1;RanBP9+/- slices may be explained by lower probability of Ca2+- evoked neurotransmitter release as evidenced by somewhat reduced fEPSP slope per stimulus amplitude (input/output curve). However, the second paired Ca2+ signal appears to overcompensate to release a larger pool of neurotransmitter. We hypothesize that RanBP9 reduction enhances synaptic associativity and cooperativity perhaps at the level of synapse

formation and maintenance. Indeed, RanBP9+/- slices produced significantly enhanced LTP compared to WT and did not differ significantly from APP/PS1;RanBP9+/- slices.

Overexpression of RanBP9 significantly reduces axon elongation in the chick spinal cord via

Plexin-A (H. Togashi, E. F. Schmidt, & S. M. Strittmatter, 2006), and we also showed in this study that reduction in RanBP9 increases neurite arborization/complexity in mature neurons and protects against both Aβ oligomer-induced growth cone collapse and dendritic postsynaptic protein depletion. Previous studies have also shown the involvement of RanBP9 in negatively regulating β1-integrin function (J.A. Woo et al., 2012) and a role in the nucleocytoplasmic regulation of cell morphology via its interaction with muskelin (Valiyaveettil et al., 2008a).

Therefore, RanBP9 may be involved in regulating neuritogenesis, synaptic maintenance, and synaptic plasticity changes via these pathways in addition to Cofilin activation.

RanBP9 reduction protected against both AβO- and hydrogen peroxide-induced Cofilin-

Actin rod formation in primary neurons, thereby indicating a direct effect of RanBP9 and SSH1 reduction on Cofilin-Actin rods. Indeed, SSH1 reduction per se significantly mitigated Aβ42O- induced Cofilin rod/aggregate formation and collapse of growth cones, indicating that SSH1 reduction, at least in part, is responsible for such effects of RanBP9. In vivo, however, RanBP9 reduction mitigated both Cofilin-Actin rod/aggregate formation and Aβ accumulation/plaques.

Therefore, it is likely that RanBP9 reduction lowers Cofilin-Actin rod/aggregate formation both directly via SSH1 regulation and indirectly by reducing Aβ accumulation. Cofilin-Actin rods are thought to be an initial protective response to oxidative insults (B.W. Bernstein, Chen, Boyle, &

Bamburg, 2006); however, persistent accumulation of rods physically blocks neuritic transport of cargo and impairs synaptic activity (Cichon et al., 2012; Mi et al., 2013). Inhibition of RanBP9 and SSH1 may not be sufficient to fully block Cofilin-Actin pathology, as ROS pathways acting

directly on Cofilin may also promote Cofilin-Actin pathology. In this light, it is intriguing that

Tau overexpression induces Cofilin-Actin pathology and potently increases F-Actin bundling

(Fulga et al., 2007) while disrupting mitochondrial fission via an Actin-dependent mechanism

(DuBoff, Gotz, & Feany, 2012).

Conclusion

Taken together, this study demonstrated the crucial involvement of endogenous RanBP9 in Aβ- induced Cofilin activation via SSH1 and downstream consequences on Cofilin-Actin rod formation, Aβ accumulation, synaptic plasticity, and contextual memory. These findings underscore the role of RanBP9 and its associated molecular pathway as critical mediators of AD pathology and potential therapeutic targets.

Figure 3-1. RanBP9 mediates Aβ oligomer-induced translocation of cofilin to mitochondria and lowers cofilin activation via SSH1. a) Aβ1-42 monomers (Aβ42M) or Aβ1-42 oligomer preparation (Aβ42O) subjected to SDS-PAGE and immunoblotted for Aβ. Note the monomer (M), dimers (Di), trimmers (Tri), and tetramemers (Tet) in the oligomer preparation. b & c) Hippocampus- derived HT22 cells transiently transfected with control or RanBP9 siRNA for 48 h, treated with/without Aβ oligomers for 2 hours, separated for mitochondrial and cytosol fractions, and subjected to immunoblotting for the indicated proteins. A representative experiment is shown. Notice the reduction phospho-cofilin upon Aβ oligomers treatment but mitigated response in RanBP9 siRNA knocked down cells. c) Quantitation of mitochondrial cofilin (ANOVA, post-hoc Tukey, **P<0.005, n=3 replicates). d & e) Hippocampal extracts from 3-month-old APP/PS1 and APP/PS1;RanBP9+/− mice subjected to separation of mitochondrial and cytosol fractions and immunoblotted for the indicated proteins. Note the reduced level of cofilin in mitochondrial fraction of APP/PS1;RanBP9+/− brain. e) Quantitation of mitochondrial cofilin (t-test, **P<0.005, n=4 mice per genotype). f-h) Hippocampal extracts from 3-month-old WT and littermate RanBP9+/− (Ran9+/−) mice subjected to immunoblotting for the indicated proteins. g) Quantitation of P-cofilin in hippocampus (t=2.74, P=0.022, *P<0.05, n=5 mice per genotype). h) Quantitation of SSH1 (t-test, **P=0.0016, n=4 mice per genotype). i & j) HT22 cells transiently transfected with vector control (Vec) or Flag-RanBP9 and immunoblotted for the indicated proteins. Note the increase in SSH1 but not LIMK1 by RanBP9 overexpression. j) Quantitation of SSH1 (t-test, ***P<0.0001, n=4 replicates). Error bars represent S.E.M. on graphs. k) Cortex (CTX) and hippocampus (HIPP) homogenates of 6-month-old WT or littermate Flag-RanBP9 transgenic (TG) mice immunoprecipitated for SSH1 and/or immunoblotted for the indicated proteins. Note the increase in SSH1-RanBP9 complex in Flag-RanBP9 transgenic mice (TG). l) HT22 cells transiently transfected with/without RanBP9 siRNA and treated with/without Aβ oligomers (1 μM) for 2 hours followed by immunoprecipitation for SSH1 and/or immunoblotting for the indicated proteins. Note that RanBP9 siRNA reduces SSH1 levels and decreases Aβ oligomers-induced enhancement of cofilin–SSH1 interaction. m) Quantitation of SSH1 protein levels with/without RanBP9 siRNA transfection in HT22 cells (t-test, ***P=0.0008, n=4 replicates). n) DIV18 cortical primary neurons treated with or without Aβ42O (1 μM) for 2 hours, subjected to immunoprecipitation for SSH1, and/or immunoblotting for the indicated proteins. Note the increase in RanBP9-SSH1 and cofilin-SSH1 complex formation with Aβ42 oligomers treatment. o) HT22 cells transiently transfected with control, RanBP9 siRNA, or RanBP9 and subjected to cycloheximide (CHX) treatment for the indicated times followed by immunoblotting for SSH1 or actin. Representative blots adjusted to show similar signal at time 0. Note that RanBP9 siRNA and overexpression accelerates and delays SSH1 turnover, respectively

Figure 3-2. RanBP9 mediates cofilin–actin rod formation and Aβ oligomer-induced collapse of growth cones in primary hippocampal neurons. a & b) DIV8 hippocampal neurons from WTand RanBP9+/− mice transiently transfected with cofilin-GFP and treated with hydrogen peroxide (1 μM) for 30 min and subjected to F-actin staining (Rhodaminephalloidin). Representative images showing marked reduction in cofilin-GFP rods in RanBP9+/− neurons, despite strong GFP cofilin fluorescence. b) Quantitation of cofilin- GFP rods per neuron (t-test, **P= 0.0029, n= 6 replicates from three pups per genotype) or per NCA (t-test, **P =0.0009, n=6 replicates from three pups per genotype) after treatment of hydrogen peroxide (1 μM) for 30 min. c-e) DIV8 hippocampal neurons from WTand RanBP9+/− mice treated with or without Aβ oligomers for 24 hours and subjected to staining for F-actin (Rhodamine-phalloidin) and cofilin. c) Representative images showing no cofilin rods observed without Aβ oligomers treatment but 16.3% and 6.9% of WTand RanBP9+/− neurons, respectively, contain cofilin rods. Also note the near absence of F-actin containing growth cones in WT neurons treated with Aβ42O but not in RanBP9+/− neurons. d) Quantitation of Cofilin rods per neuron (t-test, *P=0.0329, n=6 replicates) or per NCA (t-test, *P =0.030, n=6 replicates from three pups per genotype). e) Quantitation of F-actin containing growth cones in WT and RanBP9+/− (Ran9+/−) neurons with or without Aβ oligomers treatment (ANOVA, post-hoc Tukey, ***P<0.0001, n=8 replicates from four pups per genotype). f & g) Sholl analysis of neurite arborization/elongation in WT and RanBP9+/− hippocampal neurons on DIV9. f) Representative images of saturated F-actin stain. g) Quantitation of neurite intersections on concentric circles from the soma in 10μm increments (10–200 μm) (two-way ANOVA, post-hoc Bonferroni, *P<0.05, ^P<0.005, #P<0.0005, n=3 replicates from two mice per genotype). Error bars represent S.E.M. in graphs. H-J) DIV8 hippocampal neurons from WT mice with or without control or SSH1 siRNA transfection treated with or without Aβ oligomers for 24 hours and subjected to staining for F-actin and cofilin. h) Representative images showing cofilin and F-actin stains. i) Quantitation of cofilin rods per NCA (t-test, ***P<0.0001, n=6 replicates from three pups per genotype). j) Quantitation of F-actin containing growth cones in neurons transfected with control or SSH1 siRNA with or without Aβ42O treatment (ANOVA, post-hoc Tukey, ***P<0.0001, n=8 replicates from four pups per genotype)

Figure 3-3. RanBP9 mediates the depletion of postsynaptic proteins and F-actin induced by Aβ oligomers in mature primary hippocampal neurons. a-d) DIV21 primary hippocampal neurons derived from P0 RanBP9+/− and WT littermate mice treated with or without Aβ oligomers (1 μM) for 2 hours and subjected to immunocytochemistry for Drebrin, PSD95, and F-actin (Rhodamine-phalloidin). a) Representative images showing Aβ42 oligomer-induced depletion of Drebrin, PSD95, and F-actin in WT neurons but not in RanBP9+/− neurons. b) Quantitation of Drebrin intensity in secondary and tertiary spine-containing dendrites (ANOVA, post- hoc Tukey, ***P<0.0005, n=9 replicates from three pups per genotype). c) Quantitation of PSD95 intensity in secondary and tertiary spine-containing dendrites (ANOVA, post-hoc Tukey, ***P<0.0005, n=10 replicates from three pups per genotype). d) Quantitation of F-actin (Rhodamine-phalloidin) intensity in secondary and tertiary spine-containing dendrites (ANOVA, post-hoc Tukey, **P<0.005, *P<0.05, n=4 replicates from two pups per genotype). Error bars represent S.E.M. on graphs. e & f) Sholl analysis of neurite arborization/elongation in WT and RanBP9+/− hippocampal neurons on DIV21. e) Representative images of saturated F-actin stain. f) Quantitation of neurite intersections on concentric circles from the soma in 10μm increments (10–220 μm) (two-way ANOVA, post-hoc Bonferroni, *P<0.05, #P<0.0005, n=3 replicates from two pups per genotype). Error bars represent S.E.M. in graphs.

Figure 3-4. Increased endogenous RanBP9 levels in APP/PS1 mice a & b) Hippocampal extracts from 8-month-old WT and littermate APP/PS1 mice immunoblotted for the indicated proteins. a) Representative blots showing a robust increase in endogenous RanBP9 protein together with overexpression of human APP (6E10 antibody). b) Quantitation of endogenous RanBP9 protein levels in WT and APP/PS1 hippocampus normalized to WT (t- test, **P=0.0016, n=4 mice per genotype, 2 F and 2 M). c) Representative immunohistochemical images of endogenous RanBP9 immunostaining within the dentate gyrus (DG) and CA3 of the hippocampus.

Figure 3-5. RanBP9 reduction rescues neuroinflammation and synaptic protein depletion in APP/PS1 transgenic mice. a-e) Brains from 8-month-old WT, APP/PS1, and APP/PS1;RanBP9+/− mice subjected to immunohistochemistry for GFAP (activated astrocyte marker), Iba1 (activated microglia marker), Synapsin I (presynaptic marker), and PSD95 (postsynaptic marker) in the anterior cortex (CTX) and/or hippocampus (HIPP). A) Representative images showing RanBP9 reduction rescues neuroinflammation (GFAP and Iba1) and synaptic protein loss (Synapsin I and PSD95) in APP/PS1 mice. b) Quantitation of PSD95 intensity within the SL of CA3 (ANOVA, post-hoc Tukey, *P<0.05, **P<0.005, n=4 mice per genotype, 2 F and 2 M). Quantitation of synapsin I intensity within the SL of CA3 (ANOVA, post-hoc Tukey, *P<0.05, **P<0.005, n=4 mice per genotype, 2 F and 2 M). Quantitation of GFAP intensity in the hippocampus (ANOVA, post-hoc Tukey, *P<0.05, n=4 mice per genotype, 2 F and 2 M). Quantitation of GFAP intensity in the anterior cortex (ANOVA, post-hoc Tukey, **P=<0.005, ***P<0.0005, n=4 mice per genotype, 2 F and 2 M). Quantification of Iba1 intensity in the hippocampus (ANOVA, post-hoc Tukey, ***P<0.0005, n=4 mice per genotype, 2 F and 2 M). Error bars represent S.E.M. on graphs.

Figure 3-6. Mitigation of Aβ accumulation by RanBP9 reduction in APP/PS1 mice. a) Representative immunohistochemical images showing reductions in Aβ deposits (red) in the anterior cortex and hippocampus of 8-month-old APP/PS1;RanBP9+/− mice compared with littermate APP/PS1 mice. b) Representative images showing reductions in thioflavin S-positive fibrillar Aβ deposits in the anterior cortex and hippocampus of 8-month-old APP/PS1;RanBP9+/− mice compared with littermate APP/PS1 mice. c) Quantitation of area covered by Aβ deposits normalized to APP/PS1 mice in the anterior cortex and hippocampus (Cortex: t-test,*P=0.0155, n=4 each; Hippocampus: t-test, **P=0.0067, n=4 mice per genotype, 2 F and 2 M). Quantitation of area covered by thioflavin S-positive fibrillary Ab deposits in the anterior cortex and hippocampus (t-test, ***P<0.0005, n=4 mice per genotype, 2 F and 2 M). Error bars represent S.E.M. in graphs. d) Hippocampal extracts from 6-month-old WT, APP/PS1, and APP/PS1;RanBP9+/− mice immunoblotted for human APP (6E10— Triton-X100 extract) and Aβ (PBS extract). e) Quantification of soluble Aβ levels (PBS extract) from 6-month-old APP/PS1 and APP/PS1;RanBP9+/− hippocampus (t-test, *P=0.0167, n=3 mice per genotype)

Figure 3-7 Accumulation of cofilin–actin rod/aggregate structures in APP/PS1 mice. a-c) Nine-month-old WT, APP/PS1, and APP/PS1;RanBP9+/− littermate brains subjected to immunohistochemistry for cofilin and actin in the anterior cortex and hippocampus using Sudan black to quench background signal. a) Representative images showing increased cofilin–actin rods in APP/PS1 mice and reduction in APP/PS1;RanBP9+/− mice. b & c) Quantitation of cofilin–actin rods in the hippocampus and anterior cortex normalized to WT (ANOVA, post-hoc Tukey, **P<0.005, ***P<0.0005, n=4 mice per genotype). Error bars represent S.E.M. in graphs. d) Representative images of cofilin–actin rods and neuropil threads (phospho-Tau 12E8 antibody) using the same Sudan black quenching immunohistochemical method in the entorhinal cortex from a cognitively normal control (NC) and AD. Note the marked accumulation of cofilin–actin rods (green) and neuropil threads (red) in AD but few in NC.

Figure 3-8 Rescues of synaptic plasticity and contextual memory impairments in APP/PS1 mice by RanBP9 reduction. a-c) Stimulating electrode placed in the Schaffer collaterals of the hippocampus and recording glass electrode positioned at the CA1 stratum radiatum below the pyramidal cell layer. a) Input–output analysis performed by stepping up stimulation amplitude from 1–15mV in WT, APP/PS1, RanBP9+/−, and APP/PS1;RanBP9+/− acute slices. No significant differences observed. Slices from WT n=45, RanBP9+/− n=31, APP/PS1 n=19, APP/PS1;RanBP9+/− n=25; slices derived from four to six mice per genotype. b) PPF showing significant differences among all interstimulus intervals (two-way ANOVA, genotype: P<0.0001; interstimulus interval: P<0.0001; interaction: P=0.0122). Post-hoc Tukey test shows significant increases in fEPSP slope in APP/PS1;RanBP9+/− slices compared with WT, RanBP9+/−, and APP/PS1 slices at nearly all interstimulus intervals (P<0.05 to P<0.0001). Slices from WT n=49, RanBP9+/− n=33. APP/PS1 n=31, APP/PS1;RanBP9+/− n=25; slices derived from four to six mice per genotype. c) No significant changes between genotypes at baseline before LTP but LTP induced by theta burst stimulation showing significant differences in fEPSP slope among WT, APP/PS1, RanBP9+/−, and APP/PS1;RanBP9+/− slices (two-way ANOVA, genotype: P<0.0001; time: P<0.0001; interaction: P<0.0001). Post-hoc Tukey test shows significant differences between all genotypes at all-time points after theta burst stimulation (P<0.0001) except between RanBP9+/− versus APP/PS1;RanBP9+/−. Slices from WT n=41, RanBP9+/− n=28, APP/PS1 n=33, APP/PS1;RanBP9+/− n=29; slices derived from four to six mice per genotype. d) Quantitation of contextual fear conditioning (FC) freezing times (sec) after training session 24 hours earlier across genotypes. Kruskal–Wallis ANOVA, post-hoc Dunn’s, *P<0.05, WT n=12 (5 F and 7 M), APP/PS1 n=8 (4 F and 4 M), APP/PS1;RanBP9+/− n=6 (3 F and 3 M). e) Quantitation of cued fear conditioning (FC) freezing times (sec) across genotypes (no significant differences by Kruskal–Wallis test or one-way ANOVA). Same number and gender as contextual FC. f) Quantitation of open field activity test (total distance traveled over two- day training sessions) across genotypes (no significant differences by Kruskal–Wallis test or one-way ANOVA). Same numbers and gender as contextual FC. g) Quantitation of rotarod test (time staying on rotarod) across genotypes (no significant differences by Kruskal–Wallis test or one- way ANOVA). Error bars represent S.E.M. in graphs. Same number and gender as contextual FC

Supplementary Figure 3-1 Short-term Aβ Monomer Exposure Does Not Alter Cofilin Translocation to Mitochondria or Drebrin/F-Actin Levels

(A) Hippocampus-derived HT22 cells treated with/without A42 monomers for 2h, separated for mitochondrial and cytosol fractions, and subjected to immunoblotting for the indicated proteins. A representative experiment shows no effect of A42 monomers in Cofilin translocation to mitochondria or activation status. (B) DIV21 primary hippocampal neurons derived from P0 RanBP9+/- and wild type (WT) littermate mice treated with or without A42 monomers (1M) for 2h and subjected to immunocytochemistry for Drebrin and F-Actin (Rhodamine-phalloidin). Representative images show no effect of A42 monomers on Drebrin or F-Actin levels.

CHAPTER 4

DISCUSSION

Summary

The major defining pathological hallmarks of Alzheimer’s disease (AD) are the accumulations of amyloid β (A) and hyperphosphorylated Tau, associated with neuroinflammation, mitochondrial dysfunction, cytoskeletal aberrations, and synaptic loss.

Multiple studies have shown that A-induced neurotoxic signals require Tau, since the loss of

Tau abrogates many deleterious effects of A (Morris et al., 2011; Vossel et al., 2015; Vossel et al., 2010). Despite the clear pathogenic link between A and Tau, a large knowledge gap exists in how A pathogenically impinges on Tau. In the preceding studies, we have addressed some of the early and as yet unknown gaps in the neurotoxic signaling pathways resulting from the accumulation of A oligomers at the level of the cell surface and subsequent downstream signaling pathways. Our findings demonstrate the critical involvement of 1-integrin receptor complexes on the neuronal surface and subsequent activation SSH1, resulting in hyperactivation of Cofilin. Such activation leads to neurodegenerative changes in integrin-dependent focal complexes, destabilization of F-Actin, and mitochondrial dysfunction. Specifically, we found that Aβ42 oligomers bind with high affinity to low or intermediate activation conformers of β1- integrin, resulting in the loss of surface β1-integrin and activation of Cofilin via Slingshot homology-1 (SSH1) activation. Conditional loss of β1-integrin prevented Aβ42 oligomer-

induced Cofilin activation, and allosteric modulation or activation of β1-integrin significantly reduced Aβ42 binding to neurons while blocking Aβ42 oligomer-induced reactive oxygen species (ROS) production, mitochondrial dysfunction, depletion of F-actin/focal Vinculin, and apoptosis. Cofilin, in turn, was required for Aβ42 oligomer-induced loss of cell surface β1- integrin, disruption of F-actin/focal Talin–Vinculin, and depletion of F-actin-associated postsynaptic proteins. SSH1 reduction, which mitigated Cofilin activation, prevented Aβ42 oligomer-induced mitochondrial Cofilin translocation and apoptosis, while AD brain mitochondria contained significantly increased activated/oxidized Cofilin. In mechanistic support in vivo, AD mouse model (APP/PS1) brains contained increased SSH1/Cofilin and decreased SSH1/14-3-3 complexes, indicative of SSH1–Cofilin activation via release of SSH1 from 14-3-3. Finally, genetic reduction in Cofilin rescued APP/Aβ-induced synaptic protein loss and gliosis in vivo as well as deficits in long-term potentiation (LTP) and contextual memory in

APP/PS1 mice.

Our findings indicated that activation of SSH1 and Cofilin is not only achieved A- induced oxidation of 14-3-3 but also is mediated by RanBP9. Specifically, we found that endogenous RanBP9 positively regulates SSH1 levels and mediates Aβ-induced translocation of cofilin to mitochondria and induction of cofilin–actin pathology in cultured cells, primary neurons, and in vivo. Endogenous level of RanBP9 was also required for Aβ-induced collapse of growth cones in immature neurons (days in vitro 9 (DIV9)) and depletion of synaptic proteins in mature neurons (DIV21). In vivo, APP/PS1 mice exhibited 3.5-fold increased RanBP9 levels, which is consistent with that observed in brains of AD patients. Accordingly, RanBP9 reduction in APP/PS1 mice protected against cofilin–actin pathology, synaptic damage, gliosis, and Aβ accumulation associated with APP/PS1 mice. Brains slices derived from APP/PS1 mice showed

significantly impaired long-term potentiation (LTP), and RanBP9 reduction significantly enhanced paired pulse facilitation and LTP, as well as partially rescued contextual memory deficits associated with APP/PS1 mice.

Conclusion

These novel findings therefore implicate the essential involvement of the β1-integrin–

SSH1–Cofilin pathway in mitochondrial and synaptic dysfunction in AD and underscore the critical convergence of endogenous RanBP9 not only in Aβ production but also in mediating the neurotoxic actions of Aβ at the level of synaptic plasticity, mitochondria, and cofilin–actin pathology via control of the SSH1-cofilin pathway in vivo. As such, targeting SSH1, a relatively druggable target with a rigid catalytic pocket, may be an attractive and novel therapeutic strategy for AD.

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APPENDIX A

IACUC APPROVAL FOR ANIMAL RESEARCH

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APPENDIX B

The information in chapter 2, “Slingshot-Cofilin Activation Mediates Mitochondrial and

Synaptic Dysfunction” has been legally reproduced under the Creative Commons Attribution

(CC-BY) license. This means the publication is accessible online without any restrictions and can be re-used in any way subject only to proper citation.

Woo, J. A., Zhao, X., Khan, H., Penn, C., Wang, K., Joly-Amado, A., Weeber, E., Morgan, D.,

Kang, D. E (2015). “Slingshot-Cofilin activation mediates mitochondrial and synaptic

dysfunction via Aβ ligation to β1-integrin conformers.” Cell death and differentiation

(2015) 22, 921-934

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APPENDIX C

The information in chapter 3, “RanBP9 at the intersection between Cofilin and Amyloid β pathologies” has been legally reproduced under the Creative Commons Attribution (CC-BY) license. This means the publication is accessible online without any restrictions and can be re- used in any way subject only to proper citation.

Woo, J. A., Boggess, T., Uhlar, C., Wang, X., Cappos, G., Joly-Amado, A., Majid, S., Minamide,

L. S., Bamburg, J. R., Morgan, D., Kang, D. E (2015). “RanBP9 at the intersection

between cofilin and Aβ pathologies: rescue of neurodegenerative changes by RanBP9

reduction.” Cell death and disease (2015) 6, 1676; dio:10.1038

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