Investigating the Role of LIMK1 Signaling in PKA-Dependent LTP in the Hippocampus

Investigating the Role of LIMK1 Signaling in PKA-dependent LTP in the Hippocampus

Sammy Cai

A thesis submitted in conformity with the requirements for the degree of Master of Science Department of Physiology University of Toronto

Investigating the Role of LIMK1 Signaling in PKA-dependent LTP in the Hippocampus

Sammy Cai

Master of Science

Department of Physiology University of Toronto

2019 Abstract Long-term potentiation (LTP) in the hippocampus is the most extensive form of long-lasting synaptic plasticity and is widely regarded as the cellular basis of memory formation. The canonical

NMDA-receptor (NMDAR)-dependent form of LTP can be differentiated into protein kinase A

(PKA)-dependent and -independent forms by the spacing between theta burst stimuli (TBS) induction. Key features of PKA-dependent LTP includes insertion of Ca2+ permeable AMPA receptors (CP-AMPARs) and initiation of de novo protein synthesis. However, the molecular mechanisms that elicit these processes remain unknown. Previous studies suggest PKA can regulate LIM-domain kinase (LIMK) 1, but this regulation has not been demonstrated in synaptic plasticity. Using a combination of electrophysiology, genetic mouse models, and pharmacology, I show that CP-AMPAR-dependent LTP requires PKA regardless of TBS spacing. Furthermore, I demonstrate that LIMK1 is required for PKA-dependent LTP. These findings provide insight into the molecular mechanisms underlying LTP and long-term memory formation.

Acknowledgments

The past two and a half years have been the most challenging and interesting years of my life that I will always remember. I would like to thank my supervisor, Dr. Zhengping Jia, for waking me up from an afternoon nap in a hotel room in Brussels via email on May 23, 2016 – little did I know that this would have been the beginning of my graduate school career. His patience and guidance have made me a better scientist and overall, a more resilient person. I am grateful for my co-supervisor, Dr. Graham L. Collingridge, for his willingness to accept a Type 2 Diabetes researcher into his first cohort of neuroscientists at the University of Toronto. His optimism and enthusiasm being ever so contagious, is what fueled my interest in the neurosciences. I am thankful for having Dr. Kenichi Okamoto and Dr. William Trimble for being part of my supervisory committee. They provided invaluable insight and helped develop both my project and my professional skills for a successful career.

Although I was primarily secluded to the electrophysiology room (partly due to my experiments, but mainly due to the tropical climate), I will never forget the fun I had with each of the Jia lab members – they never failed to make me laugh. Before I begin, I would like to offer my sincerest apologies to all those who experienced my terrible taste in music upon stepping foot in

05.9475. At the very least, I hope it blocked out the horrid sound of the peristaltic pumps and carbogen perfusion tubes. Thanks to (in alphabetical order): Amir, for always giving me another

“perspective” on things; Catherine, for your never-ending supply of kindness; Celeste, for being alright*; Feng, for being my partner in crime in the electrophysiology room; Kate, for reminding me to appreciate the little things life; Neil, for having all the answers when I did not; Nicole, for being delightful during the (cumulative) three-hour period we interacted; Sarah, for reminding me that I am not as young as the others make it out to be; Susan, for being one of the first and last

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person to see throughout my day; and Youssif, for inspiring me to dream big. To past members of the Jia lab: Amy, for giving me a source of enthusiasm when I did not; Joyce, for teaching me the importance of phrasing; Shuting, for teaching me what I believe, is one of the most complicated techniques in neuroscience; Tony, for being my mentor during the two weeks our careers actually overlapped.

Of course, I would like to mention the wonderful people I have met and worked with during my involvement in two major extracurricular activities that have shaped my career. Firstly, to the friends I have met through the Graduate Association of Students in Physiology; it is heartwarming to know that there will always be someone I know no matter where I am on campus. Secondly, to

Team ACES for showing me it is possible to have fun while doing work. Thanks to: Ankur, for being by my side throughout all the crazy adventures/projects and overall being the older brother that I never had; Celeste, *for believing in me when I did not, and encouraging me to go beyond what I thought was capable; Erika, for inspiring me to follow my passion and showing me that no obstacle is too great.

A shout-out to Frankie, for starting it all – if it were not for you, I would not have considered research as part of my career and this manuscript probably would not exist. I am so glad to have met you during my undergraduate years and my only regret was not meeting you earlier.

This one is to my brothers: Ben, Kevin, Jamie, Matthew Zhang, Michael, Nathan,

Thanvin. Thank you for being with me as early as preschool. Thank you for being with me through all the highs and lows. Thank you all for always being there for me no matter where I go and wherever you are. You guys were always my excuse for being “busy” and there does not go a day where I do not reference at least one of you in one of my conversations.

Lastly, I would like to thank my family: my mother Doris, my father Shawn, and my

(actual) brother Jason, for the love and support they have provided - there does not go a day where

I do not think about any of you. Words, unfortunately, cannot describe how thankful I am to have my friends and family support me through this journey. Everyone has taught me something valuable and I hope I have returned the favour to everyone I have met throughout my graduate school career.

Table of Contents

ACKNOWLEDGMENTS ...... III

TABLE OF CONTENTS ...... VI

LIST OF FIGURES ...... IX

LIST OF ABBREVIATIONS ...... XII

INTRODUCTION ...... 1 1.1 TYPES OF MEMORY ...... 1 1.2 THE HIPPOCAMPUS AND MEMORY FORMATION ...... 4 1.3 THE HIPPOCAMPAL CIRCUITRY ...... 5 1.4 THE SYNAPSE ...... 7 1.4.1 Synaptic Architecture ...... 7 1.4.2 NMDARs ...... 11 1.4.3 AMPARs ...... 14 1.4.4 CP-AMPARs ...... 17 1.5 SYNAPTIC PLASTICITY AND LTP ...... 18 1.5.1 Discovery of LTP ...... 18 1.5.2 General Biochemical Mechanism ...... 19 1.5.3 Early-Phase LTP ...... 23 1.5.4 Late-Phase LTP ...... 24 1.5.5 CP-AMPARs in LTP ...... 28 1.6 PKA ...... 31 1.6.1 General Overview ...... 31 1.6.2 Structure and Activation ...... 32 1.6.3 Role of PKA ...... 33 1.7 THE CORRELATION BETWEEN STRUCTURAL PLASTICITY AND SYNAPTIC PLASTICITY ...... 35 1.7.1 Structural Plasticity ...... 35 1.7.2 Cytoskeletal Structure ...... 37 1.7.3 Actin-mediated Receptor Trafficking ...... 39 1.7.4 Receptor-mediated Actin Stabilization ...... 41 1.7.5 Rho-GTPases ...... 41 1.7.6 Downstream Effectors: PAK, ROCK ...... 43 1.8 A COMMON EFFECTOR: LIMK ...... 45 1.8.1 General Overview ...... 45

1.8.2 Structure and Activation ...... 46 1.8.3 Downstream Effectors ...... 48 1.9 CLINICAL RELEVANCE ...... 50 1.9.1 Synaptic Plasticity in Health and Disease ...... 50 1.9.2 Williams Syndrome ...... 50 1.9.3 Alzheimer’s Disease ...... 52

RATIONALE, HYPOTHESIS, AND OBJECTIVES ...... 55 2.1 RATIONALE ...... 55 2.2 HYPOTHESIS ...... 57

2.3 OBJECTIVES ...... 57

MATERIALS AND METHODS ...... 59 3.1 ANIMAL COLONY ...... 59 3.1.1 GluA2 and LIMK1 Mutant Mice ...... 59 3.2 GENOTYPING ...... 59 3.2.1 DNA Extraction ...... 59 3.2.2 Polymerase Chain Reaction ...... 60 3.2.3 Gel Electrophoresis ...... 61 3.3 ELECTROPHYSIOLOGY ...... 61 3.3.1 Acute Hippocampal Slice Preparation ...... 61 3.3.2 Extracellular Electrophysiology ...... 62 3.4 COMPOUNDS ...... 63 3.5 ANALYSIS ...... 63

RESULTS ...... 64 4.1 CTBS AND STBS INDUCED ENHANCED LTP IN GLUA2 KO MICE ...... 64 4.2 CTBS AND STBS INDUCED LTP USING ONLY CP-AMPARS IN GLUA2 KO MICE ...... 67 4.3 STBS-INDUCED CP-AMPAR-DEPENDENT LTP REQUIRED DE NOVO PROTEIN SYNTHESIS ...... 73 4.4 INHIBITION OF PKA TRANSIENTLY ABOLISHED CP-AMPAR-DEPENDENT LTP ...... 75

4.5 CP-AMPAR-DEPENDENT LTP REQUIRES PKA ...... 78 4.6 LIMK1 IS REQUIRED FOR STBS-INDUCED LTP ...... 81 4.7 LIMK1 IS REQUIRED FOR PKA-DEPENDENT LTP ...... 84 4.8 LIMK1 IS REQUIRED FOR DE NOVO PROTEIN SYNTHESIS IN PKA-DEPENDENT LTP ...... 89

DISCUSSION ...... 94 5.1 CP-AMPAR-DEPENDENT LTP IS MEDIATED BY PKA ...... 95 5.2 A ROLE FOR LIMK1 IN STBS-INDUCED PKA-DEPENDENT LTP ...... 97 vii

5.3 PKA-MEDIATED PROTEIN SYNTHESIS-DEPENDENT LTP REQUIRES LIMK1 ...... 98

FUTURE DIRECTIONS ...... 101

CONCLUSION ...... 103

REFERENCES ...... 104

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List of Figures

Figure 1. Subdivisions of memory...... 3

Figure 2. Schematic of the hippocampal formation...... 6

Figure 3. Schematic of an electrical synapse...... 8

Figure 4. Schematic of a chemical synapse ...... 10

Figure 5. NMDAR structure and subunit composition...... 13

Figure 6. AMPAR structure and subunit composition...... 16

Figure 7. Potentiation requires delivery of tetanic electrical stimuli...... 20

Figure 8. LTP induction...... 22

Figure 9. PKA-mediated changes in late-phase LTP...... 26

Figure 10. CP-AMPARs in LTP...... 30

Figure 11. Activation of PKA. PKA is a tetrameric enzyme that is composed of two regulatory and two catalytic subunits...... 32

Figure 12. Synaptic plasticity is correlated with structural plasticity...... 36

Figure 13. Spine structure and morphology is dependent on actin cytoskeleton dynamics ...... 40

Figure 14. Rho GTPase activation and inactivation...... 42

Figure 15. LIMK1 is the common downstream target for RhoA, Rac, and Cdc42 ...... 43

Figure 16. LIMK1 phosphorylation sites...... 47

Figure 17. Cofilin activity is regulated by LIMK1 ...... 49

Figure 18. Enhanced cTBS-induced LTP in GluA2 KO mice...... 65

Figure 19. Enhanced sTBS-induced LTP in GluA2 KO mice...... 66

Figure 20. NMDARs are required for cTBS-induced LTP in GluA2 WT mice...... 69

Figure 21. NMDARs are required for sTBS-induced LTP in GluA2 WT mice...... 70

Figure 22. GluA2 KO mice exhibit cTBS-induced LTP independent of NMDARs ...... 71

Figure 23. GluA2 KO mice exhibit sTBS-induced LTP independent of NMDARs ...... 72

Figure 24. sTBS-induced LTP requires protein synthesis in GluA2 KO mice...... 74

Figure 25. Inhibition of PKA transiently abolishes cTBS-induced LTP in GluA2 KO mice ...... 76

Figure 26. Inhibition of PKA transiently abolishes sTBS-induced LTP in GluA2 KO mice...... 77

Figure 27. Blockade of sTBS-induced LTP is dependent on the duration of PKA inhibition in GluA2 KO mice...... 79

Figure 28. PKA is required for sTBS-induced LTP in GluA2 KO mice ...... 80

Figure 29. LIMK1 is not required for cTBS-induced LTP...... 82

Figure 30. LIMK1 is required for sTBS-induced LTP ...... 83

Figure 31. PKA is not required for cTBS-induced LTP ...... 85

Figure 32. PKA is required for sTBS-induced LTP...... 86

Figure 33. LIMK1 is not required for PKA-independent LTP ...... 87

Figure 34. LIMK1 is required for PKA-dependent LTP ...... 88

Figure 35. De novo protein synthesis is not required for PKA-independent LTP ...... 90

Figure 36. De novo protein synthesis is required for PKA-dependent LTP ...... 91

Figure 37. De novo protein synthesis and LIMK1 is not required for PKA-independent LTP.. .. 92

Figure 38. PKA-dependent LTP requires LIMK1-mediated de novo protein synthesis ...... 93

Figure 39. sTBS-induced LTP via NMDAR or CP-AMPAR requires PKA-LIMK1-mediated de novo protein synthesis...... 100

List of Abbreviations

AD Alzheimer’s Disease AMPAR α-amino-5-hydroxy-3-nethyl-4-isoxazole propionic acid receptor Aβ amyloid β BDNF Brain-Derived Neurotrophic Factor C Catalytic Subunit CA Cornu Ammonis CaM Calmodulin CaMKII Ca2+/calmodulin-dependent protein kinase II cAMP Cyclic Adenosine Monophosphate Cdc42 Cell Division Cycle 42 CI Ca2+ Impermeable CNQX 6-cyano-7-nitroquinoxaline-2,3-dione CNS Central Nervous System CP Ca2+ Permeable CREB cAMP Response Element Binding Protein cTBS Compressed Theta Burst Stimuli CTD C-Terminal Domain DG Dentate Gyrus DNA Deoxyribonucleic Acid EC Entorhinal Cortex E-LTP Early-Phase LTP ERK/MAPK Extracellular Response Kinase/Mitogen Activated Protein Kinase F-actin Filamentous Actin G-actin Globular Actin GAPs GTPase Activating Proteins GDP Guanosine Diphosphate GEFs Guanine Nucleotide Exchange Factors GRIP Glutamate Receptor-Interacting Protein GTPases Guanosine Triphosphate Hydrolases HFS High Frequency Stimulation HSP90 Heat Shock Protein 90 IEGs Immediate Early Genes KO Knockout LBD Ligand Binding Domain LIMK LIM-domain-containing Protein Kinase L-LTP Late-Phase LTP LTM Long-Term Memory LTP Long-Term Potentiation MAPKAPK2 MAPK Activated Protein Kinase II MTOCs Microtubule Organizing Centres N Number of Active Channels NMDAR N-methyl-D-aspartic Acid Receptor NSF N-ethylmaleimide Sensitive Factor NTD N-Terminal Domain PAK p21-Activated Kinase

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PI3K Phosphoinositide 3-Kinase PICK1 Protein Interacting With C Kinase-1 PKA Protein Kinase A Po Open Probability PSD Postsynaptic Density Rac Ras-related C3 Botulinum Toxin Substrate RhoA Ras Homologous Member A RNA Ribonucleic Acid ROCKs Rho-Associated Kinases sTBS Spaced Theta Burst Stimuli STM Short-Term Memory TARPs Transmembrane AMPA Regulatory Proteins TBS Theta Burst Stimulation WS Williams Syndrome γ Conductance

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Chapter 1 Introduction 1.1 Types of Memory Who are we? The answer to one of the fundamental questions of life comes from the collection of unique individual experiences stored as memories within the brain – it is from these memories that we learn who we are and our ultimate purpose. Failure to form memories results in the inability to learn from past experiences, develop relationships, or have a sense of personal identity. The memories that govern these processes are synthesized from complex sensory information received by our brains. How this information is encoded is complex; ensembles of processes are required to be fluid and adaptable for efficient and effective recollection and alteration, but stable and specific for accurate integration of information. This dichotomy underlies the differences between short-term memory (STM) and long-term memory (LTM), respectively

(Kandel, 2001).

In the context of neurobiology, memories can be divided into two distinct temporal categories, with STM being comprised of two subcategories (Figure 1) (Aben, Stapert, &

Blokland, 2012). The first of which is immediate memory – the ability of the mind to store and catalogue information within an extremely brief timeframe, typically in the order of fractions of a second. This form of memory has a large capacity but is extremely limited in duration. It is continuously active to perceive sensory modalities such as, the visual, auditory, olfactory, and tactility of surrounding objects and environment. An example of immediate memory includes registering the colour or texture of an object (Aben et al., 2012; Purves et al., 2012). The second temporal category includes working memory – the ability of the mind to store information for a

brief timeframe, typically in the order of seconds, to achieve a particular task (Purves et al., 2012).

An example of working memory is recalling the name of a new acquaintance. The difference between immediate memory and working memory is that the former recognizes sensory inputs whereas the latter requires internalization (Aben et al., 2012). Thus, working memory has been suggested to be the fundamental process for reasoning and problem-solving. Although working memory is limited in duration and capacity, it is possible to convert working memory into a LTM with active rehearsal and persistent reactivation of memories (Purves et al., 2012).

The second temporal category is LTM – the ability to retain information in the order of days, weeks, or a lifetime. LTM can be subdivided into implicit and explicit memories (Purves et al., 2012). Implicit memory, sometimes referred to as procedural memory, is a form of unconscious processes that provide us with the ability to use objects and navigate our body for a particular task, such as using a pencil or riding a bicycle (Anderson, Morris, Amaral, Bliss, & O’Keefe, 2007).

Explicit memory is a form of memory that is consciously available and can be verbalized with relative ease compared to implicit memories. Explicit memories can be subdivided into three categories: episodic memory – the ability to refer to specific memories in time, such as recalling an interaction with a colleague; semantic memory – the ability to recollect factual information, such as remembering information for tests; autobiographical memory – the ability to recall experiences pertaining to an individual’s own life, such as recalling “first-time” experiences

(Figure 1) (Anderson et al., 2007). What is unique about LTM, more specifically explicit memories, is that these memories are encoded, but not consolidated, by a region in the brain known as the hippocampus. This information is typically dispersed to different regions of the brain for consolidation and storage. However, since most processes required for explicit memories are

routed through the hippocampus for encoding, this suggests the hippocampus has a central role in learning and memory.

Figure 1. Subdivisions of memory. Memory can be subdivided into two temporal categories: Short-term memory (STM) and long-term memory (LTM). STM is comprised of two subcategories: immediate memory and working memory. LTM is comprised of two subcategories: implicit memory and explicit memory, with explicit memory being further subdivided into episodic, semantic, and autobiographical memories.

1.2 The Hippocampus and Memory Formation Over centuries, the hippocampus has been proposed to have a wide host of functions, ranging from olfaction, emotion, and memory (Anderson et al., 2007; Papez, 1937). However, the importance of the hippocampus in memory formation was highlighted by William Beecher

Scoville and Brenda Milner and their epileptic patient Henry Molaison. To relieve his drug- resistant seizures, Scoville removed the medial aspect of Henry Molaison’s temporal lobe, which included the hippocampal formation and adjacent structures. While this reduced the severity of his seizures, Henry Molaison was left with global amnesia that persisted throughout the days following his operation. It was interesting to note that his STM and most LTM prior to the surgery was relatively intact (Scoville & Milner, 1957). Although this provides evidence for the importance of the hippocampus in encoding memories, it raises several questions: where does the hippocampus receive incoming information from? Where is this information relayed to? Which regions of the brain is this information consolidated? How does the hippocampus encode memories for consolidation?

Following the seminal study by Scoville and Milner, a plethora of studies began investigating the functional role of the hippocampus and related structures in memory formation.

Extrinsically, the hippocampus has connections with numerous subcortical inputs, such as the amygdala and hypothalamus, which primarily support functions pertaining to emotion, behavior, and motivation, and the thalamus, which relays sensory and motor signals (Aggleton et al., 2010;

Pitkanen, Pikkarainen, Nurminen, & Ylinen, 2000). More recently, there has been evidence that suggests that reciprocal communication exists between the hippocampus and various neocortical areas. Specifically, the hippocampus can receive sensory information from various neocortical areas and encode and project LTM for consolidation back to its respective input regions (Lavenex

& Amaral, 2000). Regardless of the sensory inputs and outputs for consolidation, the hippocampus has been widely regarded as a necessary component for encoding STM into LTM (Lynch, Rex, &

Gall, 2007). This suggests that the architecture of the hippocampus, namely the internal circuitry, contains the necessary molecular processes for encoding to occur.

1.3 The Hippocampal Circuitry The hippocampus is part of a compound structure known as the hippocampal formation, which consists of the entorhinal cortex, dentate gyrus, the hippocampus proper, and the subiculum

(Schultz & Engelhardt, 2014). Unlike the reciprocal nature of the connections between neocortical structures, the projections within the hippocampal formation is unidirectional. Beginning with the entorhinal cortex (EC), cells in the superficial layer project to the dentate gyrus (DG) to form the perforant path, with no projections from the DG returning back to the EC (Figure 2A). Likewise, the DG sends projections from granule cells, which comprises the mossy fibres, to the hippocampus proper, specifically the region Cornu Ammonis (CA) 3 (Figure 2B). In turn, the CA3 field sends its projections to the CA1 field via the Schaffer collateral axons (Figure 2C). The CA1 field then projects to both the subiculum and EC (Figure 2D). The hippocampal circuitry is then closed by having the CA1 and majority of the subiculum project to the EC, with parts of the connections from the subiculum projecting to the presubiculum and the parasubiculum (Figure 2E)

(Anderson et al., 2007).

In the context of learning and memory, the Schaffer collateral pathway between the CA3 and CA1 field regions have been of particular interest. This is in part due to the high density of excitatory connections within the CA1 pyramidal neurons and relative ease to perform extracellular and intracellular electrophysiological recordings (Megias, Emri, Fruend, & Gulyas,

2001). The robust nature of this model has contributed to tremendous advances in understanding the synapse, synaptic transmission, and synaptic plasticity within the central nervous system.

Figure 2. Schematic of the hippocampal formation. A. The circuit of the hippocampal formation begins in the entorhinal cortex (EC) and projects to the dentate gyrus (DG). B. The DG sends its projections, which form the mossy fibres, unilaterally to the CA3 region of the hippocampus proper. C. The CA3 region sends its projections unilaterally to the CA1 region via the Schaffer Collaterals. D. The CA1 region primarily projects its axons to the subiculum. E. The hippocampal circuitry is completed by receiving projections from both the CA1 and the subiculum. Parts of the subiculum project to the presubiculum and the parasubiculum.

1.4 The Synapse

1.4.1 Synaptic Architecture Originally introduced by Charles Sherrington, the “synapse” is an organelle in the nervous system that permits the transmission of signals from one neuron to another or a target efferent cell.

In the nervous system, the synaptic connection is comprised of a pre- and postsynaptic neuron

(Sherrington, 1906). Fundamentally, there are two types of synapses: electrical and chemical. In an electrical synapse, the pre- and postsynaptic membranes are interconnected by specialized intracellular molecules called gap junctions that connect the cytoplasm between the two cells

(Figure 3). This allows for direct flow of molecules, ions, and electrical impulses from one cell to another. One of the consequences of this direct connection is that communication between these two cells can be bidirectional, hence the terms “pre-”and “postsynapse” in an electrical synapse refer to the origin and recipient of the electrical impulse, respectively (Purves et al., 2012).

Figure 3. Schematic of an electrical synapse. An electrical synapse is composed of two neurons interconnected by specialized intracellular molecules called gap junctions. Gap junctions connect the cytoplasm between two cells and allow for direct ion flow from one cell to another; thus, communication can be bidirectional. The terms pre- and postsynaptic neurons refer to the origin and recipient of the electrical impulse, respectively.

The other type of synapse is a chemical synapse, which is comprised of three components: the presynaptic neuron, the synaptic cleft, and the post synaptic neuron (Figure 4). Compared to an electrical synapse, the space between the pre- and postsynaptic neuron is substantially larger in a chemical synapse (Hormuzdi, Filippov, Mitropoulou, Monyer, & Bruzzone, 2004). In a chemical synapse, the presynaptic neuron contains membrane-bound organelles called “synaptic vesicles”, each containing one or more neurotransmitters (Volknandt, 1995). These neurotransmitters act as a mode of communication between pre- and postsynaptic neurons. The release of neurotransmitters

is an intricate process; it is initiated when an action potential, a phenomenon that results in the rapid depolarization and repolarization of the membrane potential of an axon, propagates towards the axon terminal of the presynaptic neuron. This depolarization results in activation of voltage- gated Ca2+ channels and subsequent influx of Ca2+. This elevation of Ca2+ concentration allows for fusion of synaptic vesicles to the membrane of the presynaptic neuron and results in the exocytosis of neurotransmitters into the synaptic cleft. The neurotransmitters then diffuse across the synaptic cleft and bind to specific receptors located on the membrane of the dendritic spine of a postsynaptic neuron (Purves et al., 2012). This postsynaptic region is known as the postsynaptic density (PSD) and contains a high concentration of neurotransmitter receptors and associated kinases and phosphatases critical for signal transduction. Together, the site of neurotransmitter release, synaptic cleft, and the PSD forms an active zone (Okabe, 2007). Depending on the neurotransmitters and the receptors, the receptors will either open or close and change the flow of ions in or out of the postsynaptic neuron, all of which can either increase or decrease the probability of that neuron to fire an action potential across the dendrite. To ensure transmission of information from one neuron to another is transient and accurate, the neurotransmitters are subsequently removed to terminate the signal, either via reuptake through neighbouring glial cells or enzymatic degradation. Neurotransmission at chemical synapses is the predominant means of neuronal communication in the mammalian brain and many other nervous systems. Therefore, I will focus my discussion on this type of synapse.

Figure 4. Schematic of a chemical synapse. A chemical synapse is composed of two neurons separated by a large space known as the synaptic cleft. Following excitation, the presynaptic neuron will release vesicles containing neurotransmitters. The neurotransmitters will diffuse across the synaptic cleft and bind to its respective postsynaptic receptors for signal transduction.

The principal excitatory neurotransmitter in the hippocampus, as elsewhere in the mammalian central nervous system (CNS), is glutamate. Furthermore, in the hippocampus, the chemical synapse has been the most characterized in the context of learning and memory. Upon excitation, the axon terminals of the presynaptic pyramidal neurons from the CA3 field region release glutamate upon the dendrites of the postsynaptic pyramidal neurons from the CA1 field region (Meldrum, 2000). In glutamatergic neurotransmission, glutamate binds to and activates a wide host of receptors. These include the ionotropic N-methyl-D-aspartic acid receptor (NMDAR) and α-amino-5-hydroxy-3-nethyl-4-isoxazole propionic acid receptor (AMPAR).

1.4.2 NMDARs Discovered over half a century ago, NMDARs are glutamate-gated ion channels that mediate long lasting synaptic changes within the CNS (Paoletti, Bellone, & Zhou, 2013).

NMDARs are heterotetrameric assemblies consisting of four subunits that vary throughout development. Seven subunits have been identified and is further subdivided into three subfamilies according to sequence homology: the GluN1 subunit, four GluN2 subunits (GluN2A, GluN2B,

GluN2C, and GluN2D), and two GluN3 subunits (GluN3A and GluN3B). All of these subunits are encoded by separate genes and can impart unique biophysical and pharmacological properties, dictate molecular interactions, and regulate subcellular localization (Paoletti & Neyton, 2007). As heterotetrameric assemblies, NMDARs are always comprised of GluN1 subunits with the addition of either GluN2 subunits alone or with a mixture of GluN2 and GluN3 subunits (Figure 5B).

Throughout development, the GluN1-containing NMDARs are ubiquitously expressed throughout the CNS, with increasing heterogeneity until adulthood is reached. However, in the hippocampus, the NMDARs are typically comprised of GluN1/2A and GluN1/2B subunits. All of these subunits consist of four discrete regions: an extracellular-facing globular N-terminal domain (NTD) that is

involved in subunit assembly and allosteric regulation; a ligand binding domain (LBD) that binds glycine or D-serine (found on GluN1 and GluN3 subunits) and glutamate (found on GluN2 subunits); a transmembrane domain that acts as an ion selectivity filter; and an intracellular C- terminal domain (CTD) that allows for intracellular interactions, such as receptor trafficking, anchoring, and signaling (Figure 5A) (Paoletti et al., 2013).

The complex interaction between subunit composition and molecular partners of NMDARs allows it to act as a coincidence detector for membrane depolarization and synaptic transmission.

This is particularly important in the context of learning and memory, as the combination of the following properties of the NMDARs allow postsynaptic neurons to differentiate between spontaneous firing and correlative firing from the presynaptic neuron (T. V Bliss & Collingridge,

1993). At basal activity levels, the NMDARs are blocked by extracellular Mg2+, therefore, do not contribute to synaptic transmission. However, with sufficient postsynaptic depolarization, the

Mg2+ blockade is rapidly displaced. The activation of NMDARs also requires the binding of two molecules of glutamate, acting as the agonist, and two molecules of glycine (or D-serine), acting as a co-agonist (Vyklicky, Korinek, Smejkalova, Balik, & Krausova, 2014). Since glycine is abundantly found in the surrounding extracellular environment, the co-agonist binding sites are naturally occupied. Upon the release of glutamate from the presynaptic terminal and the binding of glutamate on the NMDAR, the NMDAR channel opens and allows for the selective intake of cations, namely Na+, Ca2+, and K+ (Paoletti et al., 2013).

Although the NMDAR allows for the influx of Na+, Ca2+, and K+, the relative permeability for the ions are different. More specifically, the differential permeability is pronounced with respect to Ca2+, with permeability of Ca2+ being 10 times higher than that of Na+ (Paoletti &

Neyton, 2007). This influx of Ca2+ results in widespread signal transduction, which consists of activation of Ca2+-dependent enzymes, second messengers, protein kinases, phosphatases, scaffolding proteins, and changes to cytoskeletal structure, many of which are required for long lasting synaptic changes following learning and memory formation (Paoletti et al., 2013). These signaling pathways can then result in the recruitment and/or modification of AMPARs.

Figure 5. NMDAR structure and subunit composition. A. Each NMDAR subunit is comprised of four distinct regions: an extracellular-facing globular N-terminal domain (NTD) involved in subunit assembly and allosteric regulation; a ligand binding domain (LBD) that binds glycine or D-serine (found on GluN1 and GluN3 subunits) and glutamate (found on GluN2 subunits); a transmembrane domain (TMD) that acts as an ion selectivity filter; and an intracellular C-terminal domain (CTD) that allows for intracellular interactions, such as receptor trafficking, anchoring, and signaling. B. There are many different populations of NMDARs that are di- or tri-heteromeric.

1.4.3 AMPARs AMPARs are glutamate-gated ion channels that mediate fast synaptic transmission within the CNS, more specifically in neurons and glia. Similar to NMDARs, AMPARs are heterotetrameric assemblies consisting of four subunits that also vary throughout development.

Four subunits have been identified: GluA1, GluA2, GluA3, and GluA4. These subunits form

“dimer of dimers” that are symmetrical, typically consisting of GluA2 and either GluA1, GluA3, or GluA4 subunits, although there have been studies reporting the presence of homomeric dimers

(W. Lu et al., 2009; Wenthold, Petralia, & Niedzielski, 1996). Each of these subunits are encoded by separate genes, and each subunit can not only regulate AMPAR trafficking and regional expression, but more importantly control synaptic development and plasticity (Henley &

Wilkinson, 2016). In the adult hippocampus, majority of the synaptic AMPARs are comprised of

GluA1 and GluA2 subunits (Henley & Wilkinson, 2013). Each subunit has similar membrane topology and is comprised of four discrete regions that parallel that of the NMDAR: an extracellular NTD, a LBD, a transmembrane domain that acts as an ion selectivity filter, and an intracellular CTD. The CTD varies between each of the AMPAR subunits and determines the intracellular trafficking, channel open probability, and channel conductance; all of which are features that change the efficacy of synaptic transmission following patterned neuronal activity

(Figure 6A) (Ziff, 2007).

AMPARs can be found at postsynaptic membranes of glutamatergic synapses at basal activity levels. One of the unique properties of AMPARs that differentiates it from NMDARs is its high mobility (Borgdorff & Choquet, 2002). The dynamic nature of the AMPAR allows it to not only mobilize laterally across the surface of the cell membrane, but also undergo constitutive trafficking to and from the cell surface. In the hippocampus, these changes are largely mediated

by the Ca2+/calmodulin-dependent protein kinase II (CaMKII)-dependent phosphorylation of the

GluA1 subunit. Upon arrival at the cell membrane, the AMPARs can diffuse laterally towards the

PSD. Although AMPARs bind to glutamate, it interestingly has a low affinity for it (Chater &

Goda, 2014). Thus, in order for AMPAR to bind and respond to glutamate, AMPARs need to be located within the PSD, which is relatively in close proximity to the presynaptic active zone.

During synaptic transmission, the active zone will become highly saturated with glutamate and ensures binding of glutamate to AMPARs (Lisman, Ysuda, & Raghavachari, 2014). Following the binding of glutamate to each of the four subunits, the pore opens and allows for the influx of Na+ and efflux of K+, which rapidly depolarizes the postsynaptic neuron. Majority of the AMPARs in the adult brain are Ca2+-impermeable (CI), however, with the proper subunit composition and RNA editing, AMPARs can become Ca2+-permeable (CP) and facilitate long lasting changes that were previously primarily associated with NMDARs (Figure 6B) (Chater & Goda, 2014).

Figure 6. AMPAR structure and subunit composition. A. Each AMPAR subunit has similar membrane topology and is comprised of four discrete regions: an extracellular NTD, an LBD, a transmembrane domain that acts as an ion selectivity filter, and an intracellular CTD. The CTD varies between each of the AMPAR subunits and determines the intracellular trafficking, channel open probability, and channel conductance; all of which are features that change the efficacy of synaptic transmission following patterned neuronal activity. The glutamine/arginine (Q/R) editing site is present on the GluA2 subunit. Typically, Ca2+-impermeable (CI), if AMPARs contain an unedited GluA2 (Q) subunit, it will be Ca2+-permeable (CP). B. There are many different populations of AMPARs. Majority of AMPARs are comprised of GluA1 and GluA2 subunits. GluA2-lacking AMPARs are CP.

In addition to direct phosphorylation, protein-protein interactions at AMPARs are also important for alterations in receptor channel properties and trafficking. A large number of proteins interact with AMPARs via individual subunits, particularly the GluA2 subunit (Hanley, 2014).

The three most characterized mediators associated with the GluA2 subunit include glutamate receptor-interacting protein (GRIP), protein interacting with C kinase-1 (PICK1), and N- ethylmaleimide sensitive factor (NSF) (Isaac, Ashby, & Mcbain, 2007). In particular, PICK1 triggers selective internalization of GluA2-containing AMPARs, which subsequently results in overall increases in CP-AMPAR expression within the hippocampus. In contrast, GRIP and NSF promote expression of GluA2-containing AMPARs as they are involved in anchoring and disrupting PICK1 interactions, respectively (Hanley, Khatri, Hanson, Ziff, & Louis, 2002).

Together, these protein-protein interactions enhance the number of CI-AMPARs to the synapse.

This then brings to question, which protein-protein interactions are involved in CP-AMPAR trafficking? Of interest are transmembrane AMPA regulatory proteins (TARPs) as they play a role in lateral receptor trafficking, a property that is exhibited by CP-AMPARs following specific stimulation parameters (Hanley, 2014; Park et al., 2016).

1.4.4 CP-AMPARs The permeability of AMPARs to Ca2+ depends on the presence of the GluA2 subunit and whether it has undergone RNA editing (Figure 6A). This post-transcriptional process modifies the

GluA2 mRNA from its original CP glutamine (Gln; Q) encoding codon into a CI arginine (Arg;

R). Thus, CP-AMPARs either lack GluA2 subunits or contain unedited GluA2 (Q) subunits, whereas CI-AMPARs contain edited GluA2 (R) subunits (Isaac et al., 2007). Although studies have shown strong evidence that majority of the CP-AMPARs are GluA2-lacking, with less than

1% of RNA across the gray matter containing unedited GluA2 subunits, current electrophysiological techniques are unable to differentiate the two types of CP-AMPARs at the synapse (Wright & Vissel, 2012). Because of this differential expression, the general assumption is that observed CP-AMPARs are GluA2-lacking rather than the unedited isoform. Regardless of the type of CP-AMPARs, the evidence thus far suggests that the two are functionally identical.

CP-AMPARs are important in learning and memory as they offer an alternative mechanism for

Ca2+ influx and contribute to NMDAR-independent forms of synaptic plasticity and long-term potentiation (LTP) (Jia et al., 1996; Wright & Vissel, 2012).

1.5 Synaptic Plasticity and LTP

1.5.1 Discovery of LTP Synaptic plasticity is the ability of synapses to strengthen or weaken in response to increases or decreases in activity over time. These changes are not random; the modifications in synaptic efficacy are specific to the two cells that have been activated simultaneously, an idea refined by Hebb and Konorski in the late 1940s. Colloquially, this idea is known as “neurons that fire together, wire together” (Shatz, 1992). Since connections between neurons are predominantly formed via synapses, synaptic plasticity is believed to be the fundamental mechanism for learning and memory (Kandel, Dudai, & Mayford, 2014; Mayford, Siegelbaum, & Kandel, 2012).

Following the initial observations made by Lomo in 1966, the persistent strengthening of synapses was termed LTP (Lomo, 2003). LTP was later described in detail by Bliss and Lomo in

1973 following the experiments on the hippocampi of anaesthetized rabbits, specifically investigating the synapses between the perforant path and the dentate gyrus (T. V. P. Bliss &

Lomo, 1973; Lomo, 2003). Since then, LTP has been discovered across all excitatory pathways

within the hippocampus, and in several other brain regions to a certain extent, and thus hippocampal LTP has been used as the primary experimental model for understanding the synaptic basis of learning and memory in vertebrates (Nicoll, 2017).

1.5.2 General Biochemical Mechanism Potentiation is activity-dependent and is typically recorded from individual cells or a population of neurons following the delivery of tetanic electrical stimuli. While many stimulation protocols exist, the two most characterized methods are high frequency stimulation (HFS), which typically consists of an “episode” of 50-100 stimuli at a frequency of 100 Hz (Figure 7A), or theta burst stimulation (TBS), where an episode is subdivided into several bursts of stimuli at 100 Hz, with an inter-burst interval of 200 ms (Figure 7B) (Grover, Kim, Cooke, & Holmes, 2009). The synchrony of these firing patterns mirror those observed in the hippocampus following learning and memory formation. If the electrical stimuli are below the threshold for induction, LTP is not triggered. However, if the threshold for LTP induction is met, it activates the two main molecular mechanisms that underlie hippocampal LTP: the NMDAR and AMPAR.

Figure 7. Potentiation requires delivery of tetanic electrical stimuli. A. High frequency stimulation (HFS) consists of episodes of tetanic electrical stimuli, with each episode comprised of 50-100 stimuli at a frequency of 100 Hz. B. Theta burst stimulation (TBS) consists of episodes of tetanic electrical stimuli that are subdivided into several bursts of stimuli at 100 Hz with an inter-burst interval of 200 ms.

The mechanism underlying LTP can be described by three fundamental properties: cooperativity, associativity, and input specificity (T. V Bliss & Collingridge, 1993). At resting membrane potential, the presynaptic released glutamate only activates AMPARs at the postsynaptic membrane, resulting in Na+ influx and postsynaptic depolarization as NMDAR pores are blocked by Mg2+ (Figure 8A). A strong stimulus, such as TBS, is required to activate a population of neurons in synchrony to reach the cooperativity threshold. This causes greater depolarization sufficient to remove the Mg2+ blockade and activate the NMDAR (Figure 8B). The activation of the NMDAR and subsequent Ca influx is the key trigger to induce LTP. If the stimulus is insufficient, strong stimulation of associated neighbouring fibres can promote depolarization of a neuron of interest. LTP is input specific because the terminal of the presynaptic neuron must release a sufficient concentration of glutamate to activate an adequate number of postsynaptic NMDARs (Figure 8C). Altogether, the activation of NMDAR and subsequent insertion of AMPARs results in the persistent potentiation of the postsynaptic neuron (Figure 8D)

(T. V. P. Bliss, Collingridge, & Morris, 2014). It is important to note that the pattern of HFS or

TBS can affect the properties of LTP. If the pattern of HFS or TBS is separated in the order of seconds, one can elicit a form of LTP known as early-phase LTP. Conversely, if the pattern of separation is in the order of minutes, one can elicit late-phase LTP (E. P. Huang, 1998).

Figure 8. LTP induction. A. At resting membrane potential, Na+ only flows through AMPAR as NMDAR pores are blocked by Mg2+. B. Sufficient depolarization results in removal of the Mg2+ blockade and release of glutamate from the presynaptic to postsynaptic neuron. C. Removal of the Mg2+ blockade results in influx of Na2+ and Ca2+ via NMDARs. D. The resultant Ca2+ entry activates signaling cascades that can promote the insertion of additional AMPAR to the postsynaptic membrane and increase existing AMPAR permeability to Na+.

1.5.3 Early-Phase LTP Early-phase LTP (E-LTP) is characterized by the persistent activation of AMPARs via single or trains of HFS or TBS compressed in the order of seconds. As previously described, the voltage-dependent removal of the Mg2+ blockade and subsequent entry of Ca2+ via NMDAR can activate various protein kinases. One of which is calmodulin (CaM), which subsequently phosphorylates and activates CaMKII, which in itself can induce numerous changes that induce and maintain LTP (Lisman et al., 2014). Phosphorylation of AMPAR residues, namely serine 831

(S831) and 845 (S845) of the CTD of the GluA1 subunit, has been an important mediator of LTP.

These postsynaptic modifications to AMPARs can change its properties, such as single-channel conductance (γ), open probability (Po), and/or number of active channels (N) (Gouaux, 2003).

One of the changes often associated with LTP at CA1 synapses is the changes in γ of

AMPARs. Previous studies have suggested that this molecular process is mediated by CaMKII- dependent phosphorylation of S831. This was demonstrated by single-channel recordings in cultured cells, where infusion of CaMKII or mutation of S831 to Asp increased γ of existing

AMPARs or recruiting AMPARs with higher γ (Derkach, Barria, & Soderling, 1999). However, it is important to note that these posttranslational modifications were only found on CP-AMPARs containing GluA1 homomers, rather than the more ubiquitous GluA1/GluA2 heteromers (Oh &

Derkach, 2005). In a separate study, protein kinase A (PKA) was associated with phosphorylation of GluA1-S845 and subsequent increases in Po (Banke et al., 2000). While these studies provide evidence that suggests that modulation of individual AMPARs via residue phosphorylation can mediate LTP, it still remains unclear as to the extent of contribution of γ and Po with respect to synaptic potentiation (Andrasfalvy & Magee, 2004).

One property that is of particular interest is the changes in N, specifically GluA1-S845- mediated AMPAR trafficking. Due to the dynamic nature of AMPARs, they are constantly inserted and removed from the synapse at basal conditions. The key difference underlying the insertion of

AMPARs in LTP versus basal conditions is the activity-dependent nature of the former process.

Surface expression of AMPARs is mediated by phosphorylation of GluA1-S845. This promotes surface expression of AMPARs in regions outside the PSD, known as extrasynaptic sites (Esteban et al., 2003). Substantial evidence supports the importance of the phosphorylation of GluA1-S845, with mutations replacing the serine with alanine preventing PKA-dependent trafficking of

AMPARs (H. K. Lee et al., 2003). This is consistent with previous findings highlighting the importance of PKA in facilitating LTP by increasing the extrasynaptic pool of AMPARs via phosphorylation of GluA1-S845 (Oh, Derkach, Guire, & Soderling, 2006). Although PKA can insert AMPARs to extrasynaptic sites, PKA alone cannot traffic AMPARs to the synapse, rather, this process is mediated by CaMKII (Gao, Sun, & Wolf, 2006). Interestingly, these CaMKII and

PKA-dependent mechanisms are also required for late-phase LTP.

1.5.4 Late-Phase LTP Initial tetanic stimuli result in activation of mechanisms associated with E-LTP, such as increases in N, however subsequent delivery of episodes spaced, in the order of minutes, can activate mechanisms that underlie late-phase LTP (L-LTP). Thus, L-LTP has been widely regarded as an extension of E-LTP (E. P. Huang, 1998). The mechanism underlying the expression and maintenance of L-LTP is a subject of much debate. While there is evidence that suggests L-LTP requires gene transcription and protein synthesis in conjunction with posttranslational modifications to AMPARs seen in E-LTP, there is also evidence that supports the contrary

(Deadwyler, Dunwiddie, & Lynch, 1987; Villers, Godaux, & Ris, 2012). In the context of protein

synthesis-dependent L-LTP, one molecule of particular interest for the activation of L-LTP is cyclic adenosine monophosphate (cAMP). The increase in concentration of postsynaptic cAMP is mediated by CaM-dependent activation of adenylyl cyclase. The rise in cAMP can then bind to and activate PKA. In addition to PKA, increases in postsynaptic concentration of cAMP can also activate extracellular response kinase/mitogen activated protein kinase (ERK/MAPK) (Impey et al., 1998). The PKA and ERK/MAPK molecular pathways have implications in the activation of transcription factors associated with new gene expression, and thus, both have been associated with the induction and maintenance of L-LTP (Figure 9).

Figure 9. PKA-mediated changes in late-phase LTP. Spacing of tetanic stimuli can induce late- phase LTP, which requires upregulation of cAMP and activation of PKA. PKA can then subsequently activate CREB directly or through MAPK. This signaling cascade can activate gene transcription and protein synthesis mechanisms.

A common target for both PKA and ERK/MAPK and their signaling cascades is the cAMP response element binding protein (CREB); a cascade that has been well preserved throughout evolution, with many observations being made in invertebrates and vertebrates alike (Silva, Kogan,

& Frankland, 1998) One of the best characterized transcription factors involved in synaptic plasticity, CREB has been shown to play a role in integrating extracellular stimuli by activating transcription of genes critical for growing new synaptic connections or inducing structural changes in existing synapses (Benito & Barco, 2010). Activation of PKA and/or ERK/MAPK results in their translocation to the nucleus, which results in the subsequent phosphorylation and activation of CREB. Following this phosphorylation event, CREB together with CREB family of transcription factors will bind to specific regions on the DNA containing the CRE sequence. These target genes are heterogeneous and include a variety of genes with different functions, ranging from transcription, metabolism regulation, cell structure and/or signaling (Benito & Barco, 2010).

The importance of CREB is further highlighted in several studies examining the effects of inactivating CREB. For example, various studies using mice with inactive CREB isoforms revealed deficits in L-LTP and LTM behavioural paradigms (Bourtchuladze et al., 1994; Hummler et al., 1994). Furthermore, electrophysiological studies investigating the inhibition of processes downstream of CREB, such as protein synthesis, found selective deficits in L-LTP, suggesting that protein synthesis is a critical component for L-LTP. Despite the many advancements made within the field of synaptic plasticity, the complete set of CREB-activated genes have not been exhausted

(Kida, 2012).

1.5.5 CP-AMPARs in LTP While differences in the stimulation protocol can affect the properties of LTP, namely E-

LTP and L-LTP, the type of AMPARs that are recruited to the synapse can also vary depending on the stimulation protocols. Indeed, NMDARs are necessary for most forms of synaptic plasticity in the CA1 region of the hippocampus, however, it is important to note that CP-AMPARs can mediate NMDAR-independent LTP (Jia et al., 1996; Wiltgen et al., 2010). Interestingly, this form of LTP differs from NMDAR-dependent LTP because CaMKII, the protein kinase that is indispensable for NMDAR-dependent LTP, is not required for CP-AMPAR-dependent LTP

(Asrar, Zhou, Ren, & Jia, 2009). Rather, the underlying mechanism is phosphoinositide 3-kinase

(PI3K) and ERK/MAPK-dependent, with the latter being involved in NMDAR-dependent LTP as previously described (Asrar, Zhou, et al., 2009). However, because of the low distribution and expression of CP-AMPARs across the brain, whether and how much these CP-AMPAR-dependent mechanisms contribute to E-LTP or L-LTP is unknown.

Unlike NMDARs, which are present throughout development, the expression of CP-

AMPARs during LTP appears to be age-dependent, transient and present only in certain scenarios

(Man, 2011). The requirement of CP-AMPARs is apparent in young (within 2 weeks old) and mature (more than 8-week-old) mice, but not in ages in between, following an E-LTP stimulation protocol (Figure 10A) (Y. Lu et al., 2007; Shepherd, 2012). Interestingly, these changes in CP-

AMPAR dependence parallel the dependence of PKA following LTP induction, suggesting that

PKA activity is coupled with CP-AMPAR expression (Y. Lu et al., 2007). In neuronal synapses,

CP-AMPARs seem to only be required for LTP induction rather than maintenance and seem to require initial NMDAR activity under control conditions (Figure 10B) (Plant et al., 2006).

Following the initial activation of NMDARs, CP-AMPARs are quickly recruited to the synapse

and participate in LTP. However, their expression is transient as they are eventually replaced with

GluA2-containing CI-AMPAR via a process that remains relatively unclear (Figure 10C) (Liu &

Cull-candy, 2000). In contrast, CP-AMPARs have been shown to mediate neuron-glia transmission. Unlike the synapses within hippocampal synapses, glial cells do not express

NMDARs, thus glial-mediated LTP is CP-AMPAR-dependent (Ge et al., 2006). Interestingly, activation of CP-AMPAR-dependent LTP in glia results in subsequent insertion of additional CP-

AMPARs (Ge et al., 2006; Man, 2011). Similarly, the same phenomenon was observed in the basolateral amygdala, where potentiation was NMDAR-independent and largely mediated by CP-

AMPARs (Mahanty & Sah, 1998). Despite the evidence that support the involvement of CP-

AMPARs in LTP, there are studies that suggest the contrary. However, this disparity may be due to subtle differences in electrophysiology recording conditions and more importantly, age of the animals (Adesnik & Nicoll, 2007).

It is apparent in animals with sole expression of GluA2-lacking CP-AMPARs that the levels of LTP are enhanced compared to wild-type littermates (Gerlai, Henderson, Roder, & Jia,

1998; Jia et al., 1996). Interestingly, while gross neuroanatomy is similar across the animals, several behavioural abnormalities exist. Namely, in GluA2-lacking mice, behavioural deficits include increased passivity, impaired motor performance, and impaired spatial and non-spatial behavioural paradigms. These findings strongly suggest that brain function, primarily mechanisms pertaining to learning and memory, is not necessarily correlated with the magnitude of LTP.

Rather, these observations outline the necessity for LTP to be a tightly regulated process for normal brain, behavioural, and motor function (Gerlai et al., 1998).

Figure 10. CP-AMPARs in LTP. A. Initial depolarization results in activation of NMDARs and upregulation of CI-AMPARs. B. After specific patterns of activity, CI-AMPARs are replaced with CP-AMPARs. C Insertion and expression of CP-AMPARs are transient, as they are replaced overtime with CI-AMPARs.

Asides from age, it appears that the timing between the delivery of TBS can affect the type of AMPARs that are recruited to the synapse. As previously described, if tetanic stimuli are delivered in quick succession (hereinafter known as compressed), with an inter-burst interval in the order of seconds, E-LTP is induced with recruitment of GluA2-containing AMPARs. Whereas if tetanic stimuli are spaced, with an inter-burst interval in the order of minutes, L-LTP is induced, which is sensitive to both inhibitors of PKA and de novo protein synthesis (Abel et al., 1997; Frey,

Krug, Reymann, & Matthies, 1988). Intriguingly, L-LTP is also sensitive to inhibitors of CP-

AMPARs, which suggest that membrane insertion of CP-AMPARs require PKA (Park et al.,

2016). This comes as no surprise as translocation of GluA1 homomers and GluA1/GluA3 heteromers require PKA-dependent phosphorylation of S845 (Banke et al., 2000). While the evidence provides insight as to the purpose of compressed and spaced tetanic stimulation, what is

then the purpose of each episode of tetanic stimulation? It has been proposed that the first episode of tetanic stimuli induces a PKA-independent form of LTP, such as CaMKII-mediated changes in

AMPARs. Additionally, the first episode primes the neuron for spacing of stimulation that involves

PKA-dependent membrane insertion of CP-AMPARs. If this were the scenario, then why is it that inhibitors of PKA and CP-AMPARs have no effect on E-LTP? It is proposed that PKA-dependent translocation of CP-AMPARs is directed towards extrasynaptic sites, with subsequent stimulation, spaced in the order of minutes, results in trafficking of CP-AMPARs to the synapse (Park et al.,

2016). This model suggests that there is a greater role for PKA and CP-AMPARs in LTP.

1.6 PKA

1.6.1 General Overview Protein phosphorylation is mediated by protein kinases and thus, can regulate many key regulatory processes in neurons, ranging from neuronal development, growth, and plasticity. A substantial amount of these intracellular signal transduction cascades use cAMP as a secondary messenger to target PKA (Walaas & Greengard, 1991). Due to the ubiquity of PKA, it is involved in various cellular processes. While it is apparent that PKA can modulate chemical synaptic transmission by modifying properties of AMPARs and by altering gene transcription, it should be known that PKA can regulate long-lasting morphological alterations in an activity-dependent manner (Nguyen & Woo, 2003).

1.6.2 Structure and Activation The mammalian PKA family consists of four regulatory (R) subunits: RIα, RIβ, RIIα, RIIβ; and three catalytic (C) subunits: Cα, Cβ, Cγ, where each is encoded by a unique gene and all are expressed within the mammalian brain. Two predominant forms of PKA have been characterized, type I (RIα and RIβ-containing dimers) and type II (RIIα and RIIβ-containing dimers). PKA is a tetrameric enzyme composed of two R subunits bound to two C subunits and is inactive in the absence of cAMP. Each R subunit contains binding sites for cAMP and both must be bound to dissociate the C subunits (Nguyen & Woo, 2003). Dissociation of C subunits can result in the subsequent phosphorylation of serine and threonine residues on various proteins (Figure 11). There are two principal mechanisms for activating PKA, both of which require Ca2+ influx and stimulation of adenylyl cyclase. Activation of adenylyl cyclase and subsequent increase in cAMP can be achieved via two ways: one of which, as previously described, is mediated by CaM, whereas another route is through coupling of adenylyl cyclase to guanine nucleotide-binding regulator proteins (G-proteins) (Impey et al., 1998; Nguyen & Woo, 2003).

Figure 11. Activation of PKA. PKA is a tetrameric enzyme that is composed of two regulatory and two catalytic subunits. PKA is inactive in the absence of cAMP. However, upon binding of cAMP, the catalytic subunits will dissociate and subsequently phosphorylate serine or threonine residues present on a target substrate.

An area of emerging interest is regulating the PKA-dependence of LTP through different stimulation parameters. One key characteristic of stimulation protocols is the total number of electrical impulses, where single episodes of HFS are insensitive to genetic or pharmacological inhibition of PKA whereas multiple episodes show the contrary (Abel et al., 1997; Y. Huang &

Kandel, 1994). It is interesting to note that intermediate forms of PKA-dependent LTP exist. This is evident from studies using hippocampal slices from mice expressing a mutant form of PKA that exhibits 50% of normal activity. The authors reported that LTP elicited by two episodes require less PKA than LTP induced by four episodes, with much of the PKA activity being dependent on the activity of Ca2+ activated protein phosphatases, such as calcineurin or protein phosphatase-1

(Woo, Abel, & Nguyen, 2002). Needless to say, LTP is dependent on the activity of not one, but several proteins from interdependent signaling cascades. In addition to the total amount of electrical stimulation, the temporal spacing between successive episodes can regulate PKA- dependent LTP. While maintaining the number of episodes and electrical pulses, if the inter-burst interval is spaced, a PKA-dependent form of LTP will be elicited, whereas compressed inter-burst intervals are PKA-independent. Interestingly, the authors reported that both forms of LTP are protein synthesis-dependent (Scharf et al., 2002). Given the aforementioned evidence, it is reasonable to hypothesize that perhaps different isoforms of PKA are recruited in response to different stimulation parameters.

1.6.3 Role of PKA The differences in R subunits result in PKA isoforms with different sensitivities to cAMP.

For example, PKA isoforms containing the RIβ subunit have a higher sensitivity to cAMP (Cadd,

Uhler, & McKnight, 1990). Furthermore, each C subunit can regulate distinct sets of genes, thus

different isoforms of PKA may control certain types of synaptic plasticity (R. C. Morris, Morris,

Zhang, Gellerman, & Beebe, 2002).

As previously mentioned, one of the main targets for PKA is CREB, a nuclear protein that regulates transcription of CRE-containing genes critical for plasticity. Of these genes, the best characterized is brain-derived neurotrophic factor (BDNF) (Tao, Finkbeiner, Arnold, Shaywitz, &

Greenberg, 1998). Similar to LTP, its expression is activity-dependent and studies have suggested it is important for L-LTP within the hippocampus. More specifically, it appears that activity- dependent increase in cAMP upregulates TrkB receptors for BDNF, which contributes to the maintenance of L-LTP. Although majority of the CREB-targeted genes require CRE sequences, not all genes upregulated during LTP have an identified CRE sequence. A family of genes known as immediate early genes (IEGs) respond rapidly to synaptic stimulation. One such example of

IEGs include Arc, an IEG that is input specific and localizes only to stimulated synapses.

Comparable to other signaling cascades, Arc transcription is PKA and MAPK/ERK-dependent

(Waltereit et al., 2001). It is unclear as to whether transcription of certain genes, whether CRE-or

IEG-containing, are mutually exclusive to certain PKA isoforms. Thus, further understanding the role of specific PKA isoforms will be essential to understanding the process underlying activity- dependent plasticity. Many of these protein changes result in changes in synaptic plasticity, many of which reshape the synaptic structure of the dendritic spine.

1.7 The Correlation Between Structural Plasticity and Synaptic Plasticity

1.7.1 Structural Plasticity It is believed that reversible physiological changes in synaptic transmission, most of which consist of transient molecular changes, comprise STM, whereas stable and persistent changes are

LTM. It has been widely accepted that creation of LTM requires gene expression and protein synthesis, which run in parallel with resulting structural changes in synaptic morphology (Goelet,

Castellucci, Schacher, & Kandel, 1986). The first studies showing changes in synaptic architecture, such as alterations in size, shape, or number of synapses, occurred in Aplysia (Bailey & Chen,

1988a, 1988b). Following these studies, dendritic spines of excitatory synapses have been the focus of recent work in the mammalian brain. Experiments using two-photon microscopy has shown that LTP induction in hippocampal slice cultures result in the formation of new spines in a

NMDAR-dependent manner, as inhibition of NMDARs prevented structural changes (Engert &

Bonhoeffer, 1999). Other studies have also reported an increase in spine size as well as shortening and widening of the spine neck following LTP (Figure 12) (Lamprecht & LeDoux, 2004).

Figure 12. Synaptic plasticity is correlated with structural plasticity. Induction of LTP can result in growth of existing dendritic spines or formation of new dendritic spines in an NMDAR- dependent manner.

How do these structural modifications facilitate synaptic potentiation? Many of these alterations modulate synaptic transmission, and to a certain extent, biochemical events, that occur at dendritic spines following LTP. In one such example, widening and shortening of the neck allows for greater and quicker Ca2+ influx into the dendrite (Volfovsky, Parnas, Segal, &

Korkotian, 1999). In addition, enlargement of the spine head leads to a multitude of postsynaptic changes that facilitate potentiation. Using glutamate uncaging, authors reported greater sensitivity of larger spine heads to glutamate (Matsuzaki, Honkura, Ellis-Davies, & Kasal, 2004). Moreover, within the Schaffer collateral synapses, it has been demonstrated that larger spines have a greater

AMPAR to NMDAR ratio, which increases the strength of synaptic transmission through

AMPARs (Takumi, Ramírez-León, Laake, Rinvik, & Ottersen, 1999). Furthermore, larger spines receive input from larger presynaptic terminals, which typically contain more vesicles (Schikorski

& Stevens, 1997). The enlargement of spine heads also allows for preferential incorporation of local protein synthesis machinery, such as polyribosomes (Ostroff, Fiala, Allwardt, Harris, &

Street, 2002). Finally, increases in number of spines could enhance synaptic transmission by increasing connections with presynaptic neurons.

1.7.2 Cytoskeletal Structure Spine structure and morphology is dependent on the underlying cytoskeletal filaments; a complex of interlinking proteins that extend throughout the intracellular space that underlies the dynamic nature of cells. The cytoskeleton is not a single component, rather, it is composed of three filaments that differ in size and composition: microtubules, intermediate filaments, and actin filaments.

Microtubules are the largest class of filament (diameter of 25 nm) and are primarily composed of a protein called tubulin (Blanchoin, Boujemaa-paterski, Sykes, & Plastino, 2014).

Together, tubulin can dimerize and form long strands known as protofilaments, which can then subsequently come together to form a hollow filament that consists of thirteen protofilaments.

Microtubules are dynamic, however the rate of change at each end differs. There is a plus end, a region of rapid growth, and a minus end, a region of slower growth. In cells, the minus end is typically anchored to a region adjacent to the nucleus known as the microtubule organizing centres

(MTOCs). These properties of microtubules provide cellular structure and basic organization of the cytoplasm, such as positioning of organelles (Herrmann, Bär, Kreplak, Strelkov, & Aebi,

2007).

There are several types of intermediate filaments, however in general, they are smaller than microtubules but larger than actin filaments (diameter of 10 nm) and are less dynamic than microtubules. Typically strong and ropelike, intermediate filaments provide structural support to existing fragile tubulin structures seen in microtubules (Herrmann et al., 2007). Intermediate filament protein subunit composition is associated with specific cell types; however, each cell can have multiple types of intermediate filaments. For example, neurons consist of intermediate filaments known as neurofilaments and primarily exist in axonal structures (M. K. Lee & Don,

1996).

Actin filaments are the smallest of the three filaments (diameter of 6 nm) and consist of identical actin proteins arranged in a long spiral chain-like structure. Similar to microtubules, actin filaments have a plus and minus end, with the plus end having greater ATP-powered growth for faster dynamics (Revenu, Athman, Robine, & Louvard, 2004). Polymerization of actin requires

ATP hydrolysis inside the actin monomer, formally known as globular actin (or G-actin), to form dimers, trimers, and longer filaments. Because of the abundant and dynamic nature of actin, it can associate with various membrane-bound proteins, and form specialized structures such as dendritic spines (Blanchoin et al., 2014). For example, actin interacts closely with the PSD, with many studies demonstrating that changes in spine mobility and stability is dependent on actin polymerization (Fischer, Kaech, Knutti, & Matus, 1998; Halpain, Hipolito, & Saffer, 1998). Thus, much of the structural changes seen in spines following LTP induction is mediated by actin reorganization. On one hand, this hypothesis is consistent with findings from pharmacological studies, where inhibition of actin polymerization (F-actin) attenuated LTP in the rodent hippocampus (Krucker, Siggins, & Halpain, 2000). On the other, increased F-actin was observed in dendritic spines following LTP induction in rodent hippocampi (Fukazawa et al., 2003).

1.7.3 Actin-mediated Receptor Trafficking Intracellular trafficking mechanisms, such as vesicle transportation and receptor membrane insertion, is an active process that requires actin dynamics. Because dendritic spines are highly dynamic in nature it is enriched with dynamic F-actin, which serves to regulate AMPAR localization within the synapse (Figure 13) (Porat-Shliom, Milberg, Masedunskas, & Weigert,

2013). While it is generally understood that AMPAR localization is actin-dependent, the exact mechanism by which AMPARs are trafficked to the membrane and how AMPARs interact with actin remains unclear (Hanley, 2014a). Several studies highlight the involvement of actin in

AMPAR trafficking. In cultured hippocampal neurons, disruption of F-actin resulted in reduction in GluA1-containing AMPARs in dendritic spines, whereas stabilization of F-actin blocked

AMPAR internalization. Suggesting that actin polymerization is required for AMPAR insertion and actin depolymerization is required for AMPAR internalization (Allison, Gelfand, Spector, &

Craig, 1998; Zhou, Xiao, & Nicoll, 2001). In addition, it was found that LTP is attenuated following the blockade of actin depolymerization, suggesting that the dynamic nature of actin is required for potentiation and subsequent AMPAR trafficking (Kim & Lisman, 1999).

Figure 13. Spine structure and morphology is dependent on actin cytoskeleton dynamics. A. Globular (G)-actin is abundant and can dimers and trimers in the dynamic state. B. Following LTP induction, G-actin will polymerize and stabilize to form filamentous (F)-actin. Formation of F- actin can increase dendritic spine width and stabilize AMPAR localization at the synapse.

1.7.4 Receptor-mediated Actin Stabilization The changes in actin dynamics, orientation, and assembly kinetics is activity-dependent, thus extracellular stimulation and subsequent LTP induction can contribute to spine formation

(Matus, 2000). Results from in vivo studies indicate that inhibition of NMDARs attenuate F-actin, and further blockade of F-actin prevents the development of L-LTP in adult rodents (Lamprecht

& LeDoux, 2004). Together, these findings suggest that NMDAR-dependent mobilization of F- actin is important for consolidating E-LTP into L-LTP. Although NMDARs are important for initiating changes in spine dynamics, it appears that AMPARs are required for the stabilization of spines to ensure long-lasting synaptic changes. Several studies support this hypothesis: blockade of AMPARs with CNQX (antagonist 6-cyano-7-nitroquinoxaline-2,3-dione) resulted in attenuation of F-actin (Lamprecht & LeDoux, 2004). In a separate study, application of small amounts of AMPA prevented decreases in spine density in CA1 pyramidal cells in hippocampal slice cultures. Furthermore, blockade of vesicular glutamate release via botulinum toxin A or C resulted in lower spine density, suggesting that spontaneous glutamate release is sufficient for maintaining dendritic spine heads (Mckinney, Capogna, Dürr, Gähwiler, & Thompson, 1999).

Together, these results support the hypothesis that AMPAR insertion to the PSD following LTP induction is critical for LTP maintenance and for LTM formation.

1.7.5 Rho-GTPases While the previous studies highlight the importance of extracellular stimulation and the requirement for glutamate receptors to trigger and stabilize actin dynamics, how then, do these changes result in rearrangements in the cytoskeletal structure and subsequent spine morphogenesis? A common target for cytoskeletal rearrangement following LTP induction is the activation of Rho-family of guanosine triphosphate hydrolases (Rho GTPases). Together, Rho

GTPases and their downstream effectors have been implicated in several studies investigating their role in synaptic plasticity (Luo, 2002).

Figure 14. Rho GTPase activation and inactivation. Activation of Rho GTPases is mediated by the exchange of GDP to GTP via guanine nucleotide exchange factors (GEFs). Inactivation of Rho GTPases is mediated by the hydrolysis of GTP to GDP via GTPase activating proteins (GAPs).

Rho GTPases are a family of proteins that serve as an intracellular signaling switch and can exist in two states: a GTP-bound active state or a guanosine diphosphate (GDP)-bound inactive state. This state is mediated by guanine nucleotide exchange factors (GEFs) by facilitating the exchange of GDP to GTP. In contrast, GTPase activating proteins (GAPs) facilitating the hydrolysis of GTP to GDP (Figure 14) (Luo, 2000). In the active form, Rho GTPases bind to and activate downstream effectors that regulate actin cytoskeleton dynamics (Lamprecht & LeDoux,

2004). The best characterized members of the Rho GTPase family includes: Ras homologous member A (RhoA), Ras-related C3 botulinum toxin substrate (Rac), and cell division cycle 42

(Cdc42), all of which are essential for regulating dendritic spine formation during development

(Figure 15). Interestingly, there is significant crosstalk between the Rho GTPases; Ras and Cdc42 can activate Rac, whereas Rac can activate RhoA (Hall, 1998). It appears that dominant-negative

Rac reduces the number of dendritic spines, whereas hyperactivation of RhoA results in simplification of dendritic arborization in hippocampal slices from young rats (Nakayama, Harms,

& Luo, 2000). Furthermore, in a separate study, overexpression of RhoA in cortical neurons resulted in reduction of number and length of dendritic spines (Tashiro, Minden, & Yuste, 2000).

Figure 15. LIMK1 is the common downstream target for RhoA, Rac, and Cdc42. Rho GTPases serve as intracellular signaling switches, however they do not directly interact LIMK1, a key regulator for actin cytoskeleton dynamics. RhoA primarily regulates LIMK1 activity via ROCK, whereas Rac and Cdc42 regulate LIMK1 via PAK.

1.7.6 Downstream Effectors: PAK, ROCK Several studies have implemented Rho GTPases in synaptic plasticity and memory formation. While Rho GTPases serve as intracellular signaling switches, they do not actually directly interact with the actin cytoskeleton. Rather, activation of Rho GTPases results in

subsequent phosphorylation and activation of numerous downstream effectors that regulate actin dynamics (Vega & Ridley, 2008). One important family of effectors downstream of Rac and

Cdc42 GTPase is p21-activated kinase (PAK) (Figure 15). Part of the serine/threonine kinase family, PAKs have been shown to regulate many aspects of cellular function, such as cell motility, gene expression, and cytoskeletal dynamics. The PAKs can be subdivided into groups, with group

I being comprised of PAK1-3, and group II being comprised of PAK4-7 (Zhao & Manser, 2012).

Of these PAK isoforms, only PAK1 and PAK3 have been implemented in synaptic plasticity

(Asrar, Meng, et al., 2009; J. Meng, Meng, Hanna, Janus, & Jia, 2005). Previously, our lab has shown that genetic knockout (KO) of PAK3 resulted in deficits in hippocampal L-LTP without affecting basal neuronal activity. These deficits stemmed from dramatic reduction in active CREB, which is consistent with previous studies investigating the signaling processes underlying L-LTP

(J. Meng et al., 2005). In a follow up study, our lab has shown that genetic KO of PAK1 resulted in impaired hippocampal LTP, while preserving basal synaptic transmission and presynaptic function. Although neuronal ultrastructure, such as spines and synapses, appeared normal, cultured hippocampal neurons showed decreased F-actin compared to wild-type neurons. These deficits were tied with the activity-dependent regulation of cofilin, a protein that binds to and regulates the polymerization and depolymerization of actin filaments. Despite this deficit, the gross anatomy of the brain of PAK1 KO mice remained normal. This inconsistency could be the result of compensation from other members of the PAK family, namely PAK3 due to structural similarity

(Asrar, Meng, et al., 2009). This is supported by the fact that mice lacking both PAK1 and PAK3 have reduced brain size and simplified dendritic arbors (W. Huang et al., 2011).

Although actin dynamics can be regulated by PAK1, changes to the actin cytoskeleton is also in part mediated by Rho-associated kinases (ROCKs) (Figure 15). Also a serine/threonine

kinase, ROCK is known to regulate cell contractility, cell adhesion, and actin dynamics

(Schmandke, Schmandke, & Strittmatter, 2007). Two isoforms of ROCK exist in rodents: ROCK1 is mainly expressed in lung, liver, and spleen, whereas ROCK2 is mainly expressed in the brain and the heart (hereinafter, ROCK will refer to ROCK2) (Nakagawa, Fujisawa, Ishizaki, Saito, &

Nakao, 1996). Recent studies have linked ROCK to synaptic plasticity and memory formation.

Pharmacological inhibition of ROCK prior to fear conditioning impaired LTM but not STM.

Additionally, inhibition of ROCK approximately 22 hours following fear conditioning had no effect on memory retrieval. Furthermore, a separate study showed that inhibition of ROCK effectively blocked spine enlargement. Together, these findings suggest a role for ROCK in consolidating STM into LTM and provides a link between structural morphology and LTM formation. Similar to PAK, it appears that ROCK regulates spine morphology via activity- dependent regulation of cofilin and subsequent polymerization or depolymerization of actin filaments (Bosch & Hayashi, 2012).

1.8 A Common Effector: LIMK

1.8.1 General Overview Much of the cofilin-dependent actin reorganization is not mediated by PAK and ROCK directly. Rather, these kinases activate another important effector known as the LIM-domain- containing protein kinase (LIMK) (Figure 15) (Lamprecht & LeDoux, 2004). An important effector of the Rho GTPase pathways, LIMKs are expressed ubiquitously and are primarily involved in regulating cofilin-dependent reorganization of the actin cytoskeleton (Cuberos et al.,

2015). However, more recent studies have suggested a role for LIMKs in CREB function

(Todorovski et al., 2015; E. J. Yang, Yoon, Min, & Chung, 2004). Together, the functional role of

LIMKs underlie many neuronal functions such as: neuronal development, cell migration, and synaptic plasticity (Y. Meng et al., 2002, 2004).

1.8.2 Structure and Activation Two isoforms of LIMKs exist in mammals, with LIMK1 being most abundant in neuronal tissue whereas LIMK2 expression is ubiquitous. The structure of the two LIMK genes is identical, which includes: C-terminal serine/threonine kinase domain and three protein-protein interaction domains comprised of one central PDZ domain and two N-terminal LIM domains (Bernard, 2007).

The LIM acronym is derived from the three genes that it is a product of: lin-11, isl-1, and mec-3; all of which contribute to a double zinc finger motif that is enriched with cysteine. The structure of both LIMK isoforms are identical and share approximately 50% of the amino acid sequences

(Bernard, 2007). The PDZ domain facilitates protein-protein interactions and often regulates protein localization, with LIMK1 being preferentially located in the cytoplasm. Despite this preferential localization, there is evidence that suggests that LIMK1 can shuttle between the nucleus and the cytoplasm (N. Yang, Higuchi, & Mizuno, 1998; N. Yang & Mizuno, 1999).

The level of activity of LIMKs are regulated by a variety of proteins as well as microRNAs.

LIMK activity requires phosphorylation, however, studies have suggested that the efficacy of

LIMK catalytic activity is differential depending on the phosphorylation site of the multiple serine or threonine residues (Figure 16) (Nadella et al., 2009). LIMK1 activity is primarily dependent on

PAK or ROCK-mediated phosphorylation of threonine 508 (T508) located in the kinase domain, with studies showing that a LIMK1 T508V mutation prevented PAK1-mediated activation

(Edwards, Sanders, Bokoch, & Gill, 1999). Likewise, a LIMK1 T508A mutation prevented

ROCK-mediated phosphorylation (Ohashi et al., 2000).

Figure 16. LIMK1 phosphorylation sites. LIMK1 has multiple phosphorylation sites: Ser323, which is regulated by MapkAP2 and pKA; Thr 508 is regulated by ROCK and PAK; Thr 596 is regulated by PKA. While LIMK1 activity only requires one site to be phosphorylated, studies have suggested that the efficacy and stability of LIMK1 catalytic activity is dependent on the number of phosphorylated sites.

Other phosphorylation sites are present in the kinase domain, such as S323 and S596, and there is growing evidence suggesting that other signaling pathways are capable of phosphorylating and activating LIMK1 (Nadella et al., 2009). Studies suggest that activation of the ERK pathway can mediate phosphorylation of S323 in LIMK1 via MAPK activated protein kinase II

(MAPKAPK2) (Figure 16) (Hogg, Müller, & Corrêa, 2016; Muto et al., 2012). Recent work has suggested that mice lacking MAPKAPK2 have less F-actin and greater concentration of G-actin, suggesting that phosphorylation of S323 in conjunction with T508 are important mediators of structural plasticity (Hogg et al., 2016). Several studies have also suggested a role for PKA in cytoskeletal reorganization (Howe, 2004). More interestingly, recent work has suggested that PKA can directly activate LIMK1 (Nadella et al., 2009). In addition to S323, it appears that PKA can

also directly phosphorylate S596, with S323A and S596A mutations showing substantial reduction in LIMK1 activity. Interestingly, it appears that T508 phosphorylation is not necessary as previously described, with T508V mutations not affecting the PKA-LIMK signaling pathway

(Nadella et al., 2009). This then raises the question as to why do multiple phosphorylation sites exist on LIMK1? Studies have suggested that phosphorylation of LIMK1 can result in dimerization and subsequent transphosphorylation. Phosphorylation of additional sites promotes LIMK1 stability by promoting the binding of heat shock protein 90 (HSP90), which increases LIMK1 half- life from 5 hours to 20 hours (Dong, Ji, Cai, & Chen, 2012; Li et al., 2006).

The absence of LIMK1 in mice manifests in enhanced locomotive activity and contextual fear responses. This is in contrast to the observed impaired spatial memory as indicated by Morris

Water Maze tests. Interestingly, these behavioural abnormalities, namely visuospatial deficits and hyperactivity, are hallmark features of the cognitive profile of patients with William Syndrome, a neurodevelopmental disorder caused by hemizygous deletion of 28 genes, one of which being

LIMK1 (Y. Meng et al., 2002; Todorovski et al., 2015).

1.8.3 Downstream Effectors One of the primary downstream effectors of LIMKs is through phosphorylation and inactivation of cofilin, a protein that mediates polymerization/depolymerization of F-actin, as previously mentioned. In the dephosphorylated state, cofilin binds to the side of F-actin to induce a conformational change that favors actin depolymerization (Figure 17) (Pavlov, Muhlrad, Cooper,

Wear, & Reisler, 2007). However, once phosphorylated, cofilin becomes inactive and subsequently reduces actin turnover to promote F-actin assembly (Ohashi, 2015). Thus, the activity of LIMK1 and the subsequent inactivation of cofilin has a major role in both synaptic and structural

plasticity. Previous studies from our lab using LIMK1 KO mice have demonstrated abnormal spine morphology and abnormal clustering of cofilin. Results from biochemical analysis suggests that there was reduced phosphorylation of cofilin, but not the total protein level, in LIMK1 KO mice

(Y. Meng et al., 2002). Furthermore, in a separate study, it appears that LIMK1 KO mice have impaired L-LTP (Todorovski et al., 2015). Together, these results suggest that these changes to cofilin activity and subsequent structural morphology may account for the observed deficits in L-

LTP. Interestingly, despite the widespread expression of LIMK2, genetic KO of LIMK2 resulted in minimal abnormalities in the CNS (Y. Meng et al., 2004).

Figure 17. Cofilin activity is regulated by LIMK1. In the unphosphorylated state, cofilin is active and promotes actin destabilization and turnover from F-actin to G-actin. However, when phosphorylated by LIMK1, cofilin becomes inactive which results in F-actin assembly.

In addition to cofilin, there is evidence to support that LIMK1 can directly phosphorylate and activate CREB. More specifically, treatment of cells with growth factors stimulated direct binding of LIMK1 with CREB. Furthermore, a S133A mutation in CREB resulted reduced

LIMK1-mediated phosphorylation (E. J. Yang et al., 2004). In conjunction with this data, our lab has also demonstrated the interactivity between LIMK1 and CREB. Although LIMK1 contains domains that promotes cytoplasmic localization, LIMK1 was shown to also colocalize with CREB in the cell bodies. Indeed, brain lysates from LIMK1 KO mice showed reduced phosphorylation of CREB. Additionally, the same study found that impaired L-LTP could be rescued with pharmacological activation of CREB in LIMK1 KO mice, thus suggesting that LIMK1-mediated activation of CREB is an important regulator of L-LTP (Todorovski et al., 2015).

1.9 Clinical Relevance

1.9.1 Synaptic Plasticity in Health and Disease It is believed that aberrant activation of signaling pathways associated with LTP can result in deficits in structural and synaptic plasticity. Thus, it is believed that abnormalities in synaptic plasticity may be the fundamental cause for many brain disorders (T. V. P. Bliss et al., 2014;

Henley & Wilkinson, 2013). In Canada alone, approximately 3.7 million people live with an intellectual disability and 618,000 live with dementia; with approximately 25% and 4% new cases, respectively, each year (Bizier, Fawcett, & Gilbert, 2015; Chambers, Bancej, & Mcdowell, 2016).

Therefore, further understanding of the physiological mechanisms that underlie synaptic deficits may offer insight to improving existing treatment regiments or to develop new novel therapeutic compounds for various neurological and psychiatric conditions.

1.9.2 Williams Syndrome Williams Syndrome (WS) is a multisystem disorder that occurs every 1/20,000 births caused by a 1.5 mega base pair deletion on chromosome 7 (C. A. Morris & Mervis, 2000). Patients affected by WS are characterized by dysmorphic facial features alongside infantile hypercalcemia

and abnormalities in connective tissue. Neurologically, these individuals exhibit a unique cognitive profile; typically displaying learning disabilities with profound deficits in visuospatial construction tasks (C. A. Morris & Mervis, 2000). Behaviourally, WS patients are highly sociable and empathetic, traits that are unlike any other developmental disorder. The neurological phenotypes are attributed to deletion of several neuron-specific genes, one of which is LIMK1. Thus, LIMK1 has been a candidate for the underlying neurobehavioural deficits see in WS (Martens, Wilson, &

Reutens, 2008).

How then, does LIMK1 deletion result in the cognitive and behavioural deficits observed in WS patients? Mechanistically, deletion of LIMK1 impairs cofilin-mediated actin cytoskeleton dynamics and impairs LTM via a CREB-dependent mechanism (Y. Meng et al., 2002; Todorovski et al., 2015). Specifically, our lab has shown that cofilin phosphorylation was reduced in LIMK1

KO mice, which subsequently resulted in abnormalities in dendritic spine morphology; abnormalities that are seen in human patients diagnosed with other developmental disorders, such as Down’s Syndrome and Fragile X Syndrome (Y. Meng et al., 2002; Sorra & Harris, 2000).

Behaviourally, LIMK1 KO mice demonstrated increased locomotive activity and enhanced fear response, but had impaired spatial learning performance. Electrophysiological data from LIMK1

KO mice suggests that E-LTP is enhanced but L-LTP is impaired. Together, the results are consistent with the cognitive profile that is present in WS patients; hyperactivity and impaired

LTM, with selective deficits in visuospatial cognition. These findings highlight the importance of assessing the role of LIMK1 as a molecular mediator that underlies cognitive deficits.

1.9.3 Alzheimer’s Disease Alzheimer’s disease (AD) is the cause of approximately 60-80% of dementia and manifests itself as a progressive loss of mental and behavioral function (Bizier et al., 2015). Eventually, AD results in overall functional decline. Several hypotheses have been put forward to explain the pathophysiology associated with AD, with the most common being the amyloid β (Aβ) hypothesis

(Kumar & Singh, 2015). According to the Aβ hypothesis, amyloid precursor protein (APP), an endogenous neuronal protein, is cleaved by α-secretase in healthy individuals, but aberrantly processed by β- and γ-secretase in patients with AD. This results in an imbalance between nontoxic and toxic forms being Aβ40 and Aβ42, respectively (the toxic peptide will hereinafter be referred to as Aβ). As a consequence, increases in Aβ peptides result in aggregation and formation of soluble oligomers, which in turn form fibrils that comprise insoluble β-sheets. Proliferation of insoluble

β-sheets results in formation of senile plaques which contribute to neuronal death and eventual cognitive decline (Cras et al., 1991; Kumar & Singh, 2015). Indeed, formation of insoluble plaques result in many of the cognitive deficits associated with AD, however, many of these hallmark pathophysiological events appear in the late stages and are typically untreatable (Cras et al., 1991).

Many studies have suggested that the early cognitive impairments seen in AD are correlated with loss of synaptic function (Terry et al., 1991). Therefore, it is worthwhile to investigate mechanisms that prelude plaque formation in hopes to find potential therapeutic strategies.

It has been proposed that many of the early toxic effects begin at the level of the synapse, with studies suggesting that elevation of Aβ peptides impair synaptic function. Shankar and colleagues demonstrated that administration of soluble Aβ dimers inhibit LTP and LTD, and mechanistically, the spine loss associated with LTD was NMDAR-dependent (Shankar et al.,

2008). The effects of Aβ can be attenuated as co-administration of monoclonal antibodies targeting

the N-terminus of Aβ peptides prevented LTP and LTD deficits in rats (Klyubin et al., 2005;

Shankar et al., 2008). These findings suggest that Aβ can disrupt synaptic function and structure through an NMDAR-dependent mechanism. Although studies have highlighted the importance of

NMDARs in pathological states, it is important to note that signaling pathways downstream of

NMDARs, namely the regulation and trafficking of AMPARs, can have profound effects in AD as well.

Of the AMPARs subtypes, CP-AMPARs have been of particular interest as they have established roles in epilepsy, traumatic brain injury, and in substance addiction and withdrawal

(Arundine & Tymianski, 2003; Grooms, Opitz, Bennett, & Zukin, 2000; Volkow & Morales,

2015). Because the role of CP-AMPARs in these disease etiologies have been thoroughly investigated, there has been postulation that CP-AMPARs are involved in neurodegeneration.

Indeed, there is growing evidence that CP-AMPARs are enriched in patients with AD, with exposure to Aβ resulting in rapid insertion of CP-AMPARs in the hippocampus (Whitcomb et al.,

2015). Furthermore, increased PKA-mediated expression of CP-AMPARs resulted in synaptic deficits in AD transgenic mouse models prior to apparent development of neuropathological symptoms (Megill et al., 2015). Together, these findings suggest that aberrant CP-AMPAR insertion may be involved in the early stages of AD pathogenesis. While the role of CP-AMPARs is apparent in cognitive deficits in early stage AD, how the expression of CP-AMPARs results in neurodegeneration is relatively unclear. At the fundamental level, it is hypothesized that CP-

AMPARs facilitate Ca2+-mediated synaptic degeneration resulting in synaptic weakening and elimination of synaptic connections (Laferla, 2002). With these processes being a hallmark feature of several neuropathological conditions, signaling pathways surrounding cytoskeletal

rearrangement have become an area of increasing interest for further study (Collingridge, Peineau,

Howland, & Wang, 2010).

Chapter 2 Rationale, Hypothesis, and Objectives 2.1 Rationale Postnatal glutamatergic principal neurons are mainly comprised of GluA2-containing CI-

AMPARs. However, in certain forms of synaptic plasticity events and neurodegenerative and developmental disease states, there is upregulation of GluA2-lacking CP-AMPARs (T. V. P. Bliss et al., 2014). CP-AMPARs have an important role in synaptic plasticity as they are capable of eliciting LTP, a process previously thought to be exclusive to NMDARs. When CP-AMPARs are engaged, it has been found that their activation alone is sufficient to induce NMDAR-independent

LTP in animals lacking the GluA2 subunit (Jia et al., 1996). In a separate study, it was found that

LTP results in rapid insertion and incorporation of CP-AMPARs to the synapse under normal physiological conditions. However, the presence of these CP-AMPARs are transient, as they are replaced by CI-AMPARs ~25 min after LTP induction (Plant et al., 2006). Although CP-AMPARs are capable of inducing LTP, it is interesting to note that the canonical CaMKII pathway in

NMDAR-dependent LTP is not required for CP-AMPAR-dependent LTP. Rather, there are distinct signaling pathways that are exclusive to CP-AMPARs (Asrar, Zhou, et al., 2009)

The elusive and controversial CP-AMPARs have been implicated in some (Jia et al., 1996;

Plant et al., 2006), but not all LTP phenomena (Adesnik & Nicoll, 2007), and this discrepancy is thought to be due to differences in experimental conditions. A recent study has suggested that the involvement of CP-AMPARs in LTP is determined by the timing of stimuli, with sTBS eliciting a

PKA-dependent form of LTP that recruits CP-AMPARs (Park et al., 2016). The differential recruitment of CP-AMPARs in cTBS and sTBS provides evidence that different stimulation

parameters can elicit different molecular mechanisms. If so, what are the molecular mechanisms underlying each form of LTP elicited from cTBS and sTBS?

The central underlying difference between cTBS and sTBS is that the latter involves the activation of PKA and subsequent recruitment and insertion of CP-AMPARs. However, this process is dependent on the initial activation of NMDARs. The first question is then, if CP-

AMPARs can elicit LTP in an NMDAR-independent manner, are CP-AMPARs alone capable of activating PKA-dependent mechanisms? To address this question, I will perform extracellular field recordings using stimulation parameters such as cTBS and sTBS to activate PKA-independent and

–dependent LTP, respectively, in GluA2 KO mice, a murine model that exclusively expresses CP-

AMPARs.

While PKA-dependent LTP has been demonstrated extensively in previous literature, the ubiquity of PKA and its involvement in various molecular pathways has made the exact mechanisms surrounding sTBS-induced LTP unclear (Abel et al., 1997; Bolshakov, Golan,

Kandel, & Siegelbaum, 1997; Esteban et al., 2003; Park et al., 2016). A recent study has suggested that PKA directly interacts with LIMK1, a potent regulator of actin dynamics, however whether this interaction occurs during synaptic plasticity is unknown (Nadella et al., 2009). The second question is then, does PKA require LIMK1 during synaptic plasticity? To address this question, I will perform extracellular field recordings using cTBS and sTBS stimulation parameters on

LIMK1 KO mice.

Because synaptic recruitment of AMPARs are linked with structural changes, the final question is then, is structural change required for PKA-dependent LTP? To address this question,

inhibitors targeting pathways related to structural changes, such as protein synthesis, will be used following sTBS-induced PKA-dependent LTP in GluA2 and LIMK1 KO mice.

2.2 Hypothesis My hypothesis is that CP-AMPARs can trigger PKA-dependent LTP independent of

NMDARs and that PKA-dependent de novo protein synthesis requires LIMK1.

2.3 Objectives To address this hypothesis, my objectives are as follows:

1. To establish cTBS and sTBS protocols in GluA2 and LIMK1KO mice

Our lab has predominantly used HFS in previous studies, however recent studies have shown

that molecular mechanisms can differ between HFS and TBS (Hernandez, Navarro, Rodriguez,

Martinez, & Lebaron, 2005; Zhu, Liu, Wang, Bi, & Baudry, 2015). Thus, I aim to determine

whether previous findings surrounding GluA2 and LIMK1 KO mice from our lab can be

replicated using cTBS and sTBS protocols

2. To determine if CP-AMPAR-dependent LTP require PKA

NMDAR activation is believed to be the primary trigger for PKA-dependent LTP (Park et al.,

2016). Our lab has previously shown that CP-AMPARs are capable of inducing LTP

independent of NMDARs (Asrar, Zhou, et al., 2009; Jia et al., 1996). Because the molecular

mechanisms downstream of CP-AMPARs remain unknown, I aim to determine if CP-

AMPAR-dependent LTP require PKA.

3. To determine if PKA-mediated activity dependent de novo protein synthesis require

LIMK1

Studies have shown that PKA can mediate structural changes following LTP induction by activating protein synthesis pathways via CREB. Recently, a study has shown that PKA can activate LIMK1 and furthermore, a study from our lab has shown that LIMK1 localizes with

CREB (Abel et al., 1997; Todorovski et al., 2015). Therefore, I aim to determine if PKA- mediated activity dependent protein synthesis require LIMK1.

Chapter 3 Materials and Methods 3.1 Animal Colony

3.1.1 GluA2 and LIMK1 Mutant Mice The initial generation and characterization of GluA2 and LIMK1 mutant mice were as described previously (Jia et al., 1996; Y. Meng et al., 2002). The mice were housed under a standard 12 h light/12 h dark cycle condition. All studies with mutant LIMK1 animals were performed alongside C57BL/6 animals as controls. Studies with mutant GluA2 animals were performed alongside wild-type littermates as controls. All the procedures used for this study were approved by the Animal Use Committee at the Hospital for Sick Children, Toronto, Ontario,

Canada.

3.2 Genotyping

3.2.1 DNA Extraction DNA was extracted from tail biopsies. The tip of the tail (~0.5 cm) was cut and digested in a 500 µL cocktail containing Proteinase K (BioShop) and lysis buffer solution (20 mg/mL) for a minimum of 6 h at 60°C. Samples were mixed via inversion (5 times) until the solution was homogenous. Afterwards, 600 µL of chloroform (Caledon) and 200 µL of 5 M NaCl (Sigma

Aldrich) was added and mixed via inversion (5 times). Following inversion, samples underwent centrifugation at 10,000 RPM for 10 min allowing for phase separation. The solution separates into three phases: 1. Aqueous phase containing DNA 2. Interphase containing cell debris 3.

Organic phase from top to bottom, respectively. 400 µL of the aqueous phase was transferred to a clean 1.5 mL tube, followed by addition of equal volume 100% ethanol (Commercial Alcohols).

The solutions were mixed via inversion to precipitate the DNA. The samples were then centrifuged at 14,000 RPM for 5 min to pellet the DNA. The supernatant was then discarded and the pellet was washed with 300 µL of 70% ethanol (Commercial Alcohols). Following the wash, the pellet underwent centrifugation at 14,000 RPM for 5 min. Following centrifugation, the 70% ethanol was removed via a p200 pipette and the pellet was left to dry for a minimum of 30 min. The DNA pellet was then dissolved in 120 µL of water.

3.2.2 Polymerase Chain Reaction A standard protocol for Taqman Polymerase Chain Reaction (PCR) was used for determining the genotype of GluA2 KO mice. The following primers (ACGT Corp.) were used:

GluA2 WT Primer 1: 5′-CTCAGAAGTCCAAACCAGGAGTG-3′

Primer 2: 5′-GCAGGCATGGAATGATAGGAAC-3′

GluA2 KO Primer 1: 5′-CTCAGAAGTCCAAACCAGGAGTG-3′

Primer 3: 5′-GGGGGAACTTCCTGACTAGG-3′

LIMK1 WT Primer 4: 5′-CCAGACCGTGGTAACTCCAG-3′

Primer 5: 5′-CTCTTCCCCACACAGGTTG-3′

LIMK1 KO Primer 6: 5′-GACCATGATGGAAGGAGAG-3′

Primer 7: 5′-GGGGGAACTTCCTGACTAGG-3′

2.2 µL of the dissolved DNA was added to 13.5 µL of a PCR Master Mix containing: 0.04 µM primer 1, 0.02 µM primer 2 and primer 3 for GluA2 animals, whereas 0.02 µM primer 4, primer

5, primer 6, and primer 7 for LIMK1 animals, 1.15x PCR buffer with HCl, 1.81 mM MgCl2 (Sigma

Aldrich), 1.45 x 10-2 u/µL Taq Polymerase (New England BioLabs). PCR amplification was performed on an Eppendorf Mastercycler using the following protocol:

Initial Denaturation: 94°C for 3 min

Followed by 33 Cycles of:

Denaturation: 94°C for 30 sec

Annealing: 58°C for 1 min

Extending: 72°C for 1 min

Final Extension: 72°C for 10 minutes

3.2.3 Gel Electrophoresis Each PCR product was mixed with 3 µL of 1x DNA loading buffer and then loaded into a

1.2% agarose gel containing 0.04 uL/mL RedSafe (iNtRON Biotechnology). The gel underwent electrophoresis at 100 V for 60 min and then imaged using ultraviolet light to detect DNA bands.

Sizes of WT and KO PCR products for GluA2 were 600 base pairs (bp) and 500 bp, respectively.

Sizes of WT and KO PCR products for LIMK1 were 500 bp and 300 bp, respectively.

3.3 Electrophysiology

3.3.1 Acute Hippocampal Slice Preparation Sagittal hippocampal slices (350 µm) were obtained from 4 to 6-week old wild-type or mutant mice. The brain was isolated rapidly from the skull using a curved dissection spatula following decapitation and was submerged for 30 sec in ice-cold artificial cerebrospinal fluid

(ACSF). The brain was then blocked by removing the cerebellum, the prefrontal cortex, and removing ~1 mm of the temporal lobe from both hemispheres. The brain was then separated into two halves by cutting along the longitudinal fissure and placing both halves on the lateral face.

The brain was mounted via tissue glue (3M) and sliced using a vibratome (Leica 1200 S). Slices were transferred to a submerged incubation chamber with ACSF saturated with 95% O2 and 5%

CO2. Slices were allowed to recover at 32–34°C for 30 min and then at room temperature for a minimum of 1 h before proceeding to extracellular electrophysiology recordings. ACSF contained the following (mM) (all purchased from Sigma Aldrich unless stated otherwise): 10 glucose, 124

NaCl, 26 NaHCO3, 3 KCl, 1.25 NaH2PO4, 2 MgSO4 (Fisher Scientific), 2 CaCl2

3.3.2 Extracellular Electrophysiology Extracellular recordings were performed in an interface chamber (Warner Instruments) maintained at 30-32°C and perfused at 2 ml/min with ACSF. Synaptic responses were evoked using bipolar platinum/iridium electrodes and fEPSPs were placed 200-400 µm from the cell body layer and recorded from the CA1 region using borosilicate glass electrodes (2-5Ω) (World

Precision Instruments) filled with ACSF. Recordings were monitored using pCLAMP 7 and fEPSPs were analyzed with pCLAMP 10 (Axon Instruments) by taking the slope of the rising phase between 10-90% of the peak response. Field recordings were recorded using an AxoClamp

2B (Axon Instruments) and LPF202A (Warner Instruments) amplifiers. Each experiment was conducted on slices from separate animals, therefore the n value represents both the number of slices and animals used. SCCPs were stimulated at a frequency of 0.067 Hz at 40% of max response. After a stable baseline of at least 30 min, LTP was induced using TBS delivered at basal stimulus intensity. An episode of TBS is comprised of five bursts at 5 Hz, with each burst composed of five pulses at 100 Hz. For cTBS, three episodes were given with an inter-episode interval of 10 s. For sTBS, three episodes were given with an inter-episode interval of 10 min.

Representative sample traces are an average of five consecutive responses, collected from typical experiments (stimulus artifacts were blanked for clarity)

3.4 Compounds Drugs were prepared as frozen stock solutions (stored below -20°C) and dissolved into

ACSF at least 30 min before their bath application. The dissolved compounds were perfused 15-

20 min prior to LTP induction. Vehicle treatments were performed with either <0.1% DMSO

(anisomycin and KT5720) or distilled water (D-AP5) after being added to ACSF. Compounds used were as follows (all purchased from Hello Bio): Anisomycin (2R,3S,4S)-2-[(4-

Methoxyphenyl)methyl]-3,4-pyrrolidinediol 3-acetate, KT5720 [(9R,10S,12S)-2,3,9,10,11, 12- hexahydro-10-hydroxy-9-methyl-1-oxo-9,12-epoxy-1H-diindolo[1,2,3-fg: 3′,2′,1′-kl]pyrrolo[3,4- i][1,6]benzodiazocine-10-carboxylic acid, hexyl ester (KT)], D-AP5 (D-(-)-2-Amino-5- phosphonopentanoic acid)

3.5 Analysis All treatment groups were interleaved with control (i.e. vehicle) experiments. Data were normalized to baseline and for the comparison of the magnitude of LTP, the last 10 min of recordings were compared statistically, with values being presented as mean ± SEM. Statistical significance was assessed using paired and unpaired Student’s t test; the level of significance is denoted as follows: *p < 0.05 and **p < 0.01.

Chapter 4 Results 4.1 cTBS and sTBS induced enhanced LTP in GluA2 KO mice Previous work has shown that activation of PKA-dependent LTP is dependent on induction parameters; where a single episode or multiple episodes compressed in time induce a form of PKA- independent LTP, whereas multiple episodes spaced in time induce PKA-dependent LTP

(Bortolotto & Collingridge, 2000; Y. Huang & Kandel, 1994; Park et al., 2016; Woo et al., 2002).

Previously, our lab has shown that HFS is able to produce enhanced LTP in GluA2 KO mice

(Asrar, Zhou, et al., 2009; Jia et al., 1996). However, studies have demonstrated that potentiation magnitude, as well as molecular pathways, can differ between HFS and TBS (Hernandez et al.,

2005; Zhu et al., 2015). Thus, before investigating the mechanism surrounding PKA-dependent

LTP, I wanted to determine whether the mechanisms observed in GluA2 KO mice following HFS are preserved in TBS.

To test this, stimuli were delivered as three episodes, with each episode comprised of five bursts (with each burst consisting of five shocks at 100 Hz, with an interburst interval of 200 ms).

The inter-episode interval was varied, with cTBS having a 10 s interval whereas sTBS had a 10 min interval. I have found that the levels of LTP following cTBS, expressed as a percentage of the baseline, were 150 ± 8% (n = 5) and 206 ± 13% (n = 5, p = 0.0069) for GluA2 WT and KO groups respectively (Figure 18A, B). Similarly, the levels of LTP following sTBS were 145 ± 6% (n = 4) and 261 ± 32% (n = 5, p = 0.0169) for GluA2 WT and KO groups respectively (Figure 19A, B).

Together, these results suggest that GluA2 KO mice express enhanced LTP regardless of stimulation parameters.

Figure 18. Enhanced cTBS-induced LTP in GluA2 KO mice. A. TBS was delivered 3 times with a compressed protocol (cTBS, 10 sec inter-episode interval) to induce LTP in GluA2 WT (black; n=5) and KO (black, open; n=5) mice. Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between GluA2 WT (black; 150 ± 8%) and KO (black, open; 206 ± 13%) in the last 10 min. **p = 0.0069 vs WT.

Figure 19. Enhanced sTBS-induced LTP in GluA2 KO mice. A. TBS was delivered 3 times with a spaced protocol (sTBS, 10 min inter-episode interval) to induce LTP in GluA2 WT (black; n=4) and KO (black, open; n=5) mice. Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between GluA2 WT (black; 145 ± 6%) and KO (black, open; 261 ± 32%) in the last 10 min. *p < 0.0169 vs WT.

4.2 cTBS and sTBS induced LTP using only CP-AMPARs in GluA2 KO mice The current model for sTBS suggests that the first episode can activate CaMKII-mediated mechanisms found in cTBS-induced LTP, such as alterations in the number or function of

AMPARs (Chater & Goda, 2014). The second episode involves the activation of Ca2+-sensitive adenylyl cyclase and PKA, and the subsequent insertion of CP-AMPARs. All these changes require the initial activation of NMDARs, however, whether these forms of LTP be elicited independent of NMDARs remains unknown. More specifically, can cTBS- and sTBS-induced LTP use pre-existing CP-AMPARs? To isolate the contributions of CP-AMPARs in these different forms of LTP, D-AP5 was used to block NMDARs. As expected, D-AP5 completely blocked LTP in GluA2 WT mice regardless of cTBS (150 ± 8%, n = 5, vs 110 ± 2%, n = 2, p = 0.0290; vehicle control and D-AP5 treatment respectively) (Figure 20A, B) and sTBS stimulation parameters (145

± 6%, n = 4, vs 103 ± 1%, n = 2, p = 0.0097; vehicle control and D-AP5 treatment respectively)

(Figure 21A, B). Rather than completely blocking LTP, D-AP5 was only able to attenuate LTP in

GluA2 KO mice following cTBS (206 ± 13%, n = 5, vs 142 ± 12%, n = 4; vehicle control and D-

AP5 treatment respectively) (Figure 22A, B). Despite the reduction in LTP, there was significant level of potentiation compared to baseline in the D-AP5 treated GluA2 KO mice (142 ± 12%, n =

4, vs 100 ± 2%, n = 4; *p = 0.0476). Similarly, treatment with D-AP5 in GluA2 KO mice following sTBS reduced the levels of LTP (236 ± 27%, n = 4, vs 168 ± 21%, n = 4; vehicle control and D-

AP5 treatment respectively) (Figure 23A, B). Comparable to cTBS, the level of potentiation compared to baseline in the D-AP5 treated GluA2 KO mice was trending towards significance

(168 ± 21%, n = 4, vs 104 ± 2%, n = 4; p = 0.059). This trend may be attributed to low sample size. These findings are in accordance with previous studies and suggest that cTBS and sTBS can induce LTP using only CP-AMPARs in GluA2 KO mice (Asrar, Zhou, et al., 2009; Jia et al.,

1996). The observed reduction in LTP is supported by previous data, as CP-AMPAR-dependent

LTP does not activate CaMKII-dependent mechanisms (Asrar, Zhou, et al., 2009).

Figure 20. NMDARs are required for cTBS-induced LTP in GluA2 WT mice. A. Treatment of slices with D-AP5 blocks cTBS-induced LTP in GluA2 WT (brown; n=2) compared to vehicle control (black; n=5). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between GluA2 WT vehicle control (black; 150 ± 8%) and D-AP5 (brown; 110 ± 2%) in the last 10 min. D-AP5 was applied 15 min prior to the first episode until 5 min following the last episode. *p < 0.0290 vs vehicle control.

Figure 21. NMDARs are required for sTBS-induced LTP in GluA2 WT mice. A. Treatment of slices with D-AP5 blocks sTBS-induced LTP in GluA2 WT (brown; n=2) compared to vehicle control (black; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between GluA2 WT vehicle control (black; 145 ± 6%) and D-AP5 (brown; 103 ± 1%) in the last 10 min. D-AP5 was applied 15 min prior to the first episode until 5 min following the last episode. **p = 0.0097 vs vehicle control.

Figure 22. GluA2 KO mice exhibit cTBS-induced LTP independent of NMDARs. A. Treatment of slices with D-AP5 attenuates, but does not block, cTBS-induced LTP in GluA2 KO (brown, open; n=4) compared to vehicle control (black, open; n=5). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at baseline (black, open; 100 ± 1%) versus during LTP (black, open; 206 ± 13) in GluA2 KO vehicle control and D-AP5 (brown, open; 101 ± 4% vs 142 ± 12%, baseline and LTP, respectively); D-AP5 was applied 15 min prior to the first episode until 5 min following the last episode. **p = 0.0011 vs vehicle control baseline, *p = 0.0476 vs D-AP5 baseline.

Figure 23. GluA2 KO mice exhibit sTBS-induced LTP independent of NMDARs. A. Treatment of slices with D-AP5 attenuates, but does not block, sTBS-induced LTP in GluA2 KO (brown, open; n=4) compared to vehicle control (black, open; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at baseline (black, open; 100 ± 2%) versus during LTP (black, open; 236 ± 27%) in GluA2 KO vehicle control and D-AP5 (brown, open; 104 ± 2% vs 168 ± 21%, baseline and LTP, respectively); D-AP5 was applied 15 min prior to the first episode until 5 min following the last episode. *p = 0.0183 LTP vs vehicle control baseline, p = 0.059 LTP vs D-AP5 baseline.

4.3 sTBS-induced CP-AMPAR-dependent LTP required de novo protein synthesis One of the hallmark features of sTBS-induced LTP is the requirement of PKA and related molecular pathways, of which includes de novo protein synthesis – a process critical for the consolidation of STM into LTM (Abel et al., 1997; Scharf et al., 2002). Previous literature has demonstrated that these mechanisms require initial activation via NMDARs (Lüscher & Malenka,

2012; Park et al., 2014). However, since sTBS is capable of eliciting LTP via CP-AMPARs, is it possible that CP-AMPARs can activate protein synthesis-dependent pathways as seen with

NMDARs? To investigate this phenomenon, I continued to use D-AP5, however, in conjunction with anisomycin (ANI), a protein synthesis inhibitor, to isolate the effect of CP-AMPARs on de novo protein synthesis. The results suggest that sTBS-induced CP-AMPAR-dependent LTP is capable of eliciting long-lasting forms of plasticity that require de novo protein synthesis, as administration of D-AP5 and ANI did not elicit significant potentiation compared to baseline (116

± 8% vs 101 ± 3%, n = 4, p = 0.1171 vs baseline) in GluA2 KO mice (Figure 24A, B).

Figure 24. sTBS-induced LTP requires protein synthesis in GluA2 KO mice. A. Treatment of slices with D-AP5 + Anisomycin (ANI) blocks sTBS-induced LTP in GluA2 KO (purple, open; n=4) compared to D-AP5 treated slices (brown, open; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at baseline (brown, open; 104 ± 2%) versus during LTP (brown, open; 168 ± 21% in GluA2 KO D-AP5 vs D-AP5 + ANI (purple, open; 101 ± 3% vs 116 ± 8%, baseline and LTP, respectively). p = 0.059 LTP vs D-AP5 baseline. p = 0.1171 LTP vs D-AP5 + ANI baseline.

4.4 Inhibition of PKA transiently abolished CP-AMPAR-dependent LTP The results thus far suggest that CP-AMPAR-dependent LTP requires processes downstream of PKA, such as de novo protein synthesis. However, it is unknown if PKA is required by CP-AMPARs following LTP induction. Furthermore, it is unknown if different stimulation parameters result in differential requirement of PKA in animals with widespread expression of CP-

AMPARs. To investigate the requirement of PKA in CP-AMPAR-dependent LTP, KT5720, a

PKA inhibitor, was used in conjunction with D-AP5. D-AP5 + KT5720 had no effect on LTP following cTBS (133 ± 13%, n = 3) versus D-AP5 control (142 ± 12%, n = 4) (Figure 25A, B).

Interestingly, while D-AP5 + KT5720 had no effect on the last 10 min of LTP following cTBS, the observed levels of LTP during application of the treatment (15 min) were no different compared to that of baseline (89 ± 20%, n = 3, and 106 ± 5%, n = 3, respectively; p = 0.4672)

(Figure 25B). However, following washout (60 min), the level of potentiation in the D-AP5 +

KT5720 treated group was trending towards significance compared to baseline (133 ± 13% vs 106

± 5%, 60 min vs -20 min respectively, n = 3; p = 0.1196) (Figure 25B). A similar phenomenon occurred during application of D-AP5 + KT5720 during sTBS-induced LTP; no differences in the levels of LTP were observed in the D-AP5 + KT5720 group versus D-AP5 control (155 ± 18%, n

= 4 and 168 ± 21%, n = 4, respectively) (Figure 26A, B). Likewise, following sTBS induction, there were no observable differences between baseline at -20 min (96 ± 2%, n = 4) and application of D-AP5 + KT5720 at 30 min (111 ± 6%, n = 4, p = 0.0675) (Figure 26B). However, levels of

LTP following washout at 80 min was trending towards a significant increase compared to baseline

(155 ± 18% vs 96 ± 2%, respectively, n = 4; p = 0.0541) (Figure 26B). The trends observed in cTBS and sTBS groups treated with D-AP5 + KT5720 may be attributed to low sample size. The results suggest that CP-AMPAR-dependent LTP is transiently abolished following the inhibition

of PKA. More importantly, it appears that removal of PKA inhibition results in potentiation in

GluA2 KO mice.

Figure 25. Inhibition of PKA transiently abolished cTBS-induced LTP in GluA2 KO mice. A. D-AP5 + KT5720 completely blocks LTP during treatment, however cTBS-induced LTP is restored following washout (green, open; n=3) and is indistinguishable from D-AP5 treated groups (brown, open; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at -20 min, 15 min, and 60 min for GluA2 mice treated with D-AP5 (brown, open; 104 ± 2%, 143 ± 19%, 142 ± 12%, respectively) and D-AP5 + KT5720 (green, open; 106 ± 5%, 89 ± 20%, 133 ± 13%, respectively). D-AP5 + KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. *p = 0.0489 for 15 min vs -20 min in D-AP5 treatment group. *p = 0.0476 for 60 min vs -20 min in D- AP5 treatment group. p = 0.12 for 60 min vs -20 min in D-AP5 + KT5720 treatment.

Figure 26. Inhibition of PKA transiently abolishes sTBS-induced LTP in GluA2 KO mice. A. D-AP5 + KT5720 completely blocks LTP during treatment, however sTBS-induced LTP is restored following washout (green, open; n=4) and is indistinguishable from D-AP5 treated groups (brown, open; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at -20 min, 30 min, and 80 min for GluA2 mice treated with D-AP5 (brown, open; 104 ± 2%, 148 ± 16%, 168 ± 21%, respectively) and D-AP5 + KT5720 (green, open; 96 ± 2%, 111 ± 6%, 155 ± 18%, respectively). D-AP5 + KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.06 80 min vs -20 min in D-AP5 treatment. p = 0.0541 for 80 min vs -20 min in D-AP5 + KT5720 treatment.

4.5 CP-AMPAR-dependent LTP requires PKA The present observations suggest that inhibition of PKA transiently abolishes CP-AMPAR- dependent LTP, as washout of D-AP5 + KT5720 restored levels of LTP regardless of stimulation parameters. This may suggest that PKA is required throughout the maintenance of CP-AMPAR- dependent LTP. To test this, D-AP5 + KT5720 was perfused for different lengths of time (30 min and 1 hr) following the third TBS episode. There were no significant levels of potentiation during at 40 min compared to baseline (-20 min) (101 ± 8% vs 102 ± 3%, respectively, n = 3) (Figure

27A, B). This is unlike the D-AP5 control, where levels of potentiation at 40 min trended towards significance compared to -20 min (141 ± 13% vs 104 ± 2%, respectively, n = 4; p = 0.0762) (Figure

27B). Similar to the previous experiments, the D-AP5 + KT5720 group presented a trend towards significant increase in level of potentiation following washout at 80 min compared to baseline (137

± 12% vs 102 ± 3%, respectively, n = 3; p = 0.1418) (Figure 27B). Interestingly, if D-AP5 +

KT5720 was perfused for the entire LTP period (1 hr following the third TBS episode), the levels of LTP were completely blocked as levels of potentiation at 80 min were not significantly different compared to baseline (101 ± 3% vs 100 ± 5%, respectively, n = 4) (Figure 28A, B). All observed trends in the sTBS groups treated with D-AP5 + KT5720 may be attributed to low sample size.

Altogether, the results suggest that PKA is required throughout CP-AMPAR-dependent LTP as potentiation could only be blocked via continuous application of D-AP5 + KT5720.

Figure 27. Blockade of sTBS-induced LTP is dependent on the duration of PKA inhibition in GluA2 KO mice. A. Prolonged treatment of D-AP5 + KT5720 only blocks LTP during the treatment period, as sTBS-induced LTP is restored following washout (green, open; n=3) and is indistinguishable from D-AP5 treated groups (brown, open; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at -20 min, 40 min, and 80 min for GluA2 mice treated with D-AP5 (brown, open; 104 ± 2%, 141 ± 13%, 168 ± 21%, respectively) and D-AP5 + KT5720 (green, open; 102 ± 3%, 101 ± 8%, 137 ± 12%, respectively). D-AP5 + KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.1418 for 80 min vs -20 min in D-AP5 + KT5720 treatment.

Figure 28. PKA is required for the maintenance of sTBS-induced LTP in GluA2 KO mice. A. Treatment of D-AP5 + KT5720 throughout the entire recording period completely blocked sTBS-induced LTP (green, open; n=4) compared to D-AP5 treated groups (brown, open; n=4). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the fEPSP at -20 min, 40 min, and 80 min for GluA2 mice treated with D-AP5 (brown, open; 104 ± 2%, 141 ± 13%, 168 ± 21%, respectively) and D-AP5 + KT5720 (green, open; 101 ± 3%, 109 ± 11%, 100 ± 5%, respectively). D-AP5 + KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.06 80 min vs -20 min in D-AP5 treatment.

4.6 LIMK1 is required for sTBS-induced LTP While there is overwhelming evidence for the involvement of PKA in long-lasting activity- dependent synaptic changes, such as AMPAR trafficking and morphological alterations, how PKA exerts these biochemical changes remain relatively unknown. One protein of interest is LIMK1, a protein critical in actin cytoskeleton dynamics and structural plasticity, as studies have shown it directly interacts with PKA. However, there is no evidence of this interaction in the context of synaptic plasticity. Thus, to investigate the role of LIMK1 in the PKA signaling pathway, stimulation parameters activating PKA-independent (cTBS) and –dependent (sTBS) LTP were used on LIMK1 KO mice. cTBS revealed no significant differences in the levels of LTP between

LIMK1 WT (162 ± 5%, n = 6) and KO (151 ± 7%, n = 4, p = 0.2053) (Figure 29A, B). Intriguingly, induction of PKA-dependent LTP via sTBS revealed significant deficits in LIMK1 KO (123 ±

10%, n = 5) compared to LIMK1 WT (164 ± 10%, n = 5, p = 0.0185). These results indicate that

LIMK1 is required for sTBS-induced LTP (Figure 30A, B)

Figure 29. LIMK1 is not required for cTBS-induced LTP. A. TBS was delivered 3 times with cTBS to induce LTP in LIMK1 WT (black; n=6) and KO (black, open; n=4) mice. Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 WT (black; 162 ± 5%) and KO (black, open; 151 ± 7%) in the last 10 min. p = 0.2053 vs WT.

Figure 30. LIMK1 is required for sTBS-induced LTP. A. TBS was delivered 3 times with sTBS to induce LTP in LIMK1 WT (black; n=5) and KO (black, open; n=5) mice. Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 WT (black; 164 ± 10%) and KO (black, open; 123 ± 10%) in the last 10 min. *p = 0.0185 vs WT.

4.7 LIMK1 is required for PKA-dependent LTP Although the current observations imply that LIMK1 KO mice have impaired sTBS- induced PKA-dependent LTP, there is no evidence to suggest the reduction in levels of LTP are caused by impaired PKA activity. However, prior to investigating this phenomenon, I had to examine whether cTBS or sTBS could elicit PKA-sensitive forms of LTP. Using LIMK1 WT mice, I used KT5720 to examine whether cTBS or sTBS stimulation parameters induced PKA- sensitive forms of LTP. Application of KT5720 following cTBS in LIMK1 WT mice revealed no significant differences compared to the vehicle control group (151 ± 11%, n = 4, and 162 ± 5%, n

= 6, respectively; p = 0.0743) (Figure 31A, B). In accordance to previous studies, application of

KT5720 in LIMK1 WT mice following the sTBS stimulation parameter resulted in significant reduction in the levels of LTP compared to vehicle controls (139 ± 5%, n = 5, and 164 ± 10%, n =

5, respectively; p = 0.0460) (Figure 32A, B). Indeed, these findings suggest that cTBS and sTBS stimulation parameters elicit PKA-independent and –dependent forms of LTP, respectively. Since the results suggest that sTBS induces a form of PKA-dependent LTP, this then brings to question: is PKA activity impaired following sTBS-induced LTP LIMK1 KO mice? To examine this question, I used KT5720 to determine whether PKA activity is impaired following cTBS or sTBS- induced forms of LTP in LIMK1 KO mice. Application of KT5720 following cTBS resulted in no significant difference in the level of LTP compared to the vehicle control group (157 ± 14% n =

4, and 162 ± 5%, n = 6, respectively; p = 0.6712) (Figure 33A, B). Likewise, application of KT5720 following sTBS resulted in no significant difference in the level of LTP compared to the vehicle control group (130 ± 13%, n = 7, and 123 ± 10%, n = 5, respectively; p = 0.7179) (Figure 34A, B).

Because KT5720 did not further reduce the level of sTBS-induced LTP, this suggests that PKA activity is impaired in LIMK1 KO mice. Thus, the results suggest that LIMK1 is required for PKA- dependent LTP.

Figure 31. PKA is not required for cTBS-induced LTP. A. Treatment of LIMK1 WT slices with KT5720 (green; n=4) has no effect on the levels of cTBS-induced LTP compared to vehicle control (black; n=6). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 WT vehicle control (black; 162 ± 5%) and KT5720 (green; 151 ± 11%) in the last 10 min. KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.0743 vs vehicle control.

Figure 32. PKA is required for sTBS-induced LTP. A. Treatment of LIMK1 WT slices with KT5720 (green; n=5) reduced the levels of sTBS-induced LTP compared to vehicle control (black; n=5). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 WT vehicle control (black; 164 ± 10%) and KT5720 (green; 139 ± 5%) in the last 10 min. KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. *p = 0.0460 vs vehicle control.

Figure 33. LIMK1 is not required for PKA-independent LTP. A. Treatment of LIMK1 KO slices with KT5720 (green; n=4) had no effect on cTBS-induced LTP compared to vehicle control (black; n=6). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 KO vehicle control (black; 162 ± 5%) and KT5720 (green; 157 ± 14%) in the last 10 min. KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.6712 vs vehicle control.

Figure 34. LIMK1 is required for PKA-dependent LTP. A. Treatment of LIMK1 KO slices with KT5720 (green; n=7) failed to further reduce sTBS-induced LTP compared to vehicle control (black; n=6). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 KO vehicle control (black; 123 ± 10%) and KT5720 (green; 130 ± 13%) in the last 10 min. KT5720 was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.7179 vs vehicle control.

4.8 LIMK1 is required for de novo protein synthesis in PKA- dependent LTP The ubiquitous nature of PKA allows it to modulate long-lasting structural synaptic changes through various cellular processes, one of which includes activity-dependent gene transcription. To identify the components that contribute to the selective deficits observed in

LIMK1 KO mice in PKA-dependent LTP, I used ANI to probe the requirement of protein synthesis following the induction of PKA-independent and –dependent LTP using the cTBS and sTBS stimulation parameters, respectively. In LIMK1 WT mice, ANI had no effect on PKA-independent

LTP compared to the vehicle control group (150 ± 2%, n = 5, and 162 ± 5%, n = 6, respectively; p = 0.0743) (Figure 35A, B). Conversely, ANI attenuated PKA-dependent LTP compared to the vehicle control group (132 ± 2%, n = 6, and 164 ± 10%, n = 5, respectively; p = 0.0057) (Figure

36A, B), as supported by the literature. Similar to LIMK1 WT mice, ANI had no effect on PKA- independent LTP in LIMK1 KO mice compared to vehicle control (157 ± 13%, n = 7, and 162 ±

5%, n = 6, respectively; p = 0.7494) (Figure 37A, B). Likewise, ANI had no effect on PKA- dependent LTP in LIMK1 KO mice compared to vehicle control (126 ± 6%, n = 6, and 123 ± 10%, n = 5, respectively; p = 0.8316) (Figure 38A, B), however the levels of LTP were reduced compared to LIMK1 WT. Together, these data suggest that LIMK1 is required for protein synthesis in PKA-dependent LTP.

Figure 35. De novo protein synthesis is not required for PKA-independent LTP. A. Treatment of LIMK1 WT slices with ANI (blue; n=5) has no effect on the levels of cTBS-induced LTP compared to vehicle control (black; n=6). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 WT vehicle control (black; 150 ± 2%) and ANI (blue; 162 ± 5%) in the last 10 min. ANI was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.0743 vs vehicle control.

Figure 36. De novo protein synthesis is required for PKA-dependent LTP. A. Treatment of LIMK1 WT slices with ANI (blue; n=6) reduced the levels of sTBS-induced LTP compared to vehicle control (black; n=5). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 WT vehicle control (black; 164 ± 10%) and ANI (blue; 132 ± 2%) in the last 10 min. ANI was applied 15 min prior to the first episode until 15 min following the last episode. **p = 0.0057 vs vehicle control.

Figure 37. De novo protein synthesis and LIMK1 is not required for PKA-independent LTP. A. Treatment of LIMK1 KO slices with ANI (blue, open; n=7) has no effect on the levels of cTBS- induced LTP compared to vehicle control (black, open; n=6). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 KO vehicle control (black, open; 162 ± 5%) and ANI (blue, open; 157 ± 13%) in the last 10 min. ANI was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.7494 vs vehicle control.

Figure 38. PKA-dependent LTP requires LIMK1-mediated de novo protein synthesis. A. Treatment of LIMK1 KO slices with ANI (blue, open; n=6) has no effect on the levels of sTBS- induced LTP compared to vehicle control (black, open; n=5). Traces are averages of 5 consecutive sweeps for baseline (1) and the last 10 min of LTP (2). B. Summary graph comparing the levels of LTP between LIMK1 KO vehicle control (black, open; 123 ± 10%) and ANI (blue, open; 126 ± 6%) in the last 10 min. ANI was applied 15 min prior to the first episode until 15 min following the last episode. p = 0.8316 vs vehicle control.

Chapter 5 Discussion In the present study, I set out to determine if PKA-dependent LTP can be triggered by CP-

AMPARs and if PKA-dependent protein synthesis is mediated by LIMK1. To address this hypothesis, my objectives were three-fold. Firstly, cTBS and sTBS protocols in GluA2 and LIMK1

KO mice needed to be established. I found that cTBS and sTBS protocols could elicit LTP in both

GluA2 and LIMK1 KO with varying magnitudes, suggesting alterations in the molecular pathways. Secondly, a combination of methods in genetics, electrophysiology, and pharmacology were used to determine if PKA is required for CP-AMPAR-dependent LTP. Indeed, I have found that CP-AMPARs can induce PKA-dependent forms of LTP in the absence of NMDAR activity regardless of stimulation protocol. This conclusion is based on experiments that revealed significant reduction in the levels of CP-AMPAR-dependent LTP during perfusion of distinct antagonists targeting NMDARs and PKA following cTBS and sTBS. Finally, I employed the same methodology to determine if LIMK1 is required for PKA-mediated activity dependent protein synthesis. I have found that LIMK1 is required for PKA-dependent LTP as sTBS-induced LTP revealed reduced levels of LTP compared to cTBS stimulation parameters. Furthermore, application of a PKA antagonist had no effect on cTBS stimulation parameters and failed to further reduce sTBS-induced LTP. Moreover, my study has shown that activity dependent de novo protein synthesis requires PKA and LIMK1. These findings are supported by results showing complete inhibition of LTP following treatment with a protein synthesis inhibitor in PKA/CP-AMPAR- dependent LTP in GluA2 KO mice. Moreover, the protein synthesis inhibitor did not further reduce levels of PKA-dependent LTP in LIMK1 KO mice. Interestingly, these mechanisms are PKA-

dependent because altogether, the inhibitors targeting PKA and protein synthesis had no effect whatsoever on cTBS-induced LTP in WT controls.

5.1 CP-AMPAR-dependent LTP is mediated by PKA I have found that animals solely expressing CP-AMPARs are capable of expressing a form of PKA-dependent LTP that is supported by some (Park et al., 2016; Plant et al., 2006) but not all studies (Adesnik & Nicoll, 2007). While this study utilized animals expressing GluA2-lacking CP-

AMPARs, it is unknown if the same mechanisms can be activated with unedited GluA2(Q) subunits.

Interestingly, GluA2 KO mice can express PKA-dependent LTP regardless of cTBS or sTBS stimulation parameters. This is supported by my observations of inhibition of LTP throughout the duration of the PKA inhibitor treatment regardless of cTBS- or sTBS-induced LTP

(Figure 27). While this study utilized animals expressing GluA2-lacking CP-AMPARs, it is unknown if the same mechanisms can be activated with unedited GluA2(Q) subunits. However, since the majority of expressed CP-AMPARs are GluA2-lacking, it is reasonable to assume that the activation of PKA-dependent LTP is primarily mediated by GluA2-lacking CP-AMPARs.

Previous studies have suggested that a critical time window exists for the activation of

PKA-dependent LTP, as application of PKA or CP-AMPAR inhibitors during or immediately following the third TBS episode can reduce the levels of LTP whereas no effect is seen when applied to the maintenance phase of LTP (Park et al., 2016; Plant et al., 2006). However, results from my study suggest that activation of CP-AMPAR-dependent LTP constantly requires PKA, as LTP was only inhibited during the presence of PKA inhibitors and washout of the inhibitor

restored levels of LTP (Figure 25, 26). This is in contrast to other CP-AMPAR specific molecular pathways, such as the PI3/MAPK signaling cascade, where its required for only the induction but not maintenance phase. This then, prompts two questions: firstly, how do different stimulation patterns activate PKA? Second, how does PKA facilitate potentiation despite pharmacological blockade during electrical stimulation?

One of the mechanisms underlying the stimulation pattern-dependent activation of PKA is that cTBS and sTBS protocols differentially change intracellular Ca2+ concentrations. Previous reports have suggested that HFS can raise intracellular Ca2+ by fivefold compared to baseline levels

(Bonsi et al., 2004; Cooper, Mons, & Karpen, 1995). It has been suggested that differences in the frequency of stimulation patterns, such as those seen in HFS and LFS, results in different changes in intracellular Ca2+, where HFS can induce a much greater increase in Ca2+ than LFS (Bonsi et al., 2004). Thus, in the case of cTBS and sTBS, it is possible that sTBS is capable of increasing intracellular Ca2+ concentrations to a critical level required for PKA activity. Interestingly, in

GluA2 KO mice, LTP required PKA regardless of stimulation pattern. This may be attributed to the widespread expression of CP-AMPARs at baseline conditions, resulting in elevated intracellular Ca2+ compared to WT control animals following any stimulation parameter. Further studies are required to determine if intracellular concentrations of Ca2+ in GluA2 WT and KO animals differ, as well as whether or not these differences are observed in cTBS and sTBS stimulation patterns.

It was observed that PKA can facilitate potentiation despite pharmacological blockade during electrical stimulation. This suggests that although PKA can drive CP-AMPARs to the synapse, CP-AMPARs are capable of triggering and maintaining PKA activity. This then brings

to question, when and how do CP-AMPARs activate PKA? During sTBS, the first stimulus activates NMDARs and induces cTBS-dependent mechanisms, such as CaMKII-associated pathways, and primes for sTBS-induced LTP. This priming process includes activation of PKA and insertion of CP-AMPARs to the plasma membrane, but not the synapse itself. Subsequent spaced stimuli then result in the translocation of CP-AMPARs to the synaptic space (Park et al.,

2016). However, once at the synapse, Ca2+-influx from CP-AMPARs maintain PKA activity and trigger mechanisms required for long-lasting plasticity, such as protein synthesis and changes to structural morphology, until they are internalized following a period of time. This model supports the notion that synaptic expression of CP-AMPARs are transient (Plant et al., 2006).

5.2 A role for LIMK1 in sTBS-induced PKA-dependent LTP Previously, our lab has shown that LIMK1 KO mice have selective deficits in L-LTP.

Furthermore, we have shown that LIMK1 interacts with and regulates CREB activity, a critical effector downstream of PKA (Todorovski et al., 2015). Additionally, PKA has been suggested to interact with LIMK1, as PKA null mouse embryonic fibroblasts demonstrated lowered phosphorylation levels of LIMK1 and altered cell shape and size (Nadella et al., 2009). Together, these studies suggest that PKA requires LIMK1 in certain forms of long-lasting plasticity. Indeed, my studies suggest that PKA requires LIMK1 as I observed selective deficits for the levels of LTP following the activation of the PKA-dependent pathway via sTBS in LIMK1 KO mice (Figure 29).

This is in contrast to cTBS-induced PKA-independent LTP, where levels of LTP were unaffected

(Figure 28). In accordance to a previous study, I observed no effect on the levels of cTBS-induced

LTP following the pharmacological inhibition of PKA (Figure 30) (Park et al., 2016). Interestingly,

PKA inhibition did not further attenuate levels of sTBS-induced LTP in LIMK1 KO animals

(Figure 33). Together with the previous findings, my results provide evidence that LIMK1 is

downstream of PKA (Nadella et al., 2009). Furthermore, my findings suggest that LIMK1 is critical for PKA-dependent LTP.

5.3 PKA-mediated protein synthesis-dependent LTP requires LIMK1 Past studies have shown that PKA can modulate LIMK1 activity via phosphorylation to enhance actin filament turnover, a process that critically regulates spine structure and structural plasticity, subsequently (Nadella et al., 2009). Similarly, data from our lab has demonstrated that

LIMK1 KO mice have abnormal spine morphology due to altered cofilin activity, which resulted in abnormalities in actin distribution (Y. Meng et al., 2002). However, in a separate study, our lab has shown that regulation of L-LTP in LIMK1 KO mice is independent of cofilin activity, as manipulation of cofilin activity did not rescue L-LTP deficits in LIMK1 KO mice (Todorovski et al., 2015). Interestingly, in the same study, our lab has shown that LIMK1 can activate CREB, a transcription factor that is also downstream of PKA and critical for the establishment of L-LTP, in an activity-dependent manner. Because PKA-dependent LTP is a form of L-LTP, this brings to question, rather than regulating structural plasticity via actin cytoskeleton dynamics, does PKA- dependent activation of LIMK1 regulate this process via de novo protein synthesis?

Evidence from my study showed that inhibitors of NMDARs and protein synthesis could completely block LTP following CP-AMPAR-dependent activation of PKA (Figure 23). This suggests that PKA is required for de novo protein synthesis, which is in accordance to earlier studies, however it is unclear whether or not LIMK1 is required for this process (Abel et al., 1997;

Asrar, Zhou, et al., 2009; Scharf et al., 2018). Based on my experiments, it appears that LIMK1 is required for de novo protein synthesis, as application of an inhibitor of protein synthesis did not further reduce levels of LTP in LIMK1 KO mice following activation of PKA-dependent LTP

(Figure 37). In conjunction with previous findings, my study provides evidence that PKA is required to maintain CP-AMPAR-dependent LTP, and PKA-dependent mechanisms require

LIMK1 (Nadella et al., 2009; Park et al., 2016). My study provides evidence that LIMK1 has two roles: firstly, LIMK1 regulates actin dynamics via cofilin and secondly, LIMK1 is required for

PKA-dependent mechanisms, such as de novo protein synthesis (Y. Meng et al., 2002; Todorovski et al., 2015).

Figure 39. sTBS-induced LTP via NMDAR or CP-AMPAR requires PKA-LIMK1-mediated de novo protein synthesis. A. Following sTBS, NMDAR-mediated influx of Ca2+ activates the cAMP signaling cascade, which activates molecular pathways downstream of PKA, such as recruitment of CP-AMPAR to the synapse and LIMK1-mediated de novo protein synthesis. B. CP- AMPAR-dependent LTP is also mediated by PKA-dependent mechanisms as described in A.

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Chapter 6 Future Directions Although my study further elucidates the molecular mechanisms downstream of CP-

AMPAR-dependent LTP, it presents itself with limitations in methodology, namely the extensive use of in vitro electrophysiology and pharmacology. To broaden the scope of this study and to further elaborate the molecular mechanisms, I wish to diversify the techniques for future investigation by combining the use of biochemical and imaging techniques.

For this study, I have assumed that the only difference between GluA2 WT and KO mice is the widespread expression of CP-AMPARs and capability to produce LTP independent of NMDARs in the latter animal model. However, based on the findings in my current study, if CP-AMPARs are capable of activating PKA-dependent pathways, then we would expect phosphorylation levels of associated proteins, such as LIMK1, cofilin, and CREB, to be elevated (Y. Meng et al., 2002,

2004; Todorovski et al., 2015). By using biochemical techniques such as Western blot analysis on brain lysates from GluA2 KO mice prior to and after LTP, I would be able to test whether these molecular pathways are altered. Since these pathways critically regulate dendritic spine morphology, if they are indeed altered in GluA2 KO mice, then I would expect the shape and size of the spines to be enlarged (Y. Meng et al., 2002, 2004; Todorovski et al., 2015). Technological advancements have made two-photon microscopy techniques a valuable tool for determining spatiotemporal changes in spine morphology. In particular, spine morphology can be directly monitored within GluA2 KO mice during LTP.

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While the current study assumed that the selective deficits observed in sTBS-induced LTP in LIMK1 KO mice was primarily due to protein synthesis-dependent structural plasticity, the effects of receptor trafficking should not be neglected. Although our lab has previously shown that modulation of cofilin failed to rescue L-LTP deficits, this may be in part due to the treatment paradigm. Because AMPAR trafficking is activity-dependent, treatment of slices with peptides that modulate cofilin activity should occur throughout LTP experiments rather than prior

(Todorovski et al., 2015). If this were to have an effect, it would be of interest to test whether there is a difference in the number and type of AMPARs trafficked prior to and following LTP induction in LIMK1 KO mice. By differentially tagging GluA1 and GluA2 subunits with pHluorin and pHuji, pH-sensitive green and red fluorescent proteins respectively, I would be able to visually determine whether or not CP-AMPARs are trafficked to the synapse (Mahon, 2011; Rathje et al.,

2013; Shen, Rosendale, Campbell, & Perrais, 2014; Zhang, Cudmore, Lin, Linden, & Huganir,

2015). Furthermore, I would also be able to determine whether or not CP-AMPAR trafficking requires LIMK1.

Understanding the role of CP-AMPARs in various pathological conditions should be of great importance as many neurological and psychiatric conditions are presented with increased expression of CP-AMPARs. The use of GluA2 KO mice can provide valuable insight as to whether reduction of Ca2+-influx is sufficient to restore aberrant signaling pathways. As demonstrated in this study, signaling pathways downstream of PKA, such as LIMK1, appear to be promising therapeutic targets. Thus, the GluA2 KO animal model can be used to probe the efficacy of other therapeutic candidates.

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Chapter 7 Conclusion Ca2+ signaling is a tightly regulated process as it underlies many critical physiological functions. However, in certain neurodegenerative and developmental disease states, Ca2+ homeostasis is lost and results in aberrant signaling. Despite the relative low abundance of CP-

AMPARs within the hippocampus, many disease states, such as Alzheimer’s Disease and epilepsy, result in upregulation of CP-AMPARs and activation of molecular pathways that unique and not seen in NMDARs. In this study, I have further elucidated the molecular pathways downstream of

CP-AMPAR-dependent LTP. Namely, I have demonstrated that CP-AMPARs require PKA independent of NMDAR activity. I have also highlighted that PKA-dependent LTP requires

LIMK1, and that this signaling pathway is a critical regulator of protein synthesis-dependent mechanisms that may be related to structural plasticity, in addition to its existing role in regulating actin cytoskeleton dynamics. The mechanisms I have highlighted in this study provide us with a better understanding of the role of CP-AMPARs in synaptic plasticity in health and disease, and provides valuable insight with respect to potential therapeutic targets.

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Figure 2 adapted by permission from Oxford University Press: The Hippocampus Book; Per Andersen, Richard Morris, David Amaral, Tim Bliss, and John O'Keefe, 2006.

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