Regulation of Dendritic Spine Proliferation Orenda Lyons Johnson

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THE FLORIDA STATE UNIVERSITY

COLLEGE OF ARTS AND SCIENCES

REGULATION OF DENDRITIC SPINE PROLIFERATION

ORENDA LYONS JOHNSON

A Dissertation submitted to the Department of Psychology Program in Neuroscience in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Degree Awarded: Summer Semester, 2005

The members of the Committee approve the Dissertation of Orenda L. Johnson defended on June 17, 2005.

Charles C. Ouimet Professor Directing Dissertation

Paul Q. Trombley Outside Committee Member

Colleen M. Kelley Committee Member

Cathy W. Levenson Committee Member

Zuoxin Wang Committee Member

Approved:

Janet A. Kistner, Chair, Department of Psychology

Joseph A. Travis, Dean, College of Arts and Sciences

The Office of Graduate Studies has verified and approved the above named committee members.

This body of work is dedicated to Carolyn Anagnos, my grandmother.

iii

ACKNOWLEDGEMENTS

Very special thanks…

To the following professors and staff members whose guidance, assistance and technical support have been invaluable during my tenure at Florida State: Charles Badland (Neuroscience Graphics), Karen Berkley, Ellen Berler, Laura Blakemore, Rani Dahnarajan (Molecular Cloning), Jon Ekman, Debbie Fadool, Jim Fadool, Tom Fellers, Cheryl Fitch, Mark Freeman, Chris Gillis, Dick Hagen, Tom Houpt, Rick Hyson, Frank Johnson, Colleen Kelley, Cathy Levenson, David Lowe, Mike Overton, Mike Rashotte, Kim Riddle (Biological Sciences Imaging Resources), Jim Smith, Paul Trombley, and Zuoxin Wang. Also to the administrative staff of the Psychology Department and the Program in Neuroscience: Cherie Dilworth, Sunsan Gay, Janice Parker, Sandy Saunders, Sharon Weaver and Earline Whitley.

To my fellow students with whom I grew academically, professionally and personally: Brandon Aragona, Heather Bradshaw, Jesse Brann, Angie Cason, Brad Chambers, Beverly Colley, Mike Darcy, David Dietz, Karen Dietz, Christie Fowler, Glen Golden, Bumsup Kwon, Andrea Leishman, Denessa Lockwood, Micky Messina, Seth Norrholm, Chad Samuelsen, Mike Sellix, Patrick Smith, Derek Snyder, Carrie Stafstrom, Jake VanLandingham, Jennifer Wersinger, Jenne Westberry and Brandy Wilkinson. As Dr. Carter said, I feel like I grew up with you.

To the Ouimet Lab technicians, who taught me many of the techniques used in this dissertation, who worked tirelessly by my side or in my place, and who all became dear friends: Katie Cavnar, Amanda Clark and Lisa Sander.

To Dr. Ouimet, whose kindness, respect, intellect, encouragement, mentorship, direction, wit and friendship are unparalleled in a “boss,” and have likely spoiled me forever.

To my wonderful devoted friends, for their tireless support and constant understanding: Bay Abreu, Sheyna Carroccio, Kristin Dozier, Kristen Holland, Gwen Hooper, Melanie Jones, LoriAnn Kehler, Eirin Lombardo and Shea Zahner.

To my husband, Eric Johnson, my mother, Judith Lyons, my father, Tom Lyons, my sister, Halle Lyons, and to my extended family, whose amazing support and encouragement and faith in my abilities played an essential role in the completion of this dissertation, and in all of my academic, professional and personal accomplishments. Also to my son Logan Johnson, who has given me unparalleled perspective and incredible joy. And again to Eric, for his infinite patience.

TABLE OF CONTENTS

List of Tables ...... viii List of Figures ...... ix Abstract ...... x

1. Introduction ...... 1 Composition of dendritic spines ...... 1 Dendritic spine development ...... 3 Function of dendritic spines...... 4 Dendritic spine calcium dynamics...... 6 Local protein synthesis ...... 8 Plasticity of dendritic spines: an essential role for actin...... 9

2. Protein synthesis is necessary for dendritic spine proliferation in adult brain slices ...... 12 Introduction ...... 12 Methods ...... 13 Results ...... 17 Discussion ...... 27 Conclusions ...... 30

3. A regulatory role for actin in dendritic spine proliferation...... 31 Introduction ...... 31 Methods ...... 32 Results ...... 37 Discussion ...... 45 Conclusions ...... 51

4. Discussion ...... 52 Methodological considerations...... 52 A common pathway for spine growth?...... 54 Actin upregulation during spinogenesis ...... 54 Proteins involved in spine growth ...... 55 Possible involvement of dendritic protein synthesis...... 59 Actin: a limiting resource for spine density...... 60 Actin treadmilling as a delivery system...... 61 Conclusions ...... 63

REFERENCES ...... 64

BIOGRAPHICAL SKETCH ...... 81

vii

LIST OF TABLES

Table 1: Heterogeneous protein distribution in dendritic spines ...... 2

Table 2: Degree of analysis for counting spines...... 18

Table 3: Degree of analysis for measuring spine length and dendritic width...... 19

Table 4: Morphological consequences of protein overexpression or constitutive expression in neurons...... 56

Table 5: Spine-localized proteins with actin binding domains or motifs ...... 62

viii

LIST OF FIGURES

Figure 1: Transfer of slices does not affect dendritic spine density ...... 15

Figure 2: Spine density increases on secondary apical dendrites of CA1 as a function of time (minutes in vitro)...... 20

Figure 3: Protein synthesis inhibitors block spine proliferation ...... 21

Figure 4: Representative dendritic twigs from each experimental group ...... 23

Figure 5: Early treatment with PUR does not prune spines ...... 24

Figure 6: Mean spine length increases as a function of time, an effect that is blocked by protein synthesis inhibition ...... 25-26

Figure 7: Spine density as assessed by Golgi staining increases during the acute slice preparation in P21 rats...... 38

Figure 8: Spine density as assessed by phalloidin staining increases during the acute slice preparation in P21 rats ...... 39

Figure 9: Actin expression increases during acute slice preparation...... 40

Figure 10: Spine density increases after actin-overexpression in organotypic cultures ...... 42

Figure 11: Increase in density of elongated, filopodial and branched spines on apical dendrites of actin-EGFP-transfected pyramidal neurons ...... 43

Figure 12: Increase in density of elongated and filopodial spines on basal dendrites of actin-EGFP-transfected pyramidal neurons...... 44

Figure 13: Elongated spines and filopodia are apposed by axon terminals ...... 46

Figure 14: Exogenous actin enters all spines ...... 47

ABSTRACT

Dendritic spines are small protrusions from dendrites on which the majority of excitatory synapses occur (Gray, 1959; Peters et al., 1976; Harris and Kater, 1994). Dendritic spines serve to compartmentalize second messenger systems (Koch, Zador, 1993) and are fundamental to synaptic transmission, long-term potentiation, and long-term depression ((Moser et al., 1994; Stewart et al., 1992). As such, spines are thought to be the locus of memory formation ((Moser et al., 1994; Stewart et al., 1992), and their pathology is seen in diseases characterized by a decrease in mental capacity such as Alzheimer’s Disease (Ferrer and Gullotta, 1990) and Fragile X Syndrome (Hinton et al., 1991; Wisniewski et al., 1991; Irwin et al., 2000; Nimchinsky et al., 2001). Thus, dendritic spines are critically important to neuronal function (reviews by (Harris, Kater, 1994; Shepherd, 1996). Spines have a highly dynamic actin cytoskeleton. In the past, this cytoskeleton has been regarded as merely a support structure. However, recent work from this and other laboratories identifies a much more functional, even regulatory role for actin within the spine. In the current series of experiments, we demonstrate the following aspects of spine formation and actin’s role in said process: (1) spines proliferate in acute adult hippocampal slices (2) protein synthesis is essential for slice spine proliferation, (3) actin is upregulated during spine formation in the juvenile acute slice, and is thus regulated as opposed to constitutively expressed, (4) actin overexpression leads to an increased spine density in the organotypic slice culture, largely the result of an increased number of elongated spines (5) all spines have a large degree of actin turnover in less than twenty-four hours.

CHAPTER 1

INTRODUCTION

Composition of Dendritic Spines

First identified and described by Santiago Ramon y Cajal in 1891, dendritic spines are specialized protrusions from dendrites that are now known to receive 90% of cortical excitatory connections (Gray 1959; Peters et al., 1976; Harris and Kater, 1994). The morphology of spines varies, but under normal conditions is mushroom-shaped (with a thin neck and bulbous head), short and stubby, branched, or long and thin (reviewed in Koch and Zador 1993). The morphology of each spine is derived from the differential organization of the actin filaments forming its cytoskeleton; actin filaments are organized in longitudinal bundles in the spine neck and as a meshwork in the head (Fifkova and Delay, 1982; Landis and Reese, 1983; Hirokawa, 1989). This actin organization is achieved by actin nucleating, bundling and crosslinking proteins that are enriched in spines. In addition to actin and actin-organizing proteins, spines can also contain other actin binding proteins (ABP), neurotransmitter receptors, ion channels, signaling molecules, a spine apparatus (SA; sacs of specialized smooth endoplasmic reticulum that may play a role in buffering calcium or making, storing, transporting, and/or modifying synaptic proteins; Fifkova and Delay, 1982), polyribosomes, multivesicular bodies and mitochondria (Reviews Harris and Kater, 1994 and Shepherd, 1996). Individual spines differ in their specific composition, sometimes by brain region, and sometimes on a single dendrite; not all spines have an SA, polyribosomes or mitochondria (Cameron et al., 1991; Spacek and Harris, 1997; Reviewed by Shepherd, 1996). Furthermore, the specific protein content of spines differs (Table 1). The heterogeneous distribution of proteins involved in actin organization and neurotransmitter receptor regulation could be a way to regulate spine motility and excitability in a synapse-specific manner.

Table 1. Heterogeneous protein distribution in dendritic spines

Protein Location & relative Function References amount α-Actinin 2 - high in cerebellum - crosslinks actin Wyszynski et al., - low cortex & hippocampus - links NMDAR to 1997, 1998; Smart - dentate & CA2 > CA1 actin cytoskeleton and Halpain, 2000 - within dendrite - actin-binding heterogeneity inhibited by Ca2+ Profilin - cortex & hippocampus: - helps add Carlsson et al., dendrite & subset of monomers to 1977; Kang et al., spines (head > neck) barbed end; 1999; Pollard et al., - cerebellum: presynaptic assists 2000; Neuhoff et terminal in actin filament al., 2005 growth PSD-95 - hippocampal primary - major component Takahashi et al., culture: in half of of the post- 2003 filopodia & most mature synaptic density spines SPAR (spine- - hippocampus: 70% of - Rap-specific Pak et al., 2001 associated excitatory synapses on GTPase activating RhoGAP) spiny neurons (determined protein (rap may by colocalization with have role in PSD-95, so could be in cytoskeletal even fewer spines) regulation) - binds PSD-95 Synaptopodin - hippocampus: CA2 > CA1 - actin binding Mundel et al., 1997; & CA3 - spine motility & Deller et al., 2000 - most likely in spines with elongation (review) spine apparatus & smooth - calcium release endoplasmic reticulum

2 Dendritic Spine Development

The initial formation of dendritic spines in early postnatal development begins with the extension and retraction of transient and motile filopodia, which are gradually replaced by more persistent yet highly dynamic protrusions, and finally, by persistent and stable mature dendritic spines (Daily and Smith, 1996; Ziv and Smith, 1996). During the latter part of development, stable spines are occasionally reabsorbed into the parent dendrite, and rapidly emerging spines and transient filopodia are sometimes observed (Daily and Smith, 1996). In the most general terms, the transition from filopodia to spine occurs when an appropriate presynaptic contact is made and sustained (Ziv and Smith, 1996). When contact between a filopodium and an axon occurs, a functional presynaptic bouton can develop on the axon (Ziv and Smith, 1996; Ahmari et al., 2000), and the long, thin filopodium can transform into a mature spine (Ziv and Smith, 1996). It has also been postulated that once a filopodium contacts an axon, it recruits the terminal to the parent dendrite, forming a shaft synapse, and a spine subsequently emerges (Fiala et al., 1998). While the latter theory is not in full agreement with the notion of filopodia developing into spines, it could explain the observation of mature spines emerging directly from dendrites. Regardless of the sequence of events, exactly how the connection between axon and filopodia leads to spine maturation is not fully understood. Prior to filopodial extension and spine development, actin filaments must form and gather the propulsive strength needed to push out the cell membrane. Much of our knowledge regarding this initial phase of spine development arises from studies in non- neuronal cells, in which filopodial outgrowth is examined and extrapolated, to a proper degree, to spine growth in neurons. The sequence of events has been summarized by Svitkina and colleagues (2003). A new actin oligomer is nucleated, or actin polymerizes onto an existing oligomer. This process is initiated by certain actin binding proteins such as the Arp 2/3 complex, which nucleates the growth of daughter branches that project at an angle of about 70 degrees from the mother branch. Actin fibers continue growing at the barbed ends until a capping protein attaches at the barbed end and blocks further polymerization. Some fibers, however, associate with tip complex proteins at the barbed ends and are therefore not capped and continue to grow. When the tip complexes of two

3 or more actin fibers collide, they bind to each other. This leads to a gradually enlarging tip complex and associated bundle of actin fibers. The resulting column of parallel actin fibers has the propulsive strength needed for filopodial extension. What other proteins are involved in spine development? Actin binding and bundling proteins such as drebrin, α-actinin 2 and PSD-95, and neurotransmitter receptors, such as α-amino-3-hydroxy-5-methylisoxazolepropionic acid (AMPA) receptors (which are seen in spine synapses prior to N-methyl-d-aspartate (NMDA) receptors), are among the first to arrive in developing spines (Rao et al., 1998; Matsuzaki et al., 2001; Takahashi et al., 2003). Furthermore, overexpression, constitutive expression and knockout studies have implicated some of the same proteins and numerous others in spine development (these are discussed in depth Chapter 4).

Function of Dendritic Spines

From synaptic transmission to global learning and memory, dendritic spines are fundamental to cognitive function (reviews by Harris and Kater, 1994, Shepherd, 1996, Matus, 2000 and Fiala et al., 2002). Induction of tetanus and long-term potentiation (LTP) lead to an increase in spine size and density, and LTD induction leads to spine shrinkage that is reversed by high frequency stimulation (Desmond and Levy, 1988; Hosokawa et al., 1995; Engert and Bonhoeffer, 1999; Maletic-Sevatic, et al., 1999; Toni et al., 1999; Matsuzaki et al., 2004; Zhou et al., 2004; reviews by Fifkova and Morales, 1992 and Smart and Halpain, 2000). Various learning paradigms are associated with increases in spine density (Brandon and Coss, 1982; Mahajan and Desiraju, 1988; Lowndes and Stewart, 1994; Moser et al., 1994; Stewart and Rusako, 1995; O’Malley et al., 1998) and rearing in or exposure to an enriched environment leads to increased spine density (Turner and Greenough, 1985; Comery et al., 1996; Johansson and Belichenko, 2002). In contrast, numerous diseases and injuries are characterized in part by decreased mental capacity, and involve the malformation of spines, either in shape, density or molecular composition. These include Creutzfeldt-Jacob disease, Rett syndrome, epilepsy, Fragile X syndrome, Patau’s syndrome (trisomy 13), Down’s syndrome, schizophrenia, Huntington’s disease, HIV-related dementia, hypoxia, hypoglycemia,

4 ischemia, Alzheimer’s disease and traumatic brain injury (Ferrer and Gullotta, 1990; Reviews by Kaufmann and Moser, 2000; Smart and Halpain, 2000 and Fiala et al., 2002). The predominating theory as to why spines are so critically important is that each spine provides a synapse-specific compartmentalization of calcium entry, biochemical reactions and second messenger cascades (Gamble and Koch, 1987; Holmes, 1990; Reviews by Koch and Zador, 1993; Shepherd, 1996, Segal and Andersen, 2000, Hering and Sheng, 2001, Sabatini et al., 2001 and Blanpied and Ehlers, 2004). This is achieved because the thin neck of the spine results in a separation of the tiny volume of the spine head and the immense volume of the parent dendrite. Studies using fluorescence recovery after photobleaching (FRAP) indicate that diffusion from dendrites into spines is 100 times slower than would be expected for free diffusion (Svoboda et al., 1996; Majewska et al., 2000). One consequence of this compartmentalization is that spines can act autonomously, strengthening or weakening overall neural connectivity one synapse at a time (reviewed by Shepherd, 1996). A second result of biochemical compartmentalization is that relatively few ions or molecules are needed to enact dramatic changes within the spine. This is especially important in the case of calcium, the concentration of which regulates the state of actin polymerization as well as enzymes and transcription factors that are necessary for the induction and maintenance of LTP and LTD (see further discussion of spine calcium below; Reviewed by Sabatini et al., 2001). In addition to biochemical compartmentalization, the spine may also provide some degree of electrical compartmentalization. The spine neck attenuates current as it leaves the spine and enters the parent dendrite (although some reports suggest that the amount of attenuation is negligible; Reviews by Shepherd, 1996 and Segal, 2005). Conversely, there is no decrement of current from the dendrite into the spine. Spines, therefore, are very sensitive to both their own synaptic input as well as the integrated potential of the parent dendrite. The importance of spine geometry in electrical compartmentalization has resulted in an attempt to understand the specific relationship between spine morphology and synaptic currents (reviewed by Segal, 2005). The size of mEPSCs recorded at the soma has been positively correlated with spine size, and it was proposed that the increase in synaptic potential was a result of more glutamate receptors

5 in the larger spines. However, changes in the size, density or shape of spines brought on by transfection of various proteins have led to a range effects on mEPSPs that are not directly or obviously correlated with the accompanying morphological change. Interestingly, comparisons of spines from different brain areas suggests regional differences in the type of compartmentalization provided by spines; CA1 pyramidal cell spines act as biochemical compartments, while Purkinge cell spines act as both biochemical and electrical compartments (Reviewed by Sabatini et al., 2001).

Dendritic Spine Calcium Dynamics

Calcium concentration, [Ca2+], within dendritic spines can reach as high as 20-40 µM (Petrozzino et al., 1995), and the various sources of this calcium have been examined. Spines contain 1-20 voltage-sensitive calcium channels (VSCC), which open with about 50% probability in response to back-propagating action potentials (Yuste and Denk, 1995; Sabatini and Svoboda, 2000; reviewed by Sabatini et al., 2001). Calcium also enters spines through NMDA receptors and, to a lesser degree, through AMPA receptors (reviewed by Sabatini et al., 2001). Finally, calcium can be released from internal stores, in particular smooth endoplasmic reticulum (SER). Not all spines in hippocampal pyramidal cells contain SER, however, and the role of internal calcium release is not fully understood. Calcium transients have been reported, however, that are NMDA receptor-dependent and almost completely produced by calcium-induced calcium release from internal stores (Emptage et al., 1999). The regulation of calcium once it has entered the spine is affected by a variety of 2+ 2+ factors. Internal [Ca ] ([Ca ]i) is impacted by endogenous calcium buffers such as calmodulin, calcium extrusion mechanisms such as SER calcium ATPase (in pyramidal cells) and transport or diffusion of calcium between the spine and dendrite (Reviewed by Sabatini et al., 2001). Transport and diffusion are dependent on three main factors: spine neck length, dendritic diameter, and the strength of spine calcium pumps (Majewska et al., 2000; Holthoff et al., 2002). Dendritic diameter and strength of calcium pumps change with respect to position on the dendritic tree. Spines, therefore, have different calcium kinetics based on their position on the cell; spines in basal and distal apical

6 dendrites have faster decay and undershoot than spines on proximal apical dendrites (Holthoff et al., 2002). Functionally, slower calcium decay makes spines more susceptible to LTD. This differential handling of calcium could also have an affect on calcium-induced morphological plasticity, as actin dynamics are readily influenced by 2+ [Ca ]i (discussed below), although such an affect has not been measured. 2+ Transient increases in [Ca ]i are thought to be important for LTP; titanic stimuli 2+ lead to [Ca ]i in spines up to 10 µM, while stimuli that generate LTD cause smaller and longer calcium accumulations (reviews by Sabatini et al., 2001 and Segal 2005). An especially interesting aspect of spine calcium and its potential role in LTP is the phenomenon of supralinear summation (reviewed by Sabatini et al., 2001). Supralinear summation describes an event wherein synaptic activity (EPSPs) and a back-propagating 2+ action potential result in [Ca ]i that is greater than the sum of what would be produced by each stimulation alone. The supralinear increase in calcium occurs via the unblocking of NMDA receptors by the action potential, and is greater when the action potential follows synaptic input (Yuste and Denk, 1995). Because of NMDA receptors and the property of supralinear summation, spines function as the substrate for coincidence detection, with millisecond shifts in the input and output of the cell translating into a 2+ specific [Ca ]i (Yuste et al., 1999). 2+ Spine [Ca ]i is critically important for actin dynamics. AMPA receptor blockade rapidly and reversibly blocks spine motility (Fischer et al., 2000; Brunig et al., 2004) in a manner that is dependent upon Ca2+ entry through VSCCs (Fischer et al., 2000). NMDA receptor activation leads to a delayed and irreversible suppression of actin dynamics (Ackermann and Matus, 2003; Brunig et al., 2004), and this is dependent upon calcium entry through the receptor-associated channel. The rapidity of the AMPA receptor affect is likely due to an immediate affect on the spine cytoskeleton, while the delay in the NMDA receptor affect is thought result from the need to build up depolarization sufficient to remove the magnesium block. Action potential trains lead to calcium transients in spine heads and fast contraction of spines, the latter blocked by Latrunculin B (Korkotian and Segal, 2001). Caffeine-induced calcium release from internal stores leads to the growth of new spines and the enlargement of established spines (Korkotian and Segal., 1999), but has no affect on spine motility (Brunig et al., 2004). Finally, the

7 blockade of calcium reuptake into internal stores, resulting in influx of external calcium, leads to a delayed blockade of spine motility (Brunig et al., 2004). Taken together, these 2+ data suggest that calcium influx and the resultant spine [Ca ]i, but not factors specific to the mode of Ca2+ entry, arrest actin dynamics, and thus spine motility (Brunig et al., 2004).

Local Protein Synthesis

Protein synthetic machinery including ribosomes, polyribosomes, nissel bodies, endoplasmic reticulum, golgi apparatus and spine apparatus exist in dendrites and/or spines, and function in protein translation, translocation, folding and glycosylation (Torre and Steward, 1996; reviews by Job and Eberwine, 2001 and Glazner and Eberwine, 2003). The mRNAs for many proteins, including but not limited to MAP2, CaMKIIα, arc, calmodulin 1, NMDAR1 and β-actin have also been identified within dendrites. The high levels of these mRNAs within dendrites suggest that active transport and not diffusion are responsible for their presence. Indeed, sequences that are necessary for dendritic targeting have been identified in some mRNAs. For example, the 3’ UTR of both β-actin and CaMKIIα contain a sequence that is necessary for localization to dendrites (Mayford et al., 1996; Tiruchinapalli et al., 2003). Dendritic localization of mRNA containing the appropriate targeting sequence is then achieved by RNA binding proteins and the microtubule system (reviewed by Glanzer and Eberwine, 2003). The level of mRNA for some proteins is much higher in development than in adulthood, likely reflecting an increased need for local protein synthesis during periods of increased synaptogenesis. Localization and translation of dendritic mRNA is activity-dependent; localization can be altered by electrical or NMDA-induced stimulation, and translation can be stimulated by neurotrophins (Kang and Schuman, 1996; Steward et al., 1998). Local protein synthesis is especially germane to the study of dendritic spines because it is likely involved in spine development. This is exemplified well by the disruption of the fragile X mental retardation protein (FMRP). FMRP is an RNA binding protein that is believed to be involved in the localization and/or translation of proteins important for spine development (Reviewed by Glazner and Eberwine, 2003). In Fragile

8 X syndrome, the most common form of inherited mental retardation (Turner et al., 1996), there is a significant knockdown of FMRP, as well as significant alterations in dendritic spine density and shape (Irwin et al., 2000, 2001). Local protein synthesis is also important for dendritic spines because it allows rapid synthesis and specific localization of post-synaptic components upon a brief stimulation, a proposed mechanism of synapse-specific plasticity. Pierce and colleagues (2000) demonstrated that active protein translation sites, ribosomes, and lumenal ER proteins are present in the endomembrane systems of dendrites and spines in the hippocampus. The authors interpret this to support a model wherein proteins are synthesized in dendrites and spines in a regulated fashion, and immediately gain entry to the secratory pathway. Therefore, proper stimulation could induce local synthesis of integral membrane receptors in a synapse-specific fashion, and these would be immediately put in place, thus contributing to long-term synaptic plasticity.

Plasticity of Dendritic Spines: An Essential Role for Actin

Actin exists in a monomeric (G-actin) and filamentous (F-actin) form, the latter of which forms a double helix. Each G-actin monomer is bound to either ATP or ADP, and G-actin bound to ATP polymerizes more quickly than that bound to ADP (Cooper, 1988). At the barbed end of a growing actin filament, G-actin-ATP is added, while at the

pointed end ATP is dephosphorylated to ADP+Pi and the actin monomer is released, usually to be bound again by free ATP. This constant polymerization at one end and depolymerization at the other is known as treadmilling. Growth can actually occur at either end, but occurs faster at the barbed end due to a lower critical concentration for actin monomer addition as compared to the pointed end (reviewed by Cooper and Schafer, 2000). F-actin forms the cytoskeleton of each dendritic spine. Dendritic spines are known for their high rate of elongation, retraction, proliferation, head morphing and membrane ruffling. These morphological changes are possible because of the highly dynamic nature of actin filaments. Drugs that interfere with spine actin polymerization, either by depolymerizing or stabilizing actin filaments, arrest spine motility (Fischer et

9 al., 1998; Dunaevsky et al., 1999; Korkotian and Segal, 2001), block the late phase of LTP (Fukazawa et al., 2003), block the reversal of LTD (de-depression; Chen et al., 2004), and disrupt AMPA receptor mediated synaptic transmission (Kim and Lisman, 1999), illustrating the importance of actin not only in spine development, but also in morphological and functional plasticity. Because actin filaments are critical to the structural stability of the spine, as well as the ability of the spine to undergo morphological alterations, two populations of spine actin fibers were hypothesized to exist within single spines (Halpain, 2000). In this model, there is a large stable group of actin filaments in the core of the spine as well as a smaller, more dynamic group around the edge. The relative amounts and locations of the two populations were determined partly in order to explain the ability of the edges of spines to ruffle and wiggle, as described by Dunaevsky et al. (1999), while the spine itself remains intact. This prediction was based on multiple studies of spine stability and plasticity, and since it was made, studies using FRAP have presented direct evidence of multiple actin filament populations. Fluorescence recovery time for actin momoners is shorter than for treadmilling filaments, and there seems to be a third pool of actin with exceptionally long recovery time, suggesting that it is very stable (Star et al., 2002; Zito et al., 2004). In the FRAP studies, 85% of spine actin was shown to turn over in 44 seconds (Star et al., 2002). Why would such a large percent of fibers be dynamic? As previously described, edges of spines ruffle, perhaps in order to challenge weak connections (the functions of spine motility are not entirely clear). Additionally, it was recently reported that spines can adjust diffusion between the head and the parent dendrite by very subtle motility in the spine membrane, regulated by AMPA receptor activation (Richards et al., 2004). As highly regulated diffusion between the spine head and the parent dendrite is a critical element in spine function, the devotion of multiple actin fibers to this task while maintaining the spine core would not be surprising. Importantly, studies utilizing FRAP on spine actin necessarily visualize only those spines that incorporate exogenously introduced enhanced green fluorescent protein (EGFP)-tagged actin. Therefore, a population of spines with exceptionally low actin turnover could be unaccounted for.

10 In this dissertation, two main questions were addressed. Neurons have the ability to rapidly produce dendritic spines. Many proteins are involved in spine growth, but the immediate origin of these proteins was not known. Did they already exist within the cell, or did they have to be generated upon an activating stimulus? The first question we asked was: Is protein synthesis necessary for dendritic spine development? Once we established that protein synthesis is indeed required, we shifted our focus to the specific protein(s) needed for spine growth. Concentrating on β-actin, we then asked: Do actin levels increase in parallel with spine proliferation, and can β-actin expression itself regulate spine development?

11 CHAPTER 2

PROTEIN SYNTHESIS IS NECESSARY FOR DENDRITIC SPINE

PROLIFERATION IN ADULT BRAIN SLICES

Introduction

Spines and filopodia can change shape, disappear, and grow de novo both spontaneously (Daily and Smith, 1996; Fischer et al., 1998; Dunaevsky et al., 1999; Lendvai et al., 2000) and after various manipulations including depression (Norrholm and Ouimet, 2001), estrogen administration (Wooley and McEwen, 1992; Pozzo-Miller et al., 1999; Calizo and Flanagan-Cato, 2000), morphine administration (Robinson and Kolb, 1999), learning paradigms and environmental enrichment (discussed and sited in Chapter 1), and brain slice preparation (Kirov et al., 1999). Such changes in spines occur on a time-scale of minutes to hours (Dailey and Smith, 1996; Kirov et al., 1999; Lendvai et al., 2000). Given the degree of plasticity inherent to dendritic spines, the variety of stimuli that elicit spine growth, and the rapidity with which dendritic spines are formed, recent research has focused on the mechanism of spine growth (reviews by Matus, 2000 and Segal, 2001). While this research has yielded data regarding proteins that are involved in spine growth and maturation, it is not known whether protein synthesis is necessary for new spine growth, or if existing proteins are recruited for the generation of new spines. Recruitment of pre-existing proteins has been documented in the formation of presynaptic active zones (Ahmari et al., 2000) and developing synapses (Prang and Murphy, 2001). Since spines can grow rapidly, it is possible that prefabricated units of proteins necessary for spine growth are activated by appropriate stimuli, and spines are made without requiring new protein synthesis. Other data, however, suggest that protein synthesis could be necessary for spine growth. Synapse formation (even in the absence of cell bodies; Schacher and Wu, 2002), BDNF-induced growth cone stabilization after nitric oxide (Gallo et al., 2002), and glutamate receptor-induced dendritic spine

12 elongation (Vanderklish and Edelman, 2002) are dependent upon protein synthesis. The rapid and efficient signaling capabilities between nucleus and dendrite, as well as the abundance of mRNAs and protein synthetic machinery present in dendrites, make the fast generation and localization of newly synthesized proteins possible (Gao, 1998; Job and Eberwine, 2001; Steward and Schuman, 2001). In the present study, we sought to determine whether dendritic spine formation in adult animals requires protein synthesis. To answer this question, we used an acute slice preparation in which marked spine proliferation is known to occur in neonatal and preadolescent animals (Kirov et al., 1999). Our data demonstrate that dendritic spines also proliferate in adult brain slices and that this proliferation requires protein synthesis.

Methods

Slice Preparation. Hippocampal slices were prepared from a total of 68 4-month old male Sprague- Dawley rats (Charles River; 51 in Experiment 1 and 17 in Experiment 2). Rats were group-housed in a temperature-controlled room with constant access to food and water. Each rat was anesthetized with a single dose of pentobarbital sodium (60 mg/Kg), decapitated, and the brain removed and cut into 400 µm sections (in the coronal plane) using a McIllwaine tissue chopper. The sections were then incubated in 150 ml artificial cerebrospinal fluid (aCSF) containing 117 mM NaCl, 5.3 mM KCl, 26 mM NaHCO3, 1 mM NaH2PO4, 2.5 mM CaCl2, 1.3 mM MgSO4, and 10 mM glucose, equilibrated with

95% O2-5% CO2, pH 7.4, at 32°C. In experiment 1, to determine the time period during which new spines were generated, slices were removed from aCSF after 5, 10, 30, 60, 90 and 120 minutes and immersion-fixed with microwave enhancement (8 seconds, 700 watts, after Kirov et al., 1999) in 8 ml of 4% formaldehyde containing 0.03% glutaraldehyde. The tissue was further fixed in the same fixative for 20 minutes at room temperature. Statistical analysis revealed that there were significantly more dendritic spines at 60 MIV as compared to perfused, and this difference persisted through 120 MIV (see Results section for details). It was therefore concluded that new spine development was observable by 60 MIV.

13 In experiment 2, designed to determine the role of protein synthesis in spine proliferation, protein synthesis inhibitors were used at concentrations known to block synthesis (90% blockade of tritiated amino acid incorporation in 20 minutes) in similar systems: cycloheximide (CLX; 400 mM final concentration; Feig and Lipton, 1993), or puromycin (PUR; 25 mM final concentration; Jones et al., 1992). Both drugs were dissolved in dH2O. To block protein synthesis during spine development (not before or after, when protein synthesis inhibition might affect only existing spines), some slices were incubated in aCSF for 40 minutes, at which time they were transferred to aCSF containing PUR or CLX for an additional 20 minutes. Immediately following this treatment (i.e., at 60 MIV), slices were immersion fixed as described above. Slices were transferred quickly and gently in a small amount of aCSF without exposure to air. In addition, a control group of slices (n=5 rats) was transferred at 40 MIV to aCSF containing no drug to make sure that changes in spine density were not related to the transfer itself. There was no difference in spine density between the 60 MIV group in the timecourse experiment and the 60 MIV transfer group (Figure 1). These two groups were therefore combined (total n=12). To determine the effect of protein synthesis blockade prior to new spine growth, another group of slices was treated immediately with PUR from 5-25 MIV (this early PUR group was called ePUR) and immersion fixed without further incubation. The purpose of this experiment was to determine whether acute PUR treatment caused dendritic spine loss independent of its effect on blocking spine growth. Following fixation, slices were transferred to 0.01 M phosphate buffered saline and sectioned at 100 µm using a vibratome. Sections from the middle 200 µm (previously shown to be the healthiest part of the tissue; Kirov et al., 1999) were prepared for Golgi staining as described by Izzo et al. (1987). In brief, sections were incubated in

1% aqueous osmium tetroxide (OsO4) for 30 min. OsO4 was washed out in three 15- minute 0.1 M phosphate buffer rinses, and the slices were then incubated in aqueous 3.5% potassium dichromate for three hrs. Next, sections were placed between two microscope slides in a "sandwich" configuration that was then submerged in aqueous 1% silver nitrate for 48 hrs. Sections were subsequently dehydrated in a graded ethanol series and mounted in DPX.

) 2.5

1.5

0.5

Spine Density (spines/micron 0 60 MIV 60 MIV-transferred

Figure 1. Transfer of slices does not affect dendritic spine density. Graph shows mean spine density in 60 MIV group from time-course experiment and in control transfer group. The transfer of slices at 40 MIV to a second well of aCSF had no effect on spine density (p=0.671). The 60 MIV transfer group (N=5) was therefore included in the 60 MIV group (total N=12).

15 Perfusions. Baseline dendritic spine density was determined in tissue from perfused animals. Rats were anesthetized with pentobarbital sodium (60 mg/kg) and transcardially perfused with a mixture of 4% formaldehyde (freshly prepared from paraformaldehyde) and 0.03% glutaraldehyde. The brains were post-fixed overnight at 4°C and then sectioned at 100 µm using a vibratome. Microscopic Analysis. Dendritic spines on the secondary branches of CA1 apical dendrites in stratum radiatum were counted using a light microscope with a camera lucida attachment. Analyzing these dendrites as opposed to the thicker primary dendrites reduces the possibility that spine length would affect detectability using the Golgi technique. All analysis was done blind with regard to experimental condition. For each rat, an average of 5 neurons was analyzed and spines were counted over an average dendritic length of 713 µm (Table 2). The average total dendritic length analyzed for each group was 5520 µm. No differentiation was made among spine types during analysis. Stubby spines, mushroom-shaped spines with thin necks and bulbous heads, branched spines, and thin spines without heads (possibly filopodia) were all counted. Statistical analysis. One-way ANOVAs, with least significant difference (LSD) post hoc analysis when necessary, were performed (a) between the perfused and six time point groups to determine time-course of proliferation, (b) between perfused, 60, CLX and PUR groups to determine the effect of protein synthesis inhibition, (c) between 30 MIV and ePUR groups to determine if the drug caused loss of existing spines, and (d) between the PUR and ePUR groups. Length Measurements. Because it appeared that a subpopulation of spines was longer after 60 MIV, spine length was measured in 60 and 120 MIV slices, in perfused tissue and in PUR- and CLX- treated slices. Image stacks of Golgi-stained secondary dendritic twigs were made using a Zeiss 510 laser confocal microscope. These stacks consisted of 3-36 optical slices taken in 0.5 µm intervals in the z-plane. A total of 50 twigs (each approximately 50 µm long) were examined, on which a total of 3050 spines (392-749 spines per group) were

16 measured (details of sample size and data are shown in Table 2). A one-way ANOVA with LSD post hoc analysis was performed among the five groups described above. Because a positive correlation has been found between spine density and dendritic diameter (Bannister and Larkman, 1995), and in order to rule out the possibility that a difference in spine length could be attributed to dendritic width (in other words, what appeared to be shorter spines were actually spines whose necks or bases were obscured by the girth of the parent dendrite itself), the widths of the dendrites on which spines were counted were also measured. For each twig, an average of 14 (SES=0.5958) width measurements were taken along its length (Table 3). A mean twig width was calculated from these measurements, and a one-way ANOVA among the 5 groups was performed.

Results

Dendritic spine density increases during the first hour of the acute slice preparation It was previously reported that the density of dendritic spines in slice preparations increases within two hours (Kirov et al., 1999). To determine the time course for spine proliferation, slices were fixed after 5, 10, 30, 60, 90 or 120 MIV (Figure 2). At 60, 90 and 120 MIV, there were approximately 12% more spines than in perfused tissue. One- Way ANOVA among the 6 time-point groups and the control perfused group showed a significant difference in spine density (p=0.028). LSD post hoc analysis indicated that at 90 and 120 MIV, spine density was significantly higher than in the perfused tissue or in the slice preparation tissue at 10 MIV. Additionally, spine density at 120 MIV was significantly higher than at 5 MIV. A planned comparison between perfused and 60 MIV revealed a significant difference at this time point as well (p=0.038). Based on these data, we determined that the bulk of the increase in spine density begins later than 30 MIV and is significant at 60 MIV. Therefore, CLX and PUR were administered beginning at 40 MIV and ending with fixation at 60 MIV. Protein synthesis inhibitors block spine proliferation Twenty minute incubation of slices in CLX or PUR, starting at 40 MIV, completely blocked slice-induced spine proliferation (p=0.004; Figure 3). There was no difference in spine density between CLX- and PUR-treated groups (p=0.725).

Table 2. Degree of analysis for counting spines.

______Group Animals (n) Number of Dendritic length cells analyzed (µm) analyzed ______Mean per rat Group Mean per rat Group + SES total + SES total ______Perfused 7 4.143 + 0.143 29 921 + 125 6450 5 MIV 8 6.375 + 1.051 51 1047 + 129 8375 10 MIV 8 5 + 1.018 40 689 + 93 5515 30 MIV 8 5.75 + 0.75 46 897 + 234 7173 60 MIV 12 4.917 + 0.743 59 541 + 85 6490 90 MIV 8 6 + 1.165 48 728 + 166 5825 120 MIV 7 4 + 0.218 28 693 + 145 4850 CLX 7 4.125 + 0.978 33 815 + 248 6520 PUR 5 4.2 + 0.583 21 406 + 33 2030 ePUR 5 4.4 + 1.249 22 394 + 117 1970___

Table 3. Degree of analysis for measuring spine length (µm) and dendrite width (µm).

______Group No. of rats No. of twigs Total spines Mean spine Width measurements Mean dendrite sampled sampled measured length + SES per twig + SES width + SES ______Perfused 2 7 392 0.776 + 0.013 12.333 + 0.333 0.977 + 0.036 60 MIV 2 10 617 0.922 + 0.015 12.8 + 1.2 0.919 + 0.078 120 MIV 4 11 721 0.920 + 0.013 16.143 + 0.857 0.849 + 0.038 CLX 3 9 571 0.853 + 0.012 16.5 + 1.258 0.826 + 0.027 PUR 4 13 749 0.775 + 0.010 14.2 + 1.241 0.860 + 0.018

2.5 abc ab ad

2.25

1.75

Spine Density (spines/micron)

1.5 perfused 5 10 30 60 90 120 in vitro Minutes

Figure 2. Spine density increases on secondary apical dendrites of CA1 as a function of time (minutes in vitro). A one-way ANOVA was performed on all groups, yielding p=0.028, with LSD post hoc test yielding a significant difference (p<0.05) as indicated by "a" (greater than perfused), "b" (greater than 10 MIV) and "c" (greater than 5 MIV). While perfused and 60 MIV were not significantly different in this test due to the necessary adjustment of alpha, a planned comparison between these groups yielded p=0.038, as indicated by "d". N=7, 8, 8, 8, 12, 8, 7, respectively.

2.5 * 2.25

1.75

Spine Density(spines/micron) 1.5

1.25

perfused 60 MIV CLX-60MIV PUR-60MIV

Figure 3. Protein synthesis inhibitors block spine proliferation. Drugs were applied 20 min. prior to the 60 MIV time point at which spine proliferation is first detectable (beginning at 40 MIV and ending with fixation at 60 MIV). Thus, CLX- and PUR-treated slices are compared to 60 MIV treated slices. A one-way ANOVA on these four groups yields p<0.001. Spine density is significantly higher at 60 MIV (indicated by asterisk) as compared to perfused (p=0.013), CLX and PUR (p=0.002). There is no significant difference between perfused and CLX- or PUR-treated tissue (p=0.142, p=0.101, respectively) or between the two inhibitors (p=0.755). N=7, 12, 7, 5, respectively.

21 Representative examples of dendritic segments of untreated slices at the different time- points and of CLX- and PUR-treated slices are shown in Figure 4. Short-term protein synthesis blockade did not reduce spine number to a lower value than that observed in perfused tissue, suggesting that the doses used were not simply toxic to the cells, causing general spine loss. In addition, RNA synthesis rate has been reported to be normal in primary neuronal culture after treatment with 20 µg/ml CLX and 200 µg/ml PUR, suggesting that these cells are still metabolically active (Kleiman et al., 1993). Early protein synthesis inhibition does not prune spines To determine whether short-term protein synthesis blockade caused dendritic spine loss independent of its effect on blocking spine growth, a group of slices was treated with 25 µM PUR as soon as the slices were prepared (i.e., for 5-25 MIV). These slices were fixed at the end of their incubation and spine density was compared to the 30 MIV group. No significant change in spine number was observed (p=0.171; Figure 5). Additionally, there was no significant difference in spine density between slices treated with PUR from 40-60 MIV and the early PUR-treated group (p=0.238). Spine proliferation is accompanied by an increase in mean spine length Because it appeared that some spines were longer after 60, 90 and 120 MIV, spine length was measured in perfused tissue, in 60 and 120 MIV slices and in PUR- and CLX- treated slices (Figure 6). While the average spine length of perfused and PUR-treated tissue was 0.77 + 0.01 µm, the average spine length of 60- and 120-MIV slices was 0.92 + 0.014 µm (spine length of CLX treated tissue fell between these values at 0.85 + 0.012 µm). A one-way ANOVA among the 5 groups examined yielded p<0.001. LSD post- hoc analysis confirmed that the spines in 60 and 120 MIV slices were significantly longer compared to perfused tissue (p<0.001). Thus, not only are there more spines at 60 and 120 MIV, but the new and/or existing spines are longer. In addition, PUR treatment blocked the increase in spine length. As shown in Figure 6c, approximately 12% of spines at 60 and 120 MIV are longer than spines in perfused tissue. Because there was also a 12% increase in spine density, it is possible that the new spines are longer than previously existing spines.

Figure 4. Representative dendritic twigs from each experimental group. Note the increase in spine density and spine length in 60, 90 and 120 MIV slices. Closed arrows indicate headless spines (filopodia). Open arrows indicate elongated spines with clearly delineated heads.

) 2.5

1.5

0.5

Spine Density (spines/micron 0 30 MIV early PUR

Figure 5. Early treatment with PUR does not prune spines. There was no significant difference in spine density between slices treated with PUR from 5 to 25 MIV and time- matched, non-treated tissue (30 MIV; p=0.171 by one-way ANOVA), indicating that acute protein synthesis inhibition does not disassemble existing spines. N=8, 6, respectively.

1 * * 0.9 0.8 0.7 0.6

0.5 0.4

0.3

0.2

spine length (microns) Mean 0.1 0 perfused 60 MIV 120 MIV

1 * 0.9 # 0.8 0.7 0.6 0.5

0.4

0.3 0.2

Mean spine length (microns) 0.1 0 perfused 60 MIV CLX-60MIV PUR-60MIV

Figure 6. Mean spine length increases as a function of time, an effect that is blocked by protein synthesis inhibition. (A) Mean spine length for perfused tissue and slices after spine density has increased. A one-way ANOVA was performed on perfused, 60 MIV and 120 MIV groups, yielding p<0.001. An LSD post hoc analysis confirmed that both 60 and 120 MIV slices have greater mean spine length as compared to perfused (p<0.001), as indicated by the asterisks. N=392, 617, 721, respectively. (B) Mean spine length for perfused tissue, 60 MIV slices (after spine density has increased) and CLX- and PUR-treated tissue. A one-way ANOVA on perfused, 60 MIV, CLX- and PUR- treated tissue yielded p<0.001. LSD post hoc analysis revealed that spines were longer in 60 MIV slices than in perfused or drug-treated tissue (p<0.001). In addition, spines in CLX-treated slices were significantly longer than those in perfused and PUR-treated slices, yet shorter than those in 60 MIV tissue (p<0.001), as indicated by the pound sign. N=392, 617, 749, 571, respectively.

18 Perfused 16 60 MIV 120 MIV 14 CLX 12 PUR 10

(percent) Spines 6 4 2

0 9 9 9 9 59 39 .79 39 -.19 -.3 -. 1.5 .1 .3 .5 .7-.79 .9-.99 -1. . -1 -2. .3 .5 .3 1.1-1.1 1 1 1.7 1.9-1.992.1-2.1 2 Spine length (microns)

Figure 6. (C) Distribution of spine lengths. 12% of spines in the 60 and 120 MIV groups are equal to or greater than 1.4 µm long (arrow); almost no spines in the other three groups reach or exceed that length.

26 To rule out the possibility that the measured dendritic spine length was associated with the width of the dendrites analyzed, dendritic width was also measured on the sample dendritic twigs. A one-way ANOVA yielded p=0.188 indicating no significant difference in mean dendritic width between the perfused, 60 and 120 MIV groups (data not shown).

Discussion

Slices from neonatal and periadolescent rat brains have been shown to contain significantly more spines than age-matched perfused brains (Kirov et al., 1999). This increase in spine density, observed by electron microscopy, was noted in area CA1 after a period of at least 1.75 hours. In the present study of older animals, we analyzed the time course of spine proliferation in the slice preparation and report that spine density begins to increase by 60 MIV, and that this increase in spine density is dependent on protein synthesis. Treatment of slices with the protein synthesis inhibitors CLX or PUR during the time when spines proliferate blocked new spine growth; spine density in CLX-and PUR-treated slices did not differ from that found in perfused tissue. Early incubation with PUR prior to spine proliferation had no net effect on spine density. Methodological considerations Golgi staining of perfused tissue yielded the same spine density as reported previously for perfused tissue analyzed by electron microscopy (~2 spines/micron; Kirov et al., 1999). We observed, however, only a 12% increase in spine density during the acute slice preparation. The Kirov study reported a decline in the degree of spine proliferation from 90% in weanling rats (21 days old) to 40% in periadolescent/young adult rats (55-68 days old) and it is likely that this decline continues throughout adulthood. In addition to age, there were differences in the preparation and method of analysis. First, Long-Evans rats were used in the Kirov study; Sprague-Dawley rats were used in the present study. Second, slices were maintained briefly in ice-cold aCSF in the Kirov study; in the present study, brains were blocked in ice-cold aCSF, and then chopped and immediately transferred to 32°C. Third, the plane of section in the Kirov study was 70° to the long axis; our brains were cut coronally. Fourth, in the Kirov study

27 slices were maintained in an interface chamber, while ours were maintained submerged in an aCSF bath. In addition, electron microscopy would resolve spines, especially small spines, with greater efficiency than would the Golgi technique. Finally our counting was done exclusively on secondary dendrites while the Kirov study counted a sample of all spines in the neuropil. Significance of spine length In addition to an increase in spine density, an increase in spine length was identified at 60 MIV and persisted to 120 MIV. This increase in spine length was completely abolished by PUR treatment, and only partially blocked by CLX treatment. Given a greater n, an effect closer to that observed after PUR treatment may have been observed for CLX treatment. On the other hand, the data could reflect a sufficient n, and demonstrate a differential effect of the two protein synthesis inhibitors. There was a 12% increase in spine density at 60 and 120 MIV and 12% of spines in the 60 and 120 MIV groups were longer than 1.4 µm. Very few spines in the PUR or perfused groups that were that long. Some of the longer spines had clearly delineated heads whereas others resembled headless spines (possibly filopodia). Taken together, these data strongly suggest that newly developed spines are longer than previously existing spines in the slice preparation. These elongated spines/filopodia may represent a transitional state between immature and mature spines. Kirov et al. (1999) reported that slices contain more stubby and mushroom-shape spines than perfused tissue. Because they looked primarily at spines in older acute slices (up to 10 hours in vitro), it is possible that long, thin filopodia had developed their mature spine phenotype. Role of Filopodia in Spine Development The function of filopodia in synapse formation is unclear; some filopodia may arise under synaptic contacts on shafts (Harris et al., 1989; Harris et al., 1992; Westrum et al., 1980) and others may extend from dendritic shafts in search of axon terminals (Ziv and Smith, 1996). Such filopodia could guide the terminal to the parent dendrite, creating a shaft synapse where a spine may develop in the future (Fiala et al., 1998). The progression from long, filopodial protrusions to mature spines has been observed during development. Time-lapse photography of young DiO-stained hippocampal neurons in primary culture revealed (a) dynamic, short-lived filopodia extending from dendrites, (b)

28 enough filopodia to account for the number of spines found on a mature dendrite, (c) a loss of filopodia coinciding with the appearance of more stable spines (also reported in neonatal brain slices by Dailey and Smith, 1996), and (d) the formation of functional synapses on newly-stabilized spines (Ziv and Smith, 1996). Possible Involvement of Local Protein Synthesis It is not known whether gene activation, translation of pre-existing mRNA, or both are necessary to create the new proteins required for spine development. Multiple mRNAs have been identified within dendrites; protein synthesis occurring on dendritic polyribosomes located at the base of spines is hypothesized to confer synapse specificity (Gao, 1998; Job and Eberwine, 2001; Steward and Schuman, 2001). Because the growth of spines in the slice preparation is rapid, it is possible that protein synthesis is occurring in association with existing dendritic mRNA. In culture of the Aplysia presynaptic sensory neuron and postsynaptic L7, synaptic connections are formed even after the divorce of either or both neurites from their cell bodies, as long as protein synthesis is not blocked (Schacher and Wu, 2002). Protein synthesis itself seems to play a role in mRNA localization. For example, protein synthesis is required for transport of microtubule- associated protein 2 RNA to the dendrites (Kleiman et al., 1993). Another function of dendritic protein synthesis (i.e., protein synthesis occurring at the base of or within spines) could be in the maintenance of mature dendritic spines. In the present study, it was essential to demonstrate that acute protein synthesis blockade did not lead to a loss of spines. Protein synthesis, however, is likely critical in the long-term maintenance of spines and synapses. Protein synthesis within dendrites is involved in activity-dependent synaptic modification. One example of this is the role of the protein Arc in LTP and memory (reviewed in Steward and Schuman, 2001; Steward and Worley, 2001). The immediate early gene Arc is induced by electrical stimulation or behavioral experience (Steward and Worley, 2001). The newly generated mRNA is then delivered to dendrites, where it is targeted to active areas of the dendrite and depleted from inactive regions. An accumulation of Arc protein is also noted in these active regions, and the protein is assembled into the post-synaptic density-NR2 complex. Finally, behavioral studies show that interference with Arc (by antisense oligonucleotide infusion) blocks consolidation of hippocampus-dependent memory (Steward and Worley, 2001).

29 Conclusions

In conclusion, we have shown that dendritic spines in adult hippocampal slices begin to proliferate within one hour, and that protein synthesis is necessary for this proliferation. In addition, the population of elongated spines produced during spine proliferation may represent immature filopodia and newly formed spines. The ability of brain slices from 4-month-old rats to produce these changes demonstrates the retention of plastic capabilities well into adulthood.

30 CHAPTER 3

A REGULATORY ROLE FOR ACTIN IN DENDRITIC SPINE

PROLIFERATION

Introduction

Dendritic spines are supported by a dynamic actin cytoskeleton. Differential crosslinking of actin filaments within spines yields various spine types (reviewed by Rao and Craig, 2000): mushroom-shaped spines have stalks with bulbous heads, stubby spines are thick and have no necks, and filopodia are long, thin, and headless. Filopodia are prominent in development (Papa et al., 1995; Ziv and Smith, 1996; Daily and Smith, 1996; Fiala et al., 1998), and are likely immature spines that evolve into mushroom or stubby spines if appropriate axonal contact is made and sustained (Ziv and Smith, 1996; Daily and Smith, 1996). We previously reported that spine proliferation in the acute slice preparation is protein synthesis-dependent (Chapter 2; Johnson and Ouimet, 2004). Overexpression studies have implicated multiple proteins in spinogenesis, including actin binding proteins, receptor subunits, and small GTPases (Zito et al., 2004; Passafaro et al., 2003; Choi et al., 2005; Tashiro et al., 2000; Gartner et al., 2005). Ultimately, all of the signaling pathways regulating spinogenesis must converge on the polymerization of globular actin (G-actin) to fibrous actin (F-actin), since this is the final step causing the protrusion of a filopodium (a proto-spine) from a dendritic shaft. In neurons, it is not clear if actin can be up- or down-regulated in response to physiological pressures, and if so, whether such regulation could be used to adjust spine density. This study was undertaken to determine whether neurons elevate actin expression during reactive spinogenesis, whether actin overexpression per se can increase spine density, and whether newly generated actin preferentially enters newly generated spines. We show that neurons dramatically increase actin expression during reactive

31 spinogenesis, that actin overexpression leads to increased spine density, and that newly generated actin enters all spines within 24 hrs.

Methods

Slice Preparation

Postnatal day 21 (P21) rats were anesthetized with pentobarbital sodium (60 mg/Kg), decapitated, and the brain was removed and cut into 400 µm sections (in the coronal plane) using a McIllwaine tissue chopper. Slices from one hemisphere were fixed immediately in 4% paraformaldehyde and 0.03% glutaraldehyde as previously described (Kirov et al., 1999; Johnson and Ouimet, 2004). Slices from the other hemisphere were incubated on porous Millicell membranes (0.4um, Millipore Corporation, Bedford MA, 2-4 slices per membrane), in a 6-well plate with 750 µl of artificial cerebrospinal fluid (ACSF-117 mM NaCl, 5.3 mM KCl, 26 mM NaHCO3, 1 mM NaH2PO4, 2.5 mM CaCl, 1.3 mM MgSO4, and 10 mM glucose, equilibrated with

95% O2-5% CO2, pH 7.4, at 32°C) beneath the membrane. Plates were maintained at

32°C with 95% O2-5% CO2. After 90 minutes in vitro (MIV), slices were collected and fixed in the same manner.

Golgi Staining and Phalloidin with Silver Enhancement Two techniques were used to measure spine density in stratum radiatum of CA1: Golgi staining, to assess spine density on the secondary dendrites, and phalloidin staining, to assess spine density in the entire neuropil. Following fixation, the slices were transferred to 0.01 M phosphate buffered saline (PBS) and sectioned at either 100 µm (to be processed for Golgi) or 50 µm (to be processed for phalloidin-gold staining) using a vibratome (Leica). In either case, only sections from the middle 200 µm, previously shown to be the healthiest part of the slice (Kirov et al., 1999), were prepared for analysis. Golgi staining for brain slices (Izzo et al., 1987) was modified as described in Norrholm and Ouimet, 2001. In brief, sections were incubated in 1% aqueous osmium

tetroxide (OsO4) for 30 minutes. OsO4 was washed out in 0.1 M phosphate buffer, and

32 the slices were then incubated in aqueous 3.5% potassium dichromate. Next, sections were placed between two microscope slides in a "sandwich" configuration that was then submerged in aqueous 1% silver nitrate for 48 hrs. Sections were subsequently dehydrated in a graded ethanol series and mounted in DPX (BDH Laboratory Supplies, England). For phalloidin staining, 50 µm sections were washed in PBS, followed by a 10 min. incubation in 0.1% triton and 2x10 min. PBS washes. Non-specific binding was blocked by a 30 min. incubation in PBS with 5% bovine serum albumin (BSA) and 0.1% cold water fish skin gelatin (Electron Microscopy Sciences, Ft. Washington, PA). Tissue was then washed for 5 min. with incubation buffer (IB; PBS with 0.2% BSAc; Electron Microscopy Sciences), followed by a 1 hr. incubation in 5% biotinilated phalloidin (Molecular Probes, Eugene, OR). This was followed by 4x10 min. IB washes and a 2 hr. incubation in 1% streptavadin gold (Molecular Probes) in IB. Tissue was then washed in PBS (3x5 min.), 2% glutaraldehyde in PBS (5 min.), and distilled water (2x5 min.). Developer and enhancer (Electron Microscopy Sciences) were combined 1:1 and put on tissue for approximately 15 min. Following enhancement, tissue was washed in distilled water (3x5 min.), mounted, air-dried overnight, dehydrated, and coverslipped in DPX. All steps were carried out at room temperature. Protein extraction

Actin expression was measured during rapid spinogenesis in hippocampal slices from P21 rats. 400 µm thick brain slices were made as described above. Slices from one hemisphere were flash frozen immediately, while slices from the other hemisphere were

maintained for 90 minutes at the ACSF-air interface at 32°C with 95% O2-5% CO2, after which, the slices were flash frozen. The protocol for extraction of detergent-soluble proteins and membrane-bound proteins has been previously described (Murphy et al., 2001). Tissue was manually homogenized on ice in homogenization buffer containing 0.32 M sucrose, 10 mM tris base, 50 mM KCl, 1mM EDTA (pH 7.8) and the protease inhibitors phenylmethylsulfonyl, leupeptin, pepstatin A, A-protinin and sodium orthovanadate (Sigma, St. Louis, MO). For extraction of detergent-soluble proteins, Nonidet P40 was added to the homogenate, and the mixture was kept on ice and vortexed every 15 min. for 4 hr. The sample was then centrifuged at 4°C (30 min.), and the supernatant was stored at -80°C for SDS-PAGE and western blotting. For extraction of

33 membrane-bound proteins, homogenized tissue was centrifuged for at 4°C (30 min.), and the supernatant transferred to another tube and centrifuged again. The supernatant was collected and centrifuged at 45K RPM (4°C, 90 min.). The supernatant was discarded and the pellet resuspended by brief sonication in homogenization buffer. A Bradford protein assay (Bio-Rad Laboratories, Hercules, CA) was used to determine protein concentration in each sample. The samples were stored at -80°C for SDS-PAGE and western blotting.

SDS-PAGE and Western blots Purified hippocampal protein samples (detergent-soluble or membrane enriched) were separated on 12% acrylamide gels by SDS-PAGE and electro-transferred to nitrocellulose membranes. The nitrocellulose membranes were blocked with 5% blotting grade blocker (Bio-Rad) and incubated overnight at 4°C in rabbit polyclonal antibody against actin (Sigma; 1:2000). Membranes were then incubated with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (1:3000, 90 min., room temperature; Jackson ImmunoResearch Laboratories Inc). Enhanced chemiluminescence (Amersham Biosciences, England) exposure on Kodak Rx film (Fisher Scientific, Suwanee, GA) was used to visualize immunolabeled proteins. Six 15-min. washes at 45°C in 0.01M tris buffer (pH 8.8; also containing SDS and β-mercaptoethanol) and six 15-min. washes at 45°C in sodium citrate buffer (pH 3.0; also containing SDS and β-mercaptoethanol) were used to strip antibodies from the nitrocellulose for reprobing with neuron specific enolase (NSE; Chemicon International Inc, Temecula, CA). Band density of actin was normalized to that seen with neuron specific enolase (NSE diluted 1:6000, with secondary goat-anti-mouse diluted 1:3000).

Band density on films was analyzed by quantitative densitometry using a Hewlett- Packard Photosmart Scanner (model 106816; Hewlett Packard, San Diego, CA, USA) in conjunction with Quantiscan software (Biosoft, Cambridge, UK). A paired one-tailed t- test was used to assess the change from 5 to 90 MIV.

Organotypic cultures

Hippocampal slice cultures were prepared from 6-day-old rat pups following the protocol of Stoppini et al. (1991). Pups were decapitated and brains placed in chilled

34 dissection medium (1mM CaCl2, 5mM MgCl2, 10mM glucose, 4mM KCl, 26mM

NaHCO3 and 40mM sucrose), oxygenated with 95% O2/5% CO2. The hippocampi were carefully removed and cut coronally into 400 um slices using a McIllwaine tissue chopper. Two to four slices were transferred onto each porous Millicell membrane in a 6- well plate with 750 µl of MEM (Gibco, Grand Island, NY) with 25% horse serum, 13mM glucose, 5mM NaHCO3, 30mM HEPES, 1mM CaCl2, 2mM MgSO4, 1mM L-glutamine, 0.00125% ascorbic acid, 1µg/mL insulin, pH=7.28) below each membrane. Slices were

maintained at this liquid-air interface for six days at 37°C with 95% O2-5% CO2 before being transfected.

Gene Cloning and Transfection To determine if actin overexpression could drive spinogenesis, organotypic hippocampal cultures were transfected with the genes for β-actin and enhanced green fluorescent protein (EGFP). This expression construct was termed actin-EGFP, and was generated by subcloning the Xho1-Xba1 fragment of the actin gene from pEGFP actin (Clontech, Mountain View, CA) into Xho1-Xba1 digested pCMS GFP (Clontech). The genes were driven by the SV40 and CMV promoters, respectively, and their expression resulted in diffusion of EGFP throughout the cell with targeting of actin within it (Fischer et al., 1998). As a control, pCMS EGFP (Clontech) alone was expressed in some cultures. Spine density of neurons over-expressing actin and EGFP (actin-EGFP) was compared to that of neurons expressing EGFP only. To determine if exogenously introduced actin entered all spines, cultures were co-transfected with both the actin-EGFP vector in its original conformation (the two proteins were fused, termed fused actin- EGFP) and pDsRed-C1 (Clontech) for 48 hours, or fused actin-EGFP and DsRed-E (Clontech) for 24 hours. DsRed filled the entire cell, and fused actin-EGFP was trafficked within it. This allowed visualization of the entire cell as well as precise subcellular actin localization. Transfection was accomplished using biolistic gene transfer (Helios gene gun, Bio-Rad; McAllister, 2000). 25 µg Of plasmid DNA (1. pEGFP, or 2. pEGFP and actin, or 3. fused pEGFP-actin and pDsRed-C1 or 4. fused pEFGP-actin and pDsRed-E) was coated onto 8 µg of gold beads (1.6 µm diameter), which were then coated on the inside of tezfel tubing, following the company protocol. The gold particles were shot into tissue

35 using the gene gun with helium propulsion at 80-100 psi. To determine the effect of actin overexpression on spine density, slices were transfected at 6 days in vitro (DIV), given 48 hours to express the proteins, and fixed at 8 DIV, as described below. To determine if fused actin-EGFP entered all spines, slices were transfected at 6 or 7 DIV and fixed at 8 DIV to provide expression at 24 and 48 hours. Slices were immersion-fixed for one hour at room temperature, transferred to PBS, and stored at 4ºC. Sucrose (final concentration 3.4%) was added to the fixative to maintain osmotic balance. Immunocytochemistry To determine if filopodia and elongated spines in actin-overexpressing cells (transfected with actin and EGFP, i.e., actin-EGFP) made synaptic contacts, fixed cultures were cut on a vibratome (Leica) into 50 µm sections. Sections were washed in PBS (3x10 min), and non-specific binding was blocked by the same treatment described in the phalloidin procedure, with the addition of 5% normal goat serum, followed by 5 min. incubation in IB. Sections were then incubated overnight in rabbit-anti-synapsin-I antibody (1:1000; Santa Cruz Biotech, Santa Cruz, CA). Tissue was washed (3x10 min.) in IB, and incubated in cy3 goat-anti-rabbit antibody (1:500; Jackson ImmunoResearch Laboratories Inc, West Grove, PA) for one hour. Tissue was washed in PBS (3x10 min.) and mounted in vectashield (Vector Laboratories, Burlingame, CA) for confocal microscopy. Antibodies were diluted in IB and all steps described above were carried out at room temperature. Laser Scanning Confocal Microscopy To assess spine density and examine spine morphology in transfected cells in organotypic culture, confocal microscopy was employed. Image stacks of slices were collected along secondary apical dendrites and along basal dendrites using a Zeiss LSM 510 Laser Scanning Confocal Microscope with LSM software and a 40X IR-Achroplan 0.8 NA water immersion objective. Images were acquired using a 3x scan zoom. Steps in the z plane were taken at 0.5 µm. Images were saved digitally and spines counted using Neurolucida (MicroBrightField Inc, Williston, VT). Density differences were analyzed by 2-sample 1-tailed t-test.

36 Results

Spine Density Increases in Hippocampal Slices from P21 Rats Two independent staining procedures were employed to assess spine density in the acute slice preparation, Golgi impregnation and silver-enhanced phalloidin staining. Golgi impregnation fills cells and allows measurement of spine density per unit length of dendrite. Phalloidin staining labels F-actin in dendritic spines, staining puncta that can be used for estimates of spine density in the neuropil. The latter was assessed using the computer program Stereo Investigator (MicroBrightField Inc, Colchester, VT) to perform unbiased stereology. In this method, the computer program randomly selects numerous small sample sites for analysis. Sites are analyzed for the number of puncta (presumptive spines) contained within them, and the data can be used to determine the average number of puncta per sample site, or the density of puncta in a given volume. 2-3 Slices were analyzed per rat, and in each slice 100 1µm3 sample areas within the inner 2/3 of area CA1 of stratum radiatum were analyzed. The average number of puncta per site was then determined. After 90 minutes in vitro, spine density on secondary dendrites in area CA1 of hippocampus increased by 23% (p<0.01) as determined by Golgi staining (Figure 7). In this experiment, no differentiation was made among spine types during analysis. Spine density within the neuropil of CA1 increased by 28% (p=0.01) as determined by phalloidin labeling (Figure 8). Actin Expression Increases During the Period of Rapid Spinogenesis

The acute slice preparation described above was then used to assess changes in actin expression during spinogenesis. Over 90 min., actin expression in the detergent- soluble fraction of hippocampus increased 28% (p=0.03) as determined by Western blots. Furthermore, when only the membrane-enriched fraction was analyzed, actin expression increased 81% (p=0.01; Figure 9).

Actin Overexpression Increases Dendritic Spine Density

The fact that actin expression was upregulated during reactive spinogenesis does not prove that actin overexpression per se caused an increase in spine density;

2.5 * 2

1.5

0.5

Spine Density (spines/micron) 0 5 MIV 90 MIV

5 MIV

90 MIV

Figure 7. Spine density as assessed by Golgi staining increases during acute slice preparation in P21 rats. (Top) Spine density on apical secondary dendrites of CA1 pyramidal cells, as assessed by Golgi staining, increased 23% over the first 90 minutes in vitro (p<0.01). (Bottom) Example Golgi-stained dendrites from acute slice preparation (top, 5 MIV; bottom, 90 MIV). Calibration bars indicate 10 µm. N=8, 6 respectively.

1.6 * 1.4 1.2 1

0.8

0.6

0.4

0.2

Spine Density (spines/sample site) 0

5 MIV 90 MIV

Figure 8. Spine density as assessed by phalloidin staining increases during the acute slice preparation in from P21 rats. (Top) Spine density in CA1 stratum radiatum neuropil, as assessed by phalloidin staining, increased 28% over the first 90 minutes in vitro (p=0.01). (Bottom) Example images of phalloidin staining with silver enhancement. Large square within each image indicates area of 10µm x 10µm. That area is shown at higher magnification at the bottom right of each image, and the small square within it indicates size (1µm3) of sample sites used in analysis. N=5 per group.

2.5

* 2

1.5 * 5 MIV 90 MIV 1

0.5

Pixel Density (Actin/NSE)

0 Detergent-soluble Membrane-enriched fraction fraction

NSE, 47 kDa

Actin, 43 kDa

Figure 9. Actin expression increases during acute spice preparation. (Top) Graph of Western blot data demonstrating a relative increase in pixel density from 5 minutes in vitro (MIV) to 90 MIV for detergent-soluble fraction (left) and for the membrane- enriched fraction (right). Actin levels increased significantly (p=0.03, p=0.01, respectively) during the first 90 minutes in vitro, corresponding to the period of spinogenesis. (Bottom) Panels with example bands from Western blots. Each panel shows a band from 5 MIV (left) and from 90 MIV (right). Corresponding to the graphs, panels on the left show bands from the detergent-soluble fraction and panels on the right show bands from the membrane-enriched fraction. Top panels show the housekeeping protein, neuron specific enolase; bottom panels show actin. N=4 (detergent-soluble fraction), 7 (membrane-enriched fraction).

40 upregulation of actin expression could have occurred in non-neuronal cells. Further, it is almost certain that the expression of other proteins was upregulated as well. To determine whether actin overexpression per se could increase spine density, we transfected neurons in organotypic slice cultures with the genes for β-actin and GFP. The two genes were present in a single vector but driven by different promoters (actin-EGFP). This enabled the visualization of cells overexpressing actin.

Only pyramidal cells in area CA1 of hippocampus were analyzed. Furthermore, cells were chosen for analysis only if they appeared healthy (i.e., full apical and basal dendritic trees, no blebbing in the dendrites). An average of 273 µm of dendritic length

(with a minimum of 100 µm) was analyzed for each dendritic tree of each cell. In the EGFP group, the apical trees of 6 cells from 4 rats were analyzed and the basal trees of 5 cells from 4 rats were analyzed. In the actin and EGFP group, the apical trees of 4 cells from 3 rats were analyzed and the basal trees of 3 cells from 2 rats were analyzed.

CA1 pyramidal cells overexpressing actin (actin-EGFP) had 38% more spines on their secondary apical dendrites, and 41% more spines on their basal dendrites than controls expressing EGFP alone (p=0.008, p=0.002, respectively; Figure 10). In addition to estimating overall spine density, we assigned spines to 5 categories: (1) short stubby spines with no heads, (2) mushroom-shaped spines with short necks and bulbous heads, (3) branched spines, (4) elongated spines having thin necks at least 1.5 µm long and small heads and (5) filopodia. On apical dendrites there was a 162% increase in the number of elongated spines (p=0.009), a 38% increase in the number of filopodia (p=0.02), and no significant change in the number of mushroom or stubby spines (Figure 11). There was also a significant increase in the number of branched spines (p=0.03). On basal dendrites, there was a 113% increase in the number of elongated spines (p=0.04), an 88% increase in the number of filopodia (p=0.03), and no significant change in the number of mushroom, stubby or branched spines (Figure 12). These data demonstrate that the majority of additional spines are elongated spines and filopodia. Examples of dendrites from EGFP and actin-EGFP transfected cells are shown in Figures 10, 11.

) 0.8 * 0.7 * 0.6 0.5 EGFP 0.4 actin-EGFP 0.3

0.2

0.1 Spine (spines/micron Density 0

apical basal

Figure 10. Spine density increases after actin-overexpression in organotypic cultures. Overall spine density from actin-EGFP-transfected pyramidal neurons in organotypic slice culture increased 38% on apical dendrites (p=0.005) and 41% on basal dendrites (p=0.002) over EGFP-transfected controls. N=6, 4, 5, 3, respectively.

0.3

) 0.25

0.2

EGFP 0.15 * * actin-EGFP

0.1

Spine (spines/micron Density 0.05

* 0

y d m e bb h oo u c st ngated ran ushr filopodial o b m el

EGFP

Actin- EGFP

Figure 11. Increase in spine density of elongated, filopodial and branched spines on apical dendrites of actin-EGFP-transfected pyramidal neurons. (Top) There were 38% more filopodial spines, and 162% more elongated spines in actin-EGFP-transfected cells as compared to EGFP-transfected cells (p=0.02), whereas the density of stubby and mushroom-shaped spines did not change. There was also a significant increase in the density of branched spines in actin-EGFP-transfected cells (p=0.03). (Bottom) Example apical dendrites from EGFP- and actin-EGFP-transfected neurons. Closed arrow indicates elongated spine; open arrows indicate filopodia; closed arrows with balls indicate mushroom-shaped spine; ‡ indicates stubby spine. N=6 (EGFP), 4 (actin-EGFP). Scale bars indicate 10 µm.

0.25

) 0.2 *

* 0.15 EGFP

actin-EGFP 0.1

0.05 Spine Density(spines/micron

y ial ed ted h od a c p stubb shroom lo u fi bran m elong

EGFP

Actin- EGFP

Figure 12. Changes in spine density of specific spine types on basal dendrites from EGFP- and actin-EGFP-transfected pyramidal neurons. (Top) There were 88% more filopodial spines (p=0.03) and 113% more elongated spines (p=0.04) in actin-EGFP- transfected cells as compared to EGFP-transfected cells, whereas the density of branched and stubby, and mushroom-shaped spines did not change. (Bottom) Example basal dendrites from EGFP- and actin-EGFP-transfected neurons. Closed arrow indicates elongated spine; open arrow indicates a filopodium; closed arrow with ball indicate mushroom-shaped spine; ‡ indicates stubby spine. N=5 (EGFP), 3 (actin-EGFP). Scale bars indicate 10 µm.

44 Elongated Spines and Filopodia are Apposed by Axon Terminals

Organotypic cultures containing pyramidal cells expressing actin-EGFP were sectioned and stained for synapsin-I. 107 Filopodia and 207 elongated spines were analyzed for the presence of adjacent synapsin-I puncta. The heads of 93% of elongated spines were apposed by a presynaptic terminal, and 80% of filopodia were apposed by one or more axon terminals (Figure 13).

Exogenous actin enters all spines

To determine whether actin turnover occurs in all (new and established) spines, we transfected organotypic hippocampal cultures with both DsRed, which filled the entire cell, and fused actin-EGFP, which was trafficked within the cell. We assessed the number of DsRed spines that were double labeled for fused actin-EGFP. At 48 hours (the time of overexpression when density was assessed above), actin-EGFP was observed in all spines on both apical (Figure 13) and basal (Figure 13d) dendrites. We chose to look at an earlier time point as well, and at 24 hours actin-EGFP was again observed in all spines (Figure 14).

Discussion

Actin Upregulation and Increased Spine Density

In the acute hippocampal slice preparation, dendritic spine density increases over time (Kirov et al., 1999) in a protein synthesis-dependent manner (Johnson and Ouimet, 2003). In P21 rats, we found similar increases in spine density using Golgi staining and phalloidin staining with silver enhancement. The consistency in results using these two methods suggests that phalloidin labeling with silver enhancement, in conjunction with unbiased stereology, is a valid method for identifying and quantifying dendritic spines. In the future, this technique could be used without the corroboration provided by Golgi staining to determine and report spine density.

Figure 13. Elongated spines and filopodia are apposed by axon terminals. Synapsin-I staining of a culture with an actin-EGFP-transfected pyramidal cell reveals that filopodia and elongated spines are opposed by synapsin-positive terminals, suggesting the presence of a synapse. Closed arrows indicate elongated spines; open arrow indicates a filopodium.

Figure 14. Exogenous actin enters all spines. Pyramidal cells expressing DsRed and fused actin-EGFP. Each panel has 3 images: (top) fused actin-EGFP, (middle) DsRed, and (bottom) composite. (A) Apical dendrite, 24 hours post-transfection, (B) Basal dendrite, 24 hours post-transfection, (C) Apical dendrite, 48 hours post-transfection, and (D) Basal dendrite, 48 hours post-transfection. At both time points, all spines contain exogenously introduced actin.

47 Here we also show that actin expression increases during spinogenesis, raising the possibility that neurons can control overall spine density by regulating the amount of available actin. On the other hand, the increase in actin expression could simply reflect increased spine density and concomitant changes in the expression of proteins other than actin could impact spine density. Further, the increased actin expression could have occurred in non-neuronal cells. We therefore transfected neurons with the gene for β- actin and showed that overexpression of actin per se was sufficient to increase spine density. Non-neuronal cells regulate actin expression using an autoregulatory mechanism that involves the 3’ UTR of actin mRNA (Lyubimova et al., 1999), and it is thought that the presence of G-actin exerts direct negative feedback on actin gene expression (Lyubimova et al., 1997; Schmitt-Ney and Habener, 2003). In the acute slice preparation, the pool of G-actin could have become depleted as new spines developed, thus stimulating the cell to increase actin production. Taken together, these observations suggest that neurons can regulate actin expression, and that actin upregulation could be one mechanism neurons employ to increase spine density. The morphology of dendritic spines observed after transfection with EGFP alone was similar to that described previously for this type and age of culture (Dailey and Smith, 1996). In neurons overexperessing actin, there was a specific and dramatic increase in the number of elongated spines and synaptic filopodia on apical dendrites, with no change in the density of mushroom, stubby or branched spines. This suggests that actin overexpression produced a new population of elongated spines and synaptic filopodia. The morphology of this new population may reflect spine development in the presence of a mismatch between the concentration of actin and that of actin-organizing proteins. A similar mismatch could be related to the clinical observation that elongated spines and filopodia are also present during fetal development in Down Syndrome in which specific actin binding proteins are under-expressed (reviewed by Engidawork and Lubec, 2003). In addition, it is not known whether excess actin can increase the rate of monomer addition to actin filaments such that newly developed transient spines or filopodia that would have retracted now become stabilized. Synapsin-I staining revealed that the vast majority of the filopodia and elongated spines on actin-EGFP transfected cells were apposed by presynaptic terminals. Although

48 synaptic contacts may be defined only at the electron microscopic level, these observations strongly suggest that the elongated spines observed after actin overexpression form synapses. This implies that presynaptic endings in hippocampal organotypic culture are capable of sprouting to rapidly occupy newly generated spines. In contrast, dendritic protrusions in primary hippocampal tissue culture are not apposed by axon terminals in response to actin overexpression driven by the same construct (Eom et al., 2003). These primary tissue cultures were analyzed on postnatal days 10-12, and were therefore 2-4 days younger than the organotypic cultures used in the present study and this may in part account for the different results. Interestingly, when an additional zipcode nucleotide sequence was added to the construct, filopodia were likely to be apposed by axon terminals (Eom et al., 2003).

The response of dendritic spines to actin overexpression was not uniform throughout the dendritic tree in organotypic culture; although the density of elongated spines and synaptic filopodia increased on both apical and basal dendrites, the density of branched spines increased on apical dendrites only. This observation is in keeping with previous observations of a heterogeneous change in dendritic complexity and/or spine density on apical vs. basal dendrites (Santos et al., 2004). Factors intrinsic to the basal dendrites vs. apical dendrites may have accounted for the difference. Additionally, in organotypic cultures, stratification of axon terminal input is maintained and spine changes could be associated with synaptic input. For example, the increase in branched spines could have been caused by higher levels of activity in the inputs to stratum radiatum than were present in the stratum oriens. Branched spines have been observed to increase in density after high-frequency stimulation as well as following deafferentation of their parent cells (Dhanrajan et al., 2004; Chen et al., 1982). Finally, branched spines are among 3 spine types (along with thin spines and mushroom-shaped spines with perforated PSDs and SAs) that increase 4-fold from P15 to adulthood (Harris et al., 1992). The specific increases observed in the density of elongated spines, and filopodia, as well as the increase in branched spines on the apical dendrites, might reflect the beginning of this process.

49 Exogenous Actin Enters all Spines

It has been demonstrated using the FRAP technique that 85% of spine actin turns over in less than one minute (Star et al., 2002; Zito et al., 2004). FRAP studies using fused actin-EGFP, however, necessarily analyze only EGFP-labeled spines; spines with slow actin turnover would not be detected. To determine whether a population of mature spines with slow actin turnover exists, we transfected neurons with both DsRed (to fill the entire cell) and fused actin-EGFP. Twenty-four hours after transfection, the earliest time point at which EGFP and DsRed could be observed, actin-EGFP had entered all spines. These data show that exogenously introduced actin rapidly enters both new and established spines and are consistent with the earlier FRAP studies (Star et al., 2002; Zito et al., 2004) showing that actin turnover is high.

The high actin turnover rate in all spines suggests that this process is important to spine function. In addition to its roles in spine motility and structural support, actin is likely important in the transport of spine proteins. Actin fibers are known to treadmill by adding monomers at one end (the barbed end) and removing them from the other (the pointed end; reviewed by Cooper and Schaffer, 2000) Actin fibers are attached to the PSD by their barbed ends. Fibers of both polarities run between the plasma membrane and the spine apparatus and along (parallel to) the PSD (Fifkova and Delay 1982). In the spine neck, actin fibers run longitudinally, and their polarity may be mixed (Fifkova and Delay, 1982; reviews by Fifkova, 1985 and Rao and Craig, 2000). Many spine- enriched proteins have actin binding domains that are necessary for spine targeting (discussed and referenced in Chapter 4). These proteins, bound to treadmilling actin fibers, would move from the barbed to the pointed end. Actin treadmilling could therefore shuttle proteins to many destinations in the spine and the junction between spine and dendritic shaft.

50 Conclusions

Taken together, the findings presented here show that neurons may upregulate actin expression in order to increase spine density, especially during periods of rapid spinogenesis. Further, the newly generated actin enters both established and newly developed spines.

51 CHAPTER 4

DISCUSSION

The studies in this dissertation demonstrate that (1) dendritic spines proliferate in the first hour after a hippocampal slice is made, (2) protein synthesis is necessary for dendritic spine proliferation in the acute slice preparation, (3) actin expression increases concurrently with spine proliferation, (4) spine formation in organotypic cultures can be driven by actin overexpression, and (5) all spines on CA1 pyramidal cells in organotypic slice cultures have a rapid degree of actin turnover. Taken together, these data suggest that spines do not arise from pre-packaged units; rather they require protein synthesis in order to form. Additionally, actin appears to have a regulatory role in spine growth, as opposed to the conventional understanding of actin only as a structural support system.

Methodological Considerations

In any series of experiments, methods and preparations are chosen based on careful balancing of the questions they can help answer and the limitations inherent to them. It is therefore prudent to consider data and draw conclusions in this context. One of the biggest considerations in the current series of experiments is the difference between young and adult tissue, in terms of spine development and experimental possibilities. For example, spine proliferation in the acute slice preparation had already been reported in young animals. It was therefore a good model to address spine proliferation in adults, as well as the role of protein synthesis in spine proliferation. Acute slices, however, do not live long enough for transfection. In fact, no adult brain tissue can be cultured well. Overexpression studies, therefore, are done almost exclusively in young tissue or cultured neurons. Although much of the data gleaned from such studies can be reasonably extrapolated to adults, potential differences exist. Two examples of such differences are the dendritic localization of actin mRNA in developing, but not mature neurons, and the differential affect of Latrunculin A on PDS-95

52 localization in developing and mature neurons (Allison et al., 2000; Takahashi et al, 2003). Due to the existence of such differences, it was possible that when moving from adult rats, where protein synthesis is necessary for spine proliferation, to P21 rats and then to organotypic culture (ending at 8 days in vitro, which was comparable to P14), we could get to a point in development where protein synthesis was no longer necessary for spine growth. Our data, however, that actin expression increases with spine density and can drive spine proliferation, as well as data from other groups showing increases in spine density after the overexpression of proteins other than actin, indicate that spine development may well necessitate protein synthesis in young rats as well as old. The use of cultures instead of in vivo brain tissue is another important consideration. The beauty of the organotypic slice culture, especially from hippocampus or any laminar structure, is that it largely maintains the connectivity which is found in vivo. Additionally, organotypic cultures have roughly the same compliment of dendritic spines, in terms of overall density and specific spine type, as the postnatal day 15 brain (reviewed by Harris, 1999). The limitation, however, is that biolistic gene transfer, arguably the best way to transfect cells in a slice culture, has a much lower transfection rate than electroporation or calcium phosphate (Martinez and Hollenbeck, 2003; Watanabe et al., 1999), the latter of which can only be used in monolayers of cells. A final consideration is the use of enhanced green fluorescent protein (EGFP) to label transfected cells and to tag exogenous actin. The main two concerns are that (1) actin tagged with EGFP can still be localized and polymerized properly, and (2) a cell expressing and filling with EGFP will function properly. Indeed, it is now well established that that EGFP does not alter the function of the cell (reviewed by Arun et al., 2005), and it has been reported that overexpressed, fluorescence-tagged actin concentrates into puncta at newly forming synapses and in dendritic spines, supporting the notion that exogenously introduced actin indeed translocates to appropriate sites (Colicos et al., 2001; Kaech et al., 1997; Fischer et al., 1998). Furthermore, the current data show actin-EGFP in dendritic spines.

53 A Common Pathway for Spine Growth?

Dendritic spine density increased in both the acute slice preparation and after actin overexpression in organotypic culture. In the acute slices there was also an increase in spine length, which likely characterized the new population of spines. Likewise, in actin-overexpressing cells, the increase in spine density was due almost exclusively to an increase in elongated spines and filopodia. This similarity in spine morphology in the two preparations points to a common pathway for spine growth occurring both in cases of injury (the acute slice) as well as in actin overexpression. It is not very surprising that the two preparations would look similar; filopodial extension is widely believed to be the first step in spine formation (discussed and referenced in Chapter 1), and the elongated spines fit in well to a model where filopodia mature into spines. In the overall context of ‘how spines grow’ it is important and encouraging that two models which differ significantly in age of tissue and type of preparation seem to follow a similar sequence of events leading to spine formation.

Actin Upregulation During Spinogenesis

We demonstrated a concomitant increase in spine density and actin expression in the acute slice preparation. Are those two changes causally related, and if so, which occurs first? Data from non-neuronal cells suggests that a net loss of G-actin within the cell signals an increase in actin expression (Schmitt-Ney and Habener, 2003). In the context of the present studies, an increase in spines could begin a positive feedback loop of actin expression and spine formation, wherein spine proliferation depletes the G-actin pool, which in turn signals the cell to upregulate actin expression. Alternatively, the physiological quiescence that occurs after a slice is made, and which is the driving force behind spine proliferation in acute slices (Kirov et al., 1999), could somehow signal the cell to increase actin production, resulting in spine formation. The observation that actin overexpression led to an increase in spine density in the organotypic cultures demonstrates that actin upregulation can increase spine density. It is therefore possible that the events in the acute slices began with actin upregulation and resulted in spine

54 proliferation. In this case, what was the specific signal to the cell to increase actin expression? Does this unknown signal occur in all situations where spine density increases? These questions have not been answered, but could begin to be addressed by looking at actin levels in other models of spine proliferation.

Proteins Involved in Spine Growth

Spinogenesis is a highly regulated process that requires multiple proteins at each step, from actin polymerization to filopodial extension to spine maturation. Many proteins play important roles in filopodial initiation and spinogenesis. Overexpression of VASP, fascin, N-WASP and drebrin cause filopodial initiation in non-neuronal cells (Svitkina et al., 2003; Miki et al., 1998; Shirao et al., 1994), and α-actinin 2, dynamin 3 (splice variant), Rac 1, β-actin and cortactin lead to the formation of non-synaptic filopodia in neurons (Nakagawa et al., 2004; Gray et al., 2005; Tashiro et al., 2000; Eom et al., 2003; Hering and Sheng, 2003). Most relevant to the data presented in this dissertation, however, are the proteins whose overexpression or constitutive expression increase dendritic spine density; neurabin 1, glutamate receptor 2 subunit of AMPA receptors (GluR2), insulin receptor substrate 53 (IRS-53), rac1, ras, and now β-actin have all been reported to drive new spine formation in spine-bearing neurons (Zito et al., 2004; Passafaro et al., 2003; Choi et al., 2005; Tashiro et al., 2000; Gartner et al., 2005; Johnson and Ouimet, under review; Table 4), and GluR2 overexpression causes aspiny interneurons to form dendritic spines (Passafaro et al., 2003). Furthermore, proteins that cap, nucleate, sever and otherwise bind actin filaments, and that until recently have been studied only in non-neuronal cells, are now being recognized for their functions in neuronal filopodial extension (Svitkina et al., 2003; Vignjevic et al., 2003; reviews by Rao and Craig, 2000 and Cooper and Schaffer, 2000). If one posits that multiple proteins are necessary for normal spine development, then how can experimentally increasing the level of just one drive spine proliferation? This is especially perplexing when the protein in question is not, to our knowledge,

Table 4. Morphological consequences of protein overexpression or constitutive expression in neurons.

Increased Increased Spine/filo Morphological Protrusions in spine loss after Protein spine # filopodia # elongation changes (other) aspiny neurons knockdown

α-actinin-2 x x . cortactin x x x drebrin x . dynamin 3 sv x GluR2 x x x x . IRS 53 x Neurabin 1 x x . Rac1 x x x x Ras x . Rnd1 x x Kalirin 7 x x . β-actin x x

56 involved in filopodial initiation during normal development. One possibility is that an increase in the level of one protein recruits other spine-related proteins that already exist in excess within the cell. In this case, the cell maintains an optimal level of reserve proteins that are capable of responding quickly to a signal to make more spines. A second possibility is that an increase in just one spine related protein signals the cell to upregulate all spine related proteins. This could be tested by overexpressing one protein that is known to drive spine formation, and then measuring the levels of other spine- related proteins. A third possibility is that each of the proteins that drive spine formation has the capability, directly or indirectly, to signal the cell to synthesize more actin. Obviously, each of these proteins can signal actin polymerization, as this is the final common step toward forming a spine, but there is only so much actin available in the cell at any given time, and a ceiling effect could occur quickly. Upregulation of actin expression would increase the level of monomers within dendrites, shifting the G:F-actin ratio, and likely initiating filopodial formation and possibly increasing the length of existing spines. Additionally or alternatively, an increase in monomers could block the retraction of otherwise transient filopodia, thus increasing spine density. (Interestingly, no study to date has addressed the origin of new spines driven by experimental manipulations). The increase in filopodia and thus in synaptic contacts could signal the cell to make or recruit other spine-related proteins to assist in the maturation and stabilization of the new spines/synapses. Each protein, when overexpressed, initially drives spine formation in a different way, but ultimately must initiate actin polymerization, which could result in actin upregulation, as the G-actin pool within the cell becomes depleted. Exactly how each protein leads to actin polymerization is not entirely clear, but parts of the pathways are known. Neurabin 1 is an F-actin and PP1 binding protein that is enriched in spines (Nakanishi et al., 1997, Zito et al., 2004; McAvoy et al., 1999 ). It was recently reported that the actin binding domain of neurabin 1 bundles actin and promotes filopodia (Terry- Lorenzo et al., 2005). Interestingly, in neurons overexpressing neurabin 1, and thus making more spine synapses, the total synaptic input is constant (Zito et al., 2004). By decreasing the amplitude of mEPSCs, the cell, despite additional synapses, is able to regulate its overall excitability (Zito et al., 2004).

57 Cortactin is an F-actin binding protein that is involved in the stabilization and branching of actin via its interaction with the Arp2/3 complex (Weaver et al., 2001). Overexpression of cortactin leads to spine formation in aspiny neurons and spine elongation in spiny neurons (Hering and Sheng, 2003). The interaction of cortactin with Arp2/3 is significant because Arp 2/3 nucleates new actin filaments and filament branches, and is implicated in filopodial initiation in both neuronal and non-neuronal cells (Reviews by dos Remedios et al., 2003, Pollard and Beltzner, 2002, and Millard et al., 2004). GluR2 is a subunit of the AMPA receptor, and it’s presence in AMPA receptors blocks them from fluxing calcium. GluR2 might initiate filopodial extension by binding to the extracellular domain of a presynaptic membrane protein, thereby aligning receptors with terminals (Passafaro et al., 2003). The state of actin polymerization is regulated, in part, by calcium, which stabilizes actin filaments, and at high concentrations causes depolymerization (discussed and referenced in Chapter 1). GluR2-contaning AMPA receptors may function to enhance the efficacy of synaptic transmission by aligning receptors with their ligands, but not allow an overabundance of calcium into the postsynaptic cell. Ras is involved in neurotrophic signal transduction downstream of Trk receptors, where it activates MAPK (Reviewed by Huang and Reichardt, 2003 and Gartner et al., 2005). This is significant because MAPK is involved in activity-induced formation of dendritic filopodia (Wu et al., 2001; Alonso et al., 2004). The exceedingly complex pathways activated by Trk receptors involve a multitude of proteins and signaling molecules. What is particularly germane to the present studies is the ability of MAPK to affect the actin cytoskeleton, as well as the ability of neurotrophins to regulate local translation of mRNA including that for actin (reviewed by Huang and Reichardt, 2003). The small GTPase Rac1 is believed to locally regulate the actin cytoskeleton, but exactly how this occurs is unknown (Nakayama et al., 2000). Related to Rac1, kalirin 7 is a splice variant of the Rho GEF kalirin (a GDP/GTP exchange factor for Rho family GTPases) that is enriched in dendritic spines (Penzes et al., 2001), and that likely increases spine-like protrusions via Rac1 (Penzes et al., 2001).

58 Finally, α-actinin 2 could form spines by linking actin filaments to receptor complexes (Nakagawa et al., 2004). In spines, α-actinin 2 links the NMDA receptor to the actin cytoskeleton (Wyszynski et al., 1997; Dunah et al., 2000). It therefore seems possible that α-actinin 2 could assist in accumulation of F-actin at sites ripe for filopodial formation. Furthermore, α-actinin 2 might affect actin polymerization in a manner similar to GluR2; by modulating calcium entry via NMDA receptors, α-actinin 2 could affect actin polymerization. However, calcium entry through the NMDA receptor leads to a displacement of α-actinin 2 from the NMDA receptor and an inhibition in receptor activity (Ehlers et al., 1996; Wyszynski et al., 1997; Zhang et al., 1998). Additionally, α- actinin 2 does not colocalize with NMDA receptors on dendritic shaft synapses (Rao et al., 1998).

Possible Involvement of Dendritic Protein Synthesis

An interesting aspect of the studies presented in this dissertation is the rapidity by which dendritic spines were formed in the acute slice preparation, and the fact that this proliferation was blocked with protein synthesis inhibitors. Because of the critical role of spines in learning and memory, cells need to be capable of making new spines quickly in response to a brief and transient stimulation. Given that de novo protein synthesis is a way for the cell to control spine proliferation, and that spines must grow quickly at a particular location, it seems possible that the mRNA for the protein(s) which must be made is located in dendrites. Intradendritic translation of the spine-inducing proteins may be a way to both specify exactly where the spine is made and quicken the process by negating the step of protein transport. The question thus arises, where are the mRNAs for each of the spine-inducing proteins? Rac1, neurabin 1, rnd1 and drebrin mRNAs are found in the cell layers of the hippocampus and dentate gyrus, but do not appear to be localized to the dendrites (Nakayama et al., 2000; Burnett et al., 1998; Ishikawa et al., 2003; Ooe et al., 2004). The localization of mRNA for α-actinin 2, kalirin 7, IRS 53, cortactin, dynamin and ras has not been identified. GluR2 mRNA (as well as GluR1 mRNA) is found in dendrites of hippocampal pyramidal cells, and the protein is locally synthesized and inserted into

59 the membrane (Ju et al., 2004). β-Actin mRNA has also been identified within dendrites (Tiruchinapalli et al., 2003), and when the amount dendritic actin mRNA is increased, BDNF-induced filopodial formation increases (Eom et al., 2003). Conversely, by knocking down dendritic actin mRNA, BDNF-induced filopodia formation is impaired. The authors speculated that this was directly related to the level of G-actin monomers within the dendrites. These data, however, were from very young neurons in primary culture. Dendritic actin mRNA is not found in neurons from adult rats (Kaech et al., 1997). This suggests that the spine proliferation observed in the acute slices from adult rats in the present study was not due to translation of dendritic actin mRNA (although in non-neuronal cells actin mRNA is translocated in response to injury; Hoock et al., 1991). Furthermore, the role of dendritic actin translation in the organotypic cultures was probably not significant. Actin mRNA is localized to dendrites via zipcode binding protein 1, a protein that interacts with a 3’ UTR region of β-actin mRNA and translocates it to dendrites. Although there may have been endogenous dendritic actin mRNA in the organotypic cultures, the β-actin gene that we overexpressed did not contain that zipcode. Therefore, the actin protein that caused the increase in spine density in the organotypic cultures was likely made in the cell body. It is still quite possible that there exists a role for locally produced proteins in spine proliferation, but the present experiments and the pertinent literature do not identify it.

Actin: A Limiting Resource for Spine Density

The findings presented here, that an increase in actin expression leads to an increase in spine density, is indicative of actin being a limiting resource for spine density. This is different from stating the obvious, that actin is a necessary resource for spinogenesis. As a limiting resource, cells can be driven to make more spines simply by increasing actin levels. There are several lines of data in addition to what is reported in this dissertation to support this. Increasing the level of actin mRNA in dendrites has been reported to increase BDNF-induced filopodial formation, likely by increasing the level of available G-actin (Eom et al., 2003). Using in situ hybridization, it has been demonstrated that actin mRNA levels are highest in large spine-bearing neurons such as

60 cortical and hippocampal pyramidal cells and cerebellar Purkinje cells (Kaech et al., 1997). On the other hand, immunostaining in sections of perfused adult rat brain show that actin is present in aspiny cholinergic neurons of the striatum (unpublished data). This last piece of data could still fit with the limiting resource hypothesis, as the exact level of actin was not measured. All cells need actin for functions other than making spines, such as structural support, motility, and endocytosis (Example reviews by Etienne- Manneville, 2004 and Schafer, 2002). Aspiny neurons could have a very low level of actin, too low to make spines. It would be interesting to measure levels of actin in cells with different spine characteristics, and to determine if overexpression of actin could stimulate spine formation in aspiny neurons, as has been seen for other proteins (Passafaro et al., 2003; Hering and Sheng, 2003).

Actin Treadmilling as a Delivery System

It was previously shown that most spine actin turns over in less than one minute (Star et al., 2002; Zito et al., 2004). In the present studies, we report that exogenous actin enters all spines within 24 hours, suggesting that the rapid turnover of spine actin is not limited to a subset of spines. If stable actin filaments provide adequate structural support, and a high degree of motility is not necessary in mature spines, then why do all spines have a high rate of actin turnover? An intriguing possibility is that treadmilling actin filaments are used to transport molecules into, out of and within spines. Many spine proteins contain an actin binding domain or motif, and for some of these proteins, this area is necessary for correct spine targeting (Table 5). If this is indeed a function of actin, it could have implications for other areas of cell biology. For example, myosin-V is the primary actin-based motor used for dendritic transport, and it also has a role, along with kinesin, in the transition of vesicles from microtubules (in dendrites) to actin filaments (in spines; Tabb et al., 1998; Molyneaux et al., 2000; Brown et al., 2004). Myosin motors “walk” from the pointed end to the barbed end along polymerized actin, carrying vesicles of molecules to be delivered at the end of the filament (for a review of this process by myosin V, see Tyska and Mooseker, 2003). It is not understood, however,

Table 5. Spine localized proteins with actin binding domains or motifs. Symbols represent: * Deletion of actin binding motif disrupts spine targeting; ^ Highly likely, but not definite, that binding to actin is direct; # There is no immunocytochemical or EM evidence of the protein in spines, but based on its function and ability to bind other spine proteins, it is highly predicted to be in spines.

Protein G/F Reference (for direct actin binding)

ADF G & F Giuliano et al., 1988 . α-Actinin 2* F Mimura and Asano, 1986; Nakagawa et al., 2004* α-Adducin F Mische et al., 1987 . Arp2/3# F, G? Beltzner and Pollard, 2004; Mullins et al., 1998 Calponin F Takahashi et al 1986 . CaMKIIB F Shen et al., 1998; Fink et al., 2003 Cofilin F Nishida et al., 1984; Abe et al., 1989; . . Renoult et al 1999 . Cortactin* F Wu and Parsons, 1993; Weed et al., 2000 Hering and Sheng, 2003* Drebrin ? Shirao, 1995; Hayashi et al., 1999 . Filamin F Hartwig and Stossel, 1975 Gelsolin G & F Yin et al., 1988; Renoult et al., 2001; . . Van Troys et al., 1996; Puius et al., 2000 . Myosin V F Cheney et al., 1993 Neurabin I* F Nakanishi et al., 1997; Zito et al., 2004* . NF-L G Hao et al., 1997 # N-WASP G Egile et al., 1999 Profilin G Carlsson et al., 1977 SPAR*^ F Pak et al., 2001* Spectrin (fodrin) G & F Cohen et al., 1980 Spinophilin (neurabin II)* F Satoh et al., 1998; Barnes et al., 2004* Synaptopodin F Kremerskothen et al., 2005; Mundel et al., 1997

62 what happens to myosin when it reaches the barbed end of an actin filament and releases its cargo. Perhaps myosin-V simply “rides” an actin filament back to the pointed end where it can bind another vesicle. Similarly, it is possible that many proteins that are enriched in spines ride a monomer or a small portion of actin filament into the spine.

Conclusions

These experiments indicate a role for protein synthesis and actin expression in dendritic spine proliferation. The acute slice preparations from mature rats demonstrate that neurons maintain plastic capabilities well into adulthood, especially in response to injury. The role of protein synthesis in spine proliferation could reflect a need for specific spine components, the result of a disruption of the steady-state of the neuron, or an increase in actin production which occurs whenever spines begin to grow. The ability of actin to drive spinogenesis is especially exciting because such regulation can rapidly generate new spines, negating all other signaling pathways within the cell. Finally, a novel function for actin is proposed, in which treadmilling actin transports proteins into, out of and within spines.

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80 BIOGRAPHICAL SKETCH

Orenda L. Johnson was born Orenda Lyons in Tallahassee, FL where she lived until graduation from high school in 1993. She attended Stetson University, earning a Bachelor of Science degree in 1997 with a major in Psychology and a minor in Biology. During this time she was awarded a research fellowship under the direction of Dr. Camille T. King, and participated in a summer research program at the University of Florida under the direction of Dr. Alan Spector. Orenda returned to Tallahassee and began graduate school at Florida State University in the fall of 1997. In 2001 Orenda earned a Master’s Degree in Psychology under the direction of Dr. Karen J. Berkley. In January of 2000, Orenda began work on her doctoral research in the laboratory of Dr. Charles C. Ouimet. During her 5 ½ years in the Ouimet lab, Orenda taught undergraduate psychology labs, an undergraduate psychology lecture class, and the medical school neuroanatomy lab. Orenda was awarded two Bryan Robinson Fellowships for her research in the Ouimet lab, and will have two publications based on her doctoral research (listed below).

Publications

Johnson OL and Ouimet CC (under review with J Neurosci) A regulatory role for actin in dendritic spine proliferation.

Johnson, OL and Ouimet CC (2004) Protein synthesis is necessary for dendritic spine proliferation in adult brain slices. Brain Res 996: 89-96.

Johnson, OL and Berkley, KJ. (2002) Estrous influences on micturition thresholds of the female rat before and after bladder inflammation. Am J Physiol Regul Integr Comp Physiol 282: R289-R294.

Dmitrieva N, Johnson OL, Berkley KJ. (2001) Bladder inflammation and hypogastric neurectomy influence uterine motility in the rat. Neurosci Lett. Nov 2;313(1-2):49-52.

81 Seminars

“Reproductive status-dependent variations in turpentine-induced bladder hyper-reflexia in the female rat” Baker Seminar, Florida State University, 1999

“Effect of chorda tympani and glossopharyngeal neurectomy on quinine-induced activation of the NST and PBN” NSF Summer Research for Undergraduates, University of Florida, 1996, and Psy Chi Conference, University of Georgia, 1997

Posters

Johnson O, Dietz K and Ouimet C. Actin has a regulatory role in dendritic spine morphogenesis. Society for Neuroscience Abstracts 2005.

Johnson O and Ouimet C. Protein synthesis is necessary for dendritic spine proliferation in vitro. Society for Neuroscience Abstracts 2002.

Dmitrieva N, Johnson OL and Berkley KJ. Viscero-viseral interactions between the lower urinary and reproductive tracts. Society for Neuroscience Abstracts, 2001. 818.3

Kohut B, Johnson OL, Dmitrieva N, Elam JS, Berkley KJ. Influence of reproductive status on bladder capacity of the inflamed and uninflamed bladder in the female rat. Society for Neuroscience Abstracts, 2001. 842.14

Lyons, O.L. and Berkley, K.J. (1999) Influence of reproductive status on lower urinary tract pathophysiology in the female rat. Society for Neuroscience Abstracts, 674.7.

Lyons, O.L. and Berkley, K.J. (1998) Estrous variation in bladder hyper-reflexia induced by bladder inflammation in the rat. Society for Neuroscience Abstracts, 635.3

Teaching Experience

Sensation and Perception (2005)

Medical Neuroanatomy Laboratory (2002)

Guest lecturer in Medical Neuroanatomy (2002)

Conditioning and Learning Laboratory (2000)

82 Teaching Experience (cont.)

Physiological Psychology Laboratory (1999-2001)

Sensation and Perception Laboratory (1999-2001)

Wrote lab and teaching manuals for Sensation and Perception Laboratory (2000)