Scaffolding Mechanisms Supporting Fast Neurotransmitter Release at the Presynaptic Active Zone

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Citation Held, Richard Guy. 2018. Scaffolding Mechanisms Supporting Fast Neurotransmitter Release at the Presynaptic Active Zone. Doctoral dissertation, Harvard University, Graduate School of Arts & Sciences.

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Scaffolding mechanisms supporting fast neurotransmitter release at the presynaptic active zone

A dissertation presented

Richard Guy Held

The Division of Medical Sciences

in partial fulfillment of the requirements

for the degree of

Doctor of Philosophy

in the subject of

Neurobiology

Harvard University

Cambridge, Massachusetts

August 2018

Dissertation Advisor: Dr. Pascal S. Kaeser Richard Guy Held

Scaffolding mechanisms supporting fast neurotransmitter release at the presynaptic active zone

Abstract

Synaptic vesicle exocytosis from presynaptic nerve terminals occurs with extreme temporal and spatial precision, on the order of microseconds and nanometers. Within the presynaptic terminal, a protein complex known as the active zone enables this and serves as the site of vesicle fusion. Extensive research has identified individual protein components of the active zone and the important roles they play in controlling fusion. Active zone proteins are responsible for ‘docking’ vesicles to the plasma membrane, for generating fusion competent

‘readily releasable’ vesicles, and for positioning those vesicles in close proximity to Ca2+ channels to increase their release probability. Despite this functional knowledge, major questions remain about the structure of the active zone protein complex itself. First, the molecular mechanisms that establish and maintain the structural integrity of the active zone have remained unclear. Second, the scaffolding interactions that are responsible for localizing essential proteins to the active zone are incompletely understood.

This work was designed to address these two questions using gene knockouts of important active zone protein families. I first analyzed the effects of knocking out ELKS, a protein family that may mediate active zone scaffolding. While these studies revealed no independent structural roles for ELKS, I found surprising diversity in the synaptic mechanisms that underlie release deficits in ELKS knockouts. In subsequent work, both ELKS and another active zone protein, RIM, were genetically removed. This resulted in near complete disassembly of the active zone protein complex, revealing surprising, strong scaffolding redundancy between

ELKS and RIM. Finally, I determined how interactions between active zone proteins and

2+ presynaptic Ca channels contribute to active zone scaffolding. Removing all CaV2 channels

iii revealed that CaV2 channels themselves do not scaffold active zone proteins. Instead, multiple redundant interactions of the CaV2 C-terminus tether the channels to release sites. This work highlights a fundamental organizational principle of the active zone: highly redundant protein interactions ensure the structural integrity of the active zone. Such an organizational scheme can increase the functional reliability of this molecular machine and allow specific interactions and functions to be favored depending on the local context, enabling synapse-specific release properties.

Table of contents

Abstract……………………………………………………………………………………………….…...iii

Table of contents……………………………………………………………………………………….…v

Acknowledgments………………………………………………………………………………………..vi

CHAPTER ONE: Introduction………………….……………………………..…………..………....….1

CHAPTER TWO: ELKS controls the pool of readily releasable vesicles at excitatory synapses through its N-terminal coiled-coil domains……………………...………………………………..…..16

CHAPTER THREE: Fusion competent synaptic vesicles persists upon active zone disruption and loss of vesicle docking……………………………………………………………………..…...…50

CHAPTER FOUR: Redundant scaffolding mechanisms target CaV2 family voltage-gated calcium channels to the presynaptic active zone……………..……………………………………..84

CHAPTER FIVE: Discussion and Outlook………….……………………………..…………….….120

APPENDIX ONE: Supplemental Figures.………...………………………………………………...133

APPENDIX TWO: Supplemental Figures.………..…………………………………………………162

References …………………………..…………………..…………………………………………….173

Acknowledgements

I am deeply indebted to many people and institutions for the position I find myself in and any success I may have found along the way. My graduate school experience was shaped by the people that I spent it with. I was fortunate to be in an incoming class filled with people that were simultaneously talented and willing to not take themselves too seriously. In particular,

Rohan Ramesh, Andrea Yung, Chris Tzeng, Ben Andreone, Aurora Zhang, Michael Coulter,

Greg Bryman, and the adopted Brian Chow have been fantastic classmates and better friends.

Their spouses and significant others, in particular Fatima Santiano and John Sapienza, were wonderful additions to our group of friends and extremely patient with all the talk about science.

Rohan has the added distinction of sharing a disastrous apartment with me for five years, for which I am both grateful and apologetic.

I am cognizant of the fact that these people, myself included, were brought together by

Harvard University and the Program in Neuroscience. The chance to work in an environment where you are surrounded by high achieving people is a privilege. Harvard and PiN gave me that chance, a fact that I will remain grateful for. I am also grateful to Benjamin Hall, who gave me my first opportunity to work in his lab at Tulane University and encouraged me to apply to graduate school at Harvard; I thought he was crazy. Chih-Chieh Jay Wang, a graduate student in Ben’s lab and later a post-doc at Harvard, taught me how to work in a lab and let me work on his project as a lowly undergraduate. Without his hard work and accomplishments, I would never have gotten into graduate school in the first place.

I have been equally fortunate in who I shared a lab space with. My fellow Kaeser lab members make the lab a wonderful place, one where you can be both productive and social.

Shan Shan Wang in particular has been the best collaborator I could have asked for. She has graciously accepted my work on her projects and selflessly contributed her work to mine. Our collaboration has hugely enhanced my productivity and I hope she feels I have helped hers.

Sunny Nyitrai, who started at the same time as me, has been an incredibly valuable source of scientific skepticism and rigor that has improved the quality my work, regardless of the time of night. Changliang Liu has served as both a source of pragmatic realism and of creative, sometimes absurd, ideas of both a scientific and non-scientific nature. Lauren Kershberg makes me feel old but also proud that the lab I decided to join continues to attract smart people who are willing to work hard to succeed. Jennifer Wang, and before her Lydia Bickford, keep the lab running and allow us to conduct the type of mouse genetic experiments that labs twice our size would struggle to maintain in their colonies. Postdocs, past and present, including Arthur de

Jong, Man Yan Wong, Aditi Banerjee, and Javier Emperador provide the wisdom and resource of experience. I am also grateful to Shan Shan, Lauren, Lydia, and the once again adopted

Brian Chow for coming to get coffee with me at almost any hour of the day.

My advisor, Pascal Kaeser, has been a phenomenal mentor. He has supported me in different ways as I have developed as a scientist, giving me opportunities to gain experience early on and the independence to explore my ideas in later projects. I have learned a lot from him, but if I had to guess the most valuable lesson, it is that science never ends and you should always be preparing for the project that you will be working on two years from now while finishing the one you are working on today. I am sure that I have not always been an easy person to mentor but he has managed to guide me down interesting paths.

People always say that graduate school is hard and in a quantifiable sense they are right. Long hours, failed experiments, and the unpredictable nature of biology mean that balance can be a difficult thing to find. Subjectively, speaking from my own experience, they are wrong, however. I have worked hard as a graduate student because I wanted to, because I chose this path and find it fulfilling. If I was in lab late on a Saturday night, it was because there were questions to be asked and answers to be found that motivated me to be there. They were not

vii always the ones that the world and the field might find the most compelling; they were just the ones I liked. There is no hardship in that.

For the people whose lives I am a part of, however, there was considerably less choice involved. For them, far more than for myself, graduate school has been difficult at times, as they have shared my time and focus with science. First and foremost among them is my wife Anna.

She has been patient and supportive of a life that is often consumed by lab work and science talk, both in the lab and in social settings. All our friends our scientists; we will do our best to diversify. I struggle to decide whether or not I would waste away or be the size of a bus without her influence, since she encourages both the consumption of real food (with large amounts of butter) and the running of marathons. Most importantly, however, she is the balance and depth in my life. She is the reason I read good books, travel to interesting places, and dream of things like dogs and space. I would be a worse person without her.

The forwarding address for any complaints Anna may have about me directs to my family, who made me who I am before she had any choice in the matter. My mother Lani and my father Bruce are the reasons for my intellectual curiosity and my work ethic. I will let them decide who wants to take credit for each and whether that is something they should be proud of.

My older sister Nikki, my first friend, is the example of brilliance, drive, and success that I will always be forced to chase. To paraphrase another, she is better than me and I know she is better than me, but I will try to keep up regardless. John, her husband, is also acceptable. I think very highly of him, but to say more would be too earnest. My younger brother Sasha is the challenger I will always be forced to run from, a race I may already have lost. Whenever I want to be lazy, the knowledge that he is not keeps me going. They are all singular people and I am fortunate that blood and marriage has placed me in their company.

These acknowledgements could have been much shorter but equally appropriate if they had simply read, “thank you to everyone who makes it possible for me to do what I want to do.”

viii

There are many more people who fall into that category not named here. I am extremely grateful to them all. It is not an exaggeration to say that every day I feel the influence of all the people who have supported me, pushing me forward in science and in life. I am extremely privileged to have their influence in my life. The only appropriate thanks I can imagine is to keep moving forward.

— CHAPTER ONE —

Introduction

Synaptic transmission involves the transfer of information between a presynaptic neuron and its postsynaptic target cell. When a neuron is sufficiently depolarized, it fires an action potential that propagates down the axon into the presynaptic nerve terminals (Hodgkin and

Huxley 1939). Depolarization by the arriving action potential opens voltage-gated Ca2+ channels

(VGCCs), allowing influx of Ca2+ into the terminal (Schneggenburger, Han, and Kochubey

2012). In response to Ca2+ entry, membrane-bound synaptic vesicles fuse with the plasma membrane, releasing chemical neurotransmitters (Südhof 2013). Once released, they diffuse across the extracellular space to bind to receptors on the postsynaptic target. These receptors are often ligand-gated ion channels that, once bound to an appropriate neurotransmitter, allow the passage of ions into or out of the postsynaptic cell, further propagating the signal. While the exact details of this process vary widely depending on the type of synapse in question, this general sequence of synaptic transmission is broadly conserved throughout the central (CNS) and peripheral nervous system (PNS) in organisms as diverse as nematodes and humans.

Hallmarks of synaptic vesicle fusion are its speed, spatial precision, and ability to undergo diverse forms of plasticity. These properties are controlled by a specialized protein complex known as the active zone. At the active zone, synaptic vesicles are docked in close proximity to the presynaptic plasma membrane, brought into a fusion competent ‘primed’ state, and positioned in close proximity to the molecular machinery responsible for fusion (Südhof

2012). This helps to accelerate the rate of fusion and ensure that it occurs within a defined region of the plasma membrane.

The active zone has been defined in a variety of ways over time. It was first defined functionally as the site of exocytosis of synaptic vesicles, hence ‘active zone’ (Couteaux and

Pécot-Dechavassine 1970). Later, it was morphologically described as an electron dense structure on the presynaptic membrane directly opposed the the postsynaptic density

(Pfenninger et al. 1969). More recently, extensive work using biochemical purifications and genetic experiments in invertebrates and vertebrates has led to a molecular definition of the

2 active zone as a protein complex composed of a network of multi-domain proteins (Zhai and

Bellen 2004; Schoch and Gundelfinger 2006). At small synapses in the vertebrate CNS, such as those that release glutamate and GABA, this complex contains six protein families that are relatively restricted in their localization to the active zone: ELKS, RIM (Rab3 interacting molecule), RIM-BP (RIM-binding protein), Munc13 (mammalian Unc13), Liprin-α, and

Piccolo/Bassoon. In this work, I define the active zone as the protein complex that is attached to the presynaptic plasma membrane opposed to the postsynaptic density.

Biophysical descriptions of synaptic transmission

Prior to the advent of modern molecular biology, the pioneering work of Bernard Katz in the mid-twentieth century began to establish a biophysical model of synaptic transmission. Katz and colleagues, working at the neuromuscular junction (NMJ) and later at the squid giant synapse, developed key hypotheses about the nature of the communication between the presynaptic nerve terminal and the postsynaptic muscle fiber. The first was the quantal hypothesis, which theorized that the electrical response of the muscle, known as an end plate potential (EPP), to a presynaptic action potential was made up of the summation of smaller discrete ‘quantal’ events (del Castillo and Katz 1954; Fatt and Katz 1952). Before there was any knowledge of the existence of synaptic vesicles, they proposed that these quantal events reflected the release of packets of neurotransmitter and that the action-potential evoked EPP was caused by the synchronized release of many such packets. Not long after, the first electron microscope images of synapses revealed the presence of synaptic vesicles, the physical correlate of Katz’s quanta (Sanford L Palay 1955; De Robertis and Bennett 1955)}.

The second related hypothesis was that Ca2+ ions in the extracellular solution, were required to trigger release of these quanta, or vesicles (Katz and Miledi 1967; Katz and Miledi

1966; Llinás, Steinberg, and Walton 1976). By decreasing extracellular Ca2+, EPP amplitude was gradually decreased until the amplitude of the evoked EPP matched that of the miniature

3 events. Statistical analysis of the proportion of failures at low extracellular Ca2+ concentration suggested that the mean number of quanta in an EPP matched that predicted by a low probability Poisson process (del Castillo and Katz 1954). They proposed a model in which vesicles inside the presynaptic nerve terminal could only fuse at discrete ‘reactive sites’ on the plasma membrane (Katz 1969). The amplitude of the EPP would then be determined by the number of reactive sites, N, the size of an individual quanta, q, (i.e. the response to release of a single vesicle), and the average probability that a vesicle is released at each release site, p;

(EPP = Npq). Since increasing extracellular Ca2+ increased the EPP amplitude without changing the size of miniature EPPs (i.e. single quanta), either the number of release site, N, or the release probability, p, was increased by elevated Ca2+.

This model, though it has been modified in many ways, still influences how the field thinks about synaptic transmission. Synaptic vesicle fusion at many synapses is well described as a binomial process where a number of discrete release sites act independently to fuse vesicles (Stevens 2003; Malagon et al. 2016). As knowledge of the physical organization of the synapse has progressed, physical and molecular details have been fit in to this model. We know that Katz’s quanta are synaptic vesicles and that the size of the quantal response (q) is determined by the amount of neurotransmitter in each vesicle and by the number of postsynaptic receptors available to be activated by the released neurotransmitter (Takamori

2016; Raghavachari and Lisman 2004). We also know that release probability (P) is generally dictated by the probability that Ca2+ ions coming into the cell encounter the Ca2+ sensor proteins that trigger fusion (Eggermann et al. 2011; Südhof and Rizo 1996). The exact molecular definition of release sites (N), however, is still unclear.

Extensive work has demonstrated that a subset of the total number of vesicles at the synapse constitutes a readily releasable pool (RRP), which is available to be released in response to a single action potential (Kaeser and Regehr 2017). The total number of vesicles released is therefore proportional to the size of the RRP times the probability of release (P) of

RRP vesicles. This relationship, release = RRP x P, originates from Katz’s original model, but exactly how the RRP relates to Katz’s release sites requires a more detailed understanding of the molecular markers of each. The early functional descriptions and biophysical models of synaptic transmission have shaped how we study and understand the synapse for decades. We are now at the point where we can relate this work to our increasing knowledge about the proteins and structure of synapses and the active zone.

Conserved proteins for membrane fusion

At its core, presynaptic neurotransmitter release is a membrane fusion reaction.

Membrane fusion occurs in all eukaryotic cell types and in many different compartments throughout the cell (Wickner and Schekman 2008). The process of fusing two membranes requires three categories of highly conserved proteins: SNARE proteins, S/M proteins, and Rab

GTPases (Jahn, Lang, and Südhof 2003; Südhof and Rothman 2009). In addition, many fusion reactions, including synaptic vesicle fusion, are controlled and regulated by Ca2+ binding to Ca2+ sensor proteins (de Wit et al. 2009; Voets et al. 2001; Pang and Südhof 2010). This creates a need for a Ca2+ source, since intracellular Ca2+ concentrations are tightly maintained in the nanomolar range (Clapham 2007).

SNARE proteins, for Soluble N-ethylmaleimide-sensitive factor Attachment protein

Receptors, are a conserved superfamily characterized by the presence of a heptad repeat containing SNARE motif and, typically, a link to the membrane via either a transmembrane domain or post-translational membrane anchor (Kloepper, Kienle, and Fasshauer 2008).

SNARE proteins can interact with one another via their SNARE motifs, which assemble to form an extremely stable four -helix coiled-coil bundle known as a SNARE complex (Sutton et al.

1998; Söllner, Bennett, et al. 1993; Söllner, Whiteheart, et al. 1993). Each SNARE motif is classified as either a QA, QB, QC, or R-SNARE based on its contribution of either one of three glutamine (Q) residues or an arginine (R) to the core ‘0’ layer of the assembled SNARE complex

(Fasshauer et al. 1998). When SNARE proteins are oriented in a trans configuration, on opposing membranes, the correct parallel formation of this four-helix SNARE complex can provide the energy to fuse the two membranes (Hu et al. 2003; Weber et al. 1998). In the case of synaptic vesicle fusion, the synaptic vesicle SNARE synaptobrevin (or VAMP2) contributes the R-SNARE, while the plasma membrane SNAREs syntaxin-1 and SNAP-25 contribute the QA and QB/C-SNAREs, respectively (Sutton et al. 1998; Fasshauer et al. 1998).

While SNARE complex formation is thought to provide the energy for membrane fusion, other protein families are also essential. S/M (Sec1/Munc18) proteins are another conserved protein family essential for vesicle fusion (Verhage et al. 2000). Munc18-1, the main isoform involved in synaptic vesicle fusion, binds to the SNARE protein syntaxin-1 and the assembled

SNARE complex in several different conformations (Südhof and Rothman 2009). Syntaxin-1 contains an N-terminal Habc domain, which consists of a three-helix bundle that binds to the syntaxin SNARE motif to generate a ‘closed’ conformation (Dulubova et al. 1999; Richmond,

Weimer, and Jorgensen 2001). Munc18-1 has a curved, clamp-like structure that can bind to this closed conformation of syntaxin, thus stabilizing it (Misura, Scheller, and Weis 2000; Ma et al. 2013). A second interaction between an N-terminal peptide motif in syntaxin and a hydrophobic pocket on Munc18 is thought to tether Munc18-1 to syntaxin (Burkhardt et al.

2008). Finally, Munc18-1 can bind to the assembled SNARE complex in a similar manner to closed syntaxin, suggesting that a primary function of SM proteins is to stabilize four-helix bundles (Dulubova et al. 2007). Interactions between Munc18-1 and syntaxin-1 are also essential to the stability of both proteins, as knockout of either results in loss of expression of the other (Bouwman et al. 2006; Gerber et al. 2008). Munc18-1, and SM proteins more broadly, are therefore likely serve multiple important roles in the trafficking, stability, and function of

SNARE proteins.

Finally, many membrane trafficking and fusion events require a specific Rab GTPase

(Jahn and Südhof 1999). Rabs are thought to facilitate attachment and detachment of vesicles

6 and other organelles to their target membranes by cycling GTP/GDP (Geppert, Bolshakov, et al.

1994). In yeast, loss of the appropriate Rab typically blocks the fusion process (Salminen and

Novick 1987). Results in mammalian cells have been less clear, however. Many Rabs are expressed at synapses, and the primary candidate for targeting synaptic vesicles to fusion sites was long thought to be Rab3 (C. Li et al. 1994; Fischer von Mollard et al. 1994; Kobayashi et al.

2016). However, knockout of all isoforms of Rab3 does not abolish synaptic transmission, suggesting that other Rabs may be important (Schlüter et al. 2004) .

Ca2+ triggered fusion

Beyond these core components, the proteins involved in fusion begin to specialize. The characteristics of membrane fusion reactions can be markedly different depending on the cellular context in question. They can occur over time scales of minutes to milliseconds, across broad swathes of membrane or in strictly localized regions, and either as constitutive processes or highly time-locked events that can be rapidly triggered (Wickner and Schekman 2008).

Synaptic vesicle fusion in particular is exceptionally fast and spatially precise. A major reason for this is the strict requirement for Ca2+ in order to trigger fusion.

Typical intracellular Ca2+ concentrations are in the low nM range, and cells maintain strict Ca2+-ion homeostasis (Clapham 2007). Elevations in intracellular Ca2+ concentration are therefore an ideal triggering mechanism for membrane fusion, as they are both spatially and temporally limited (Meinrenken, Borst, and Sakmann 2003). At the synapse, VGCCs serve as the source of Ca2+ to trigger fusion (Llinás, Steinberg, and Walton 1976; Llinás et al. 1989;

Hirning et al. 1988; Gasparini, Kasyanov, Pietrobon, Voronin, and Cherubini 2001a).

Vertebrates have ten genes for VGCCs, but, of these, the CaV2 family are the primary channels responsible for synaptic vesicle fusion (Dolphin 2016). CaV2 channels are tightly localized to the active zone in close proximity to docked synaptic vesicles (Eggermann et al. 2011; Holderith et al. 2012; Adler et al. 1991). When they open during an action potential, Ca2+ influx through the

7 channel can trigger fusion of vesicles which are ‘coupled’ to the channel at a distance of between 30 to 100 nanometers, depending on the synapse (Arai and Jonas 2014; Vyleta and

Jonas 2014; Meinrenken, Borst, and Sakmann 2002). The number of channels required to fuse a single vesicle, known as the channel cooperativity, also varies from as few as one channel to dozens (Stanley 1993; Matveev, Bertram, and Sherman 2009; Mintz, Sabatini, and Regehr

1995). These strict spatial relationships are key determinants of the vesicular probability of release (Nakamura et al. 2015; Keller et al. 2015; Meinrenken, Borst, and Sakmann 2003).

Once VGCCs are opened, Ca2+ sensor proteins are then required to detect the influx of

Ca2+ and either initiate or permit fusion to proceed. They are often C2-domain containing proteins that bind multiple Ca2+ ions cooperatively (Rizo and Südhof 1998). This Ca2+ cooperativity leads to a steep dependence of release on the concentration of Ca2+, typically estimated at a fourth-power relationship (Dodge and Rahamimoff 1967; Schneggenburger and

Neher 2000). Synaptotagmins are the primary Ca2+ sensor of this type in synapses (Brose et al.

1992; Geppert, Goda, et al. 1994; Fernandez-Chacon et al. 2001). Synaptotagmin-1 (Syt-1), a transmembrane vesicle protein with tandem C2 domains (C2A and C2B), is the Ca2+ sensor for fast release at most CNS synapses (Südhof and Rizo 1996). The C2 domains of Syt-1 are able to bind Ca2+, membrane phospholipids, and the assembled SNARE complex (Söllner, Bennett, et al. 1993; Zhou et al. 2015; Fernandez et al. 2001). While the exact mechanism by which Syt-

1 triggers fusion is still debated, it likely involves Ca2+ induced conformational changes in the interactions between the Syt-1, SNARE proteins, and the membrane reducing the overall energy barrier to membrane fusion (Martens, Kozlov, and McMahon 2007; Vrljic et al. 2010;

Giraudo et al. 2006). Other Ca2+ sensor proteins operate at the synapse, but their affinity and cooperativity of for Ca2+ binding are tailored to their specific functions (Groffen et al. 2010;

Jackman et al. 2016).

The active zone protein complex

The core fusion machinery, VGCCs, and synaptotagmins mediate exocytosis in many secretory cells where fusion is less spatially restricted and executed on much slower time scales

(de Wit et al. 2009; Voets et al. 2001; Neher 2017; Wickner and Schekman 2008). Additional protein machinery is therefore required to account for the speed and spatial precision of synaptic signaling. Remarkably, at physiological temperatures, the latency between Ca2+ influx and the postsynaptic response, which includes vesicles fusion, neurotransmitter diffusion, and opening of postsynaptic receptors, can be as fast as 60 s (Sabatini and Regehr 1996).

Furthermore, vesicle fusion within synapses preferentially occurs within nanodomains of roughly

80 nm (Tang et al. 2016). Such precision is possible due to the proteins of the active zone:

Munc13, RIM, ELKS, RIM-BP, Liprin-α, and Piccolo/Bassoon.

Striking work from many laboratories has demonstrated that individual active zone proteins support important functions in release. Munc13, for example, is essential for generating primed RRP vesicles. Munc13 knockout neurons show a complete loss of synaptic vesicle fusion due to a loss of RRP vesicles (Augustin, Rosenmund, et al. 1999; Varoqueaux et al.

2002; Aravamudan et al. 1999; Richmond, Davis, and Jorgensen 1999). Morphologically,

Munc13 is required for tight docking of synaptic vesicles, and in its absence vesicles accumulate roughly 10 nm back from the membrane (Imig et al. 2014). Munc13 interacts with

SNARE proteins through a central motif called the MUN domain (Basu et al. 2005; Yang et al.

2015; Ma et al. 2013). This interaction is able to displace Munc18-1 from the closed conformation of syntaxin and promotes proper SNARE complex assembly (Ma et al. 2011; Lai et al. 2017). These functions combined with the loss of vesicle docking and fusion have led to a model in which primed, RRP vesicles are docked vesicles in which Munc13 has facilitated proper trans SNARE complex assembly with plasma membrane SNAREs.

RIM proteins are also critically important for vesicle fusion. RIM knockouts result in both a decrease in RRP vesicles and reductions in presynaptic Ca2+ influx (Koushika et al. 2001;

Schoch et al. 2002; Kaeser et al. 2008; Kaeser et al. 2011; Han et al. 2011; Calakos et al.

2004). RIM interacts with an N-terminal C2 domain in Munc13 to anchor it to the active zone and to activate its priming function (A. Betz et al. 1997; Andrews-Zwilling et al. 2006; Dulubova et al. 2005; Deng et al. 2011). Through its PDZ domain, RIM can also bind to the C-terminal tail of certain CaV2 VGCCs to promote recruitment of VGCCs to the active zone (Kaeser et al.

2011; Han et al. 2011). RIM has known biochemical interactions with almost every other active zone protein, including Munc13, ELKS, Liprin-, RIM-BP, and VGCCs (Yun Wang et al. 2002;

Dulubova et al. 2005; Schoch et al. 2002; Kaeser et al. 2011; Yun Wang, Sugita, and Südhof

2000). Additionally, RIM has recently been shown to bind to the membrane lipid PIP2, an interaction that is essential for its functions in release (de Jong et al. 2018). These data suggest that RIM may serve a scaffolding function at the active zone.

Similar to RIM, ELKS proteins interact with almost every other active zone protein in in vitro assays (X Wang et al. 2009; Kiyonaka et al. 2012; Ohtsuka et al. 2002; Yun Wang et al.

2002; Ko et al. 2003; Kawabe et al. 2017). Unlike RIM, ELKS proteins are coiled-coil proteins with no clear domain structure (Yun Wang et al. 2002; Ohtsuka et al. 2002; Deguchi-Tawarada et al. 2004). Knockout of the major isoforms of ELKS decreases synaptic transmission by roughly half, far more mildly than either Munc13 or RIM (C. Liu et al. 2014). Due to its plethora of protein interactions, ELKS has also been proposed as a central active zone scaffold, but there is mixed evidence for this at vertebrate synapses (tom Dieck et al. 2012; C. Liu et al.

2014). In invertebrates, ELKS is not necessary for synapse structure in C. elegans, but the T- bar structure of the D. melanogaster active zone is completely abolished by knockout of the partial ELKS homolog Bruchpilot (Deken et al. 2005; Kittel et al. 2006). The structural and functional roles of ELKS proteins are therefore incompletely understood and may vary across different synapses (see Chapter 2).

RIM-BP, together with RIM, plays a role in the positioning of Ca2+ channels. RIM-BP contains three SH3 domains and three fibronectin II domains (Yun Wang, Sugita, and Südhof

2000; Mittelstaedt and Schoch 2007). The SH3 domains can bind the PxxP motifs in both RIM and the C-terminal tail of CaV2 channels, potentially forming a tripartite complex to tether channels (Hibino et al. 2002). In vertebrates, RIM-BP knockout neurons show relatively mild effects on synaptic vesicle fusion, including an increase in the coupling distance between

VGCCs and vesicles and a decrease in the reliability of release (Grauel et al. 2016; Acuna et al.

2015). How exactly RIM and RIM-BP might work together to localize VGCCs remains an interesting question (see Chapter 4).

Bassoon/Piccolo proteins are the largest of the active zone proteins and perhaps the most difficult to study. They are massive ~400 kD multi-domain proteins which, similar to ELKS and RIM, can bind in vitro to almost every other active zone protein (tom Dieck et al. 1998; X

Wang et al. 2009; X Wang et al. 1999). Structurally, both Bassoon and Piccolo contain N- terminal Zinc finger domains and large coiled-coil stretches. In addition, Piccolo has a PDZ domain and two C2 domains. Studies of Bassoon/Piccolo have been complicated by the large size of the proteins and difficulties with gene targeting leading to partial knockout of the protein

(Altrock et al. 2003; Mukherjee et al. 2010). With those complications in mind, these studies have not shown dramatic effects on basal synaptic transmission at central synapses, but suggest that Bassoon is involved in the capture and recycling of synaptic vesicles (Hallermann,

Fejtova, et al. 2010; Mukherjee et al. 2010).

Finally, Liprin-‘s are perhaps the least well understood protein families at the vertebrate active zone. In invertebrates Liprin- controls active zone composition and vesicle accumulation in presynaptic terminals, placing it upstream of other active zone proteins in synapse assembly

(Dai et al. 2006; Kaufmann et al. 2002; Zhen and Jin 1999). Vertebrates express four Liprin- genes, of which Liprin-2 and Liprin-3 are the primary isoforms in the brain (Serra-Pagès et al.

1998; Zürner et al. 2011; Zürner and Schoch 2009; Spangler et al. 2011). They were hypothesized to be active zone proteins due to their interactions with RIM and ELKS, but their

11 expression patterns were consistent with both pre- and postsynaptic roles (Ko et al. 2003;

Schoch et al. 2002; Spangler et al. 2011; Zürner et al. 2011; Wyszynski et al. 2002). However, recent studies have demonstrated that Liprin-3 is localized at the active zone using super- resolution microscopy and revealed presynaptic roles in exocytosis for both Liprin-2 and

Lipring-3 (M. Y. Wong et al. 2018; Spangler et al. 2013). A more clear understanding of Liprin-

’s role at the vertebrate active zone will require more comprehensive genetic tools.

Scaffolding architecture of the active zone

While extensive work has gone into identifying the proteins of the active zone and elucidating their functional roles in vesicle fusion, much less is known about the scaffolding and structural organization of the active zone protein complex itself, in particular at small CNS synapses. Central questions remain unresolved, including how and where the active zone is assembled, what holds this protein complex together, what links it to the plasma membrane, and what positions it in opposition to postsynaptic receptors. These questions all relate to protein scaffolding. In the context of the active zone, I define protein scaffolding as the capture, anchoring, and spatial organization of the molecular machinery for synaptic vesicle fusion.

The highly interconnected nature of the active zone has made the study of the molecular mechanisms of protein scaffolding difficult to address. Just considering the main active zone proteins, each one binds to at least two other active zone proteins and likely many more proteins not considered part of the core active zone complex. It is perhaps not surprising then that no single protein knockout results in disassembly of the active zone at vertebrate CNS synapses. Individual knockouts may disrupt parts of the active zone; RIM knockout causes a loss of Munc13 and a slight increase in ELKS solubility, while Bassoon knockouts decrease synaptic RIM-BP levels (Deng et al. 2011; Davydova et al. 2014; Schoch et al. 2002). The overall structure of the active zone at vertebrate synapses, however, appears to be robust to

12 structural perturbations, at least on the scale at which we can detect them. The nematode C. elegans seems to share this robustness. Both RIM and ELKS are localized to the synapse independently of other active zone proteins, though expression of a mislocalized fragment of

RIM disrupts ELKS localization (Deken et al. 2005).

Some synapses, in particular the NMJ of the fly Drosophila melanogaster, appear to be easier to disrupt. The fly NMJ forms characteristic T-shaped electron-dense structures known as

T-bars which tether synaptic vesicles and mark the active zone. Drosophila expresses a partial homolog of ELKS, bruchpilot (brp), which shares sequence similarity with ELKS in its N-terminal half but has an unrelated C-terminal portion (Monier et al. 2002; Wagh et al. 2006). Loss of Brp leads to a complete loss of T-bar structures at the NMJ and a decrease in the expression of tagged, overexpressed VGCCs (Kittel et al. 2006; Fouquet et al. 2009). Drosophila RIM-BP

(DRBP) null mutants also disrupt the T-bar and almost completely abolish release, a much more severe phenotype than vertebrate RIM-BP knockouts (K. S. Y. Liu et al. 2011; Acuna et al.

2015; Grauel et al. 2016).

Ribbon synapses in photoreceptors and auditory hair cells also seem to share this sensitivity. Similar to the T-bar, synaptic ribbons are elongated electron-dense structures that sit along the plasma membrane at the active zone and accumulate vesicles (Prescott and Zenisek

2005). They are thought to facilitate high rates of vesicle fusion at these synapses, which operate in a tonic fashion rather than releasing vesicles time-locked to action potentials

(Mennerick and Matthews 1996). These synapses express a specialized active zone protein

RIBEYE, which is essential for the ultrastructure of the electron-dense ribbon (Lv et al. 2016).

Loss of CaV1.4 VGCCs or their auxiliary 24 subunits also disrupts the ribbon at rod photoreceptors, resulting in reduced ribbon density (Yuchen Wang et al. 2017). The active zone of ribbon synapses, or at least the ribbon structure itself, is therefore far more susceptible to structural perturbations.

Role of the active zone in synaptic plasticity

Strong disruptions of synaptic transmission, such as would be expected from structural disruption of the active zone, typically result in lethal phenotypes (C. Liu et al. 2014; Schoch et al. 2001). However, understanding the molecular mechanisms of protein scaffolding at the active zone of small CNS synapses has implications for our understanding of the nervous system that go beyond its necessity for survival. These synapses make up a vast majority of the synapses in the brain and are the underpinning of neuronal circuit function throughout the CNS.

Evidence for the importance of the active zone can be found in the range of developmental disorders with genetic links to active zone proteins, indicating that even perturbations that might be subtle in an experimental context can have profound impacts (Silva et al. 2014; Thevenon et al. 2013).

Synapses are far more than just information relay stations. Through a diverse array of different plasticity mechanisms, synapses can modify their strength in response to prior activity and extracellular cues (Zucker and Regehr 2002). Neurotransmitter release can be facilitated by or depressed during repetitive activity; the sensitivity of the postsynaptic cell to neurotransmitter can be up- or down-regulated; and similar processes can occur either transiently on the order of milliseconds to seconds or as lasting changes. Presynaptic plasticity can be controlled both by changing the size of the RRP and by modifying release probability, allowing synapses to tune their output to select for certain information (Fioravante and Regehr 2011). Synapses are therefore arguably the most basic computational unit in the brain (Abbott and Regehr 2004).

The active zone plays a major role in controlling many forms of plasticity. RIM is essential for a presynaptic form of long-term potentiation, for example, and expression of different isoforms of Munc13 can confer dramatically different short-term plasticity phenotypes in cultured hippocampal neurons (Rosenmund et al. 2002; Castillo et al. 2002). Munc13 isoforms are differentially expressed throughout the brain and at different synapses, indicating that

14 synapses can modify their release properties to suit their circuit function (Augustin, Betz, et al.

1999) (see Chapter 2). Postsynaptic sources of plasticity can also require active zone proteins as seen in the C. elegans where VGCCs are inhibited by a retrograde signal from the postsynaptic muscle (Tong et al. 2017). Dynamic structural processes can also play a role. The clustering and alignment of active zone proteins with postsynaptic receptors and the surface mobility of presynaptic VGCCs have both been proposed to effect synaptic plasticity (Schneider et al. 2015; Tang et al. 2016). The active zone is therefore a key locus of synaptic plastic plasticity, with both the protein composition and structural arrangement of the active zone protein complex serving as points of regulation.

The goal of the presented work was to determine how protein scaffolding at the active zones of small CNS synapses controls the functions of the active zone protein complex as a whole. To address this question, I have employed gene knockouts for several active zone protein families to determine their structural necessity at the active zone. This work has revealed that a hallmark feature of the active zone is scaffolding redundancy. This applies to both multiple protein isoforms within the same protein family but also, surprisingly, across protein families. Only using compound gene knockouts for multiple active zone proteins and entire protein families can scaffolding redundancy be circumvented. This has implications for the fidelity of synaptic transmission and the role of active zone proteins in supporting diverse types of synaptic plasticity.

— CHAPTER TWO —

ELKS controls the pool of readily releasable vesicles at excitatory synapses through its

N-terminal coiled-coil domains

Richard G. Held, Changliang Liu, and Pascal S. Kaeser.

C.L performed and analyzed calcium imaging experiments. R.G.H performed all other experiments. This chapter was reproduced with permission from eLife (2016),

DOI: 10.7554/eLife.14862.

Introduction

Within a presynaptic nerve terminal, synaptic vesicle exocytosis is restricted to sites of neurotransmitter release called active zones. The active zone is a dense protein complex that is attached to the presynaptic plasma membrane and is exactly opposed to postsynaptic receptors

(Couteaux and Pécot-Dechavassine 1970; Schoch and Gundelfinger 2006; Südhof 2012). At the active zone, a small subset of the synaptic vesicles are primed in close proximity to presynaptic Ca2+ channels such that the incoming action potential leads to neurotransmitter release with minimal delay. The proteins of the active zone control the size of this pool of primed, readily releasable vesicles (RRP) and the release probability of those vesicles in response to an action potential (Kaeser and Regehr 2014; Alabi and Tsien 2012). Vesicular release probability, referred to in this paper as P, and RRP size together act to set synaptic strength, sometimes referred to as the synaptic probability of release (Stevens 2003; Zucker and Regehr 2002). It is well known that RRP size and vesicular release probability differ across synapses, contributing to the generation of unique release properties (Abbott and Regehr 2004).

In the hippocampus, for example, excitatory and inhibitory synapses have markedly different properties. The underlying molecular mechanisms that control RRP and P are still only partially understood, and it is not known what components of the release machinery account for their synapse specific control.

ELKS, RIM, Munc13, RIM-binding protein (RIM-BP), Bassoon/Piccolo, and Liprin- proteins form a protein complex that defines the active zone (Schoch and Gundelfinger 2006;

Südhof 2012). This protein complex includes many additional proteins that are not active zone specific (Boyken et al. 2013; C. S. Müller et al. 2010; Schoch and Gundelfinger 2006). ELKS

(also called Erc, CAST, and Rab6IP2) was identified as an active zone protein through its interactions with RIM and named for its high content in the amino acids E, L, K, and S (Nakata et al. 1999; Yun Wang et al. 2002; Monier et al. 2002; Ohtsuka et al. 2002). ELKS has known in

17 vitro interactions with many active zone proteins and additional neuronal proteins (Figure 2.1 A).

It contains a C-terminal region that binds to the PDZ domain of RIM (Yun Wang et al. 2002;

Ohtsuka et al. 2002) and multiple coiled-coil stretches, which we have subdivided based on homology between the various vertebrate and invertebrate ELKS isoforms into four coiled-coil domains (CCA-CCD, Figure 2.1 A). The coiled-coil stretches bind in vitro to Liprin- (CCA-CCC,

2+ (Ko et al. 2003)), Bassoon (CCC, (Takao-Rikitsu et al. 2004)), and -subunits of Ca channels

(CCD, (Kiyonaka et al. 2012)). Vertebrate genomes contain two genes for ELKS, Erc1 and Erc2

(Yun Wang et al. 2002), whereas C.elegans expresses a single ELKS homolog (Deken et al.

2005). D. melanogaster expresses a protein called Brp with homology to ELKS in the N-terminal but not the C-terminal half (Wagh et al. 2006; Kittel et al. 2006; Monier et al. 2002). Vertebrate

ELKS proteins are expressed as predominant, synaptic -isoforms and shorter -variants, which account for less than 5% of ELKS (Kaeser et al. 2009; C. Liu et al. 2014). In addition, ELKS C- terminal variants determine RIM-binding: the B-isoforms are prominently expressed in the brain and contain the RIM binding site, whereas A-isoforms are expressed outside the brain and lack

RIM binding (Yun Wang et al. 2002; Kaeser et al. 2009).

The observation that ELKS binds to several active zone proteins has led to the hypothesis that ELKS scaffolds other active zone proteins, in particular RIM (Takao-Rikitsu et al.

2004; Ohtsuka 2013; Ohtsuka et al. 2002). Invertebrate studies offer mixed support to this hypothesis. Loss of Brp disrupts the T-bar structures at the fly neuromuscular junction (Kittel et al. 2006), but this function involves the C-terminal region of Brp (Fouquet et al. 2009). In contrast, C. elegans ELKS is not required for recruitment of other active zone proteins (Deken et al. 2005), but a gain of function mutation in syd-2, the C. elegans Liprin- homologue, requires

ELKS for its synaptogenic activity (Dai et al. 2006).

Relatively little is known about the role and molecular mechanisms of ELKS in neurotransmitter release at vertebrate synapses. Previous studies showed that ELKS1/2

18 boost Ca2+ influx at inhibitory synapses, whereas ELKS2 has a regulatory, non-essential function in RRP at these synapses (C. Liu et al. 2014; Kaeser et al. 2009). In contrast, at excitatory ribbon synapses of rod photoreceptors, ELKS2/CAST may have a structural role to enhance synaptic transmission (tom Dieck et al. 2012). These studies suggest the interesting possibility that ELKS may have differential roles in synaptic transmission between synapses.

Thus far, ELKS functions have not been studied at small, excitatory synapses, the most abundant synapses in the vertebrate brain. Here, we establish that ELKS is prominently expressed at excitatory hippocampal synapses. In contrast to its roles in enhancing Ca2+ influx and release probability at inhibitory synapses, ELKS1/2 control the size of the RRP at excitatory hippocampal synapses. These data establish that removal of ELKS has differential, synapse-specific effects on RRP size and P: boosting RRP at excitatory synapses but enhancing P at inhibitory synapses. Using structure-function rescue experiments, we then determine the sequences within ELKS required for RRP enhancement at excitatory synapses.

Surprisingly, RIM-binding sequences are dispensable for this function, but CCA-CCC, which include binding sites for Liprin- and Bassoon, control excitatory RRP size. Together, these data show that ELKS selectively controls RRP at excitatory synapses through its N-terminal protein interaction motifs.

Experimental Procedures

Mouse lines. All animal experiments were performed according to institutional guidelines at

Harvard University. Conditional double knockout (cDKO) mice that remove the ELKS1/2 proteins were generated by crossing conditional Erc1 ((C. Liu et al.

2014)RRID:IMSR_JAX:015830) and conditional Erc2 ((Kaeser et al. 2009)

RRID:IMSR_JAX:015831) mice. ELKS1/2 cDKO mice were maintained as double homozygote line.

Generation of antibodies. ELKS2 specific antibodies were raised in rabbits using an ELKS2 peptide (109LSHTDVLSYTDQ120). Peptides were synthesized and conjugated to keyhole lympet hemocynanin (KLH) via an N-terminal cysteine residue. Rabbits were inoculated with KLH- conjugated ELKS2 peptides and given booster injections every two weeks. Bleeds were collected every three weeks and screened against protein samples harvested from cultured neurons, brain homogenate, and transfected HEK cells expressing either ELKS1B or

ELKS2B. -actin was used as a loading control. The serum with the highest immunoreactivity

(rabbit 1029, bleed 5) against ELKS2 was affinity purified with the ELKS2 peptide coupled to an affinity column and used at 1:100 dilution.

Cell cultures and lentiviral infection. Primary mouse hippocampal cultures from newborn pups were generated as previously described (Kaeser et al. 2008; Maximov et al. 2007; Kaeser et al. 2011). All lentiviruses were produced in HEK293T cells by Ca2+ phosphate transfection.

Neurons were infected with viruses that express cre recombinase or an inactive cre truncation mutant under the human synapsin promoter (C. Liu et al. 2014). Neuronal cultures were infected with 125 - 250 l of HEK cell supernatant at 3 - 5 days in vitro (DIV). Infection efficiency was monitored by an EGFP tag attached to nuclear cre, and only cultures in which no non-infected cells were found were used for experiments. Expression of rescue proteins was achieved with a second lentivirus driven in neurons by a human synapsin promoter and applied to the neurons at DIV 3. Expression of rescue proteins was monitored by Western blotting and by immunostaining as described below.

Immunofluorescence stainings and confocal imaging of cultured neurons. Neurons were fixed in 4% paraformaldehyde/phosphate-buffered saline, permeabilized in 0.1% Triton X-

100/3% bovine serum albumin/phosphate-buffered saline, and incubated in primary antibodies overnight. The following primary antibodies were used: E-1 (1:500, RRID:AB_10841908), mouse monoclonal antibody, binds to ELKS1 isoforms (ELKS1B, ELKS1A); 1029 (1:100), custom rabbit polyclonal antibody generated against ELKS2B(109LSHTDVLSYTDQ120); mouse anti-RIM (1:500, RRID:AB_10611855); rabbit anti-Munc13-1 (1:5000; a gift from Dr. Nils Brose); mouse anti-Bassoon (1:500, RRID:AB_11181058); rabbit anti-RIM-BP2 (1:500, SySy,

#316103); rabbit anti-VGAT (1:500, RRID:AB_887869); guinea pig anti-VGAT (1:500,

RRID:AB_887873); mouse anti-GAD2 (1:500, RRID:AB_2107894, also called GAD65); guinea pig anti-VGluT1 (1:500, RRID:AB_887878); mouse anti-GAD1 (1:1000, RRID:AB_2278725, also called GAD67). Secondary antibodies conjugated to Alexa Fluor 488, 546, or 633 were used for detection. Images were acquired on an Olympus FV1000 or FV1200 confocal microscope with

60x oil immersion objectives with 1.4 numerical aperture, the pinhole was set to one airy unit, and identical settings were applied to all samples within an experiment. Single confocal sections were analyzed in ImageJ software (NIH). Background was subtracted using the rolling ball method with a radius of 2 m. For quantification of synaptic protein levels, regions of interest

(ROIs) were defined using VGAT, GAD2, or VGluT1 puncta and the average intensity of the protein of interest (in the 488 channel) inside those ROIs was quantified. In Figure 2.1, since differences in antibody affinity make raw fluorescence values between two different antibodies non-comparable, individual data points were calculated for each channel (ELKS1 or ELKS2) by normalizing the fluorescence intensity within a single ROI to the average intensity across all

ROIs. In Figure 2.5, the intensity of the ELKS1, RIM, Bassoon, and RIM-BP2 staining in cDKO neurons was normalized to the staining in control neurons. In all other figures where only

ELKS1 staining is quantified, data are expressed in arbitrary units (a.u.). When necessary,

21 representative images were enhanced for brightness and contrast to facilitate visual inspection; all such changes were made after analysis and were made identically for all experimental conditions. All quantitative data are derived from 3 cultures; 5 - 10 fields of view were quantified per culture per genotype. For all image acquisition and analyses comparing two or more conditions, the experimenter was blind to the condition.

Western blotting. Western blotting was performed according to standard protocols. After SDS-

Page electrophoresis, gels were transferred onto nitrocellulose membranes and blocked in 10%

(w/v) non-fat milk/5% (v/v) goat serum. Membranes were incubated with primary antibodies in

5% (w/v) non-fat milk/5% (v/v) goat serum for two hours at room temperature or overnight at 4C.

The following primary antibodies were used: mouse anti-ELKS1 (1:1000; E-1, RRID:

AB_10841908), rabbit anti-ELKS2B (1:500; custom antibody 1029), rabbit anti-ELKS1/2

(1:2000; P224, gift of Dr. Thomas Südhof), mouse anti-β-actin (1:2000; RRID: AB_476692).

After washing, membranes were incubated with HRP-conjugated secondary antibodies in 5%

(w/v) non-fat milk/5% (v/v) goat serum for one hour at room temperature, and chemiluminescence was used for detection after washing.

Electrophysiology. Electrophysiological recordings in cultured hippocampal neurons were performed as described (Kaeser et al. 2008; Maximov et al. 2007; Kaeser et al. 2011; C. Liu et al. 2014; Kaeser et al. 2009) at DIV 15 - 19. The extracellular solution contained (in mM): 140

NaCl, 5 KCl, 2 CaCl2, 2 MgCl2,10 HEPES-NaOH (pH 7.4), 10 Glucose (~310 mOsm). For evoked NMDAR excitatory postsynaptic currents (EPSCs) picrotoxin (PTX, 50 M) and 6-

Cyano-7-nitroquinoxaline-2,3-dione (CNQX, 20 M) were added to the bath. For miniature

EPSC recordings and RRP measurements tetrodotoxin (TTX, 1 M) was added to block action

22 potentials, in addition to PTX (50 M) for EPSCs or D-(-)-2-Amino-5-phosphonopentanoic acid

(APV, 50 M) and CNQX (20 M) for IPSCs. All recordings were performed in whole cell patch clamp configuration at room temperature. Glass pipettes for were pulled at 2-4 M and filled with intracellular solutions containing (in mM) for EPSC recordings: 120 Cs-methanesulfonate,

10 EGTA, 2 MgCl2, 10 HEPES-CsOH (pH 7.4), 4 Na2-ATP, 1 Na-GTP, 4 QX314-Cl (~300 mOsm) and for IPSC recordings in Figure 2.1: 120 CsCl, 5 NaCl, 10 EGTA, 1 MgCl2, 10

Sucrose, 10 Hepes-CsOH (pH 7.4), 4 Mg-ATP, 0.4 GTP, 4 QX314-Cl (~300 mOsm) and Figure

2.5: 40 CsCl, 90 K-Gluconate, 1.8 NaCl, 1.7 MgCl2, 3.5 KCl, 0.05 EGTA, 10 HEPES-CsOH (pH

7.4), 2 MgATP, 0.4 Na2-GTP, 10 Phosphocreatine, 4 QX314-Cl (~300 mOsm). Cells were held at at -70 mV for AMPAR-EPSC and IPSC recordings, at +40 mV for NMDAR-EPSC recordings and. Access resistance was monitored during recording and cells were discarded if access exceeded 15 M or 20 M during recording of evoked or spontaneous synaptic currents, respectively. Action potentials in presynaptic neurons were elicited with a bipolar focal stimulation electrode fabricated from nichrome wire. The RRP in Figures 2.4 and 2.5 was measured by application of 0.5 M sucrose in extracellular solution applied via a microinjector syringe pump for 10 s at a flow rate of 10 L/min. For Figure S6, the RRP was measured by focal application of 0.5 M sucrose with a picospritzer for 30 s in the presence of TTX (1 M), as described in Liu et al., 2014. Data were acquired with an Axon 700B Multiclamp amplifier and digitized with a Digidata 1440A digitizer. For action potential and sucrose-evoked responses, data were acquired at 5 kHz and low-pass filtered at 2 kHz. For miniature recordings data were acquired at 10 kHz. All data acquisition and analysis was done using the pClamp10. For all electrophysiological experiments, the experimenter was blind to the genotype throughout data acquisition and analysis.

Presynaptic Ca2+ imaging. All Ca2+-imaging experiments were done in cultured hippocampal neurons infected with lentiviruses (expressing active cre or inactive cre) at DIV 5. Presynaptic

Ca2+ transients were examined at DIV15-18 in whole cell patch clamp configuration at room temperature. The extracellular solution contained (in mM): 140 NaCl, 5 KCl, 2 CaCl2, 2 MgCl2,

10 Glucose, 0.05 APV, 0.02 CNQX, 0.05 PTX, 10 HEPES-NaOH (pH 7.4, ~310 mOsm). Glass pipettes were filled with intracellular solution containing (in mM) 140 K Gluconate, 0.1 EGTA, 2

MgCl2, 4 Na2ATP, 1 NaGTP, 0.3 Fluo5F, 0.03 Alexa Fluor 594, 10 HEPES-KOH (pH 7.4, ~300 mOsm). Neurons were filled for 7 minutes and axons and dendrites were identified in the red channel. Presynaptic boutons were identified by their typical bead-like morphology. Neurons in which the distinction between axons and dendrites was unclear were discarded. 10 min after break-in, presynaptic Ca2+ transients were induced by a single action potential evoked via somatic current injection (5 ms, 800-1200 pA) and monitored via Fluo5F fluorescence. Images were acquired using an Olympus BX51 microscope with a 60x, 1.0 numerical aperture objective.

Fluorescence signals were excited by a light-emitting diode at 470 nm, and were collected with a scientific complementary metal–oxide–semiconductor camera at 100 frames/s for 200 ms before and 1s after the action potential. After the experiment, coverslips were fixed using 4% paraformaldehyde/phosphate-buffered saline. Neurons were stained with GAD67 antibodies as described above and imaged neurons, identified post-hoc by their Alexa 594 filling, were identified as either inhibitory or excitatory neurons using confocal microscopy. In additional experiments (Figure S4), distinction between excitatory and inhibitory cultured neurons was further characterized by labeling excitatory neurons using a lentivirus expressing tandem dimer tomato (TdTomato) under a CamKII promoter (pFCK(1.3)tdimer2W; gift from Pavel Osten;

Addgene plasmid #27233). Ca2+ transients were quantified using ImageJ. For the analysis in boutons, ROIs were defined using pictures taken in the red channel, and 7 - 10 boutons were randomly selected from each neuron. Background was subtracted using the rolling ball method with a radius of 1.5 μm. After background subtraction, (F-F0)/F0 was calculated (F = average

24 green emission in a bouton at a given time point, F0 = average fluorescent intensity in frames 0 to 20 before action potential induction). For dendritic measurements, a second order dendrite was selected from each neuron, and the fluorescence from a nearby empty region was referred to as background and subtracted from the ROI. For all Ca2+ imaging experiments, the experimenter was blind to the genotype throughout data acquisition and analysis.

Statistics. Statistical significance was set at *p ≤ 0.05, **p ≤ 0.01, and ***p ≤ 0.001. Unless otherwise noted in the figure legends, all tests were performed using Student’s t-tests to compare means. In cases where significance was determined by one-way ANOVA, Holm-Sidak multiple comparisons tests were used to compare all conditions against the cDKO condition to assess rescue. All experiments were done with using a minimum of three independent cultures and in each culture multiple cells (typically 5-10 per culture and genotype) or images (typically

5-10 images per culture and genotype) were analyzed.

Results

ELKS1 and ELKS2 are enriched at excitatory hippocampal synapses

The distribution of individual ELKS proteins at excitatory and inhibitory synapses is not known. We generated ELKS2 specific antisera by immunization of rabbits (Figure S1 A-C) and used this, in addition to an available ELKS1 specific antibody, to determine if either or both

ELKS proteins are present at excitatory and inhibitory synapses. We employed immunostainings for ELKS1 and ELKS2 in cultured hippocampal neurons and analyzed their distribution using confocal microscopy. Both ELKS proteins were present at excitatory and

Figure 2.1. ELKS1 and ELKS2 are co-expressed at excitatory synapses. (A) Schematic of ELKS protein structure. Arrows: transcriptional start sites of - and -ELKS, CCA-D: coiled-coil regions A - D (ELKS1: CCA 1MYG…SKI208, CCB 209TIW…ENN358, CCC 360MLR…EAT696, CCD

697LEA…EEE988; ELKS2: CCA 1MYG…ARM204, CCB 205SVL…ENI362, CCC 363HLR…NIE656, CCD

656DDS…DEE917, B: PDZ-binding sequence (ELKS1: 989GIWA992, ELKS2: 918GIWA921) of the

ELKS-B C-terminal splice variant. Binding regions for interacting active zone proteins are indicated with black bars. (B) Sample images and quantification of ELKS1 (left) and ELKS2

(right) expression levels at excitatory and inhibitory synapses. VGAT or GAD2 (red, inhibitory synapses) and VGluT1 (blue, excitatory synapses) staining was used to define ROIs, respectively (control n = 4 independent cultures, cDKO n = 4, 10 images were averaged per culture). All data are means ± SEM; *p ≤ 0.05 as determined by Student's t test. (C) Sample images (top) and correlation of expression levels of ELKS1 and ELKS2 (bottom) at excitatory

(left) and inhibitory (right) synapses. Arrowheads indicate example puncta used to define ROIs.

Data points represent the fluorescent intensity of ELKS1 within an ROI plotted against the

ELKS2 signal in the same ROI. Within a single channel, individual puncta are normalized to the average intensity across all puncta (excitatory synapses: 329 ROIs/30 images/3 independent cultures; inhibitory synapses: 250/30/3). : Spearman rank correlation between

ELKS1 and ELKS2.

Figure 2.1 (Continued)

27 inhibitory synapses and we observed higher intensity staining at excitatory synapses compared to inhibitory synapses for ELKS1 and ELKS2 (Figure 2.1 B). A recent proteomic study of release site composition found that overall differences between excitatory and inhibitory release sites are small (Boyken et al. 2013). Interestingly, however, ELKS1 and ELKS2 were modestly enriched at docking sites for glutamatergic vesicles, consistent with our observation. We next examined the distribution of ELKS relative to one another to determine whether levels of individual ELKS proteins correlate positively or negatively at excitatory or inhibitory synapses.

Synaptic markers for either excitatory or inhibitory synapses were used to define regions of interest (ROI) and co-stained for both ELKS1 and ELKS2 (Figure 2.1 C). We found a strong positive correlation between ELKS1 and ELKS2 at excitatory and inhibitory synapses and both ELKS proteins showed a unimodal distribution of fluorescence intensity at each synapse

(Figure S2). Together, these data reveal that ELKS1 and ELKS2 are enriched at excitatory synapses and they suggest that synapses rich in ELKS1 are rich in ELKS2 and vice versa.

ELKS1 and ELKS2 control neurotransmitter release at excitatory hippocampal synapses

Excitatory transmission was not affected in single ELKS2 mutants (Kaeser et al. 2009) and excitatory synaptic transmission was not studied in ELKS1/ELKS2 double mutants. We thus decided to determine the function of ELKS1/ELKS2 at excitatory synapses in cultured hippocampal neurons employing mice in which the first coding exon of each gene, Erc1 and

Erc2, is flanked by loxP sites (C. Liu et al. 2014; Kaeser et al. 2009). At 5 days in vitro (DIV), we infected the neurons with lentiviruses that express a cre-recombinase tagged with EGFP driven by a neuron-specific synapsin promoter (C. Liu et al. 2014) to generate ELKS1/ELKS2 knockout neurons (cDKO). In all experiments, control neurons were genetically identical with the

Figure 2.2. ELKS1/2 control neurotransmitter release at excitatory synapses. (A-C)

Sample traces and quantification of IPSC (A), AMPAR-EPSC (B), and NMDAR-EPSC (C) amplitudes and rise times in control and ELKS1/2 cDKO neurons. Bar graphs show quantification of the peak amplitude (middle) and quantification of the rise time from 20% to 80% of the peak amplitude (right, A: control n = 18 cells/4 independent cultures, cDKO n = 18/4; B: control n = 18/4, cDKO n = 16/4; C: control n = 15/3, cDKO n = 15/3). (D) Sample traces (top) and quantification (bottom) of mEPSC frequency, amplitude, and 20-80% rise time (control n =

16/3, cDKO n = 15/3). Sample traces on the top left show 10 seconds of recording time. Sample traces on the top right are the overlayed averaged events from an individual cell in each condition. All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by

Student's t test.

Figure 2.2 (Continued)

30 exception that they were infected with lentiviruses that expressed an inactive, truncated cre protein. We only analyzed cultures in which no non-infected neurons could be detected.

Using focal stimulation, we recorded action-potential evoked excitatory postsynaptic currents (EPSCs) and directly compared effects to inhibitory PSCs (IPSCs). Similar to the effect on IPSCs (Figure 2.2 A), EPSCs were reduced to ~50% (Figures 2.2 B, C). At inhibitory synapses, removal of ELKS resulted in a 30% decrease in presynaptic Ca2+ influx which led to a reduction in P (C. Liu et al. 2014) and prolonged IPSC rise times (Figure 2.2 A). When we examined the EPSC rise times, there was no effect of ELKS1/2 cDKO (Figures 2.2 B, C), suggesting that ELKS may operate differently at excitatory synapses. To determine whether the deficit in excitatory transmission was presynaptic, we recorded miniature EPSCs (mEPSCs) in the presence of TTX. mEPSC frequency was reduced by ~50%, but there was no change in mEPSC amplitude, rise time (Figure 2.2 D), or decay kinetics (control = 5.727 ms, n = 16/3; cDKO = 6.321, n = 15/3; p > 0.05). The number and size of excitatory synapses was also unchanged (Figure S3). Thus, ELKS1/2 cDKO reduces release from excitatory presynaptic nerve terminals, but the mechanisms may be different from inhibitory synapses.

ELKS enhances the size of the RRP but not Ca2+ influx at excitatory synapses

At inhibitory hippocampal synapses, ELKS boosts presynaptic Ca2+ influx to increase P and to accelerate the IPSC rise. To directly test the prediction that ELKS functions differently at excitatory synapses, we employed imaging to measure the presynaptic Ca2+ transient in response to a single action potential (Figure 2.3 A). Neurons were filled with the Ca2+ indicator

Fluo-5F and a fixable Alexa-594 dye and Ca2+ influx into presynaptic boutons was imaged during a single action potential elicited by a brief current injection through the patch pipette. This

Figure 2.3. ELKS1/2 do not control Ca2+ influx and release probability in excitatory nerve terminals. (A) Sample images (top, imaged boutons are numbered and color coded), action potential and imaging traces (middle), and summary data (bottom) of Ca2+ influx measurements in example control (left) or cDKO (right) excitatory neurons. Summary plots of single action potential-induced Ca2+ transients in presynaptic boutons are shown, the inset shows the same plot for dendrites. Data are shown as mean (line) ± SEM (shaded area) and analyzed by two-way ANOVA: genotype, n.s.; time, ***, p < 0.001; interaction, n.s. (Boutons: control n = 60 boutons/6 cells /4 independent cultures, cDKO n = 80/8/4; dendrites: control n = 6 dendrites/6 cells/4 independent cultures, cDKO n = 8/8/4). (B) Sample traces (top) and quantification of paired pulse ratios (PPRs, bottom) of evoked NMDAR-EPSCs. Example traces showing overlayed responses to pairs of stimuli at 50, 100, 500, and 2500 ms interstimulus intervals. PPRs (amplitude 2/amplitude 1) are plotted against the interstimulus interval.

Significance as analyzed by two-way ANOVA: genotype, n.s.; ISI, ***, p < 0.001; interaction, n.s.

(control n = 17 cells/3 independent cultures, cDKO n = 19/3).

Figure 2.3 (Continued)

33 method has revealed impaired Ca2+ influx at inhibitory ELKS deficient synapses (C. Liu et al.

2014). After the experiment, cells were fixed and stained with GAD67 antibodies to exclude

GAD67 positive inhibitory neurons from the analysis. This method reliably distinguished excitatory and inhibitory neurons in hippocampal cultures (Figure S4). We found that there was no change in peak action potential evoked Ca2+ influx into boutons of excitatory neurons (Figure

2.3 A). At inhibitory synapses, decreased Ca2+ influx in ELKS deficient neurons resulted in a reduction in P. Thus, our data suggest that initial P may not be affected at excitatory

ELKS1/2 cDKO synapses. Initial P is inversely correlated with paired-pulse ratios (PPR). We measured PPRs of EPSC amplitudes by monitoring N-Methyl-D-aspartate receptor EPSCs

(NMDAR-EPSCs) to circumvent the strong reverberant activity that is present in cultured networks when α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors are not blocked (Maximov et al. 2007). Consistent with the Ca2+ imaging data, but different from inhibitory synapses (C. Liu et al. 2014), there were no changes in PPR in ELKS cDKO neurons across all tested inter-stimulus intervals (Figure 2.3 B). These experiments establish that defects in Ca2+ influx and P are unlikely to explain impaired neurotransmitter release at excitatory synapses of ELKS cDKO neurons.

We next tested whether the size of the RRP was changed at excitatory synapses. We stimulated release of the entire RRP using a hypertonic sucrose solution (+500 mOsm) and quantified RRP size by integrating the total charge transfer during the first ten seconds of the response (Rosenmund and Stevens 1996). Although the hypertonic stimulus is non- physiological, this method has been used as a snapshot measurement of RRP in cultured neurons and has been insightful for genotype comparisons and for dissecting molecular mechanisms of RRP control (Augustin, Rosenmund, et al. 1999; Deng et al. 2011; Neher 2015;

Rosenmund and Stevens 1996).At excitatory synapses, the RRP was reduced by ~40% (Figure

Figure 2.4. ELKS1/2 control RRP at excitatory synapses. (A) Sample traces showing

AMPAR-EPSCs in response to superfusion with 500 mOsm sucrose (left). The bar graph on the right shows the AMPAR-EPSC charge transfer, quantified as the area under the curve during the first ten seconds of the response (control n = 30/3, cDKO n = 29/3). (B) Sample traces (left) of NMDAR-EPSCs during a short action potential train (10 stimuli at 20 Hz). The charge transfer during the train and in the two seconds immediately after the train is quantified separately on the right (control n = 14/3, cDKO n = 14/3). All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01 as determined by Student's t test

2.4 A) in ELKS cDKO neurons, providing an explanation for a reduction in release in the absence of changes in P at excitatory ELKS1/2 cDKO synapses. Short action potential trains

(10 stimuli at 20 Hz), a more physiological stimulus, resulted in a 50% reduction in charge transfer during the stimulus train in ELKS1/2 cDKO neurons compared to control neurons

(Figure 2.4 B). The delayed charge transfer after stimulation ended was similarly reduced.

These data are consistent with a reduced number of RRP vesicles available for release in response to a brief action potential train.

One possible explanation for the phenotypic differences between inhibitory transmission as observed earlier (C. Liu et al. 2014) and excitatory transmission as described here is that removal of ELKS has effects that vary between cultures and over time, causing some cultures to have more pronounced effects on RRP size and others to exhibit changes in P. To control for this possibility, we conducted a series of experiments analyzing RRP size and P (as measured by PPRs) for inhibitory and excitatory transmission in the same cultures and matching sample size for all conditions. Both inhibitory and excitatory synapses had a reduction in action potential-evoked EPSC and IPSC amplitudes (Figure S5). At inhibitory synapses, ELKS1/2 cDKO increased the PPR at low interstimulus intervals but had no significant effect on RRP size, consistent with a reduction of P (Figures 2.5 A, C) and with our previous study (C. Liu et al.

2014). In contrast, excitatory synapses in the same cultures showed no change in PPR but a reduction in RRP size (Figures 2.5 B, D). Notably, there was a small, non-significant trend towards a reduced RRP at inhibitory synapses in this experiment. To further rule out that there is a reproducible RRP reduction at inhibitory hippocampal synapses, we conducted a second experiment measuring RRP with a slightly different protocol as described in the methods section and used in (C. Liu et al. 2014). Again, no change in RRP was detected (Figure S6). Thus, in three independent measurements (Figure 2.5 C, Figure S6, and Figure 7 F in Liu et al., 2014), no reduction in the inhibitory RRP could be detected.

Figure 2.5. Direct comparison of IPSC and EPSC phenotypes of ELKS1/2 cDKO. (A)

Example traces (top) showing overlayed IPSC responses to pairs of stimuli at 10, 20, 100, and

500 ms interstimulus intervals. Paired-pulse ratios (amplitude 2/amplitude 1) are plotted against the interstimulus interval (10, 20, 50, 100, 500, and 2500 ms intervals). Significance as analyzed by two-way ANOVA: genotype, ***, p < 0.001; ISI, ***, p < 0.001; interaction, n.s. Holm-Sidak post-hoc test: 10 ms, *, p < 0.05; 20 ms, *, p < 0.05 (control n = 15 cells/3 independent cultures, cDKO n = 15/3). (B) Example traces (top) showing overlayed EPSC responses to pairs of stimuli at 50, 100, 500, and 2500 ms interstimulus intervals. Paired-pulse ratios (amplitude

2/amplitude 1) are plotted against the interstimulus interval (50, 100, 500, and 2500 ms intervals). Significance as analyzed by two-way ANOVA: genotype, n.s.; ISI, ***, p < 0.001; interaction, n.s. (control n = 15/3, cDKO n = 15/3). (C) Sample traces showing IPSCs in response to superfusion with 500 mOsm sucrose (left) and quantification (right) of IPSC charge transfer during the first ten seconds of the response (control n = 15/3, cDKO n = 15/3). (D)

Sample traces showing EPSCs in response to superfusion with 500 mOsm sucrose (left) and quantification (right) of EPSC charge transfer during the first ten seconds of the response

(control n = 15/3, cDKO n = 15/3). Data are means ± SEM; **p ≤ 0.01 as determined by

Student's t test.

Figure 2.5 (Continued)

These data confirm our previous experiments and establish synapse-specific roles for ELKS in neurotransmitter release: at excitatory synapses ELKS1/2 primarily boost the RRP (Figures

2.1-5), whereas at inhibitory synapses ELKS1/2 mainly control action potential induced Ca2+ influx to enhance P (also see Figure 2.5; Liu et al., 2014).

Removal of ELKS1/2 does not lead to loss of presynaptic priming proteins.

Given the hypothesis that ELKS is a presynaptic scaffold, we set out to test the possibility that loss of ELKS1/2 changes presynaptic levels of proteins involved in controlling

RRP size at excitatory synapses. RIM and Munc13-1 were of particular interest, since both proteins localize to the active zone and knockouts of either protein have strong RRP impairments at excitatory hippocampal synapses (Augustin, Betz, et al. 1999; Calakos et al.

2004; Kaeser et al. 2011). Using confocal microscopy, we quantified the synaptic levels of the active zone proteins ELKS, RIM, Munc13-1, Bassoon, and RIM-BP2 in cDKO and control neurons. With the exception of ELKS, no protein was significantly reduced at excitatory or inhibitory synapses (Figure 2.6). These data are consistent with normal electron microscopic appearance of ELKS1/2 cDKO synapses (C. Liu et al. 2014) and indicate that ELKS is not essential to recruit the priming proteins RIM and Munc13-1 to the presynaptic nerve terminal.

ELKS controls RRP through the N-terminal coiled-coil domains

Since ELKS1/2 cDKO caused no detectable structural changes at the active zone, we instead turned to electrophysiological rescue experiments to determine how ELKS might control

RRP size. ELKS binds directly to the PDZ domain of RIM through its four C-terminal residues and may have roles in active zone anchoring of RIM (Lu et al. 2005; Yun Wang et al. 2002;

Figure 2.6. Active zone composition in ELKS1/2 cDKO synapses. (A) Sample images of control and ELKS1/2 cDKO neurons stained with antibodies against active zone proteins.

Inhibitory synapses were marked with VGAT (for RIM and Bassoon) or GAD2 (for Munc13-1 and RIM-BP2), excitatory synapses were marked with VGluT1. (B) Quantification of active zone proteins within ROIs defined by excitatory (left) or inhibitory (right) synaptic markers (RIM: control n = 4 independent cultures, cDKO n = 4; Munc13-1: control n = 4, cDKO n = 4; Bassoon: control n = 3, cDKO n = 3; RIM-BP2: control n = 3, cDKO n = 3; in each culture, 10 images were averaged per culture and genotype). Data are means ± SEM; ***p ≤ 0.001 as determined by paired t tests against the control.

(Lu et al. 2005; Yun Wang et al. 2002; Ohtsuka et al. 2002; Kaeser et al. 2009). Peptide injection experiments using the RIM-binding domain of ELKS support a role of the RIM-ELKS interactions in release (Takao-Rikitsu et al. 2004). The CCD domain, which is adjacent to the

RIM binding sequence, also binds to Ca2+ channel 4 subunits (Kiyonaka et al., 2012). These interactions suggest that the C-terminus of ELKS may tether ELKS to the active zone to support its functions in release.

We designed lentiviral rescue constructs in which we expressed either full length

ELKS1B or mutant ELKS1 that lacks either the entire C-terminal region including the PDZ binding motif (CCDB) or just CCD (CCD) (Figure 2.7 A). All three rescue constructs expressed at levels similar to wild type ELKS1 and localized to synapses (Figure S7), suggesting that the

ELKS C-terminal regions are not necessary for ELKS localization. Surprisingly, ELKS1B,

ELKS1-CCDB, and ELKS1-CCD were sufficient to rescue excitatory RRP size in ELKS cDKO neurons (Figure 2.7 B). Thus, the ELKS C-terminal domains that bind to RIM and other presynaptic proteins are not necessary for its control of the RRP at excitatory synapses.

Altogether, the C-terminal ELKS domains are unlikely to act as a central scaffolding hub at the active zone.

We next decided to take an unbiased approach and systematically tested all ELKS protein interaction sites covering the entire sequence of ELKS1B in rescue experiments. We generated rescue constructs lacking N-terminal coiled-coils (corresponding to ELKS1B) or the central coiled-coil region (CCc). ELKS1B and ELKS1-CCDB were used as positive rescue controls (Figure 2.8 A). All rescue constructs were successfully expressed, albeit at variable levels (Figure S8). Compellingly, neither ELKS1B nor ELKS1-CCC rescued the RRP in

ELKS1/2 cDKO neurons (Figure 2.8 B). The lack of rescue could be due to either a local

Figure 2.7. C-terminal ELKS sequences do not support RRP in ELKS1/2 cDKO neurons. (A) Schematic of ELKS1 rescue constructs; CCA-D: coiled-coil regions A-D, B: PDZ- binding motif; H: human influenza hemagglutinin (HA) tag, deleted sequences are illustrated as dashed lines. (B) Sample traces (left) and quantification (right) of the AMPA-EPSC charge in response to hypertonic sucrose application, measured as area under the curve during the first ten seconds after the start of the stimulus (control n = 26 cells/5 independent cultures, cDKO n

= 27/5, ELKS1B n = 21/5, CCDB n = 23/5, CCD n = 21/5). All data are means ± SEM; *p ≤

0.05 as determined by one-way ANOVA followed by Holm-Sidak multiple comparisons post-hoc test comparing each condition to cDKO.

42 function of the deleted coiled-coil regions or to a role for these regions in localizing ELKS to synapses. We distinguished between these possibilities by assessing the localization of each rescue construct using confocal microscopy. Since ELKS1B lacks the antigen recognized by our ELKS1 antibody we stained for either the HA tag included in all rescue proteins (Figure 2.8

C and Figure S8 B) or for ELKS1 (Figure S8 C-E). Synaptic expression of the rescue constructs ranged from 50 - 200% of wild-type levels of ELKS1. Importantly, all constructs localized to synapses. Thus vertebrate ELKS1 can be localized to synapses likely through multiple redundant interactions. While synaptic levels of rescue ELKS1B were low, it rescued entirely. In contrast, ELKS1B and ELKS1-CCC both failed to rescue, but were expressed above ELKS1B levels. Removal of ELKS reveals differential impairments of RRP and P at excitatory and inhibitory synapses, respectively (Figure 2.5). To test whether such differential effects are also reflected in the ELKS sequences that mediate rescue at each synapse, we tested whether ELKS1B, an ELKS isoform that failed to rescue the excitatory RRP, was sufficient to restore inhibitory synaptic transmission (Figure S9). ELKS1B was able to rescue action-potential evoked IPSC amplitudes, establishing that ELKS’ role in controlling P at inhibitory synapses does not require the N-terminal coiled-coil regions that control the excitatory

RRP. Together, these experiments establish that the N-terminal coiled-coil sequences of

ELKS1 are necessary for ELKS' role in enhancing the RRP at excitatory synapses, but not necessary for localizing ELKS to synapses.

Discussion

Our findings define a new functional role for ELKS in setting RRP size at excitatory hippocampal synapses. They extend beyond previous understanding of the role of ELKS in supporting presynaptic Ca2+ influx at inhibitory synapses (C. Liu et al. 2014) and demonstrate

Figure 2.8. N-terminal coiled-coil domains of ELKS control RRP size at excitatory synapses. (A) Schematic of ELKS1 rescue constructs; CCA-D: coiled-coil regions A-D, B: PDZ- binding motif; H: human influenza hemagglutinin (HA) tag, black bar: antigen recognized by the

ELKS1 antibody (E-1) used in Figure S8 C. Deleted sequences are illustrated as dashed lines.

(B) Sample traces (left) and quantification (right) of the AMPA-EPSC charge in response to hypertonic sucrose application, measured as area under the curve during the first ten seconds after the start of the stimulus (control n = 21 cells/4 independent cultures, cDKO n = 22/4,

ELKS1B n = 19/4, ELKS1B n = 18/4, CCC n = 20/4, CCDB n = 21/4). (C) Sample images

(left) of control, cDKO, and cDKO + rescue neurons stained with antibodies against HA.

Quantification (right) of HA fluorescent intensity within ROIs defined by VGluT1 (control n = 3 independent cultures, cDKO n = 3, ELKS1B n = 3, ELKS1B n = 3, CCC n = 3, CCDB n = 3,

5 - 10 images were averaged per culture and genotype). All data are means ± SEM; **p ≤ 0.01,

***p ≤ 0.001 as determined by one-way ANOVA followed by Holm-Sidak multiple comparisons post-hoc test comparing each condition to cDKO.

Figure 2.8 (Continued)

45 that this is not the primary function across all synapses. Most current molecular models of active zone function imply that active zones operate essentially identically across synapses despite the notion that the parameters they control, RRP and P, vary greatly between different types of synapses. Thus far, this assumption has proven principally true for genetic analyses of active zone protein function. For example, it is widely accepted that Munc13 is required at all synapses to prime vesicles and RIMs anchor and activate Munc13 to support priming (Varoqueaux et al.

2002; Andrews-Zwilling et al. 2006; Deng et al. 2011; Calakos et al. 2004; Augustin, Betz, et al.

1999). Our findings are a starting point for the dissection of synapse-specific architecture and function of the active zone.

One possibility to explain synapse-specific roles is that ELKS1 and ELKS2 account for different functions, and that they are localized to specific subsets of synapses. Such differences have been observed for roles of Munc13-1 and bMunc13-2 in short-term plasticity

(Rosenmund et al. 2002). Our data make this possibility unlikely in the case of ELKS, because there is a strong positive correlation between ELKS1 and ELKS2 levels at all synapses.

Furthermore, no ELKS isoform-specific protein interactions are described in the literature, and the interaction sequences are generally well conserved between ELKS1 and ELKS2 (Monier et al. 2002; Yun Wang et al. 2002).

Another possibility is that interaction partners of ELKS that mediate RRP control are distributed in a synapse specific fashion, and ELKS protein interactions at the active zone may be engaged differentially depending on the presence of specific interacting proteins. This model is supported by the observation that the sequence requirements for rescue of the excitatory

RRP or the IPSC are different. ELKS interacts in vitro with many active zone proteins (Ohtsuka et al. 2002; Yun Wang et al. 2002; Schoch and Gundelfinger 2006). We find that ELKS N- terminal, but not C-terminal, sequences are required for RRP control. These N-terminal sequences have been shown to bind to Liprin- via CCA-CCC (Ko et al. 2003), to Bassoon via

CCC (Takao-Rikitsu et al. 2004), and they may mediate binding to Rab6 (Monier et al. 2002).

Because constructs including CCC but lacking CCA/B localize to synapses but do not to rescue, one possible explanation is that Liprin- binding is required for ELKS function in controlling the excitatory RRP. Vertebrate genomes contain four genes encoding Liprin-, but it is not known whether Liprin- isoforms are differentially distributed to specific active zones. In fact, it is currently not clear for any of the vertebrate Liprin- isoforms whether there is a tight association with the presynaptic active zone (Zürner et al. 2011; Spangler et al. 2011). Functional roles of vertebrate Liprin- in synaptic transmission are not well understood, but a recent study supports presynaptic roles for Liprin-2 at hippocampal synapses (Spangler et al. 2013). In invertebrates,

Liprin-/syd-2 has synaptogenic activities, and effects of a gain of function mutation in syd-2 require ELKS (Dai et al. 2006; Patel et al. 2006), indicating important synaptic roles for Liprin-

/ELKS interactions. Thus, these previous studies are consistent with the hypothesis that ELKS controls RRP through Liprin-. Nevertheless, it is possible that binding to Bassoon, Rab6, or unknown proteins mediate the role of ELKS in enhancing RRP. A recent proteomic analysis of vesicle docking complexes revealed only modest differences in the composition of docking sites at glutamatergic compared to GABAergic synapses (Boyken et al. 2013). Interestingly, however,

Bassoon and ELKS were found to be enriched in glutamatergic synapses in this study, perhaps supporting a role for these proteins for preferentially promoting excitatory transmission.

Furthermore, a recent study showed that Bassoon regulates release indirectly via RIM-BP through a specific Ca2+ channel subunit, which could result in synapse-specific roles for

Bassoon (Davydova et al. 2014). In summary, these studies provide support for the hypothesis that ELKS may operate with Bassoon or Liprin- to promote release in a synapse-specific fashion.

Our studies also reveal that ELKS1 and ELKS2are not required for active zone assembly and maintenance at hippocampal synapses, consistent with studies in C. elegans

(Deken et al. 2005; Patel et al. 2006). At the D. melanogaster neuromuscular junction, however,

Brp is essential for the formation of T-bars. The ELKS homology of Brp is limited to the N- terminal half of the protein (Kittel et al. 2006; Monier et al. 2002), making it possible that such strong scaffolding functions are specific to Brp and absent in ELKS. Alternatively, an ELKS scaffolding function may not be detected in our experiments due to redundant scaffolding activities in other proteins, for example -ELKS or RIM (Schoch et al. 2002; Kaeser et al. 2009;

C. Liu et al. 2014).

Several studies suggested that there is a good correlation between the number of docked vesicles and the size of the RRP at hippocampal synapses (Schikorski and Stevens

2001; Imig et al. 2014), but electron microscopic analysis of ELKS1/2 cDKO synapses using conventional fixation has not revealed a deficit in vesicle docking (C. Liu et al. 2014). Recent work has achieved better resolution in the analysis of docking by combining high-pressure freezing with EM tomography (Imig et al. 2014; Siksou et al. 2009). We can currently not rule out the possibility that ELKS1/2 cDKO causes a subtle vesicle docking phenotype undetectable by the methods used in our previous work. An alternative hypothesis to a reduction of RRP at each excitatory synapse is the possibility that a subpopulation of excitatory synapses is very strongly affected by the removal of ELKS. Because the distribution of ELKS across excitatory synapses is unimodal (Figure S2), it is unlikely that ELKS only operates at a subset of excitatory synapses and that these synapses are silent in the absence of ELKS.

However, additional work at clearly defined synapse populations with less heterogeneity than mixed hippocampal cultures will be necessary to address these questions.

Finally, although ELKS1/2 cDKO neurons do not reveal an RRP deficit as measured by hypertonic stimuli at inhibitory synapses (Figure 2.5, Liu et al., 2014), ELKS is known to regulate RRP at these synapses. Enhancing P by increasing extracellular Ca2+ cannot completely rescue synaptic transmission at inhibitory ELKS1/2 cDKO synapses (C. Liu et al.

2014), and removal of ELKS2 alone leads to an increase in the RRP at inhibitory synapses

(Kaeser et al. 2009). These experiments suggest that there may be molecular regulation of the

RRP at inhibitory synapses through interplay between ELKS1 and ELKS2, which are both present at inhibitory active zones.

In the long-term, it will be important to understand how the synapse-specific molecular control of RRP and P contribute to circuit function. Human genetic experiments reveal that mutations in ERC1/ELKS1 may contribute to autism spectrum disorders (Silva et al. 2014), and it is possible that the pathophysiology arises from synapse-specific misregulation of neurotransmitter release.

— CHAPTER THREE —

Fusion competent synaptic vesicles persist upon active zone disruption and loss of

vesicle docking

Shan Shan H. Wang*, Richard G. Held*, Man Yan Wong, Changliang Liu, Aziz Karakhanyan

and Pascal S. Kaeser.

R.G.H. performed and analyzed all physiology experiments. SS.H.W. performed immunostaining and western blotting. SS.H.W. and A.K. processed and analyzed electron microscopy experiments. M.Y.W. performed pHluorin imaging. C.L. performed presynaptic calcium imaging. This chapter was reproduced with permission from Neuron 91(4) (2016), pp.

777-791, doi: 10.1016/j.neuron.2016.07.005. * S.S.H.W. and R.G.H contributed equally to this work.

Introduction

Ca2+-triggered fusion of synaptic vesicles is mediated by soluble NSF-attachment protein receptors (SNAREs) and is restricted to release sites called active zones (Couteaux and Pécot-

Dechavassine 1970; Südhof 2012). The active zone is a highly organized structure that docks synaptic vesicles close to release machinery and presynaptic Ca2+ channels (Figure 3.1 A). This establishes the tight spatial organization required for fast synaptic vesicle fusion upon Ca2+ entry and it provides molecular machinery to set and regulate synaptic strength (Kaeser and Regehr

2014). Functionally, synaptic strength is determined by two parameters that are set at the active zone. First, only a subset of vesicles can be released upon arrival of an action potential. This pool of vesicles is generated through a priming reaction and is called the readily releasable pool

(RRP). Second, an action potential releases RRP vesicles with a certain probability, called vesicular release probability (P). Synaptic strength, the amount of release from a given synapse, is proportional to the product of RRP and P (Zucker and Regehr 2002).

The active zone matrix consists of multi-domain proteins that control RRP and P, and their localization is restricted to the presynaptic plasma membrane area opposed to the postsynaptic density (PSD). These proteins include RIM, ELKS (also known as

ERC/CAST/Rab6IP2), Munc13, Bassoon/Piccolo, RIM-BP, and Liprin- (Schoch and

Gundelfinger 2006; Südhof 2012). Many additional proteins including SNAREs, ion channels, receptors, cytoskeletal proteins and adhesion molecules are also present (Boyken et al. 2013;

Morciano et al. 2009; C. S. Müller et al. 2010) but are not restricted to the active zone matrix.

Removing individual proteins of the active zone matrix has effects on release that vary in extent, and there are differences between synapses in vertebrates, C.elegans, and D. melanogaster in the reliance on specific active zone proteins (Acuna et al. 2015; Kaeser et al. 2011; Kittel et al.

2006; Koushika et al. 2001; C. Liu et al. 2014; M. Müller et al. 2012). At vertebrate synapses, release strongly depends on synaptic vesicle priming activities of Munc13s such that loss of

Munc13 leads to loss of all fusion competent vesicles (Augustin, Rosenmund, et al. 1999;

Varoqueaux et al. 2002). RIMs contribute to priming through anchoring and activation of

Munc13 (Andrews-Zwilling et al. 2006; A. Betz et al. 2001; Deng et al. 2011; Lu et al. 2006), and they tether primed vesicles to presynaptic Ca2+ channels to enhance release probability (Han et al. 2011; Kaeser et al. 2011; Kaeser et al. 2012). ELKS (Held, Liu, and Kaeser 2016; Kaeser et al. 2009; C. Liu et al. 2014), Bassoon/Piccolo (Davydova et al. 2014; Hallermann, Fejtova, et al.

2010), and RIM-BPs (Acuna et al. 2015) are also present, but knockouts for these proteins show milder impairments in release. The domain structure and the in vitro interactions of RIM,

ELKS and Bassoon/Piccolo further predicted strong scaffolding roles (Schoch and Gundelfinger

2006; Südhof 2012), but except for a partial loss of Munc13 in RIM knockout mice (Schoch et al.

2002) or of RIM-BP in Bassoon knockout mice (Davydova et al. 2014), the active zone protein complex was intact in knockout mice for individual protein families (Davydova et al. 2014; Deng et al. 2011; Held, Liu, and Kaeser 2016; C. Liu et al. 2014). Together with the notion that synaptic vesicle docking and fusion are spatially restricted to the active zone, these studies led to the hypothesis that the active zone is required to translate the incoming action potential into neurotransmitter release (Schoch and Gundelfinger 2006; Südhof 2012).

Synaptic vesicle tethering and docking are thought to precede fusion and have been studied using various methods and definitions. Docking is often defined as synaptic vesicles that are attached to the plasma membrane in electron microscopic images of glutaraldehyde fixed tissue such that the electron densities of the vesicle membrane and target membrane merge

(Acuna et al. 2015; Augustin, Rosenmund, et al. 1999; Bronk et al. 2007; Kaeser et al. 2011).

Using glutaraldehyde fixation only RIMs participate in synaptic vesicle docking without affecting total numbers of synaptic vesicles in a nerve terminal (Augustin, Rosenmund, et al. 1999; Bacaj et al. 2015; Bronk et al. 2007; Kaeser et al. 2011). Recently, the use of high pressure freezing and tomography have allowed for distinction of tight vesicle docking with a resolution of a few

52 nanometers. This has led to the discovery that SNARE proteins and Munc13 contribute to the tight attachment of synaptic vesicles to the presynaptic plasma membrane. Their genetic removal leads to a loss of tightly docked vesicles when observed in high pressure frozen tissue

(Imig et al. 2014; Siksou et al. 2009), but these phenotypes are too subtle to be uncovered using glutaraldehyde fixation (Augustin, Rosenmund, et al. 1999; Bronk et al. 2007). Overall, there is a good correlation between the number of docked vesicles observed with either fixation method, the size of the active zone, and the size of the RRP, which suggested, along with the observation that docked vesicles fuse upon stimulation, that docked vesicles are the RRP

(Holderith et al. 2012; Imig et al. 2014; Rosenmund and Stevens 1996; Schikorski and Stevens

2001; Watanabe et al. 2013). Here, we measure the RRP as vesicles that are released by the application of hypertonic sucrose, and we assume that the same vesicles are accessible to action potentials although differences may exist (Rosenmund and Stevens 1996; Schikorski and

Stevens 2001; Thanawala and Regehr 2016; Zucker and Regehr 2002). We use the term

‘fusion competent’ for vesicles in the RRP and vesicles that are released through spontaneous miniature events (Augustin, Rosenmund, et al. 1999).

To date, no knockout mutation has led to a strong structural disruption of the vertebrate active zone matrix. We set out to generate such a mutation with the goal to test whether the active zone is necessary for the structural assembly of synapses, whether it is required for fusion, and how it participates in setting RRP size and P. We produced conditional mouse mutants to simultaneously remove all active zone isoforms of RIM and ELKS in cultured

2+ hippocampal neurons. This led to loss of Munc13, Bassoon, Piccolo, RIM-BP, and CaV2.1 Ca channels. The overall synaptic assembly including the postsynaptic densities, the synaptic vesicle cluster, and the levels of SNARE proteins remained unaffected. However, we observed a near complete loss of vesicle docking and release probability was strongly decreased.

Surprisingly, a pool of fusion competent vesicles, released as spontaneous miniature events or

53 during stimulation with action potential trains or hypertonic sucrose, persisted upon strong disruption of the active zone and vesicle docking.

Experimental Procedures

The quadruple homozygote floxed mice for RIM1, RIM2, ELKS1 and ELKS2 were generated by crossing single conditional knockout mice (Kaeser et al. 2008; Kaeser et al.

2009; Kaeser et al. 2011; C. Liu et al. 2014). All experiments were performed in cultured hippocampal neurons infected at DIV 5 with lentiviruses expressing cre recombinase or an inactive mutant of cre under a synapsin promoter, and experiments were performed at DIV 15-

19. Biochemical, confocal, electron microscopic and electrophysiological analyses were performed as described (Deng et al. 2011; Kaeser et al. 2008; Kaeser et al. 2011). Quantitative

Western blotting was performed using fluorescently tagged secondary antibodies.

Electrophysiological recordings were done in whole cell patch clamp configuration, and action potentials were triggered by a focal stimulation electrode. For pHluorin imaging, the neurons were infected with lentiviruses expressing SypHy and SV2-TdTomato at DIV 3 in addition to the cre and control lentiviruses supplied at DIV 5. Experiments were performed and analyzed by an experimenter blind to the genotype and significance was determined using Student’s t-tests unless otherwise noted. Detailed descriptions of the methods are provided in the supplemental materials (Appendix Two).

Results

Genetic disruption of the presynaptic active zone.

We generated mice to simultaneously and conditionally remove all presynaptic RIM and

ELKS proteins. We targeted RIM and ELKS proteins because they are expressed at all synapses and they interact with all major active zone proteins (Figure 3.1 A). We crossed conditional knockout mice for presynaptic RIM proteins (Kaeser et al. 2008; Kaeser et al. 2011),

54 encoded by the genes Rims1 and Rims2, to conditional knockout mice for both genes encoding

ELKS proteins (Kaeser et al. 2009; C. Liu et al. 2014), Erc1 and Erc2, to generate quadruple conditional knockout mice (Figure S10 A). All analyses were done in primary hippocampal neurons from these mice or the double conditional knockout mice for either RIM or ELKS proteins. Lentivirus expressing cre recombinase or an inactive mutant of cre in neurons (C. Liu et al. 2014) was supplied at 5 days in vitro (DIV) to generate knockout neurons (cKOR+E) or control neurons (controlR+E). In cKOR+E neurons, we remove RIM and ELKS proteins as assessed by confocal microscopy (Figures 3.1 B, C) and quantitative Western blotting using fluorescent secondary antibodies (Figures 3.1 D, E). In immunostaining, 22% of RIM and 23% of ELKS signal remained upon knockout of these proteins (grey dotted line, Figure 3.1 C) despite much stronger reductions in Western blotting (Figure 3.1 E), establishing that this signal is non-specific (see Figure S10 D for background from secondary antibodies only). Genetic removal of RIM and ELKS led to a very strong reduction of interacting active zone proteins.

Munc13-1 was eliminated from synaptic puncta in an extent similar to RIM and ELKS, and synaptic Bassoon and Piccolo were reduced nearly as strongly (Figures 3.1 B, C), whereas the total protein levels of Munc13-1 and Bassoon were reduced by ~80% (Figures 3.1 D, E).

Synaptic and total RIM-BP2 was reduced by 31% and 42% respectively (Figures 3.1C-E, S10

2+ B) and synaptic CaV2.1 Ca channel levels were reduced by 29% (Figures 3.1 B, C). Because synaptic Munc13-1 levels (Figure 3.1 C) were more strongly reduced than total Munc13-1 levels

(Figure 3.1 E), we tested whether the remaining Munc13-1 was clustered at synapses. The

Munc13-1 protein that was left in the cKOR+E neurons was insoluble as measured in a fractionation experiment of the cultured neurons (Figure 3.1 D). Furthermore, when we evaluated whether the remaining Munc13 clusters co-localized with the remaining Bassoon

(Figure 3.1 G) or with synaptic vesicles (Figures S10 E, F), we found that many Bassoon puncta and synapses did not contain

Figure 3.1. Genetic removal of RIM and ELKS leads to disruption of the active zone. (A)

Schematic of the protein complex at the active zone and its connections to other important presynaptic protein families (marked in red). SAM: sterile alpha motif, MUN: Munc13 homology domain, PDZ: PSD-95/Dlg1/ZO-1 homology domain, SH3: Src homology 3 domain, PxxP: proline rich motif, FN3: fibronectin 3 repeat. (B, C) Sample images (B) and quantitation (C) of protein levels at RIM and ELKS knockout (cKOR+E) and control (controlR+E) synapses using confocal microscopy. The synaptic vesicle marker Synaptophysin-1 (Syp-1) was used to define the region of interest (ROI). The black dotted line indicates control levels and the grey dotted line non-specific staining as assessed for RIM. Example images for RIM-BP2, Piccolo, Liprin-

2, Liprin-3, SNAP-25, and quantitation of puncta number and size are in Figure S10

(controlR+E n = 3 independent cultures, cKOR+E n = 3, 10 images per culture). (D, E) Quantitative

Western blotting for presynaptic proteins using fluorescent secondary antibodies. Some cultures were fractionated into pellet and supernatant (sup.) using Triton X-100 solubilization and ultracentrifugation. Quantitation (E) of total protein levels in cKOR+E neurons normalized to protein levels in controlR+E neurons (black dotted line) are shown. Black and grey dotted as in C

(controlR+E n = 6 independent cultures, cKOR+E n = 6, except for Bassoon where n = 3 for both conditions, Syb-2: synaptobrevin/VAMP-2). (F, G) Sample images (F) and quantitation (G) of the fraction of Bassoon puncta containing Munc13-1 or RIM-BP2. The fraction of Bassoon pixels and the fraction of Synaptophysin-1 puncta containing Munc13-1 or RIM-BP2 are in Figure S10

(controlR+E n = 6 independent cultures, cKOR+E n = 6, 10 images per culture). All data are means

± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test.

Figure 3.1 (Continued)

Munc13-1, whereas the remaining RIM-BP2 co-localized well with the same markers (Figures

3.1 G and S10 E, F).

This is the most extensive genetic disruption of the vertebrate active zone protein complex to date with a near complete loss of most of the vital components. Somewhat surprisingly, synapse number and size were unchanged (Figure S10 C), and the levels and localization of SNARE proteins and synaptic vesicle markers (Syntaxin-1, SNAP-25 and

Synaptobrevin-2/VAMP-2, and Synaptophysin-1) were not affected (Figures 3.1 B-E, S10 B).

At invertebrate synapses, Liprin- controls presynaptic assembly upstream of ELKS and

RIM (Dai et al. 2006; Kaufmann et al. 2002; Patel et al. 2006; Zhen and Jin 1999). Vertebrates express Liprin- proteins from four genes (Zürner and Schoch 2009). Although it is not well understood which Liprin- isoforms localize to the active zone, and post- and extra-synaptic localization has also been observed, Liprin-2 and Liprin-3 are likely the prominent synaptic

Liprin- isoforms (Spangler et al. 2011; Wyszynski et al. 2002; Zürner et al. 2011). In cKOR+E neurons, Liprin-2 and Liprin-3 localization (Figure 3.1 C), and Liprin-3 levels (Figure 3.1 E) and biochemical solubility (Figure 3.1 D) were unaffected. Notably, Liprin-2 and Liprin-3 antibodies reveal relatively widespread labeling (Figure S10 B), compatible with additional roles for Liprin- outside active zones (Miller et al. 2005). In summary, simultaneous deletion of RIM and ELKS reveals strong, redundant, and active zone specific scaffolding functions for these proteins that were not detected when a single protein family was deleted.

Loss of synaptic vesicle docking but normal postsynaptic assembly upon active zone disruption.

To characterize effects on presynaptic and postsynaptic structure, we fixed cultures by high-pressure freezing and analyzed them using transmission electron microscopy. In

58 agreement with the immunostainings, cKOR+E synapses had normal bouton size and synaptic vesicle numbers (Figures 3.2 A, B). At cKOR+E synapses, we observed a massive, 92% reduction of docked vesicles (Figure 3.2 B). We repeated this analysis in glutaraldehyde fixed tissue (Figures S11 A-C) and saw a similarly robust 85% reduction of docked vesicles. This effect was much stronger than loss of RIMs alone, as assessed with glutaraldehyde fixed tissue

(Kaeser et al. 2011). Furthermore, in cKOR+E neurons we observe a 79% loss of vesicles within

100 nm of the target membrane (which we refer to as tethered vesicles) using high pressure frozen tissue (Figure 3.2 C) and a similar reduction using glutaraldehyde fixed tissue (Figure

S11 C). Knockout mutations for Munc13 and the SNAREs, SNAP-25 and Syntaxin-1 do not have a reduced number of docked vesicles using glutaraldehyde fixed tissue nor a reduction in tethered vesicles, but reveal their functions in tight vesicle docking only with high-pressure freezing and/or tomography (Augustin, Rosenmund, et al. 1999; Bronk et al. 2007; Imig et al.

2014; Siksou et al. 2009; de Wit et al. 2006). Thus, because the cKOR+E docking deficit is easily detected in glutaraldehyde fixed tissue and extends to distances up to 100 nm away from the target membrane, we conclude that these neurons have a very strong deficit in docking and tethering synaptic vesicles to the presynaptic plasma membrane, establishing a requirement for the active zone for these processes.

Because the active zone protein complex interacts through trans-synaptic protein complexes with the PSD (Südhof 2012), it is possible that a strong disruption of the active zone affects the integrity of the PSD. In cKOR+E neurons, the length of the postsynaptic density was not affected (Figures 3.2 B, S11 B), and levels and localization of PSD-95, N-methyl-D- aspartate (NMDA) receptors, and α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid

(AMPA) receptors were not changed (Figures 3.2 D-G). Thus, we conclude that structural effects of RIM and ELKS deletion are largely restricted to the active zone.

Figure 3.2. Disruption of the active zone leads to loss of synaptic vesicle docking but

PSDs appear normal. (A, B) Sample images (A) and quantification (B) of synaptic morphology of high-pressure frozen neurons analyzed by electron microscopy of cKOR+E and controlR+E synapses. For an identical analysis using glutaraldehyde fixed tissue, see Figure S11 (controlR+E n = 50 synapses, cKOR+E n = 50). (C) Distribution of synaptic vesicles relative to the presynaptic plasma membrane area opposed to the PSD. Vesicle distribution is shown in 100 nm bins (left) in cKOR+E and controlR+E synapses. Gaussian fits were used to model the vesicle distribution.

The two genotypes were significantly different (* p < 0.05) and could not be fit with a single distribution, requiring individual fits. Distribution of synaptic vesicles within the first 100 nm in 10 nm bins and the number of tethered vesicles (defined as vesicles within 100 nm of the presynaptic plasma membrane) are shown in the middle and on the right, respectively

(controlR+E n = 50 synapses, cKOR+E n=50). (D, E) Sample images (D) and quantification (E) of postsynaptic protein synaptic fluorescence levels at cKOR+E and controlR+E synapses using confocal microscopy as described in Figures 3.1 B, C. The black dotted line indicates control levels (controlR+E n = 3 independent cultures, cKOR+E n = 3, 10 images per culture). (F, G)

Quantitative Western blotting for PSD proteins using fluorescent secondary antibodies. Sample images (F) and quantification (G) of total postsynaptic protein levels in cKOR+E neurons normalized to protein levels in controlR+E neurons (black dotted line) are shown (controlR+E n = 3 independent cultures, cKOR+E n = 3). All data are means ± SEM; *** p ≤ 0.001 as determined by

Student's t test (B) or * p < 0.05 by extra sum of squares F test (C).

Figure 3.2. (Continued)

We monitored synaptic transmission electrophysiologically in cultured neurons and found that single action potential evoked excitatory or inhibitory postsynaptic currents (EPSCs or IPSCs, respectively) were strongly reduced (by 90% and 81%, respectively) but not abolished (Figures

3.3 A, B, E, F). The rise time of the synaptic response was slowed (Figures 3.3 C, D, G, H), and the variability in amplitude and rise time was strongly increased, suggesting increased asynchrony in cKOR+E synapses.

We next stimulated the synapses with pairs of stimuli at closely spaced time intervals and calculated the paired-pulse ratios (PPRs). PPRs are inversely correlated with initial vesicular release probability P (Zucker and Regehr 2002) and can be used as a relative measure of P when comparing genotypes. Consistent with a strong reduction in release probability, PPR was strongly increased at short inter-stimulus intervals at excitatory (Figures

3.3 I, J) and inhibitory synapses (Figures 3.3 K, L). This decrease is reminiscent of the decrease in release probability observed in RIM knockout synapses (Kaeser et al. 2008; Kaeser et al.

2011; Schoch et al. 2002), which is caused by reduced tethering of presynaptic Ca2+ channels

(Kaeser et al. 2011).

To test whether Ca2+ influx is reduced upon disruption of the active zone, we imaged single action potential evoked presynaptic Ca2+ transients (Figures 3.4 A-C). Briefly, individual neurons were patched and filled with an Alexa594 dye to identify the axon and the presynaptic boutons, and with the Ca2+ indicator Fluo5F that increases fluorescence upon Ca2+ binding (Figure 3.4 A).

After dye filling, a brief somatic current injection was used to induce a single action potential, and Ca2+ transients were recorded in individual boutons and secondary dendrites (Figure 3.4 B).

We found a 44% reduction in the peak amplitude of the Ca2+ influx in boutons, but dendritic Ca2+ transients remained unaffected (Figure 3.4 C). These data match well with the observation of a

2+ loss of CaV2.1 Ca channels (Figure 3.1 C), with the strong reduction in vesicular release probability

Figure 3.3. Single action potential evoked synaptic transmission and release probability are strongly decreased upon disruption of the active zone. (A, B) NMDAR-EPSCs were evoked by a focal stimulation electrode. Example traces (A) and quantitation of EPSC amplitudes (B) and their coefficient of variation (C.V.) in cKOR+E and controlR+E neurons are shown (controlR+E n = 24 cells/4 independent cultures, cKOR+E n = 26/4). (C, D) Sample traces

(C) and quantitation (D) of EPSC rise times and their C.V. (n as in B). Individual sweeps are shown in grey and the average of all sweeps is shown in black. Traces are normalized to the average response. (E – H) Same analysis as in A-D for IPSCs (controlR+E n = 19/3, cKOR+E n =

19/3). (I,J) Analysis of NMDAR-EPSC paired pulse ratios (PPRs) in cKOR+E and controlR+E neurons. Sample traces (I, traces normalized to the first response) and quantitation (J) of the

PPR at 100 ms interstimulus interval (controlR+E n = 23/4 independent cultures, cKOR+E n= 26/4).

(K, L) Scaled sample traces (K, traces normalized to the first response) and summary data (L) of IPSC PPRs at variable interstimulus intervals (controlR+E n = 19/3, cKOR+E n = 19/3). All data are means ± SEM; **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test in A-H, or by two- way ANOVA in J (genotype, interstimulus interval, and interaction p ≤ 0.001, p values of post- hoc Holm-Sidak tests are shown).

Figure 3.3 (Continued)

(Figures 3.3 I-L), and with the previously described roles for RIM and ELKS proteins in enhancing presynaptic Ca2+ influx in hippocampal neurons (Kaeser et al. 2011; C. Liu et al.

2014).

Persistence of release upon active zone disruption.

We next stimulated the cKOR+E neurons with short action potential trains (50 stimuli at 10

Hz). Surprisingly, we detected a strong buildup of release at excitatory cKOR+E synapses starting with the second action potential, and the increase was sustained throughout the action potential train (Figures 3.5 A, B; S12 A). Similarly, vesicles were released quite efficiently throughout an action potential train at inhibitory synapses (Figures 3.5 E, F; S12 C). When we quantified the synchronous charge component throughout the train, we observed a reduction of

50% at excitatory cKOR+E synapses (Figure 3.5 C) and of 62% at inhibitory synapses (Figure 3.5

G). The total charge, the tonic component during the train, and delayed charge starting 100 ms after stimulation ended were affected to a similar extent (Figures S12 B, D). The steady state amplitude at the end of the train was reduced by 41% and 33%, for EPSCs and IPSCs, respectively (Figures 3.5 D, H). Thus, despite loss of docked vesicles, disruption of the active zone and a strong impairment of single action potential induced release, release persisted during trains of action potentials. This finding is consistent with a strong reduction in P, but suggests that an increase in P due to Ca2+ buildup during the stimulus train releases synaptic vesicles quite efficiently. This is reminiscent of a similar electrophysiological phenotype upon deletion of RIM-BP in D. melanogaster (K. S. Y. Liu et al. 2011).

2+ 2+ We next varied the extracellular Ca concentration ([Ca ]ex) and measured the IPSC amplitude. Remarkably, despite the reduction of presynaptic Ca2+ channels and Ca2+ influx,

2+ R+E increasing [Ca ]ex from 2 mM to 5 mM strongly enhanced the IPSC in cKO neurons by a

Figure 3.4. Impaired Ca2+ influx in active zone disrupted neurons. (A) Sample images of cKOR+E and controlR+E neurons filled via patch pipette with Fluo5F and AlexaFluor594 (red, top) and enlarged view of boutons (bottom) analyzed in B. (B) Somatic action potentials (top) and presynaptic Ca2+ transients imaged via Fluo5F fluorescence (bottom) of the color coded boutons shown in A. (C) Summary plots of single action potential-induced Ca2+ transients in boutons, inset: same plot for dendrites. Data are shown as mean (line) ± SEM (shaded area). *** p <

0.001 for Ca2+ transients during the first 60 ms after the action potential as assessed by two-way

ANOVA for genotype and time; interaction n.s. (boutons: controlR+E n = 202 boutons/16 cells/3 independent cultures, cKOR+E n = 157/13/3; dendrites: controlR+E n = 148 dendrites/16 cells/3 independent cultures, cKOR+E n = 100/13/3).

Figure 3.4 (Continued)

67 factor of 2.7 to 4.5 nA (Figures 3.5 I, J). In both conditions, release saturated above 5 mM

2+ 2+ [Ca ]ex likely because Ca influx itself saturates at 5 mM (Ariel and Ryan 2010). When we expressed these data normalized to the largest response (Figure 3.5 K), we observed a right-

2+ shift in the dependence of release on [Ca ]ex (EC50 values as obtained through fitting each cell: controlR+E EC50 = 1.55 ± 0.238, cKOR+E EC50 = 2.33 ± 0.197, *, p < 0.05), confirming a reduction in P. Thus, the active zone per se is not required for fusion of synaptic vesicles, and

2+ when P is increased, for example during action potential trains or by increasing [Ca ]ex, vesicles can be quite efficiently released. Because release is proportional to the product of RRP size and

P, these data suggest that RRP vesicles remain in cKOR+E neurons, despite the loss of RIM,

Munc13, and vesicle docking.

We next assessed the frequency and amplitude of miniature excitatory and inhibitory

PSCs in the presence of tetrodotoxin (mEPSCs, mIPSCs, Figures 3.5 L-N, Figures S12 E, G, H,

I) in cKOR+E synapses, and compared these data with synapses that either lack only RIM (cKOR) or ELKS (cKOE). Simultaneous removal of RIM and ELKS led to surprisingly mild, 47% and 49% reductions in mEPSC (Figure 3.5M) and mIPSC (Figure S12 H) frequencies, respectively. The effect on spontaneous release after disruption of the active zone in cKOR+E synapses was comparable to the deletion of ELKS alone, and was milder than the reduction upon loss of RIM

(78%, Figures 3.5 M, S12 F) or Munc13 (Augustin, Rosenmund, et al. 1999; Varoqueaux et al.

2002) alone. Consistent with the normal architecture of the PSD (Figures 3.2 B, S11 B), mini amplitudes were not affected. Direct comparison of the miniature frequencies in the three lines confirmed a statistically significantly stronger reduction of mini frequency in the synapses that lack only RIM compared to the synapses in which the entire active zone is disrupted (Figure

S12 F).

Figure 3.5. Release during action potential trains and mini release are sustained upon disruption of the active zone. (A-D) Sample traces (A) and quantitation of amplitudes (B), synchronous charge (C), and steady state EPSC amplitude (D, average of the last ten EPSCs) of NMDAR-EPSCs evoked by stimulation trains (10 Hz, 50 stimuli) in cKOR+E and controlR+E neurons (controlR+E n = 17/3, cKOR+E n = 18/3). (E-H) Analysis as in A-D but for IPSCs evoked by stimulation trains (10 Hz, 50 stimuli, controlR+E n = 19/3, cKOR+E n = 19/3). (I-K) Example traces (I) of action-potential evoked IPSCs at [Ca2+]ex of 0.5, 1, 2, 5, and 7 mM in cKOR+E and controlR+E neurons. Absolute IPSC amplitudes (J) and amplitudes normalized to the response at

7 mM [Ca2+]ex (K) are shown (controlR+E n = 8/3, cKOR+E n = 8/3). (L-N) Recordings of mEPSCs in synapses lacking RIM (cKOR), ELKS (cKOE), or both (cKOR+E). In each experiment, control neurons are identical to the respective cKO neurons except that the cre lentivirus is inactive in the control neurons. Sample traces (L) and quantitative analysis of mEPSC frequencies (M) and amplitudes (N) are shown (controlR n = 20/3, cKOR n = 21/3; controlE n = 25/3, cKOE n = 24/3; controlR+E n = 32/5, cKOR+E n = 31/5). For expanded mEPSC traces and mIPSC data, see

Figure S12. All data are means ± SEM; * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001 as determined by

Student's t test (A-H, L-N) or two-way ANOVA in J (genotype: *** p < 0.001; [Ca2+]ex: *** p <

0.001, interaction: *, p ≤ 0.05. p values of post-hoc Holm-Sidak tests are shown in J). For analyses of release components during trains, see Figure S12 A-D.

Figure 3.5 (Continued)

Uniform disruption of synaptic composition and release in cKOR+E neurons.

Thus far, our data reveal a strong reduction in release probability at cKOR+E synapses and show that mini release and release in response to stimulus trains is more mildly impaired than one would predict from the strong structural disruption of the active zone. This suggests that fusion competent vesicles are present despite loss of Munc13, RIM, and vesicle docking.

An alternative explanation is that only a subset of synapses is affected in cKOR+E neurons, and that a population of near normal synapses confounds our analysis. Such heterogeneity could be due to the presence of different neuronal subtypes in our cultures and could arise at the molecular or functional level. We first excluded that the heterogeneity is derived from a population of cells that does not express cre recombinase. Consistent with the analysis of protein levels (Figure 3.1 E), all cells expressed cre (Figures S13 A, B). We then tested whether the active zone components of synapses showed a distribution consistent with heterogeneity.

We plotted the data presented in Figures 3.1 B, C and S10 B-G as a frequency distribution of the fluorescence intensity. Peaks for Munc13, Bassoon (Figure 3.6 A), Piccolo, and RIM-BP

(Figure S13 C) shifted uniformly to lower intensities. These data argue against strong molecular heterogeneity upon active zone disruption.

We next turned to presynaptic imaging in cultures expressing SypHy (Granseth et al.

2006), a version of synaptophysin coupled to an intravesicular pHluorin tag. In brief, neuronal cultures were infected with two independent lentiviruses expressing SypHy (for imaging exocytosis) and SV2-TdTomato (to identify synapse-rich areas in the cultures) at DIV3 in addition to the cre and control lentiviruses (which were supplied at DIV5). At DIV15-18, a synapse dense area was chosen based on SV2-TdTomato expression, and the neurons were stimulated with a focal stimulation electrode for 40 or 200 action potentials at 20 Hz. In this experiment, exocytosis is identified as an increase in fluorescence due to unquenching of the pHluorin when it is exposed to the neutral extracellular pH. For the analysis, only puncta that

Figure 3.6. Uniform disruption of active zone composition and function in cKOR+E neurons. (A) Histograms of the distribution of fluorescence intensity levels in cKOR+E and controlR+E synapses (normalized to the average fluorescence in control). Data are from the experiments shown in Figures 3.1 B, C. For histograms of RIM, ELKS, Piccolo, RIM-BP2, Liprin-

3, and PSD-95 see Figure S13 C. (B) Pseudocolored images of SypHy expressing cultures stimulated with 40 and 200 action potentials (APs), and dequenched with NH4Cl. Images represent peak fluorescence change. (C-E) Quantification of fluorescence changes in cKOR+E and controlR+E neurons stimulated with 40 APs, including the time course of the mean fluorescence change in active synapses as a percentage of the fluorescence increase upon

NH4Cl application (C), the peak response (D) of active synapses, and frequency distribution of the % response in active synapses at the end of the stimulus train (E; controlR+E n = 3493

NH4Cl responsive synapses/ 2486 active synapses/ 9 coverslips/3 independent cultures, cKOR+E n = 2192/1272/9/3, the number of coverslips is used as a basis for statistics). (F-H)

Quantification as in C-D but for neurons stimulated with 200 APs (controlR+E n = 3493/2640/9/3, cKOR+E n = 2291/1405/9/3). All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test.

Figure 3.6 (Continued)

73 showed at least a 2-fold increase in fluorescence upon application of NH4Cl (which uniformly raised the intravesiclular pH to unquench pHluorin fluorescence) were included. We first determined the fraction of synapses responsive to electrical stimulation at both frequencies, and observed a small but significant decrease in active synapses in the cKOR+E neurons (Figure S13

D). We next characterized release at active synapses in both genotypes and found a 67% and

68% decrease in the peak response at the end of stimulation at 40 or 200 action potentials, respectively (Figures 3.6 B-H). When we plotted a frequency histogram of the % of the total pool released at the end of the stimulus train for all active synapses, there was a prominent increase in synapses with smaller pHluorin fluorescence changes in cKOR+E synapses. This experiment supports that active synapses in cKOR+E neurons have impaired release and establishes that the secretory deficit cannot be explained by inactive synapses only. Furthermore, it establishes that vesicle recycling, which contributes strongly to the response to 200 action potentials, is reduced in the cKOR+E synapses. These experiments exclude that a large fraction of synapses has normal active zones and operates at essentially normal levels.

Comparison of docking and RRP size upon active zone disruption.

Our data thus far suggest that there may be a sizable RRP left in cKOR+E synapses even though such a pool is thought to be reflected in docked vesicles at hippocampal synapses (Imig et al. 2014; Rosenmund and Stevens 1996; Schikorski and Stevens 2001; Watanabe et al.

2013). We measured the RRP at excitatory synapses using the application of 500 mM hypertonic sucrose and compared the RRP with vesicle docking and distribution at cKOR, cKOE, and cKOR+E synapses (Figure 3.7). RIM deletion resulted in a 46% reduction in docked synaptic vesicles paralleled by a 75% reduction in the RRP, but the distribution of vesicles within a nerve terminal was normal (Figures 3.7 A-D, S11 A, B). Removal of ELKS alone did not result in a

Figure 3.7. Persistence of a readily releasable pool upon loss of synaptic vesicle tethering and docking. (A-D) Sample images (A) and quantitative analyses of synaptic vesicle docking (B) in glutaraldehyde fixed neuronal cultures, and sample traces (C) and quantitation

(D) of RRP of RIM deficient cKOR and corresponding controlR synapses. Focal application of hypertonic sucrose for 10 s was used to deplete the RRP (D). (E-H) Analyses as outlined in A-

D, but of ELKS deficient cKOE and controlE synapses. (I-L) Analyses as outlined in A-D, but of

RIM/ELKS deficient cKOR+E and controlR+E synapses. For analyses of vesicle numbers, bouton size, PSD length, and vesicle distribution, see Figure S14 A-F. All data are means ± SEM; *p ≤

0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test (analysis of vesicle docking and tethering: controlR n = 25 synapses, cKOR n = 25; controlE n = 25, cKOE n = 25; controlR+E n =

25, cKOR+E n = 25; analysis of RRP: controlR n = 20 cells/3 independent cultures, cKOR n =

20/3; controlE n = 17/3, cKOE n = 17/3; controlR+E n = 20/3, cKOR+E n = 20/3). (M) Schematic of synaptic architecture and function upon disruption of the active zone. Structures and processes that are strongly disrupted upon RIM and ELKS deletion are labeled in yellow (active zone, docking, single action potential mediated release). Synaptic structures and functions that remain at least partially intact are labeled in green (the synaptic vesicle cluster, the postsynaptic density containing neurotransmitter receptors, mini release, and release in response RRP depleting stimuli such as action potential trains or hypertonic sucrose). Our experiments indicate that at least some vesicles can be recruited from vesicle pools distant from the presynaptic plasma membrane for release, and that these vesicles may be released immediately or undergo a transient docking state (dotted arrow) that is initiated after the onset of stimulation.

Figure 3.7 (Continued)

76 detectable effect on vesicle docking or vesicle distribution (albeit there was a non-significant trend towards a small reduction in docked vesicles), but induced a 34% reduction in RRP at excitatory synapses (Figures 3.7 E-H, S14 C, D). At cKOR+E synapses the RRP was more mildly affected (Figures I-L) than one would predict from the effects observed in cKOR or cKOE synapses, from the loss of docking, and from the massive reduction in RIM, Munc13, and other active zone proteins (Figure 3.1). 42% of RRP vesicles remained despite the strong reduction in vesicle docking (89% in glutaraldehyde fixed tissue, Figure 3.7 J; 92% in high pressure frozen tissue, Figure 3.2 B). Direct comparison of the three genotypes revealed a significantly stronger loss of docking in cKOR+E synapses compared to cKOR synapses. Conversely, RRP was more strongly reduced in cKOR synapses than in cKOR+E or cKOE synapses (Figures S14 G, H).

These data suggest that at least some RRP vesicles can be recruited over distance and do not have to be stably docked at the active zone before the application of hypertonic stimulus.

Discussion

We here establish a conditional mouse mutant that strongly and specifically disrupts the active zone matrix and synaptic vesicle docking in cultured hippocampal neurons (Figures 3.1 and 3.2). We find that disruption of the active zone results in a strong impairment of vesicular release probability, but surprisingly >40% of RRP vesicles remained (Figures 3.3 to 3.7).

Redundant scaffolding functions of RIM and ELKS.

The multi-domain structure of RIM and ELKS and their extensive biochemical binding activities with other active zone proteins suggested that they operate as scaffolds (Ohtsuka et al. 2002; Schoch et al. 2002; Takao-Rikitsu et al. 2004; Yun Wang et al. 2002). However, loss of function approaches thus far provided mixed support for this hypothesis. Knockout mutants for

ELKS1 and/or ELKS2 showed no changes in active zone composition at hippocampal synapses

(Held, Liu, and Kaeser 2016; C. Liu et al. 2014), except for a small increase in the biochemical solubility of RIM (Kaeser et al. 2009). Similarly, RIM1 and/or RIM2 knockout mice revealed isolated changes in the clustering and levels of Munc13-1 (Deng et al. 2011; Schoch et al.

2002). Beyond these changes in individual active zone proteins however, the active zone protein complex was intact. Here, we reveal a strong, redundant scaffolding role for RIM and

ELKS: simultaneous removal leads to disruption of the active zone with a loss of three out of four additional active zone protein families. Our morphological and functional analyses further strongly support redundant scaffolding roles for RIM and ELKS that are similar at excitatory and inhibitory synapses. Thus, we establish an important presynaptic clustering role for RIM and

ELKS that is shared across synapses and that tethers Piccolo, Bassoon, Munc13-1, and RIM-

BP2. Because these proteins cannot be anchored and maintained in levels at mutant synapses,

RIM and ELKS are necessary and thus upstream for their tethering to the active zone.

Interestingly, levels and localization of Liprin-2 and Liprin-3, the two Liprin- isoforms that are strongly expressed in brain and are thought to be localized at the active zone (Spangler et al. 2011; Zürner and Schoch 2009; Zürner et al. 2011), are not affected in our mutants. This suggests that Liprin-2/3 are either upstream in active zone assembly and can be tethered independent of RIM and ELKS, or that Liprin-2/3 are not part of the same protein complex.

Genetic experiments have firmly established presynaptic roles for Liprin-/syd-2 in synapse assembly in C. elegans and D. melanogaster (Kaufmann et al. 2002; Zhen and Jin 1999).

Although the localization of individual vertebrate Liprin- proteins has not been conclusively solved and the available data support pre-, post-, or extra-synaptic localization of Liprin- proteins (Spangler et al. 2011; Wyszynski et al. 2002; Zürner et al. 2011), a recent study employed knockdowns for Liprin-2 and supported presynaptic scaffolding functions (Spangler et al. 2013). In vitro binding of Liprin-1 through 4 to RIM and/or ELKS (Ko et al. 2003; Schoch

78 et al. 2002) provide further support for a presynaptic scaffolding role. One possible explanation for these and our data is that Liprin- is upstream of RIM and ELKS in vertebrate active zone assembly. Importantly, invertebrate Liprin-/syd-2 mutant synapses also have reduced vesicle numbers in the nerve terminal (Kaufmann et al. 2002; Patel et al. 2006; Zhen and Jin 1999) suggesting that the active zone may recruit vesicles to a presynaptic nerve terminal. Our experiments at vertebrate synapses reveal that synaptic vesicle numbers are unchanged upon active zone disassembly, establishing that the active zone protein complex downstream of

Liprin- is not required for recruitment of vesicles to the nerve terminal. Our data are consistent with additional roles for Liprin- outside of the active zone, for example in trafficking of vesicles or active zone material, as has been shown for D. melanogaster Liprin- (Miller et al. 2005).

Further genetic experiments will be necessary to dissect the roles of Liprin- in the vertebrate brain.

Recent studies support that synaptic and network activity contribute to active zone protein turnover (Lazarevic et al. 2011; Sugie et al. 2015; Weyhersmüller et al. 2011). It is thus possible that loss of synaptic activity in the cultured neurons contributes to the strong active zone disruption that we observe upon RIM/ELKS deletion. However, reduced activity is unlikely to play an important causative role for active zone disruption in our experiments because knockouts for only RIM (Deng et al. 2011) or Munc13 (Varoqueaux et al. 2002), which have similar or more severe reductions in activity, do not lead to strong active zone disruption. In the long-term, it will be important to test causes and effects of the active zone disruption we describe here in a system that allows manipulation and characterization of a specific synapse in a defined circuit to better understand how cell-type specificity and activity contribute to the phenotypes.

The role of the active zone in synaptic vesicle docking.

Because synaptic vesicles are only docked at the active zone (Couteaux and Pécot-

Dechavassine 1970; Imig et al. 2014; Siksou et al. 2009; Südhof 2012), it has been proposed that the active zone provides the molecular mechanism for docking of synaptic vesicles to the target membrane. Consistent with this hypothesis, RIM1/2 double knockout synapses have an approximately 50% reduction in the number of docked vesicles in cultured hippocampal neurons in glutaraldehyde fixed tissue. Importantly, using the same method, no other presynaptic protein, including Munc13 (Augustin, Rosenmund, et al. 1999; Varoqueaux et al. 2002), synaptobrevin-2 (Deák et al. 2004), SNAP-25 (Bronk et al. 2007), or CAPS (Jockusch et al.

2007) have a role in synaptic vesicle docking. Compellingly, disruption of the active zone in the cKOR+E neurons leads to a near complete loss of vesicle docking in glutaraldehyde fixed tissue

(Figures S11 B, 3.7 J). Recent experiments have used high-pressure freezing and tomography, which improved the resolution in the analysis of docking to two nanometers. Using this method, it has been found that loss of function mutations for Munc13, CAPS, and the SNAREs syntaxin-

1, synaptobrevin-2, and SNAP-25 have a strong reduction in synaptic vesicles within 0-5 nm of the target membrane, but normal or increased vesicle numbers in bins at 10 and 20 nm away from the active zone (Imig et al. 2014; Siksou et al. 2009). The reduction of docked vesicles at cKOR+E synapses is apparent with both fixation methods. With high-pressure freezing, the 92% reduction at cKOR+E synapses (Figure 3.2 A-C) is similar to Munc13 null mutants, which have an

96% reduction in docked vesicles when using the same method combined with electron tomography (Imig et al. 2014). Furthermore, unlike Munc13 deficient synapses, cKOR+E synapses fail to accumulate vesicles 10-20 nm away from the target membrane, but show a

75% reduction in numbers of tethered vesicles within 100 nm of the target membrane. Thus, we conclude that the loss of docking and tethering of synaptic vesicles in the cKOR+E mutant is stronger than in previous mutants because the loss of docking is readily detected in

80 glutaraldehyde fixed tissue and there is a shift of the entire vesicle cluster away from the target membrane that has not been seen in other mutants. We conclude that the active zone is required for synaptic vesicle docking and tethering.

The relationship between synaptic vesicle docking, priming, and release.

The strong decrease in single action potential mediated release (Figure 3.3) corresponds well with the loss of vesicle docking (Figure 3.2 ) and is consistent with the hypothesis that single action potentials release docked vesicles (Rosenmund and Stevens

1996). Release from a single synapse is proportional to the RRP size and P (Zucker and

Regehr 2002). Our analysis revealed a strong reduction in P upon disruption of the active zone

(Figures 3.3 I-L, 3.4, 3.5 I-K). This observation is supported by a strong increase in PPRs, a

2+ right-shift in the [Ca ]ex dependence of release, an increase in C.V. of the PSC amplitude, and a loss of presynaptic CaV2.1 Ca2+ channels and Ca2+ influx. Surprisingly, manipulations that enhanced P (increasing [Ca2+]ex and action potential trains) or bypassed the need for Ca2+

(stimulation with hypertonic sucrose) demonstrated that a significant pool of vesicles is available for release in cKOR+E synapses.

Many studies have defined the RRP as vesicles that are either released during the transient response to hypertonic sucrose (Augustin, Betz, et al. 1999; Deng et al. 2011;

Rosenmund and Stevens 1996; Varoqueaux et al. 2002) or during brief trains of action potentials (Schikorski and Stevens 2001) and RRP size estimates determined by these methods are well correlated with the number of morphologically docked vesicles. Using hypertonic sucrose in cKOR+E neurons, 42% of the RRP remained (Figure 3.7 L), which is significantly larger than the RRP left after deletion of RIM (Figure S14 H and (Kaeser et al. 2011)) or

Munc13 (Augustin, Betz, et al. 1999; Varoqueaux et al. 2002) despite the more severe docking

81 deficit at cKOR+E synapses. This challenges the notion that the priming functions of these proteins are identical to their functions in vesicle docking. Furthermore, spontaneous miniature release (Figures 3.5 L-N, S12 G, H) and release during action potential trains (Figures 3.5 A-H,

3.6 C-H) are also more mildly reduced than expected. Due to the strong reduction in P at cKOR+E synapses, it was not possible to measure RRP size using high frequency action potential stimulation (Thanawala and Regehr 2016). Nevertheless these data reveal that fusion competent vesicles can be recruited over distance and do not require a persistently docked state. RRP vesicles may therefore be stored away from the presynaptic plasma membrane, at least in the absence of an active zone, as has been proposed based on experiments that labeled RRP vesicles after recycling (Rizzoli and Betz 2004).

It is possible that hypertonic sucrose stimulation leads to a transient increase in vesicle docking that is not captured in our electron-microscopic images. This may also be the case for

2+ some of our other manipulations, for example high [Ca ]ex or prolonged stimulus trains. The small amount of remaining Munc13 in the cKOR+E neurons may rapidly add vesicles to the RRP, but this hypothesis implies that Munc13 dependent priming in the cKOR+E neurons occurs upstream or simultaneous to contact with the target membrane and is transient, since there are no stably docked vesicles. The relatively mild reduction in mini frequency supports that docking in cKOR+E neurons is transient and not a stable state of a primed vesicle. Alternatively, massive disruption of the active zone may bypass the need for Munc13 to prime vesicles before fusion.

A long-standing question has been whether the partial assembly of SNARE complexes is required for synaptic vesicle docking and priming (Jahn and Fasshauer 2012; Südhof and

Rothman 2009). Our experiments reveal that SNARE proteins are present in the nerve terminal upon disruption of the active zone, and that synaptic vesicle fusion, which is mediated by

SNARE proteins, is not abolished. However, SNARE proteins are not sufficient to drive docking

82 at synapses in the absence of an active zone, suggesting that not all fusion competent vesicles require stable preassembly of SNARE complexes.

Finally, the molecular mechanisms that underlie docking upstream of SNARE complex assembly are poorly understood. With a gene mutation that disrupts docking, but leaves synaptogenesis and presynaptic vesicle clustering intact, analysis of the minimal protein interactions between synaptic vesicles and release sites required for docking will now be possible.

— CHAPTER FOUR —

Redundant scaffolding mechanisms target CaV2 family voltage-gated calcium channels

to the presynaptic active zone.

Richard G. Held, Changliang Liu, Shan Shan H. Wang, Arn van den Maagdenberg, Toni R.

Schneider, and Pascal S. Kaeser.

A.vdM and T.R.S. created the floxed alleles of Cacna1a and Cacna1e respectively. C.L. performed calcium imaging experiments. SS.H.W. processed and analyzed electron microscopy experiments. R.G.H. performed and analyzed all other experiments.

Introduction

Calcium is perhaps the most pervasive cellular signaling molecule and is broadly utilized by all cells to trigger key processes from gene transcription to local conformational changes

(Clapham 2007). This is facilitated by a steep calcium concentration ([Ca2+]) gradient between the cytoplasm ([Ca2+] ~ nM) and extracellular space ([Ca2+] ~ mM). Electrically excitable cells use this gradient to translate changes in membrane potential into elevations in intracellular calcium via the opening of voltage-gated calcium channels (VGCCs) on the plasma membrane.

VGCC opening results in rapid local changes in calcium concentration proximal to the pore of the channel (Neher 1998). These [Ca2+] elevations are quickly returned to resting levels by the combined action of calcium buffers, extrusion, and uptake into internal stores, making them ideal for triggering spatially and temporally precise signaling events (Tank, Regehr, and Delaney

1995).

In vertebrates, 10 genes encode the pore-forming 1 subunits of VGCCs (Dolphin

2016). They are subdivided into three gene families, CaV1, CaV2, and CaV3, which are distinguished by their sequence similarity, biophysical properties, and sensitivity to agonists and antagonists (Ertel et al. 2000; Randall and Tsien 1995). Structurally, the channel consists of four six-pass transmembrane domains that form the channel pore and voltage-sensing domains

(Tanabe et al. 1987; J. Wu et al. 2015). CaV1 and CaV2 channels also have large intracellular C- terminal tails which include an EF-hand and an IQ-motif proximal to the membrane followed by an unstructured region that includes many protein interaction motifs (Ben-Johny et al. 2014;

Kaeser et al. 2011). In addition to the 1 subunits, several auxiliary subunits are part of the

VGCC complex. CaV subunits interact with the 1 subunit by binding to the intracellular loop between domains I and II of 1 (Y.-H. Chen et al. 2004; J. Wu et al. 2015; J. Wu et al. 2016).

2 subunits are extracellular proteins, and interact with extracellular loops of 1 in addition to

85 being tethered to the plasma membrane (J. Wu et al. 2015; J. Wu et al. 2016; Davies et al.

2010).

It has been understood for decades that Ca2+ influx is essential for synaptic transmission

(Katz and Miledi 1965; Dodge and Rahamimoff 1967). The CaV2 family, which includes CaV2.1,

2+ CaV2.2, and CaV2.3, are the primary channels that support Ca influx at conventional synapses to initiate vesicle fusion (Gasparini, Kasyanov, Pietrobon, Voronin, and Cherubini 2001b; L. Li,

Bischofberger, and Jonas 2007; L.-G. Wu, Borst, and Sakmann 1998; Hirning et al. 1988; Cao et al. 2004). They open in response to action potentials, allowing Ca2+ sensor proteins on synaptic vesicles, most prominently synaptotagmins, to bind calcium and trigger SNARE- mediated vesicle fusion(Fernandez-Chacon et al. 2001). This process is spatially restricted to a region within the presynaptic terminal known as the active zone. The active zone acts as a molecular scaffold which links CaV2 VGCCs to the SNARE fusion machinery and ‘docks’ synaptic vesicles in close proximity to the plasma membrane (Südhof 2012). It is also positioned directly opposite the synaptic cleft from postsynaptic neurotransmitter receptors, resulting in a spatially aligned trans-synaptic signaling complex (Biederer, Kaeser, and Blanpied 2017; Perez de Arce et al. 2015; Couteaux and Pécot-Dechavassine 1970).

Individual gene knockouts for CaV2 channels are not sufficient to abolish neurotransmitter release and often show extensive compensation by related channels (Jun et al.

1999; Urbano et al. 2003; Inchauspe et al. 2004; Cao et al. 2004; Ino et al. 2001; Lee et al.

2002). Given their essential nature for triggering vesicle fusion, remarkably few studies have addressed this compensation and studied the functions of CaV2 channels using comprehensive gene knockouts for all relevant channels (tong nishimune**). Many studies have examined the mechanisms that localize and organize CaV2 channels at the active zone, either in wild-type synapses or using overexpression. CaV2.1 channels are clustered at the active zone and their number scales with the size of the active zone (Holderith et al. 2012). These channels are

86 between 10 to >100 nm from a docked synaptic vesicle, depending on the synapse, and in some cases as few as a single channel is required to trigger fusion (Stanley 1993; Arai and

Jonas 2014; Vyleta and Jonas 2014; Meinrenken, Borst, and Sakmann 2002; Bucurenciu et al.

2008). These spatial relationships can have significant effects on the efficacy and reliability of synaptic vesicle fusion (Nakamura et al. 2015; Keller et al. 2015). A clear understanding of the structural relationship between CaV2 channels and the protein complex at the active zone is therefore essential.

The active zone protein RIM is thought to play a major role in this relationship. RIM interacts via its PDZ domain with the distal C-terminal tail of CaV2.1 and 2.2, and knockout of major RIM isoforms leads to a dramatic reduction in CaV2.1 levels at the active zone (Han et al.

2011; Kaeser et al. 2011). Another active zone protein, RIM-BP, directly interacts via an SH3 domain with a PxxP motif also in the C-terminal of CaV2 channels (Hibino et al. 2002). Knockout of RIM-BP in invertebrate results in mislocalization of VGCCs and a dramatic loss of synaptic vesicle fusion, but vertebrate knockouts have much more subtle effects on channel to vesicle coupling distance (Acuna et al. 2015; Grauel et al. 2016; K. S. Y. Liu et al. 2011). Similarly, the vertebrate active zone protein Bassoon has a role in scaffolding CaV2.1 channels through RIM-

BPs (Davydova et al. 2014). These studies support a model in which CaV2 channels are tethered at the active zone in order to serve their function as a voltage-dependent Ca2+ source for vesicle fusion.

Alternatively, several studies have suggested that CaV2 channels and their auxiliary subunits are themselves structural organizers of the active zone. At the vertebrate NMJ constitutive knockouts of either CaV2.1, CaV2.2, or both disrupt synapse ultrastructure and active zone composition (Nishimune, Sanes, and Carlson 2004; J. Chen, Billings, and

Nishimune 2011). In the retina, both CaV1.4 and the auxiliary 24 subunit are essential for the structure of synaptic ribbons, specialized structures at the active zone of these cells (X. Liu et al.

2013; Kerov et al. 2018; Yuchen Wang et al. 2017; Mansergh et al. 2005). 2 subunits have also been implicated in synapse formation and the growth and elaboration of axons and dendrites, though these functions may or may not require the 1 subunit (Eroglu et al. 2009;

Kurshan, Oztan, and Schwarz 2009; Brodbeck et al. 2002; Tedeschi et al. 2016). Several studies have also proposed a role for the C-terminal tail of CaV2 channels in docking synaptic vesicles at the active zone (Lübbert et al. 2017; F. K. Wong et al. 2014). Taken together these studies suggest a model that places CaV2 channels as active structural organizers of the active zone. There are therefore two competing hypothesis: Ca2+ channels could either position and organize the active zone, or the active zone could control the tethering and positioning of Ca2+ channels.

In order distinguish between these two fundamentally different models, we set out to test whether CaV2 channels have a structural role in active zone assembly by genetically removing presynaptic channel proteins. To comprehensively test the functions of VGCCs at the active zone we generated conditional knockout mice in which the entire CaV2 gene family can be conditionally ablated by application of cre recombinase (CaV2 cTKO). Conditional deletion of

CaV2 channels in cultured neurons circumvented compensation by other VGCCs and massively impaired action-potential evoked synaptic transmission. We find that CaV2 channels and, remarkably, Ca2+ entry through these channels, are dispensable for the integrity of the active zone complex, for synapse formation, and for synaptic vesicle docking at the active zone.

Interestingly, expression and localization of most auxiliary CaV subunits depended on CaV2 1 expression. Strikingly, however, the extracellular 21 does not colocalize with CaV2 1 channel proteins at the active zone, and its presynaptic localization is increased in the absence of CaV2

1 subunits. Hence 21 proteins are likely not a stably associated Ca channel subunit in a presynaptic nerve terminal, but they can localize and function independent of the presence of

88 the pore-forming CaV2 1 proteins. Finally, we find that the C-terminal tail of CaV2.1 localizes the channel to the active zone likely via redundant interactions with the active zone.

Experimental Procedures

All experiments were performed in cultured hippocampal neurons as previously described

(Chapter One, Chapter Two, and Appendix Two). Briefly, neurons were infected at DIV 5 with lentiviruses expressing cre recombinase or an inactive mutant of cre under a synapsin promoter, and experiments were performed at DIV 15-18. For lentiviral expression of HA-tagged

CaV2.1, neurons were infected at DIV1. For details on the production of lentivirus, HPF electron microscopy, confocal imaging, and Western blotting see Appendix Two. Materials and methods specific to this chapter are detailed below.

Mouse lines. All animal experiments were performed according to institutional guidelines at

Harvard University. CaV2.2 (Cacna1b) knockout mice were obtained from the IMPC (IMSR Cat#

KOMP:CSD34514-1a-Wtsi, RRID:IMSR_KOMP:CSD34514-1a-Wtsi) and crossed to transgenic mice expressing Flp recombinase (Dymecki 1996) to remove the inserted neo cassette and generate a conditional allele. Conditional triple knockout (cTKO) mice that remove CaV2.1,

CaV2.2, and CaV2.3 proteins were generated by crossing conditional Cacna1a (Todorov et al.

2006), Cacna1b, and Cacna1e (Pereverzev et al. 2002) mice. CaV2 cTKO mice were maintained as triple homozygote line.

cDNA and CaV2 mutant cloning. CaV2.1 (pCR3.1) was a gift from Diane Lipscombe (Addgene plasmid # 26573; (Richards et al. 2007). The CaV2.1 open-reading frame (ORF) was subcloned

89 into the pCMV5 backbone with a Human influenza hemagglutinin (HA) tag inserted between

CaV2.1 residues V27-G28 using Gibson assembly (Gibson et al. 2008). For CaV2.1 Ct, the

PDZ entire ORF after residue I1774 was replaced with a stop codon. For CaV2.1 C , the final four

SH3 residues (2366DDWC2369) were removed, again followed by a stop codon. For CaV2.1 C ,

polyR residues 2240PQTP2243 were mutated to 2240AQTA2243. For CaV2.1 C , residues S2016-

R2060 were deleted but the remaining CaV2.1 sequence (G2061-C2368) was retained. All wild- type and mutant constructs were then subcloned into lentiviral backbones driven by the human synapsin promoter (C. Liu et al. 2014).

Electrophysiology. All electrophysiology experiments were performed as described in previous chapters (Chapter 2, Appendix Two) with two modifications. First, all experiments were conducted in 1.5 mM extracellular Ca2+ and 2 mM extracellular Mg2+ in the bath. Second, for recording AMPAR-mediated action potential and sucrose evoked EPSCs, 1 mM -DGG was added to the bath to block receptor saturation (Clements et al. 1992). This had the added benefit of reducing the overall excitation and reverberant activity in the culture on action potential stimulation (Maximov et al. 2007).

Antibodies. The antibodies and concentrations used in this study are as follows. Monoclonal mouse anti-Bassoon (N-terminal, 1:1000; RRID: AB_11181058); rabbit anti-Liprin-α3 (serum

4396, 1:5000, gift from Dr T. Sudhof); anti-ELKS2α (serum E3-1029 , 1:100; custom made); rabbit anti-RIM1 (1:1000; RRID: AB_887774); rabbit anti-RIMBP2 (1:1000; RRID: AB_2619739); rabbit anti-Munc13-1 (1:1000; RRID: AB_887735); rabbit anti-CaV2.1 (1:200; RRID:

AB_2619841); rabbit anti-CaV2.2 (1:200; custom made); monoclonal mouse anti-CaV1 (1:100;

RRID: AB_2619841); monoclonal mouse anti-CaV2 (1:100; RRID: AB_10675135); rabbit anti-

CaV3 (1:500; RRID: AB_2039787); monoclonal mouse anti-CaV4 (1:100; RRID:

AB_10671176); rabbit anti-21 (1:500; AB_2039785); monoclonal mouse anti-21 (1:250;

AB_1078663); monoclonal mouse anti-PSD-95 (1: 200; RRID: AB_10698024); guinea pig anti- vGlut1 (1:500; AB_887878); rabbit anti-Synapsin 1 (1:1000; RRID: AB_2200097); guinea pig anti-synaptophysin (1:500; AB_1210382); rabbit anti-rabphilin (1:1000; a gift from Dr. T.

Sudhof); rabbit anti-MAP2 (1:500; AB_2138183); monoclonal mouse anti-MAP2 (1:500;

AB_477193).

STED imaging. For preparing cultured neurons for STED imaging, coverslips were washed two times with warm PBS, then fixed for 10 minutes in 2% PFA + 4% sucrose (in PBS). Following fixation coverslips were rinsed twice in PBS + 50 mM glycine, then blocked/permeabilized in

PBS + 0.1% Triton X-100 + 3% BSA (TBP) for 1 hour. Primary antibodies were diluted in TBP and coverslips were stained overnight at 4C. Coverslips were then rinsed twice and washed 3-4 times for 5 minutes in TBP. Alexa Fluor 488, 555, and 633 were used as secondary antibodies at 1:200 (488 and 555) or 1:500 (633) dilution in TBP. Secondary antibody staining was done overnight at 4C followed by rinsing two times and washing 3-4 times 5 minutes in TBP. Stained coverslips were then post-fixed for 10 minutes with 4% PFA + 4% sucrose in PBS, rinsed two times in PBS + 50 mM glycine, then once in deionized water. Coverslips were allowed to air-dry, protected from light, then mounted on glass slides.

STED images were acquired with a Leica SP8 Confocal/STED 3X microscope with an oil immersion 100X 1.44 numerical aperture objective at the Harvard NeuroDiscovery Center

Enhanced Imaging Core. 46.51 x 46.51 µm2 areas were selected as regions of interest (ROIs) and were scanned at a pixel density of 4096 x 4096 (11.358 nm/pixel). Alexa 633, Alexa Fluor

555, and Alexa Fluor 488 were excited with 633 nm, 555 nm and 488 nm white light lasers respectively at 2-5% of 1.5 mW laser power in sequence by frame. The Alexa Fluor 633 channel

91 was acquired first in confocal mode using 2X frame averaging. Subsequently, Alexa Fluor 488 and Alexa Fluor 555 channels were acquired in STED mode, depleted with 592 nm (80% of max power) and 660 nm (30% of max power) depletion lasers respectively. Line accumulation (4-

10X) and frame averaging (2X) were applied during STED scanning. Identical settings were applied to all samples within an experiment. For all analyses of STED images, synapses in side view were defined as synapses that contained a synaptic vesicle cluster with an elongated PSD-

95 or Bassoon structure along the edge of the vesicle cluster. For line profile analysis, synapses were selected using only the PSD-95/Bassoon signal and the vesicle signal. An ROI was manually drawn around the PSD-95/Bassoon signal and fit with an ellipse to determine the center position and angle from normal. A ~1.3 m long, ~225 nm wide line was then drawn using these coordinates, selected perpendicular to the elongated PSD-95/Bassoon structure across its center. An intensity profile was obtained for all three channels along this region. To align individual profiles, the PSD-95/Bassoon signal was smoothed using a 5 pixel moving average and the smoothed signal was used to define the point of peak intensity. All three channels (vesicle marker, test protein, non-smoothed PSD-95/Bassoon) were then aligned to this peak position and averaged across images. All analyses were performed on raw images without background subtraction or adjustments, and were done identically for all experimental conditions. Representative images were brightness and contrast adjusted to facilitate inspection. For all image acquisition and analyses comparing two or more conditions, the experimenter was blind to the condition.

92 profiles, a Gaussian fit was made to the profile of the CaV2 control or full-length CaV2 rescue conditions and the standard deviation of the fit was used to define an analysis window around the control peak. Within that window, two-way ANOVA tests were performed with genotype and position as the two factors.

Results

Genetic removal of all presynaptic CaV2 calcium channels.

To test for structural roles of the CaV2 channel protein in synapse assembly we generated conditional triple knockout animals (CaV2 cTKO) in which the genes for all three pore- forming members of the CaV2 family (Cacna1a, Cacna1b, Cacna1e) were targeted (Todorov et al. 2006; Pereverzev et al. 2002). Using primary cultures of hippocampal neurons from these

CaV2 cTKO mice we generated control and knockout conditions via infection with a lentivirus expressing either cre-recombinase (CaV2 cTKO) or a non-functional cre as control (CaV2 control) (Figure 4.1 A). Western blotting confirmed removal of CaV2.1 and CaV2.2 channels

(Figure S15 A,B), whereas CaV2.3 channels could not be detected by Western blotting, but were removed as assessed by mass spectrometric analyses of protein composition of the cultured neurons (data not shown).

In cultures from CaV2 cTKO neurons, there was a near complete loss (~95%) of action- potential evoked synaptic transmission as measured by both AMPA receptor-mediated excitatory postsynaptic currents (eEPSCS) and GABAA receptor-mediated inhibitory postsynaptic currents eIPSCs (Figure 4.1B-E). This is consistent with an essential role of CaV2 channels in triggering vesicle fusion and demonstrates that simultaneous removal of CaV2 family channels successfully circumvents compensation by other channel types. In the presence of TTX, the frequency of miniature excitatory postsynaptic currents (mEPSCs), but not their amplitude, was significantly reduced (Figure 4.1 F-H) consistent with a dependence of

2+ Figure 4.1. Loss of Ca triggered exocytosis in CaV2 cTKO neurons. (A) Gene targeting strategy for the generation of CaV2.1 (Todorov et al. 2006), CaV2.2, and CaV2.3 (Pereverzev et al. 2002) conditional alleles. All three mouse lines were crossed to generate a homozygous

CaV2 conditional triple knockout line (CaV2 cTKO). (B,C) Sample traces (B) and quantification

(CaV2 control, n = 16 cells/3 independent cultures, CaV2 cTKO n = 15/3). (D,E) Sample traces

(D) and quantification (E) of the amplitude of GABAAR-mediated evoked inhibitory postsynaptic currents (eIPSCs) (CaV2 control, n = 20 cells/4 independent cultures, CaV2 cTKO n = 22/4). (F-

H) Sample traces (F) and quantification of miniature EPSC frequency (G) and amplitude (H)

CaV2 control, n = 24 cells/4 independent cultures, CaV2 cTKO n = 22/4). (I,J) Sample traces (I) and quantification (J) of the EPSC charge transfer during the first 10 seconds of superfusion with hypertonic (+500 mOsm) sucrose (CaV2 control, n = 19 cells/3 independent cultures, CaV2 cTKO n = 19/3). All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by

Student's t test.

Figure 4.1 (Continued)

95 spontaneous vesicle fusion on Ca2+ influx through VGCCs (Ermolyuk et al. 2013; Williams et al.

2012). When we stimulated release of the entire readily-releasable pool (RRP) of vesicles using superfusion with hypertonic sucrose, the sucrose evoked EPSC charge was unchanged in CaV2

TKO neurons (Figure 4.1 I,J). These data indicate that the loss of action-potential evoked

2+ release is likely due to the loss of Ca influx through CaV2 channels not a reduction in RRP size.

Consistent with previous reports, removal of individual CaV2 channels had no effect on eIPSCs (Figure S15 C). However, removal of either CaV2.1 (CaV2.1 cKO) or CaV2.2 (CaV2.2 cKO) abolished the sensitivity of eIPSCs to the specific antagonists -agatoxin IVA and - conotoxin GVIA respectively, confirming that loss of either CaV2.1 or 2.2 is efficiently compensated (Figure S15 D-G). Removal of only CaV2.1 and CaV2.2 (CaV2.1/2 cKO) resulted in almost complete loss of eIPSC amplitude, almost identical to removal of all three CaV2 channels (Figure S15 H), demonstrating that presynaptic CaV2.3 channels alone cannot support release at these synapses. In summary, our data establish that CaV2 channels are essential for synaptic transmission. Importantly, CaV2.3 and L- (CaV1) or T- type (CaV3) channels are not able to compensate for the combined loss of CaV2.1 and 2.2.

CaV2 channels are not required for active zone scaffolding or synapse ultrastructure.

To test the structural the role of CaV2 channels at the active zone we first looked at synaptic expression levels of other components of the active zone protein complex using confocal microscopy. Synaptic expression levels of RIM1, ELKS2, Munc13-1, RIM-BP2,

Bassoon, and Liprin-3 were all unchanged (Figure S16 A,B). Synapse density was also

2+ normal, suggesting that synaptic transmission and presynaptic Ca influx through CaV2 channels is not required for synapse formation (Figure S16 C). To assess overall cell

Figure 4.2. STED imaging of active zone composition in CaV2 cTKO neurons.

(A) Representative images of CaV2 control and CaV2 cTKO ‘side view’ synapses stained for

PSD-95 (red channel, STED), the vesicle marker VGluT (blue channel, confocal), and CaV2.1

(green channel, STED). Scale bars are 200 nm. (B) Line profiles for CaV2.1 (left) and PSD-95

(right) taken from ~225 nm wide ROIs drawn perpendicular to the center point of PSD-95 ‘bars’ and aligned to the peak of PSD-95 (see experimental methods for details). Abscissa indicate the distance from the peak of PSD-95 (marked with a dashed line). CaV2 control, n = 61 synapses/3 independent cultures, CaV2 cTKO n = 58/3. (C,D) Representative images (C) and quantification

(D) of CaV2 control and CaV2 cTKO ‘side view’ synapses stained for PSD-95 (red channel,

STED), the vesicle marker VGluT (blue channel, confocal), and target proteins of interest (green channel, STED). Scale bars are 200 nm. RIM: CaV2 control, n = 56/3, CaV2 cTKO n = 55/3;

RIM-BP2: CaV2 control, n = 61/3, CaV2 cTKO n = 60/3; Liprin-3: CaV2 control, n = 60/3, CaV2 cTKO n = 55/3; ELKS2: CaV2 control, n = 59/3, CaV2 cTKO n = 60/3; Munc13-1: CaV2 control, n

= 59/3, CaV2 cTKO n = 57/3. Statistical comparisons were performed using two-way ANOVA over a window that was one standard deviation around the peak of the control. All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001.

Figure 4.2 (Continued)

98 morphology, we filled cells with a soluble Alexa dye via a whole cell patch pipette. There were no significant changes in process length or number between CaV2 control and CaV2 cTKO neurons as assessed by Sholl analysis (Figure S16 D,E). Taken together these data indicate that removal of presynaptic CaV2 channels does not cause dramatic changes to the formation of synapses.

In order to rule out nanoscale changes in the structure of the active zone, which cannot be detected at the resolution of confocal microscopy, we turned to super-resolution stimulated emission-depletion (STED) microscopy (Hell and Wichmann 1994). We selected ‘side-view’ synapses for analysis based off staining for a vesicle marker with closely opposed, bar-like

PSD-95 staining (M. Y. Wong et al. 2018). CaV2.1 exhibited presynaptic bar-like localization directly opposed to PSD-95 (Figure 4.2 A,B). This signal was specific, as it was nearly abolished in the CaV2 cTKO, and the remaining signal is likely antibody background. We next examined the other active zone proteins RIM1, ELKS2, Munc13-1, RIM-BP2, and Liprin-3. All five proteins exhibited the same characteristic bar-like morphology (Figure 4.2 C). By making line- profiles through side-view synapses and aligning them to the peak of PSD-95, we found no significant changes in the peak intensity or position of RIM1, RIM-BP2, or Liprin-3 (Figure 4.2

D). However, Munc13-1 was significantly increased in the CaV2 TKO and there was a small but significant increase in ELKS2, possibly due to a homeostatic response to the release deficit in these neurons.

Several studies have proposed a role for CaV2 channels in the ultrastructural docking of synaptic vesicles at the active zone (Lübbert et al. 2017; F. K. Wong et al. 2014). To test this hypothesis, we evaluated the ultrastructure of CaV2 cTKO synapse using electron microscopy on samples fixed using high-pressure freezing followed by freeze-substitution (Figure 4.3 A).

There were no differences in the number of docked synaptic vesicles, the total number of vesicles, or length of the postsynaptic density, indicating that the overall ultrastructure of CaV2

Figure 4.3. Ultrastuctural analysis of CaV2 cTKO synapses. (A) Representative electron microscope images of CaV2 control and CaV2 cTKO synapses fixed using high-pressure freezing, followed by freeze substitution. (B-F) Quantification of the number of docked (B), tethered (C, defined as vesicles within 10 nm of the membrane), and total vesicles (D), as well as bouton area (E) and PSD length (F). (CaV2 control, n = 102 synapses/2 independent cultures, CaV2 cTKO n = 102/2)

100 cTKO neurons is normal (Figure 4.3 B-F). These data speak against a model in which CaV2 channels alone act to dock vesicles in close proximity to the membrane at the active zone.

Together, these experiments establish that there is, strikingly, no strong role for CaV2 channels, or Ca2+ entry through them, in synapse formation, active zone assembly, and vesicle docking.

This finding largely rejects the hypothesis that Ca2+ channels are synaptogenic molecules that anchor the active zone.

Localization of auxiliary CaV subunits.

CaV2 family channels are thought to exist as multi-protein channel complex consisting of the pore-forming 1 subunit, an intracellular auxiliary subunit, and an extracellular 2 subunit

(Dolphin 2016). In heterologous cells, co-expression of all three subunits is essential for efficient trafficking of the channel to the cell surface (Cantí et al. 2005; Davies et al. 2010). Similarly, in neurons levels of the extracellular 2 subunit are thought to control surface expression of the channel complex at the active zone (Hoppa et al. 2012; Schneider et al. 2015). It is unclear however whether the auxiliary subunits require the 1 subunits for their expression and localization.

We therefore looked at the synaptic expression levels of the four CaV subunits using confocal microscopy. CaV1, 3, and 4 were significantly reduced in the absence of CaV2 1 subunits (Figure 4.4 A,B). Interestingly, CaV2 levels were unchanged in the CaV2 cTKO, indicating they do not require 1 subunits for proper localization. This is consistent with previous reports that CaV2 can localize to the plasma membrane when expressed alone in heterologous cells due to a palmitoylation site (Leroy et al. 2005). We next tested whether the distrubution of

CaV4 was consistent with its localization at the active zone using STED microscopy. CaV4 formed bar-like structures parallel to the active zone protein Bassoon at inhibitory synapses

101

Figure 4.4. Synaptic expression of auxiliary CaV subunits. (A) Representative images of

CaV1-4 (green channel) along with synaptophysin (red channel) and MAP2 (blue channel). (C)

Quantification of synaptic levels using synaptophysin puncta to define ROIs. The black dotted line indicates control levels. Scale bar = 5 m. For all conditions: CaV2 control n = 3 independent cultures, CaV2 cTKO n = 3, 10 images per culture). (C) Representative images of

CaV2 control and CaV2 cTKO ‘side view’ synapses stained for Bassoon (red channel, STED), the vesicle marker VGAT (blue channel, confocal), and CaV4 (green channel, STED). Scale bars are 200 nm. (D) Line profiles of CaV4 signal taken from ~225 nm wide ROIs drawn perpendicular to the center point of Bassoon ‘bars’ and aligned to the peak of Bassoon (CaV2 control, n = 39 synapses/2 independent cultures, CaV2 cTKO n = 40/2). (E) Western blot showing CaV4 levels in samples prepared from CaV2 control and CaV2 cTKO cultures.

Rabphilin was used as a loading control. All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤

0.001 as determined by paired t-test. For STED analysis, statistical comparisons were performed using two-way ANOVA over a window that was one standard deviation around the peak of the control.

102

Figure 4.4 (Continued)

103

(Figure 4.4 C). This signal was absent in CaV2 cTKO synapses (Figure 4.4 C,D) indicating that

CaV4 is present at the active zone and that it requires CaV2 1 subunits for proper localization.

Qualitative Western blotting revealed a dramatic reduction in CaV4 levels, supported this conclusion and suggesting that total expression levels of CaV4 depend in part on its association with CaV2 1 (Figure 4.4 E).

The extracellular 2 subunit of the VGCC channel complex has been proposed to play a role in synapse formation, axon outgrowth, and active zone structure (Eroglu et al. 2009; Kurshan,

Oztan, and Schwarz 2009; Tedeschi et al. 2016; Yuchen Wang et al. 2017; Brodbeck et al.

2002; Fell et al. 2016). 21 and 23 are thought to be the major isoforms expressed in the hippocampus and 21 in particular has been shown to regulate presynaptic levels of CaV2 1 subunits (Klugbauer et al. 1999; Hoppa et al. 2012). We tested synaptic expression levels of

21 using confocal microscopy (Figure 4.5 A,B). Synaptic levels of 21 were unchanged in

CaV2 cTKO neurons (Figure 4.5 B). Localization of 21 using STED microscopy revealed that

21 staining was punctate and appeared to be distributed both presynaptically and postsynaptically (Figure 4.5 C,D). Interestingly, the presynaptic component was significantly increased in CaV2 TKO synapses (Figure 4.5 D). Consistent with our confocal imaging data, we saw no changes in expression levels of 21 by Western blotting (Figure 4.5 E). The increase in presynaptic 21 seen in STED may therefore reflect a change in the distribution of 21. While the exact relationship between the CaV2 1 subunits and 21 remains unclear, these data establish that 21 does not require CaV2 1 subunits for its expression. Furthermore, its broad distribution is inconsistent with other active zone proteins, CaV4, and CaV2.1 (compare Figures

4.2 and 4.4 C,D with Figure 4.5 C,D). This is surprising, given these proteins known functional relationship in channel trafficking and association as subunits of the same channel complex in

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Figure 4.5 Expression and localization of 21. (A) Representative images of the 21 subunit (green channel) along with synaptophysin (red channel) and MAP2 (blue channel). (B)

Quantification of synaptic levels using synaptophysin puncta to define ROIs. The black dotted line indicates control levels. Scale bar = 5 m. CaV2 control n = 3 independent cultures, CaV2 cTKO n = 3, 10 images per culture. (C) Representative images of CaV2 control and CaV2 cTKO

‘side view’ synapses stained for PSD-95 (red channel, STED), the vesicle marker VGluT (blue channel, confocal), and 21 (green channel, STED). Scale bars are 200 nm. (D) Line profiles of 21 signal taken from ~225 nm wide ROIs drawn perpendicular to the center point of PSD-

95 ‘bars’ and aligned to the peak of PSD-95 (CaV2 control, n = 35 synapses/2 independent cultures, CaV2 cTKO n = 39/2). (E) Western blot showing 21 levels in samples prepared from

CaV2 control and CaV2 cTKO cultures. -actin was used as a loading control. All data are means

± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001. For STED analysis, statistical comparisons were performed using two-way ANOVA over a window that was one standard deviation around the peak of the control.

105

Figure 4.5 (Continued)

106 biochemical purifications , but is consistent with a previous report that could not find stable association using proteomic assessment of CaV channel complexes (C. S. Müller et al. 2010).

The C-terminus is essential for synaptic delivery and active zone anchoring of CaV2.

CaV2 cTKO neurons appear to specifically remove CaV2 family channels with no compensation and minimal additional effects on active zone structure. We therefore set out to use structure-function rescue experiments to test the necessity of the intracellular C-terminus and its known protein interaction motifs for localization to the active zone. We first tested the importance of the CaV2.1 C-terminal tail, which contains most of the sequences proposed to be important for channel localization (Maximov and Bezprozvanny 2002; Kaeser et al. 2011; Hibino et al. 2002; Lübbert et al. 2017). Rescue CaV2.1 was HA-tagged at the N-terminal in a region that does not affect channel function (Mark et al. 2011), and expressed in CaV2 cTKO neurons using lentiviral infection at DIV1. Two rescue conditions were tested, a full-length construct

(CaV2.1) and one in which the C-terminal tail of the channel was truncated after residue I1774, just after the last transmembrane domain (CaV2.1 Ct) (Figure 4.6 A). Expression of HA-CaV2.1 was able to restore eIPSC amplitude and charge in CaV2 cTKO neurons to levels that appeared slightly below control (Figure 4.6 B-D). Expression of CaV2.1 Ct had no effect on eIPSCs however, and was not significantly different from the CaV2 cTKO condition. Confocal imaging revealed that CaV2.1 Ct was indeed expressed, but appeared to be primarily localized in the soma and proximal dendrites (Figure 4.6 E-G). This suggests that the CaV2.1 C-terminal tail is necessary for proper trafficking to nerve terminals.

We next wanted to test which protein interactions within the C-terminal are necessary for targeting CaV2.1 to the active zone. Three sequences have been suggested to be important in the literature: a PDZ-binding motif (DDWC) at the very C-terminal of the channel that binds the

107

Figure 4.6. Structure-function rescue with CaV2.1. (A) Schematic representation of CaV2 control, CaV2 cTKO, full-length CaV2.1 rescue, and CaV2.1-Ct rescue. (B) Representative traces of eIPSCs in CaV2 control, CaV2 cTKO, full-length CaV2.1 rescue, and CaV2.1-Ct rescue conditions. (C,D) Quantification of eIPSC amplitude (C) and charge transfer (D) (CaV2 control, n

= 18 cells/3 independent cultures; CaV2 cTKO, n = 15/3; + CaV2.1 n = 17/3; + CaV2.1-Ct, n =

16/3). All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by one-way

ANOVA followed by Holm-Sidak multiple comparisons post-hoc testing comparing each condition to CaV2 cTKO. (E) Representative images showing staining for PSD-95 (red channel),

HA (green channel, CaV2.1 rescue), and the synaptic vesicle marker VGluT (blue channel) in

CaV2 cTKO, full-length CaV2.1 rescue, and CaV2.1-Ct rescue neurons. (F,G) Quantification of fluorescent intensity of HA staining in ROIs containing synapses, defined by synaptophysin staining (B), or the soma, defined by nuclear GFP fluorescence/somatic background staining of

PSD-95 (C). Somatic ROIs were excluded from synaptic ROIs (CaV2 cTKO, n = 6 images/3 independent cultures; + CaV2.1 n = 6/3; + CaV2.1-Ct, n = 6/3). All data are means ± SEM; *p ≤

0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by one-way ANOVA followed by Holm-Sidak multiple comparisons post-hoc testing comparing each condition to full-length CaV2.1 rescue.

108

Figure 4.6 (Continued)

109

PDZ domain of RIM (Kaeser et al. 2011) and is conserved in Cav2 channels, a PxxP motif that interacts with an SH3 domain in RIM-BP (Hibino et al. 2002) and is conserved in CaV1 and CaV2 channels, and a recently identified region (S2016-R2060) with unknown protein interactions that contains a poly-Arginine sequence and an additional PxxP motif and is only present in CaV2.1

(Lübbert et al. 2017). We generated rescue constructs to disrupt each of these sequences: a

PDZ truncation of the PDZ-binding DDWC motif (CaV2.1 C ), a PxxP to AxxA mutation (CaV2.1

SH3 polyR C ), and a deletion mutant that selectively removes the S2016-R2060 region (CaV2.1 C )

(Figure 4.7 A). All three mutant channels were able to rescue eIPSC amplitude to a similar extent as full length CaV2.1, but there was a significant reduction in eIPSC charge transfer in the

SH3 CaV2.1 C condition (Figure 4.7 B-D).

We next used STED microscopy to assess active zone localization of the rescue channels. Similar to wild-type CaV2.1 (Figure 4.2 A,B), full length HA-tagged CaV2.1 formed bar-like structures parallel at the active zone (Figure 4.8 B,C). Compellingly, CaV2.1 Ct, which lacks the entire C-terminal, was absent from the active zone, consistent with our physiology data. The CPDZ, CSH3, and CpolyR mutant channels were able to localize to the active zone in

PDZ STED imaging, but there were significant reductions in the peak intensity of both CaV2.1 C

SH3 and CaV2.1 C (Figure 4.8 C). Synaptic expression levels, as assessed by confocal imaging, were comparable between all rescue conditions (Figure S17 A-C). These data suggest that no single protein interaction is necessary for localization of CaV2.1 channels to the active zone.

Instead multiple, redundant interactions are likely employed, with significant contributions of the

RIM and RIM-BP binding sites.

110

Figure 4.7. Structure-function rescue with CaV2.1 C-terminal binding mutants. (A)

PDZ Schematic representation of CaV2 cTKO, full-length CaV2.1, CaV2.1-C (RIM binding

SH3 polyR mutant), CaV2.1-C (RIM-BP binding mutant), and CaV2.1-C mutant rescue. (B)

Representative traces of eIPSCs in the same conditions. (C,D) Quantification of eIPSC amplitude (C) and charge transfer (D) (CaV2 cTKO, n = 15 cells/3 independent cultures; +

PDZ SH3 polyR CaV2.1 n = 18/3; + CaV2.1-C , n = 17/3; + CaV2.1-C , n = 17/3; + CaV2.1-C , n =

18/3). All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by one-way

ANOVA followed by Holm-Sidak multiple comparisons post-hoc testing comparing each condition to full-length CaV2.1 rescue.

111

Discussion

2+ Despite decades of study, complete gene knockout of presynaptic CaV2 Ca channels has never been used to assess the functions of this critically important gene family. We now

establish such knockouts and show that CaV2 channels do not play a structural role in the assembly of the presynaptic active zone at hippocampal synapses (Figure 4.2). The recruitment of auxiliary CaV subunits largely depends on the presence of CaV2 1 subunits, but the extracellular 21 subunit does not colocalize with CaV2 at the active zone and does not require

CaV2 1 for its expression and synaptic localization (Figure 4.4 and 4.5). Finally, we show that the targeting of CaV2.1 to the active zone requires the C-terminal tail and that this scaffolding likely occurs through multiple protein interactions at the active zone (Figures 4.6-8)

Interaction between CaV2 1 and auxiliary subunits

Our data demonstrate that CaV2 1 subunits are involved in the expression and localization of most CaV subunits but not of the extracellular 21 (Figure 4.4-5). The 2 subunit was initially identified as a component of the dihydropyridine receptor, the CaV1.1 channel complex, in muscle cells (Curtis and Catterall 1983; Tanabe et al. 1987). In these cells it co-purifies well CaV1.1 channel and the structure of this complex has been studied using cryoEM (J. Wu et al. 2016; J. Wu et al. 2015). 2 has also been copurified with CaV2.2 channels from the brain (Witcher et al. 1993). Furthermore, 2 subunits are important for proper trafficking of 1 pore-forming subunits to the membrane in heterologous expression systems and expression of 21 sets the synaptic abundance of CaV2 channels in hippocampal neurons (Hoppa et al. 2012; Cantí et al. 2005; Davies et al. 2010). Given these data 2

112

Figure 4.8 STED imaging of CaV2.1 rescue localiation. (A) Schematic representation of CaV2

PDZ SH3 cTKO, full-length CaV2.1, CaV2.1-C (RIM binding mutant), CaV2.1-C (RIM-BP binding

polyR mutant), and CaV2.1-C mutant rescue. (B) Representative images of ‘side view’ synapses stained for Bassoon (red channel, STED), the vesicle marker VGAT (blue channel, confocal), and HA (green channel, STED, CaV2.1 rescue). Scale bars are 200 nm. (E) Line profiles of HA

(left) and Bassoon (right) fluorescence intensity taken from ~225 nm wide ROIs drawn perpendicular to the center point of Bassoon ‘bars’ and aligned to the peak of Bassoon (CaV2 cTKO, n = 59 synapses/3 independent cultures; + CaV2.1 n = 60/3; + CaV2.1-Ct, n = 57/3; +

PDZ SH3 polyR CaV2.1-C , n = 60/3; + CaV2.1-C , n = 56/3; + CaV2.1-C , n = 57/3). Abscissa indicate the distance from the peak of Bassoon (marked with a dashed line). Statistical comparisons were performed using two-way ANOVA over a window that was one standard deviation around the peak of the full length CaV2.1 rescue. All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p

≤ 0.001.

113

Figure 4.8 (Continued)

114 subunits clearly have the potential to bind 1 subunits. However, whether they remain associated with CaV2 channels at the active zone is less clear.

Proteomic analysis of the CaV2 channel complex suggests that 2 subunits are not tightly associated with 1 (C. S. Müller et al. 2010). Gene knockouts of a1 subunits, including our current data, show variable requirements of 1 subunits for the expression of 2. In C. elegans and at rod photoreceptors, CaV1.4 and Unc-2, the CaV2.2 homologue, control 24 and

23 expression, respectively (Tong et al. 2017; Yuchen Wang et al. 2017). In contrast, we find that CaV2 channels are not required for 21 expression in hippocampal neurons and that the synaptic distribution of these two proteins is distinct (Figure 4.2 A,B compared to Figure 4.5

C,D). There are four 2 genes in vertebrates, of which 21 and 23 are the major isoforms expressed in the hippocampus at the mRNA level (Dolphin 2016; Klugbauer et al. 1999;

Klugbauer, Marais, and Hofmann 2003). Unfortunately, we were unable to find specific antibodies against 23. It remains possible that a different 2 subunit is the major isoform associated with CaV2 channels at the active zone. However, knockdown or overexpression of

21 at hippocampal synapses in culture controls surface expression of CaV2 channels suggesting functional relevance of the 21-CaV2 subunit pairing (Hoppa et al. 2012; Schneider et al. 2015).

Scaffolding roles of CaV2 channels

The role of presynaptic Ca2+ channels in synapse structure and formation has been contentious. Many synapses, including the neuromuscular junction of vertebrates and flies and the ribbon synapses of rod photoreceptors, require the presence of either the dominant 1 or

2 isoform for proper assembly of the active zone (Nishimune, Sanes, and Carlson 2004; J.

115

Chen, Billings, and Nishimune 2011; Mansergh et al. 2005; X. Liu et al. 2013; Kerov et al. 2018;

Yuchen Wang et al. 2017; Eroglu et al. 2009; Wycisk et al. 2006; Kurshan, Oztan, and Schwarz

2009; Pirone et al. 2014; Fell et al. 2016; Brodbeck et al. 2002). In contrast, structural effects of

1 knockouts on small CNS synapses have not been reported. Many synapses coexpress multiple CaV2 channels, primarily CaV2.1 and CaV2.2, leading to the possibility that compensation has masked the structural roles of CaV2 channels (L.-G. Wu et al. 1999; Cao et al. 2004; L. Li, Bischofberger, and Jonas 2007; L.-G. Wu, Borst, and Sakmann 1998). Our data now support a model in which CaV2 channels are recruited to the active zone to serve as sources of Ca2+ for triggering vesicle fusion, but are not part of the scaffolding architecture of the active zone.

There are several possible explanations for these contrasting results. First, in the case of the ribbon synapses of rod photoreceptors, the dominant channel that triggers release is CaV1.4

(Mansergh et al. 2005). It is possible that non-conserved sequences in CaV1 family calcium channels are involved in active zone structure at the synapses which employ these channels.

However, at inner hair cells, which have similar ribbon-type synapses but utilize CaV1.3 to trigger fusion, there is no structural effect of CaV1.3 knockout (Nemzou N et al. 2006). A second possibility is that synapses that employ distinctive active zone architecture, such as synaptic ribbons, the Drosophila T-bar, or the highly stereotyped peg and beam architecture of the

NMJ,are more sensitive to structural perturbations than small CNS synapses. To our knowledge these are the only types of synapses that have reported structural requirements for CaV 1 subunits. The structural defects found in these synapses are dramatic and include changes in ultrastructure, synaptic expression of active zone proteins, and trans-synaptic alignment

(Nishimune, Sanes, and Carlson 2004; J. Chen, Billings, and Nishimune 2011; X. Liu et al.

2013; Mansergh et al. 2005). It is interesting to note that such effects are rarely seen in small

CNS synapses. Comparable disruptions of active zone structure have only been detected when

116 multiple protein families were targeted, and even in those cases the effects on the postsynaptic compartment were either minimal or not present (Acuna, Liu, and Südhof 2016) Chapter 3).

A final possibility, though not mutually exclusive, is that loss of 2, rather than the 1 subunit, is the cause of synaptic structural defects at these synapses. The literature suggests a correlation between synapses at which 1 subunits are required for 2 localization and those in which 1 subunits affect synapse structure and formation. At the ribbon synapse of rod photoreceptors knockout of CaV1.4 decreases 24 expression, and vice versa, and in both mutants presynaptic proteins are lost (X. Liu et al. 2013; Mansergh et al. 2005; Kerov et al.

2018; Yuchen Wang et al. 2017). Conversely, the ducky mutation, which causes a truncation of

22, disrupts transsynaptic alignment in inner hair cells, but it is unclear whether it requires

CaV1.3 expression for its localization (Fell et al. 2016; Brodbeck et al. 2002). Loss of CaV1.3 at

IHC synapses does not disrupt transsynaptic alignment, suggesting that it 22 is unaffected

(Nemzou N et al. 2006). Similarly, loss of 23 at the NMJ of Drosophila disrupts bouton formation but loss of cacophony, the Drosophila CaV2 homolog, does not (Kurshan, Oztan, and

Schwarz 2009). Furthermore, similar to our data, the localization of overexpressed 23 does not match that of cacophony, suggesting they do not form stable interactions at the synapse

(Kurshan, Oztan, and Schwarz 2009). These data suggest that at some synapses 2may be a mandatory subunit of Ca2+ channels. At these synapses loss of either 1 or 2proteins leads to impairments in synapse assembly that are selectively mediated by the synaptogenic roles of

2. At small hippocampal synapses, however, 2proteins localize and function independent of 1, and hence structural effects are not observed in the absence of 1 subunits. These findings emphasize that phenotypes associated with loss of 2 functions cannot be assumed to be related to CaV 1 subunits.

117

Localization of CaV2 channels to the active zone

The molecular mechanisms that target and scaffold CaV2 channels at the active zone are of critical importance to the control of release probability at the synapse. The exact clustering, coupling distance, and cooperativity of calcium channels at the active zone has been studied both at the ultrastructural level using immunogold labeling, and biophysically by probing channel positioning and domain overlap using exogenous calcium buffers and calcium channel antagonists (Mintz, Sabatini, and Regehr 1995; Adler et al. 1991; Holderith et al. 2012;

Matveev, Bertram, and Sherman 2009; Vyleta and Jonas 2014). Our experiments are now able to address which sequences within CaV2 channels themselves are required for proper trafficking and localization at the active zone.

At the molecular level, it is well established that RIM knockouts disrupt CaV2 localization to decrease release probability and that RIM-BP knockouts impair the reliability of release (Han et al. 2011; Kaeser et al. 2011; Acuna et al. 2015; Grauel et al. 2016). These results are in line with work that demonstrated a role for the C-terminal portion of CaV2.2 in localizing transfected, overexpressed channels in wild-type neurons (Maximov and Bezprozvanny 2002), where channel fragments that contained RIM and RIM-BP binding motifs localized as punctate structures in the axon. We find that the while the C-terminal tail of CaV2.1 is essential for active zone targeting, multiple interactions within the C-terminal are likely to redundantly tether CaV2.1 at the active zone.

Interestingly, neither the RIM-BP or the RIM binding sites are absolutely required to hold

CaV2.1 channels at the active zone in our experiments, but instead removal of either interaction site has mild effects on channel localization (Figure 4.8). This is consistent with earlier work that found that large portions of the C-terminal must be removed before CaV2.1’s contributions to triggering vesicle fusion are reduced (Lübbert et al. 2017). Furthermore, another study found that a splice variant of CaV2.1 lacking the RIM and RIM-BP binding sites restores CaV2.1’s

118 contribution to release in CaV2.1 constitutive knockouts (Cao et al. 2004). An important difference between these previous studies and our current data, however, is that CaV2 channel redundancy was not previously addressed, since CaV2.2 channels were present. It is possible that a protein network at the active zone is established by the presence of any CaV2 channels and can tether channels even if some channels lack intracellular interactions. This could involve binding to CaV subunits, for example, which are lost in our CaV2 cTKO mutants. Alternatively, extracellular proteins such as Neurexins could establish such a scaffolding network (Nishimune,

Sanes, and Carlson 2004; Tong et al. 2017).

Taken together these data suggest that either another sequence in CaV2.1 is able to scaffold it to the active zone or that multiple redundant interactions are necessary. Interestingly, the most significant effect on CaV2.1 localization in our experiments was when we disrupted the

RIM-BP binding site. All CaV2 C-terminal regions have many PxxP motifs throughout their C- terminal sequence, suggesting that RIM-BP may have multiple interactions with the channel. It will be interesting to determine whether disrupting these sites in addition to the known RIM-BP or RIM binding sites has an additive effect on channel mislocalization.

Finally, an important next step will be to establish whether CaV2.1-Ct mutants are functional on the plasma membrane or trapped in an internal compartment. If the former is the case, we may be able to identify mechanisms by which CaV channels are targeted into the axon itself. An interesting experiment will be to domain swap the C-terminal region of CaV2.1 with a typically somatic VGCC, for example an L-type channel, and then to test whether the C- terminus is sufficient to confer active zone localization. Now that we have established this structure-function rescue system, we should be able to answer many fascinating questions about how different calcium channels are localized and function in different cellular compartments.

119

— CHAPTER FIVE —

Discussion and Outlook

120

My work, together with other investigations, establishes that protein scaffolding at the presynaptic active zone is accomplished by multiple, redundant interactions. This scaffolding strategy makes the active zone a resilient structure with built-in redundancy, as is needed for an essential but complex molecular machine. However, it complicates the study of the active zone protein complex, since removal of a single protein family is not sufficient to disrupt it.

In Chapter 2, I addressed the functional roles of the active zone protein ELKS at excitatory and inhibitory synapses in cultured hippocampal neurons. This work originated from our efforts to generate a knockout mouse line which disrupted active zone structure. Since

ELKS interacts with almost every other protein family at the active zone and its proteins sequence suggests strong scaffolding, we hypothesized that ELKS may be an essential active zone scaffold. In earlier work we found that ELKS1/2 is not required for this function, but ELKS knockout had significant effects on synaptic transmission (Kaeser et al. 2009; C. Liu et al.

2014). I explored this further and found that ELKS functions via distinct synaptic mechanisms to control vesicle fusion. At inhibitory synapses, ELKS controls presynaptic Ca2+ influx, boosting it to increase release probability. At excitatory synapses, ELKS controls the size of the RRP of vesicles. We further found that the regions within ELKS that are necessary for these two functions are distinct, supporting the idea that they operate via separable molecular mechanisms.

In Chapter 3, I described our first successful attempt to disrupt the active zone by generating compound gene knockouts for both the RIM and ELKS protein families. Loss of these two proteins led to a strong, specific loss of other active zone material, a near complete loss of synaptic vesicle docking, and a comparable loss of vesicle fusion in response to single action potentials. However, despite the loss of docked synaptic vesicles, fusion still occurred in the form of miniature release, action potential evoked release in high extracellular calcium, and release in response to stimulation with hypertonic sucrose. These data suggest that disruption

121 of the active zone protein complex and loss of vesicle docking are not equivalent to abolishing synaptic vesicle release sites. While docked synaptic vesicles are fused rapidly in response to a single action potential, an intuitive match with their proximity to the membrane with which they fuse, our data establish that a stable docked state is not required for synaptic vesicle fusion.

Vesicles can be recruited to fuse from relatively far back in the terminal and fusion can be maintained during trains, indicating that the mechanisms are still in place to mobilize vesicles and direct them to fuse at sites opposed to postsynaptic receptors. The relationship between synaptic vesicle docking and the concepts of release sites and the RRP are discussed in more detail below.

Interestingly, a study that was published at the same time as our own revealed a very similar phenotype when both RIM and RIM-BP proteins were simultaneously removed (Acuna,

Liu, and Südhof 2016). That study found a similar magnitude effect on synaptic vesicle docking but a more mild loss of other active zone proteins. Of note, Bassoon was reduced in both our

RIM/ELKS mutants and the RIM/RIM-BP mutants, perhaps supporting a model in which

Bassoon acts as a vesicle tethering protein.

Finally, in Chapter 4, I analyzed neurons in which all presynaptic CaV2 family calcium channels were removed. This project was initiated to test the hypothesis that CaV2 removal leads to a disruption in active zone structure, and to identify molecular ‘nucleation points’ of release sites. CaV2 channels are intuitively appealing as potential nucleation points for a macromolecular complex that defines release sites. They are by necessity close to the vesicles destined to fuse and, as transmembrane proteins, have the potential to provide a crucial link to the plasma membrane that other active zone proteins are noticeably missing. However, in spite of evidence from other synapses, I found that CaV2 channels are not essential organizers of the active zone. While the CaV2 1 subunits are involved in the expression and localization of most auxiliary CaV subunits, they are surprisingly non-essential for expression and localization of the

122 extracellular 21 subunit. In keeping with the model of the active zone as a highly redundant scaffolding complex, removing the interaction motifs that allow CaV2.1 to bind RIM or RIM-BP had only small effects on CaV2.1 localization at the active zone and on vesicle fusion. Instead, removal of the entire C-terminal tail of CaV2.1 was necessary.

In light of my findings, the following sections reflect my views on the purpose of redundant protein scaffolding at the active zone, the structure of active zone protein complex and how it relates to classical biophysical models of synaptic transmission, and the relationship between the functional parameters of release and the molecular mechanisms that they reflect.

Redundant mechanisms for scaffolding: fidelity and flexibility

The reasons for redundant protein scaffolding at the active zone are perhaps obvious on first glance. Synaptic transmission is essential to survival and cognitive function and therefore needs to be maintained at all costs. There are however less immediate implications of a redundant scaffolding scheme.

The work presented in Chapter 2 demonstrates that individual active zone proteins can operate via distinct mechanisms at different synapses. There are two interpretations of these data. First, specific isoforms or splice variants could be targeted to different synapses. The bMunc13-2 isoform of Munc13, for example, binds ELKS1 via its N-terminal coiled-coil domain and is localized to only a subset of synapses in the hippocampus (Kawabe et al. 2017).

Inhibitory and excitatory synapses also have different requirements for Munc13. Release can be completely abolished at excitatory synapses by knockout of Munc13-1 alone, but inhibitory synapses require the removal of both Munc13-1 and Munc13-2 for the same effect (Varoqueaux et al. 2002). In the case of ELKS, this does not seem to be the case however, as ELKS1 and

ELKS2 expressed at both excitatory and inhibitory synapses in cultured hippocampal neurons

123 and their levels co-vary. Furthermore, proteomic analyses of revealed very few differences in the composition of active zones between synapse types, supporting a model where active zone composition is largely homogenous (Boyken et al. 2013). However, it is likely that the coarse division of synapses into excitatory and inhibitory is insufficient to reveal subtle differences in active zone composition.

An alternative role for redundant scaffolding interactions would be to support the diversity of release properties and plasticity mechanisms by differentially engaging different protein interactions. This model, though not mutually exclusive, would not require different synapses to express specific isoforms or splice variants of active zone proteins. Instead, different interactions would be favored depending on the protein nano-environment and information transfer requirements of the synapse in question. Redundant scaffolding could be essential to this. As a hypothetical example, if the interaction between CaV2 channels and RIM were simultaneously essential for localizing of CaV2 and boosting release probability, one function would be difficult to separate from the other. If CaV2 localization was controlled by both

RIM and RIM-BP however, a synapse with a requirement for high release probability might employ the RIM-CaV2 interaction to decrease channel coupling distance while a synapse with low release probability could not favor that interaction without entirely losing CaV2 channels.

This might also address why some domains of active zone proteins have protein interactions with multiple active zone proteins with the potential to compete. The RIM PDZ domain, for example, binds to both the GIWA motif of ELKS and the DDWC motif of CaV2.1 and CaV2.2

(Kaeser et al. 2011). Whether such a flexible protein interaction system exists and how it might be regulated is an interesting question for future research. Redundant protein scaffolding mechanisms therefore have the potential to confer functional flexibility as well as robustness.

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Specialized active zone structures

A major unanswered question is why some synapses are more easily disrupted at the ultrastructural level than the small CNS synapses studied in this work. A common theme is apparent when looking at the instances of such disruption: specialized active zone architecture is always involved. The neuromuscular junction, the Drosophila T-bar, and ribbon synapses all exhibit characteristic ultrastructure that is not seen at the active zones of hippocampal synapses. The NMJ in particular has been shown to have beautiful, highly stereotyped morphology using EM tomography, consisting of a repetitive a central ‘beam’ with extending

‘ribs’ and multiple ‘pegs’ thought to be VGCCs (Harlow et al. 2001). Similarly, the T-bar and synaptic ribbons are relatively large and electron-dense, easily visible by EM using conventional fixation methods. The active zone of CNS synapses, in contrast, is best visualized using protein staining with phosphotungstic acid (PTA) after which it appears as a hexagonal grid of dense projections (Pfenninger et al. 1969).

Active zone proteins are all large multi-domain proteins that often have multiple functions in vesicle fusion. It is possible that scaffolding redundancy is conferred simply because of this, since a given active zone protein interacts with multiple binding partners. Nevertheless, that these various functional domains are all present in individual proteins, as opposed to being separated into a larger number of smaller proteins, suggests that ‘induced proximity’, selected for by evolutionary processes, is an essential part of their function. It is tempting to speculate that at the synapses where protein scaffolding is more easily perturbed, induced proximity is less essential and therefore redundant scaffolding mechanisms are not employed. This is at odds with both the release characteristics of these synapses and the domain structure of their essential scaffolding proteins. Ribbon synapses require tight coupling of calcium channels via

RIM-BP for efficient vesicle fusion, for example, and both BRP and RIBEYE are both large

125 proteins with multiple functional roles (Wagh et al. 2006; Hallermann, Kittel, et al. 2010; Kittel et al. 2006; Lv et al. 2016).

An interesting possibility is that specialized structures like the T-bar and synaptic ribbons do not represent the active zone per se but are distinct molecular complexes. In the case of the ribbon there is evidence to support this model. Knockout of either Bassoon or CaV1.4 leads to loss of synaptic ribbons (X. Liu et al. 2013; Mansergh et al. 2005; tom Dieck et al. 2005; Dick et al. 2003). In Bassoon knockout animals ribbons appear to be detached from the membrane,

‘floating’ in the cytoplasm. These detached ribbons segregate from active zone proteins, including CaV1.4, Munc13, ELKS, and RIM2, but remain colocalized with Piccolo and RIM1 (tom

Dieck et al. 2005). This suggests that the ribbon is a separable protein complex from the underlying active zone (Prescott and Zenisek 2005). Interestingly, in CaV1.4 and 24 knockout mice the only active zone proteins that were tested to establish structural disruption were

Bassoon and RIBEYE, leaving open the possibility that the underlying active zone structure is intact in these mutants (Yuchen Wang et al. 2017; X. Liu et al. 2013) (though see western blot levels of ELKS in (Kerov et al. 2018)).

Molecular identity of Katz’ release sites

The exact molecular definition of a synaptic vesicle release site is of great interest, since it would finally unify decades of functional biophysics and molecular biology. Katz developed the quantal hypothesis and the concept of a release site at the frog NMJ. This synapse is in fact a compound synapse, similar to the Calyx of Held, where many active zones are contained within a single nerve ending (Sätzler et al. 2002). Most small CNS synapses on the other hand have on average one active zone per bouton (Holderith et al. 2012), and in extreme cases even less

(C. Liu et al. 2018). At many such CNS synapses a maximum of one vesicle can be released

126 during a single action potential, which led to the idea that a single synapse and or active zone was a release site (Korn and Faber 1991). We now know, however, that some synapses can release multiple vesicles per action potential and that vesicles can fuse at ‘hotspots’ within a single active zone indicating the presence of multiple release sites per active zone (Tang et al.

2016; Miki et al. 2016; Wadiche and Jahr 2001; Zenisek, Steyer, and Almers 2000).

When the active zone was first visualized using PTA staining, it was postulated that the gaps between dense projections were the physical correlate of release sites (Pfenninger et al.

1969). The active zone protein complex itself was also a promising candidate since vesicles only fuse at the active zone, whereas the SNARE proteins which provide the force for fusion are localized broadly across the plasma membrane. After disrupting the active zone by knocking out

RIM and RIM-BP, PTA staining density was significantly decreased (Acuna, Liu, and Südhof

2016). Based on these data, and the data presented in Chapter three, I believe that the active zone protein complex, or at least the major elements of it that we are able to disrupt, does not constitute the molecular correlate of a release site, though it is most certainly intimately related to it. This is supported by the fact that vesicles are still able to fuse after disruption of the active zone, albeit far less efficiently. The active zone protein complex is therefore most likely involved in organizing the fusion machinery to promote efficient fusion, in addition to its role in tethering and docking synaptic vesicles. It is interesting to note however that we still observe release that is targeted to the membrane opposite postsynaptic receptors; we would not expect to see relatively synchronized events otherwise. This suggests that either a mechanism still exists to restrict fusion to the active zone or fusion can occur randomly throughout the membrane in its absence. Though we have no evidence for or against ectopic fusion outside of the synapse, I believe the former to be the more likely answer. Many potential molecular markers of release sites could still be present and it is possible that lipids rather than proteins are the important release site markers (de Jong et al. 2018; Honigmann et al. 2013; van den Bogaart et al. 2011).

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Docking and priming at the active zone

The relationship between morphological docking and functional priming of synaptic vesicles at the active zone has long been a contentious topic. Some of this stems from ambiguity over what is meant by a ‘primed vesicle’ and the relationship to the term RRP. The term ‘primed’, and the idea of an RRP of vesicles, has been widely used in relation to the pool of vesicles released in response to stimulation with hypertonic sucrose (Rosenmund and Stevens

1996). The mechanisms behind this remain unclear to this day, but they are theorized to involve physical deformation of the plasma membrane reducing the energy barrier for fusion (Basu et al.

2007; Schotten et al. 2015). The assumption is that this deformation would primarily effect those vesicles closest to the membrane. The vesicles released by sucrose are from the same pool as those released by action potentials, though sucrose evoked fusion does not require Ca2+ influx

(Rosenmund and Stevens 1996). This contributed to the widespread use of the method as a standard with which to measure the RRP since changes in Ca2+ influx and release probability could be distinguished from changes in RRP size. For many, the measurement of whether or not a vesicle is primed is whether the vesicle fuses in response to sucrose.

In parallel to the sucrose method, several methods of depleting the RRP with action potential trains were developed and are employed (Elmqvist and Quastel 1965;

Schneggenburger, Meyer, and Neher 1999; Thanawala and Regehr 2016). These methods are favored by many since they are more physiological (synapses in vivo never experience a 500 mM hyperosmotic shock). Importantly, the underlying assumption of this method remains that there is an RRP of vesicles, which if release probability were one would all be released by a single action potential. These vesicles are assumed to be those docked at the membrane, but over the course of the train there is potential for new vesicles to be mobilized and brought to the active zone (i.e replenishment occurs). The challenges associated with all RRP measurements comes down to a temporal problem: how can one deplete the RRP as fast as possible in order

128 to determine how many vesicles would have been released initially if release probability had been one. When put in this context, determining exactly how many vesicles are in the action potential accessible RRP is perhaps unnecessary, as release probability will likely never be that high in vivo. A more relevant effort is to determine how a vesicle becomes primed to be part of the RRP at the molecular level.

A great deal of work has gone into addressing this question and the dominant model now holds that a primed vesicle (i.e. one that is part of the RRP) is one in which a partially zippered SNARE complex has formed through the actions of Munc18 and Munc13 (Ma et al.

2011; Yang et al. 2015; Ma et al. 2013; Lai et al. 2017; Siksou et al. 2009; Imig et al. 2014;

Richmond, Weimer, and Jorgensen 2001). This model is supported by the fact that knockouts of

Munc13 and several SNARE proteins abolish vesicle docking, action potential evoked fusion, and the RRP as measured by hypertonic sucrose (Imig et al. 2014; Siksou et al. 2009;

Varoqueaux et al. 2002; Bronk et al. 2007; Deák et al. 2004). Disrupting the ability to form

SNARE complexes therefore has a clear functional phenotype (a loss of fusion) and a clear morphological phenotype (a loss of vesicle docking), implying that partially zippered SNARE complexes are involved in both.

SNARE assembly is not always productive however. Dead end complexes can form between syntaxin and SNAP25 and tripartite SNARE complexes can form in an anti-parallel conformation which does not promote fusion (Ma et al. 2013; Lai et al. 2017). Munc18 and

Munc13 are required to guide SNAREs into productive parallel conformations, likely starting with

Munc18 holding syntaxin in a closed conformation followed by Munc13 promoting the transition from the closed syntaxin-Munc18 complex to a parallel SNARE complex. What does this tell us about SNAREs, Munc13, and vesicle docking? First, these data suggest that the state of plasma membrane SNAREs is the key determinant of whether or not fusion will occur. If dead- end Syntaxin-SNAP25 complexes and closed syntaxin require Munc18 and Munc13 to enter a

129 fusion competent state, neither of these proteins is a synaptic vesicle protein, suggesting that whether or not a vesicle is primed is determined at the membrane. Second, it is possible that an antiparallel SNARE complex could result in the morphological docking of a vesicle with the end result of that vesicle being unable to fuse. This model is interesting to consider in light of experiments looking at fusion of dye labeled vesicles using TIRF microscopy. In these experiments, a large portion of membrane proximal, presumably docked, vesicles do not fuse

(Midorikawa and Sakaba 2015). Other vesicles ‘bounce’, they approach the membrane and apparently dock, but are later released without fusing (Zenisek, Steyer, and Almers 2000;

Midorikawa and Sakaba 2015). While these data do not have the same spatial resolution as EM images, they nevertheless suggest that membrane proximal vesicles are not all in a uniform state which leads to fusion. It will be interesting to design new approaches to assess the conformational state of SNAREs in situ in relation to synaptic ultrastructure.

The functional concept the RRP takes a very vesicle-centric view of the nature of priming. The proposed molecular mechanism for priming, on the other hand, leave open the possibility that while the final stage of molecular priming involves interactions with vesicle

SNAREs, the upstream steps can be predetermined at the membrane prior to vesicle arrival

(Kaeser and Regehr 2017). This immediately recalls the concept of a release site and relates to the data presented in Chapter 3 in several ways. A surprising observation in our data was that more vesicles than expected, given the severity of the docking phenotype, could fuse in response to sucrose. This clearly demonstrates that sucrose stimulation, the classical measure of RRP and functional priming, can fuse more than just morphologically docked vesicles. The question becomes whether these vesicles, which are primed by the functional definition, are still employing the same molecular priming mechanisms as at wild-type synapses. Munc13-1 is essential for fusion at excitatory synapses in cultured hippocampal neurons (Augustin,

Rosenmund, et al. 1999; Varoqueaux et al. 2002). Although it is dramatically reduced in

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RIM/ELKS knockouts, the small amounts of Munc13 that remain may be sufficient to support some fusion. Alternatively, other isoforms of Munc13 partially compensate. An important next step will be to determine whether or not the remaining fusion relies on Munc13.

Inhibitory functions of the active zone

Given our data, it is tempting to speculate that the active zone might have an inhibitory function, keeping synaptic vesicle from fusing until an action potential arrives. The idea of a fusion being ‘clamped’ is prominent, especially in relation to the protein complexin. When complexin is knocked out the frequency of miniature events is increased though evoked release is decreased (Giraudo et al. 2006; Huntwork and Littleton 2007; Reim et al. 2001; Xue et al.

2008). It is possible that after disruption of the active zone we lose such a clamp. This is somewhat supported by comparing the magnitude of the effect of knocking out RIM alone with knocking out RIM and ELKS together. Though direct comparison of these experiments is difficult, they suggest that removing RIM alone has a more severe effect on release. The active zone may have inhibitory functions beyond simply recruiting a fusion clamp like complexin. For example, when vesicle docking is partially intact but Munc13 function is lost, as in the case of

RIM knockouts (Kaeser et al. 2011; Deng et al. 2011), more vesicles may form anti-parallel, non-fusogenic SNARE complexes. Once vesicle docking and tethering are disrupted, vesicles that are displaced from the membrane are no longer in close enough proximity to form such complexes. A promising direction will be to attempt to restore some elements of active zone function without others, for example rescuing protein scaffolding without restoring docking. This may allow disambiguation of parameters that may be correlated but functionally separable, such as Munc13 levels and vesicle docking.

131

Outlook

This work was informed and inspired by decades of research on synapses using electrophysiology, in vitro biochemistry, genetics, and cell biology. These techniques have allowed us to define and answer a broad range of questions about how synapses work and what proteins make them work. In my view, many of the core remaining question now amount to putting what we know about synaptic proteins into context. Questions such as whether a docked vesicle is a primed vesicle and how Ca2+ channels are distributed and linked to the active zone all amount to visualizing the state of proteins in their native environments. With the advent of super-resolution imaging and the rapid development of cryoEM we now have unprecedented views into the nano-scale organization of synapses. The decades to come will hopefully allow us to leverage these powerful new techniques to see the intricate molecular machines that we have been studying.

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— APPENDIX ONE —

Supplemental Figures

133

Figure S1. ELKS antibody specificity. (A) Western blot for testing specificity of ELKS2

(1029, top) and ELKS1 (E-1, middle) antibodies against samples of HEK293T cells transfected with ELKS1Bor ELKS2B cDNAs. The ELKS2 specific antibodies were raised in rabbits to a non-conserved sequence between ELKS1 and ELKS2 (109LSHTDVLSYTDQ120), the E-1 antibody is commercially available. -actin was used as a loading control. (B) Western blot testing reactivity of ELKS2 (top) and ELKS1 (middle) in cultured cDKO and control hippocampal neurons and whole brain homogenate. -actin was used as a loading control. (C)

ELKS2 antibodies were affinity purified using the ELKS2 peptide and characterized via immunostaining in cultured control and ELKS2 cKO neurons. ELKS2 cKO neurons were generated from ELKS2floxed mice (Kaeser et al., 2009), and neurons were stained for

ELKS2 (1029), GAD2, and VGluT1.

134

Figure S1 (Continued)

135

Figure S2. Frequency distributions of ELKS1 and ELKS2 at excitatory and inhibitory synapses. (A) Histogram displaying the frequency distribution of ELKS1 intensity within VGAT

(black bars, n = 194 ROIs/40 images/4 independent cultures) or VGluT1 (grey bars, n =

207/40/4) labeled puncta. (B) Histogram displaying the frequency distribution of ELKS2 intensity within GAD2 (black bars, n = 170/40/4) or VGluT1 (grey bars, n = 205/40/4) labeled puncta. The analysis shown in this Figure uses the data presented in the Figure 3.1B.

136

Figure S3. No change in synapse number in ELKS1/2 cDKO cultures. (A) Sample images of excitatory (green) and inhibitory (red) synapses in control and cDKO neurons. (B, C)

Quantification for excitatory (B) and inhibitory (C) synapses of fluorescence intensity, synapse number (puncta/10 m), and puncta size (control n = 3 independent cultures, cDKO n = 3, 10 images per culture were averaged per condition). All data are means ± SEM.

137

Figure S4. Post-hoc identification of excitatory neurons after presynaptic Ca2+ imaging.

(A) In each presynaptic Ca2+ imaging experiment, cultured neurons were stained post-hoc with

GAD67 antibodies to label the soma and neurites of inhibitory neurons, and the imaged neurons were identified by the Alexa 594 fill (red). The left panel shows an example of non-GABA neuron

(negative for GAD67) and the right panel a GABA neuron (positive for GAD67). (B) In an independent experiment, excitatory neurons were marked with a lentivirus expressing

TdTomato (arrowheads) under a CaMKII promoter in addition to a lentivirus that labeled all neurons with nuclear EGFP (merged image). Inhibitory neurons were identified by somatic and dendritic GAD67 staining (arrow, GAD67 antibodies have a nuclear background staining in all neurons). Of the 114 neurons we analyzed, 80 were TdTomato positive (excitatory), 23 were

GAD67 positive (inhibitory), and 11 were negative for both markers. Thus, 80 out of 91 of the

GAD67 negative neurons were identified as excitatory neurons. The remaining neurons may be excitatory, inhibitory, or modulatory.

138

Figure S4 (Continued)

139

Figure S5. Direct comparison of IPSC and EPSC amplitudes of ELKS1/2 cDKO.

Quantification of IPSC (left) and NMDAR-EPSC (right) amplitudes in control and cDKO neurons from the same cultures (control n = 15 cells /3 independent cultures, cDKO n = 15/3). All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01 as determined by Student's t test.

140

Figure S6. Inhibitory RRP size in ELKS1/2 cDKO neurons. Sample traces showing IPSCs in response to superfusion with 500 mOsm sucrose (left) and quantification (right) of IPSC charge transfer during the first ten seconds of the response (control n = 9 cells/3 independent cultures, cDKO n = 9/3). All data are means ± SEM.

141

Figure S7. Expression and localization of ELKS1 C-terminal rescue constructs. (A)

Sample images of control, cDKO, and cDKO + rescue neurons stained with antibodies against

ELKS1(E-1), the inhibitory synapse marker VGAT, and the excitatory synapse marker

VGluT1. (B, C) Quantification of ELKS1 fluorescent intensity within ROIs defined by VGluT1

(B) or VGAT (C) signals (control n = 3 independent cultures, cDKO n = 3, ELKS1B n = 3,

CCD n = 3, CCDB n = 3, 5 - 10 images per culture were averaged per condition). Data are shown as means ± SEM. (D) Representative western blot of control, cDKO, and cDKO + rescue neurons used for recording in Figure 2.7. Samples were blotted using an ELKS1 specific antibody (E-1) and -actin was used as a loading control.

142

Figure S8. Expression and localization of ELKS1 full length rescue constructs. (A)

Representative western blot of control, cDKO, and cDKO + rescue neurons used for recording in Figure 2.8. Samples were blotted using a pan-ELKS antibody (P224) and -actin was used as a loading control. (B) Quantification of HA staining fluorescent intensity within ROIs defined by

VGAT signals (control n = 3 independent cultures, cDKO n = 3, ELKS1B n = 3, CCD n = 3,

CCDB n = 3, 5 - 10 images per culture were averaged per condition). Data are shown as means ± SEM. (C) Representative images of ELKS1 staining in control, cDKO, and cDKO + rescue neurons. Note that ELKS1B lacks the antigen for the ELKS1 antibody (E-1). (D, E)

Quantification of ELKS1 fluorescent intensity within ROIs defined by VGluT1 (D) or VGAT (E)

(control n = 3 independent cultures, cDKO n = 3, ELKS1B n = 3, ELKS1B n = 3, CCC n = 3,

CCDB n = 3, 5 - 10 images were averaged per culture and genotype). All data are means ±

SEM.

143

Figure S8 (Continued)

144

Figure S9. Rescue of action potential evoked IPSCs with ELKS1B. Sample traces (left) and quantification (right) of the IPSC amplitude in control, cDKO, cDKO + ELKS1B, and control

+ ELKS1B (control n = 11 cells/3 independent cultures, cDKO n = 11/3, cDKO + ELKS1B n =

10/3, control + ELKS1B n = 12/3). All data are means ± SEM; **p ≤ 0.01, ***p ≤ 0.001 as determined by one-way ANOVA followed by Holm-Sidak multiple comparisons post-hoc test comparing each condition to cDKO. Holm-Sidak multiple comparisons post-hoc tests comparing each condition to control did not reveal significant differences except for control vs. cDKO, *, p ≤

0.05.

145

Figure S10. Genetic disruption of the active zone. (A) Schematic of the RIM and ELKS proteins targeted in the conditional RIM1 (Kaeser et al., 2008), RIM2 (Kaeser et al., 2011),

ELKS1 (Liu et al., 2014), and ELKS2 (Kaeser et al., 2009) knockout mouse lines crossed to quadruple homozygosity. Arrowheads indicate transcription start sites of all known -, - and - isoforms of each gene (Kaeser et al., 2008, 2009; Liu et al., 2014; Wang and Sudhof, 2003;

Wang et al., 2002), the red bars indicate the protein sequence that is encoded by the conditionally targeted exon. The zinc finger domain (Zn), PDZ domains (PDZ), C2 domains

(C2A, C2B), coiled-coil regions (CC) and the ELKS C-terminal PDZ binding domain (B) are indicated. (B) Additional sample images for Figure 3.1 B, for the assessment of synaptic protein levels within Synaptophysin-1 (Syp-1) defined ROIs, and example images for stainings excluding the primary antibodies in control neurons. (C) Synaptophysin-1 staining was used to determine synapse density and size. The synapse density is expressed as synaptophysin-1 positive puncta per 30 m of MAP2 positive dendrite, and synaptophysin-1 levels are measured as the average arbitrary fluorescence within the puncta (controlR+E n = 3 independent cultures, cKOR+E n = 3, 10 images per culture). (D) Quantitation of immunofluorescent stainings of wild type neurons incubated with or without RIM primary antibodies (with primary n = 3 independent cultures, without primary n = 3). The same fluorescent secondary antibodies were used in both conditions and in all test proteins in Figure 3.1 A, and acquisition settings were identical for the two groups. This experiment measures the background signal from the secondary antibody alone, and it does not account for background signal contributed by the primary antibody. (E)

Quantification of the fraction of Synaptophysin-1 puncta containing active zone proteins in cKOR+E and controlR+E synapses (controlR+E n = 6 independent cultures, cKOR+E n = 6, 10 images per culture). (F) Quantification of the fraction of Bassoon pixels containing active zone proteins at cKOR+E and controlR+E synapses (controlR+E n = 6 independent cultures, cKOR+E n = 6, 10 images per culture). (G) Plot of all individual synaptic

146

Figure S10 (Continued) fluorescent intensity levels normalized to the average control (controlR+E) synaptic fluorescent intensity level. Red lines indicate mean ± SEM, data shown corresponds to Figure 3.1 C. All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test.

147

Figure S10 (Continued)

148

Figure S11. Analysis of synapse ultrastructure in glutaraldehyde fixed neurons. (A, B)

Sample images (A) and quantification (B) from glutaraldehyde fixed electron microscopic analyses of cKOR+E and controlR+E synapses; postsynaptic densities are marked with arrowheads. (controlR+E n = 50 synapses/3 independent cultures, cKOR+E n = 48/3). (C)

Distribution of synaptic vesicles relative to the presynaptic plasma membrane area opposed to the PSD. Vesicle distribution is shown in 100 nm bins (left) in cKOR+E and controlR+E synapses within 1 µm. Gaussian fits were used to model the vesicle distribution. The two genotypes were significantly different (*, p < 0.05) and could not be fit with a single distribution, requiring individual fits. Distribution of synaptic vesicles within the first 100 nm in 10 nm bins and the number of tethered vesicles (defined as vesicles within 100 nm of the presynaptic plasma membrane) are shown in the middle and on the right, respectively. In each bin within 100 nm we observed a reduced vesicle number in cKOR+E synapses (controlR+E n = 50/3, cKOR+E n = 48/3).

All data are means ± SEM; *p ≤ 0.05, ***p ≤ 0.001 as determined by Student's t test (B) or by extra sum of squares F test (C).

149

Figure S11 (Continued)

150

Figure S12. Additional electrophysiological analyses of stimulus trains and mPSCs. (A)

Plot of EPSC amplitude during a 10 Hz, 50 stimulus train normalized to the amplitude of the first response (controlR+E n = 17 cells/3 independent cultures, cKOR+E n = 19/3). (B) Analyses of total

(0 - 6.5 s), tonic (0 - 5 s) and delayed (5 - 6.5 s) charge transfer from NMDAR-EPSCs during stimulation trains in cKOR+E and controlR+E neurons (controlR+E n = 17/3, cKOR+E n = 19/3). (C, D)

Same as A, B, but for IPSCs (controlR+E n = 19/3 independent cultures, cKOR+E n = 19/3). (E)

Expanded traces showing mEPSCs from the experiment presented in Figure 3.5 L. For each condition (controlR, cKOR, controlE, cKOE, controlR+E, and cKOR+E) three consecutive seconds from a single cell are shown. (F) Statistical comparison of mEPSC frequencies between all genotypes (normalized to their respective controls). Statistical comparison between genotypes was done using one-way ANOVA (***, p<0.001) followed by Holm-Sidak post-hoc tests (cKOR+E vs. cKOR: ***, p < 0.001; cKOR+E vs. cKOE: n.s., p = 0.474; cKOE vs. cKOR: ***, p < 0.001; cKOR+E n = 31/5, cKOR n = 21/3, cKOE n = 24/3). (G, H) Sample traces (G) and quantification

(H) of mIPSCs in controlR+E and cKOR+E neurons. Quantitative analysis of mIPSC frequencies

(H, left) and amplitudes (H, right) are shown (controlR+E n = 20/3, cKOR+E n = 21/3). (I) Expanded traces showing mIPSCs from the experiment presented in G, H. Three consecutive seconds from a single cell are shown for each condition. All data are means ± SEM unless otherwise specified **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test.

151

Figure S12 (Continued)

152

Figure S13. Additional analysis for heterogeneity in cKOR+E of synapses. (A, B) Sample images (A) and quantification (B) of the number of neurons (left) and the fraction of infected neurons (right) with EGFP tagged cre (cKOR+E) or an inactive form of cre (controlR+E). Consistent with Figures 3.1 and S10, no differences in neuronal density were detected and all neurons were infected with cre viruses (controlR+E n = 752 cells/9 frames/3 independent cultures, cKOR+E n = 731/9/3). (C) Frequency distribution of synaptic protein fluorescence levels for all individual cKOR+E and controlR+E synapses (normalized to the average of controlR+E synapses), related to

Figure 3.6 A. (D-F) Quantification of SypHy infected cultures related to Figures 3.6 B-H. SypHy puncta density and the fraction of total SypHy puncta responsive to NH4Cl (%(FNH4Cl/F0)

>200%) (D), SypHy fluorescence distribution of all synapses at baseline (F0) and upon NH4Cl application (%(FNH4Cl/F0)) (E), the fraction of SypHy puncta responsive to 40 and 200 APs stimulation (defined as %(Fduring stimulation/FNH4Cl) > 0) (F). The change in SypHy density in D suggests a change in synapse/cell density in cKOR+E neurons, which was not seen in other experiments (Figures S10 C, S13 A, B). We attribute this change to the potential toxicity of triple lentivirus infection (FSW-cre or control, FSW-SypHy, FSW-SV2-TdTomato) and suspect that the cKOR+E neurons are more sensitive to multi-virus infection because of the strong genetic manipulation and loss of activity. All data are means ± SEM; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test.

153

Figure S13 (Continued)

154

Figure S14. Electron microscopic analyses of cKOR, cKOE and cKOR+E synapses. (A, C, E)

Quantitation of vesicles per bouton, bouton size, and PSD length in cKOR (A), cKOE (C), cKOR+E

(E), and respective control synapses. (B, D, F) Distribution of synaptic vesicles relative to the presynaptic plasma membrane area opposed to the PSD was analyzed in 100 nm bins for cKOR

(B), cKOE (D), cKOR+E (F), and respective control synapses. Gaussian fits were used to model vesicle distribution (left); one Gaussian fit (orange line) was used if mutant and control best-fit distributions were not significantly different, and two Gaussian fits were used if the data could not be fit with a single Gaussian (*, p < 0.05 in F). Synaptic vesicle distribution within the first

100 nm in 10 nm bins and the number of tethered vesicles (defined as vesicles within 100 nm of the presynaptic plasma membrane) are shown in the middle and on the right, respectively. We observed a reduction of vesicles in the first 100 nm in cKOR+E synapses, but not in cKOR synapses or cKOE synapses (controlR n = 25 synapses, cKOR n = 25; controlE n = 25, cKOE n =

25; controlR+E n = 25, cKOR+E n = 25 in A-F). (G) Statistical comparison of docking between all genotypes, normalized to their respective controls, using one-way ANOVA (***, p < 0.001) followed by Holm-Sidak post-hoc tests (cKOR+E vs. cKOR: **, p < 0.01; cKOR+E vs. cKOE: ***, p <

0.001; cKOE vs. cKOR: n.s., p = 0.0673; cKOR+E n = 25 synapses, cKOR n = 25, cKOE n = 25).

(H) Statistical comparison of 0 - 10 s sucrose response between all genotypes, normalized to their respective controls, using one-way ANOVA (***, p<0.001) followed by Holm-Sidak post-hoc tests (cKOR+E vs. cKOR: *, p<0.05; cKOR+E vs. cKOE: *, p<0.05; cKOE vs. cKOR: ***, p<0.001; cKOR+E n = 20 cells/3 cultures, cKOR n = 20/3, cKOE n = 17/3). Data are means ± SEM; *p ≤

0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by Student's t test (E), by extra sum of squares F test (F), or by one-way ANOVA with Holm-Sidak post-hoc test (G, H)

155

Figure S14 (Continued)

156

Figure S15. Removal of CaV2 channels and loss of toxin sensitivity. (A,B) Qualitative western blots showing loss of CaV2.1 (A) and CaV2.2 (B) in samples prepared from CaV2 cTKO cultures. Synapsin (bottom panels) was used as a loading control. (C) Quantification of eIPSC amplitude from CaV2.1 cKO, CaV2.2 cKO, and CaV2.3 cKO cultures and their respective controls

(CaV2.1 control, n = 12 cells/3 independent cultures, CaV2.1 cKO n = 9/3; CaV2.2 control n =

4/2, CaV2.2 cKO n = 4/2), CaV2.3 control n = 7/2, CaV2.3 cKO n = 4/2)). (D) Normalized eIPSC amplitude of representative cells plotted against stimulus number (0.067 Hz stimulation frequency) before and after application of 200 nM -Agatoxin IVA. Example traces on the right show the average of the first five (solid black line) and the last five (dashed red line) responses of the same cells. (E) Quantification of eIPSC amplitude (average of the last five responses) in the presence of 200 nM -Agatoxin IVA normalized to the average of the first five responses

(CaV2.1 control, n = 3 cells/2 independent cultures, CaV2.1 cKO n = 4/2). (F) Normalized eIPSC amplitude of representative cells plotted against stimulus number (0.067 Hz stimulation frequency) before and after application of 250 nM then 500 nM -Conotoxin GVIA. Example traces on the right show the average of the first five (solid black line) and the last five (dashed red line) responses of the same cells. (G) Quantification of eIPSC amplitude (average of the last five responses) in the presence of 250 nM -Conotoxin GVIA normalized to the average of the first five responses (CaV2.2 control n = 4/2, CaV2.2 cKO n = 4/2).

157

Figure S15 (Continued)

158

Figure S16. Confocal analysis of active zone composition, synapse number, and dendritic branching. (A,B) Sample images (A) and quantitation (B) of active zone protein levels at CaV2 control and CaV2 cTKO synapses using confocal microscopy. The synaptic vesicle marker Synaptophysin-1 (Syp-1) was used to define the region of interest (ROI). The black dotted line (B) indicates control levels. Scale bar = 5 m (CaV2 control n = 3 independent cultures, CaV2 cTKO n = 3, 10 images per culture). (C) Quantification of the number of synapses, defined by synaptophysin puncta, per 30 m of dendritic length (CaV2 control n = 6 independent cultures, CaV2 cTKO n = 6, 10 images per culture). (D,E) Binary images (D) of max projections z-stacks of CaV2 control and CaV2 cTKO neurons filled with Alexa 594 via a whole- cell patch pipette. (E) Sholl analysis of the number of process crossings of concentric circles originating on the cell soma at intervals of 10 m (sholl radius) (CaV2 control, n = 3 cells/1 independent cultures, CaV2 cTKO n = 4/1).

159

Figure S16 (Continued)

160

Figure S17. Expression and localization of CaV2.1 rescue constructs. (A) Example images of CaV2 cTKO and C-terminal mutant rescue conditions stained with HA (green channel, CaV2.1 rescue), synaptophysin (red channel), and MAP2 (blue channel). (B,C) Quantification of dendritic (E) and synaptic (F) HA fluorescence intensity in ROIs defined by either synaptophysin staining (synapse) or MAP2 staining excluding synaptophysin ROIs (dendrite) (CaV2 cTKO, n =

PDZ 3 independent cultures; + CaV2.1 n = 3; + CaV2.1-Ct, n = 3; + CaV2.1-C , n = 3; + CaV2.1-

SH3 polyR C , n = 3; + CaV2.1-C , n = 3, 8 images per culture). All data are means ± SEM; *p ≤

0.05, **p ≤ 0.01, ***p ≤ 0.001 as determined by one-way ANOVA followed by Holm-Sidak multiple comparisons post-hoc testing comparing each condition to full-length CaV2.1 rescue.

161

— APPENDIX TWO —

Supplemental Experimental Procedures

162

Mice. Previously generated conditional RIM1 (Kaeser et al., 2008) (RRID:IMSR_JAX:015832),

RIM2 (Kaeser et al., 2011) (RRID:IMSR_JAX:015833), ELKS1 (Liu et al., 2014)

(RRID:IMSR_JAX:015830) and ELKS2 (Kaeser et al., 2009) (RRID:IMSR_JAX:015831) mice were crossed and maintained as quadruple homozygous floxed mice. The RIM1/2 double floxed mice (Kaeser et al., 2011) and the ELKS1/2 double floxed mice (Liu et al., 2014) were described previously. All animal experiments were performed according to institutional guidelines at

Harvard University.

Cell culture and lentiviral infection. Primary mouse hippocampal cultures were generated from newborn conditional quadruple or double floxed pups as described before (Kaeser et al.,

2008, 2011; Liu et al., 2014; Maximov et al., 2007) within 24 h after birth. For the biochemical solubility experiments, neurons were plated directly on 12-well cell culture plates, for all other experiments, chemically stripped glass coverslips were used. Lentiviruses expressing EGFP- tagged cre recombinase (to generate cKO neurons) or a truncated, enzymatically inactive

EGFP-tagged cre protein (to generate control neurons) were produced in HEK293T cells by

Ca2+-phosphate transfection and neuron-specific expression was driven by a synapsin promoter

(Liu et al., 2014). Neurons were infected with HEK cell supernatant at 5 days in vitro (DIV) as described before (Kaeser et al., 2008; Liu et al., 2014). All subsequent experiments were performed at DIV 15-19. In the SypHy experiments, additional lentiviruses expressing SypHy or

SV2-TdTomato under a human synapsin promoter were produced in HEK293T cells and were applied to the cultured neurons at DIV3.

Immunofluorescence stainings and confocal imaging of cultured neurons. Neurons were washed with phosphate buffered saline (PBS), fixed in 4% paraformaldehyde in PBS for 20

163 minutes, permeabilized in 0.1% Triton X-100/3% bovine serum albumin/PBS for 1 h, and incubated in primary antibodies at 4oC overnight. Following overnight primary antibody incubation, neurons were incubated in AlexaFluor-488 (for detection of the protein of interest),

AlexaFluor-546 (for detection of MAP2), and AlexaFluor-633 (for detection of Synaptophysin-1) conjugated secondary antibodies (1:500) for 2 h at room temperature and mounted in

Fluoromount-G for imaging. Images were taken on an Olympus FV1200 confocal microscope using identical settings for each condition in a given experiment with a 60X oil-immersion objective and single confocal sections were analyzed in ImageJ software. For quantitative analyses of synaptic protein levels, the Synaptophysin-1 signal was used to define synaptic puncta as ROI, and the average intensity within the ROI was quantified. For each image, the

“rolling ball” ImageJ plugin was set to a diameter of 1.4 μm to calculate a local background value for background subtraction (Sternberg, 1983). The quantitative data for each protein per condition were derived from n ≥ 3 independent cultures, and ≥ 10 fields of view per culture. A control experiment in which wild type neurons were incubated either with primary (mouse anti-

RIM) or without primary antibodies in the AlexaFluor-488 channel was done to determine the level of non-specific staining by secondary antibodies alone for comparison (Figures S1B, D).

The average intensity per protein in cKOR+E neurons was normalized to the respective staining in the control. For co-localization of active zone proteins, the ImageJ plugin BioVoxxel was used in the puncta to puncta comparison (The BioVoxxel Image Processing and Analysis Toolbox.

Brocher, 2015, EuBIAS-Conference, 2015, Jan 5) and a custom MATLAB script was used for the pixel to pixel comparison. Student’s t-test was used to determine whether experimental and control conditions were significantly different. All experiments and analyses were performed by an experimenter blind to the genotype.

164

Biochemical solubility assay. At DIV 15 the neurons were harvested in 400 l ice-cold buffer

(25 mM HEPES, 5 mM EDTA, 0.32 M Sucrose, 7.2 pH) and homogenized with a glass-teflon homogenizer. The homogenate was solubilized with 1% Triton X-100 for 1 h rotating at 4 C and then centrifuged at 100,000 x g for 1 h. The pellet and supernatant were collected in 1X sodium dodecyl sulfate (SDS) sample buffer and quantitative Western blotting was performed on the pellet and the supernatant as described below. In each fraction and for each protein, the signal was normalized to -actin as an internal loading control resulting in values for the pellet (P) and the supernatant (S), and the experiment was expressed as the cKOR+E to controlR+E ratio.

Solubility was calculated as S/(S+P). Student’s t-test was used to determine whether experimental and control conditions were significantly different.

Western blotting. For non-quantitative assessment of cre-recombination in cultured neurons, chemiluminescence was used to detect RIM and ELKS removal in all experiments. At DIV15, cultured neurons were harvested in 25 l 1X SDS buffer and run on standard SDS-Page gels followed by transfer on a nitrocellulose membrane. Membranes were blocked in filtered 10% nonfat milk/5% goat serum for 1 h at room temperature and incubated with primary antibodies in

5% nonfat milk/2.5% goat serum for 2 hours at room temperature to overnight at 4oC, and HRP- conjugated secondary antibodies (1:10,000) were used. For quantitative assessment of protein levels in cultured neurons, fluorescently tagged secondary antibodies were used. Cultured neurons were either directly harvested in 1X SDS buffer or processed as described in the biochemical solubility assay section. Membranes were blocked in filtered 5% nonfat milk/5% goat serum for 1 h at room temperature and incubated with primary antibodies in 5% BSA overnight at 4oC. Each membrane was incubated with primary antibodies against the protein of interest and Synapsin or -actin antibodies as a loading control. Blots were scanned on a fluorescent scanner and all images were analyzed with ImageJ. The fluorescent signal in each

165 experimental condition was normalized first to either Synapsin (total culture homogenates) or - actin (biochemical solubility experiments) to account for differences in loading and then to the respective controlR+E signal. All quantitative analyses were performed in three independent cultures. For the plot of total protein levels, the experiments from total culture homogenates (n =

3) and the total protein levels from the solubility assay (n = 3, sum of the soluble and insoluble fraction) were pooled. Student’s t-test was used to determine whether experimental and control conditions were significantly different.

Electron microscopy and analysis of synaptic vesicle distribution. For high-pressure freezing: neurons cultured on 6 mm carbon-coated sapphire coverslips were frozen using the

HPM 100 high-pressure freezer in extracellular solution containing (in mM): 140 NaCl, 5 KCl, 2

CaCl2, 2 MgCl2, 10 HEPES-NaOH (pH 7.4), 10 Glucose (~310 mOsm) with picrotoxin (50M),

AP5 (50 M), and CNQX (20 M) added to block synaptic transmission. After freezing, samples were stored in liquid nitrogen and processed by the Electron Microscopy Facility at Harvard

Medical School. Samples were first freeze-substituted in 1% glutaraldehyde, 1% osmium tetroxide, 1% water, and anhydrous acetone with the following protocol: -90oC for 5 h, 5oC per hour to -20oC, -20 oC for 12 h, and 10oC per hour to 20 oC (AFS2, Leica). Following freeze substitution, samples were Epon infiltrated, and baked for 24 h at 60 oC before sectioning at 50 nm and imaging. For glutaraldehyde fixation: neurons cultured on glass coverslips were fixed with 2% glutaraldehyde in 0.1 M sodium cacodylate buffer at 37oC for 15 minutes and immediately processed by the Electron Microscopy Facility at Harvard Medical School.

Glutaraldehyde fixed samples were first stained with 1% osmium tetroxide/1.5% potassium ferrocynide for 1 h at room temperature, washed in water and then maleate buffer (pH 5.15) 3X, and then stained with 1% uranyl acetate for 1 h. Post-staining, samples were dehydrated through a series of EtOH treatments (70% for 15 min, 90% for 15 min, and 100% for 15 min 2X)

166 and propylene oxide (1 h). Samples were resin infiltrated and prepared for embedding with a 1:1

Epon, propylene oxide mixture for 2 h at room temperature. Samples were then polymerized for

24 h at 60oC and sectioned at 50 nm for transmission electron microscopy. Imaging and quantification: Images of both high-pressure frozen and glutaraldehyde fixed samples were taken with a transmission electron microscope (JEOL 1200 EX at 80 kV accelerating voltage) and processed with ImageJ. The total number of vesicles, the number of docked vesicles, the length of the PSD, the area of the presynaptic bouton, and the distance of each vesicle from the active zone were analyzed with SynapseEM, a MATLAB macro provided by Drs. M. Verhage and J. Broeke. Bouton size was calculated from the perimeter of each synapse. Docked vesicles were defined as vesicles touching the presynaptic plasma membrane opposed to the

PSD. The nearest distance of the vesicle membrane to the presynaptic plasma membrane area opposed to the PSD was measured and plotted as frequency distribution over the total number of vesicles in 100 or 10 nm bins (Figures 3.2 C, S11 C and Figures S14 B, D, and F). To test whether there was a genotype effect on vesicle distribution, Gaussian fits were performed. The extra sum-of-squares F test was used to compare whether the best-fit values of mutant and control Gaussian distributions were significantly different, and if they were not different a single

Gaussian fit for all data is shown for the mutant and control data set. Student’s t-test was used to determine whether all other experimental and control conditions were significantly different.

All experiments and analyses were performed by an experimenter blind to the genotype.

Electrophysiology. Electrophysiological recordings in cultured hippocampal neurons were performed as described (Kaeser et al., 2008, 2009, 2011; Liu et al., 2014; Maximov et al., 2007) at DIV 15-19. The extracellular solution contained (in mM): 140 NaCl, 5 KCl, 2 CaCl2, 2 MgCl2,

10 HEPES-NaOH (pH 7.4), 10 Glucose (~310 mOsm). To assess excitatory transmission in response to action potentials, NMDAR excitatory postsynaptic currents (EPSCs) were

167 pharmacologically isolated with picrotoxin (PTX, 50 M) and 6-Cyano-7-nitroquinoxaline-2,3- dione (CNQX, 20 M). NMDAR-EPSCs were recorded because analyses of AMPA-receptor mediated EPSCs in cultured neurons are limited by recurrent activity in response to action potentials. Action potential evoked inhibitory postsynaptic currents (eIPSCs) were isolated with

D-amino-5-phosphonopentanoic acid (D-APV, 50 M) and CNQX (20 M). For miniature recordings, tetrodotoxin (TTX, 1 M) was added to block action potentials, and in addition PTX

(50 M) and APV (50 M) were added for mEPSCs or APV (50 M) and CNQX (20 M) for mIPSCs, respectively. For sucrose EPSC recordings, TTX (1 M), PTX (50 M), and APV (50

M) were added. All recordings were performed in whole cell patch clamp configuration at 20 –

23º C. Glass pipettes were pulled at 2-4 M and filled with intracellular solutions containing (in mM) for EPSC recordings: 120 Cs-methanesulfonate, 10 EGTA, 2 MgCl2, 10 HEPES-CsOH (pH

7.4), 4 Na2-ATP, 1 Na-GTP, 4 QX314-Cl (~300 mOsm) and for IPSC recordings: 40 CsCl, 90 K-

Gluconate, 1.8 NaCl, 1.7 MgCl2, 3.5 KCl, 0.05 EGTA, 10 HEPES-CsOH (pH 7.4), 2 MgATP, 0.4

Na2-GTP, 10 Phosphocreatine, 4 QX314-Cl (~300 mOsm). Cells were held at +40 mV for

NMDAR-EPSC recordings and at -70 mV for mEPSC, mIPSC, eIPSC, and sucrose EPSC recordings. Access resistance was monitored during recording and cells were discarded if access exceeded 15 M or 20 M during recording of evoked or spontaneous synaptic currents, respectively. Action potentials were elicited with a bipolar focal stimulation electrode fabricated from nichrome wire. For analysis of action potential trains, 50 stimuli were provided at a frequency of 10 Hz. Individual PSCs within the train were aligned using the peak of the stimulus artifact and the baseline value set to the negative peak immediately following the artifact. Peak amplitudes and the synchronous component of charge transfer during the train were determined using these aligned events. To determine tonic charge transfer, the cumulative charge during the synchronous component was subtracted from the total charge transfer during the first 5 seconds of the train. Delayed charge transfer was taken as the area under the curve

168 for 1.5 seconds after the end of the train (i.e. 5 – 6.5 seconds). Paired-pulse ratios were calculated as the amplitude of the second PSC divided by the amplitude of the first PSC. The baseline value for the second PSC was taken as the negative peak immediately following the second stimulus artifact, as in the train analysis, and the amplitude of second PSC was measured relative to this baseline. The RRP was measured by application of 500 mM sucrose in extracellular solution applied via a microinjector syringe pump for 10 s at a rate of 10 L/min through a tip with an inner diameter of 250 m. For calcium titration experiments, the following

2+ 2+ [Ca ]ex/[Mg ] ex ratios were used (in mM): 0.5/3.5, 1/3, 2/2, 5/0.25, 7/0.25. Recordings began at

2+ 2+ 0.5 mM [Ca ]ex / 3.5 mM [Mg ] ex and with each solution exchange five chamber volumes of solution were exchanged. After each exchange there was delay of at least one minute to ensure equilibration of the new solution. During this delay action-potential evoked responses were

2+ recorded at 0.2 Hz and monitored for stability at each [Ca ]ex. Afterward, at least 10 sweeps

2+ were recorded and the average amplitude was measured at each [Ca ]ex. Only cells in which all

2+ five [Ca ]ex were recorded were included in the analysis. For the normalized data, IPSC

2+ amplitudes for each cell were normalized to the same cell’s amplitude at 7 mM [Ca ]ex/ 0.25

2+ mM [Mg ]ex. The absolute and normalized data was fit to the model I = Imin + (Imax - Imin) /

2+ (1+10^((LogEC50 - [Ca ]ex) * n)). For comparing EC50 values, normalized data from individual cells was fit to the above model and the averaged EC50 values were compared using a

Student’s t-test. Data were acquired with an Axon 700B Multiclamp amplifier and digitized with a

Digidata 1440A digitizer. For action potential and sucrose-evoked responses, data were acquired at 5 kHz and low-pass filtered at 2 kHz. For miniature recordings data were acquired at

10 kHz. All data acquisition and analysis was done using pClamp10. For all electrophysiological experiments, the experimenter was blind to the genotype throughout data acquisition and analysis.

169

Ca2+ Imaging. All Ca2+ imaging experiments were done with hippocampal cultures infected with lentiviruses (expressing active cre or inactive cre) at DIV 5. Neurons were recorded at DIV15-18 in whole-cell patch-clamp configuration at 20 – 23º C. The extracellular solution contained (in mM): 140 NaCl, 5 KCl, 2 CaCl2, 2 MgCl2, 10 Glucose, 0.05 APV, 0.02 CNQX, 0.05 PTX, 10

HEPES-NaOH (pH 7.4, ~310 mOsm). Glass pipettes were filled with intracellular solution containing (in mM) 140 K Gluconate, 0.1 EGTA, 2 MgCl2, 4 Na2ATP, 1 NaGTP, 0.3 Fluo5F, 0.03

Alexa Fluor 594, 10 HEPES-KOH (pH 7.4, ~300 mOsm). Only neurons with membrane potentials between -55 and -65 mV (junction potential uncorrected) were used. After filling for 7 min, axons and dendrites were identified in the red channel. Presynaptic boutons were identified by their typical bead-like morphology. Neurons in which the distinction between axons and dendrites was unclear were discarded. 10 min after break-in, a holding current was injected to keep the membrane potential at ~ -60 mV, and Ca2+-transients were induced by a single action potential evoked via brief somatic current injection (5 ms, 500-1500 pA). Images were acquired using an upright microscope with a 60X, 1.0 numerical aperture water immersion objective.

Fluorescence signals were excited by a light-emitting diode at 470 nm, and were collected with a scientific complementary metal–oxide–semiconductor camera (sCMOS) at 100 frames/s.

Images were collected for 0.2s before and 1s after the action potential initiation. An extra 10 frames of images were acquired for each neuron without excitation and these images were used to estimate the dark current level of the camera. Ca2+ transients were quantified using ImageJ.

7-20 boutons and 5-10 areas in the second order dendrites were randomly selected from each neuron. After background subtraction (removing the dark current and rolling ball (Sternberg,

1983) with a radius of 4 m), the (F-F0)/F0 was calculated (F = average green emission at a given time point, F0 = average fluorescent intensity in frames 0 to 20 before action potential induction). For all Ca2+ imaging experiments, the experimenter was blind to the genotype throughout data acquisition and analysis.

170

Synaptophysin-pHluorin Imaging. The SypHy A4 open reading frame (Granseth et al., 2006) was obtained from Addgene (Plasmid #24478), and was subcloned into the synapsin driven lentiviral vector FSW. FSW-SypHy A4- and FSW-SV2-tdTomato- expressing viruses were produced in transfected HEK293T cells and were applied to dissociated hippocampal cultures at

DIV3. Cultures were subsequently infected with lentiviruses to express cre or a control virus at

DIV5. Cultures were imaged at DIV14-17 with an upright microscope at 20-23 ° C in extracellular solution containing (in mM): 140 NaCl, 5 KCl, 2 CaCl2, 2 MgCl2, 10 Glucose, 10

HEPES-NaOH, 0.05 APV and 0.025 CNQX (pH 7.4, ~310 mOsm). Synapse rich areas were identified in the red channel. Images were acquired at 0.5 Hz with 2x2 binning using a sCMOS camera with a 60X, 1.0 numerical aperture water immersion objective. SypHy and SV2- tdTomato were excited with at 488 nm or 550 nm with a light-emitting diode, respectively. Light paths were split and filtered with a multiband filter set. Neurons were stimulated with a bipolar electrode made from nichrome wire at 20 Hz. For each experiment, 5 s of baseline image acquisition preceded stimulation. NH4Cl solution (extracellular solution substituted with NH4Cl for 50 mM NaCl, adjusted to pH 7.4) was applied at the end of experiments to visualize all

SypHy puncta. We did not correct for photobleaching. Images were analyzed in ImageJ, a rolling ball of 5 m was used for background subtraction for all images (Sternberg, 1983). NH4Cl responsive puncta defined as %(FNH4Cl/ F0) > 200% were included in the analysis, and were used to define ROIs. Active puncta in response to action potential stimulation (40 or 200 action potentials) were defined as those puncta in which the %(Fduring stimulation/F0) during stimulations

(2s for 40Hz, 10s for 200Hz) > 0, where Fduring stimulation is defined as the mean fluorescence of

F-F0 during stimulation. SypHy F0 was defined as the mean fluorescence during 5 s prior to stimulation. SypHy F was quantified as fluorescence intensity F at each time point minus F0.

Peak fluorescence (Fpeak) is defined as the average F during the first four imaging frames immediately following the end of the stimulation. Data were normalized to the total pool as

171 defined by SypHy response to NH4Cl application (FNH4). All experiments and analyses were performed by an experimenter blind to the genotype.

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