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Electron Tomographic Methods for Studying the Chemical Synapse
CHAPTER 10
Electron Tomographic Methods for Studying the Chemical Synapse Christopher P. Arthur, David B. Serrell, Maria Pagratis, David L. Potter, Dudley S. Finch, and Michael H. B. Stowell Department of MCD Biology University of Colorado Boulder, Colorado 80309
I. II. III. IV.
V. VI. VII. VIII.
IX. X. XI.
Introduction and Rationale Neurons, Neural Networks, and the Synapse ET and the Synapse Types of Neurons and Synapses A. Neuromuscular Junction Synapse B. Ribbon Synapses C. CNS Synapses Sources of Neurons for Structural Studies Methods for Assessing Neuron and Synapse Integrity Rapid Freezing and HPF of Neurons Modeling and Analysis of Synaptic Structures A. Size and Organization of the Vesicle Pool B. Organization of the Postsynaptic Density Proteins C. Synaptic Cleft Complexes D. Endosomal Compartments E. Microtubule and Neurofilament Arrangements Stimulation-Dependent Changes in Synaptic Structure The Future of EM Tomography and Synapses Conclusions References
This chapter focuses on the use of electron tomography (ET) to study the chemical synapse. It discusses the diVerent types of chemical synapses and their properties as well as the biological questions that are addressable using ET. We present METHODS IN CELL BIOLOGY, VOL. 79 Copyright 2007, Elsevier Inc. All rights reserved.
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methods for culturing neurons on a variety of substrates suitable for ET and discuss the development of novel tools that combine electrophysiological methods with ET methods for studying active synapses.
I. Introduction and Rationale Our current understanding of synapse formation and organization comes from a variety of studies that used immunofluorescence (or GFP-tagged proteins), fluorescent dyes, and/or electron microscopy (EM) of fixed embedded samples (Garner et al., 2002). All these methods are limited in their resolution and their inability to distinguish a functioning synapse from one that is inactive or undergoing formation or decay. In addition, both detailed structural information about the molecular and supramolecular organization of the synapse and knowledge about the dynamic events associated with synaptic transmission are lacking. Such information is important for understanding phenomena such as synaptogenesis, synaptic transmission and plasticity, and synaptic regeneration following neurological damage. During the last few years, neuroscientists have begun to dig more deeply into the structures and process of synaptic behavior by using electron tomography (ET). In this chapter, we will review these methods, recent studies of synapses, their limitations, and consider the ways in which future developments may allow for the systematic structural investigation of synaptic architecture under controlled conditions.
II. Neurons, Neural Networks, and the Synapse Living neural networks (LNN) are one of Nature’s greatest technological achievements. They can be massively parallel and dynamic computational devices with built-in plasticity and repair mechanisms. The most complex LNN, a human brain, comprises over 1014 connections, more connections than all the computing power manufactured between 1981 and 2001 (Stowell, 2003). The chemical synapse is at the heart of a functioning LNN. During brain development, chemical synapses are constructed following the establishment of contact between a growth-cone at the end of a growing axon and the cell that will receive its signal. Synaptogenesis is the process by which this complex, intercellular connection is formed. Once formed, a functional synapse has a dynamic life. Of course it transmits signals between neurons, but it is also subject to slower modulations that aVect its behavior and function. Individual synaptic signals undergo modulations, such as long-term potentiation (LTP) and long-term depression (LTD) (Bliss et al., 2003; Gilbert, 1998; Lisman, 2003), and in neurodegenerative diseases they can disintegrate. These adaptive changes in synapse function, which can occur on timescales ranging from seconds to minutes, are the foundation for more permanent changes in LNN wiring that may involve the formation of new synapses (Knott et al., 2002;
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Trachtenberg et al., 2002), giving rise to biologically indelible memory. All these phenomena are central to our understanding of memory and learning, and they also serve as a foundation for understanding biological computation in general and the development of neuromorphic technologies in the future. Naturally, a great deal of eVort has been spent on trying to understand the genetic and molecular bases of neuronal function and in particular memory and learning.
III. ET and the Synapse The last decade has seen a strong resurgence of interest in biological EM. While this trend has been led by dramatic progress in EM of macromolecules, the analysis of cellular ultrastructure has not lagged far behind (McIntosh, 2001). Several technological advances have facilitated this renaissance. On the instrumentation front, field emission guns (FEGs), which provide electron beams with good coherence and brightness, have improved the imaging of frozen-hydrated samples. Computer-controlled specimen stages have improved the regulation of sample position and automated data collection. Greatly improved CCD cameras have significantly increased the speed of image acquisition and analysis. Finally, as in almost every area of research, the dramatic decrease in price for computational speed and data storage has also played an important role. However, as in most areas of research, perhaps the most dramatic improvements have been the result of greatly improved methods for sample preparation; in particular, rapid freezing of biological specimens has opened the door for higher resolution studies of macromolecules and cells in their native states (Dubochet et al., 1983; Lepault et al., 1983). As a result, the use of ET for ultrastructural analysis has been applied extensively to a variety of cells and organisms such as Caenorhabditis elegans (Mu¨ller-Reichert et al., 2003), Saccharomyces cerevisiae kineticore distribution (O’Toole et al., 2002), and cytoplasmic membrane structures (Duman et al., 2002). The emergence and interest in cryo-ET has produced some exceptional results for small cells (Grimm et al., 1998; Nickell et al., 2003) and isolated organelles or cell components (Medalia et al., 2002; Nicastro et al., 2000). EM tomographic studies of neurons have focused primarily on the neuromuscular junction (Harlow et al., 2001; Rizzoli and Betz, 2004) and the ribbon synapse of frog saccular hair cells (Lenzi et al., 1999); for a review of ultrastructural studies of the synapse see Torrealba and Carrasco (2004). In addition, ultrastructural studies of stimulation-dependent changes in the synapse have been carried out for the ribbon synapse of saccular hair cells (Lenzi et al., 2002) as well as the frog neuromuscular junction (Rizzoli and Betz, 2004). These studies have all used chemical fixation, a method that greatly reduces the quality and reliability of the biological representation derived from ET. A major eVort of current studies is to develop tools that will allow the study of stimulation-dependent phenomena by ET, using techniques such as rapid-freezing and freeze-substitution (RF-FS),
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high-pressure freezing and freeze-substitution (HPF-FS), and potentially, cryo-ET. At present, such studies are not possible because there is no technology that can combine neuronal patterning, axon stimulation, and optical observation with ET for a single platform. This chapter focuses on the use of ET to study the chemical synapse and in particular the small synapse of the brain. The best characterized of these are the glutamatergic synapses between primary cultured hippocampus neurons. Although mention is made of other neuronal systems and types of synapses, our focus will be on the hippocampus synapses, which have functionally well-characterized plasticity events and can be regarded as the ‘‘gold standard’’ sample for studying memory and learning.
IV. Types of Neurons and Synapses While all metazoans (save for sponges) are a potential source of neurons for structural investigation, a few model organisms have been primarily utilized. These range from the invertebrate neurons of Aplysia to vertebrate hippocampus neurons. While each type of neuron and neuron source may have particular experimental advantages or disadvantages, the biological question is the main driver in neuron choice. Here we will give a brief description of three diVerent types of neurons that have been used to investigate synaptic function with structural methods. At this stage, it is important not only to distinguish between chemical and electrical synapses but also between inhibitory and excitatory synapses, as well as the type of neuronal connection that is made. Chemical synapses involve the release of neurotransmitters and require the packaging of neurotransmitter in ‘‘quanta,’’ the term used to refer to the amount of neurotransmitter in a single synaptic vesicle. Electrical or fast synapses are mediated by connexons or gap junctions; they do not involve transmitter release and are not known to exhibit plasticity in the traditional sense; however, recent studies of Mauthner cells suggest that such connections can undergo a form of plasticity (Pereda et al., 2004). In the following sections, we discuss the three main synapses that have been studied using EM and ET: neuromuscular junction synapses, ribbon synapses, and hippocampal neuron synapses. While all of these synapses have common features, they diVer in a number of important ways. For example, the number and arrangement of vesicles at the synapse show considerable variation. Neuromuscular synapses contain hundreds to thousands of vesicles; they are thought to be stable and not to undergo plasticity events, such as LTP and LTD, but they do undergo more subtle remodeling during postnatal growth (Wilson and Deschenes, 2005). Hippocampal neurons and synapses are undoubtedly the most studied because they display plasticity phenomena with great clarity. Hippocampus neurons form synapses that can undergo a variety of plasticity events, and it is for this reason that much of our knowledge about synaptic plasticity derives from data obtained on hippocampus neurons.
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A. Neuromuscular Junction Synapse The neuromuscular junction is an archetypal synapse, and many of the early structural studies of synapses used the frog neuromuscular junction (Fig. 1A). Seminal studies by Heuser and colleagues (Heuser and Reese, 1981; Heuser et al., 1971) provided much of our fundamental structural understanding of the neuromuscular chemical synapse in both the resting and active states. Recent EM tomographic studies of this system have described a series of structural features that link synaptic vesicles to the ‘‘active zone,’’ from which neurotransmitter will subsequently be secreted (Harlow et al., 2001). More recent studies by Betz and colleagues have investigated vesicle recycling at the frog neuromuscular junction using fixed samples and ET (Rizzoli and Betz, 2004). B. Ribbon Synapses Structural studies on the ribbon synapse found in auditory, vestibular, and bipolar photoreceptor cells of vertebrates (Sterling and Matthews, 2005) have demonstrated that these synapses have a remarkably regular arrangement of synaptic vesicles (Fig. 1B). Although the precise role of this regular arrangement is not fully understood, it has been conjecturally viewed as a ‘‘conveyer belt’’ for vesicle delivery; plasticity phenomena in these synapses appear to revolve around variations in the size and number of vesicles on the conveyer belt (Vollrath and Spiwoks-Becker, 1996). C. CNS Synapses The synapses of the brain can occur at a wide range of places on the neurons. The interactions can be axo-axonic, somato-axonic, somato-dendritic, dendroaxonic, and even dendro-dendritic. The types of activity at these various sites can
Fig. 1 Electron micrographs of the three main types of synapse being studied by ET. (A) Neuromuscular junction, (B) ribbon synapse, from Sterling and Matthews (2005), and (C) a small synapse of the brain, Stowell et al. (unpublished data). The main structural diVerences between these types of synapses are the number and organization of synaptic vesicles.
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be either excitatory (glutamatergic) or inhibitory (GABAergic). The earliest ultrastructural investigation of the brain described two types of synapses, type 1 or asymmetric and type 2 or symmetric (Colonnier, 1968; Gray, 1959). The vast majority of synapses of the brain are asymmetric (Fig. 1C). Ellisman and colleagues have investigated structural changes that occur in the rat hippocampus following ischemia (Martone et al., 2000) and found an intriguing vesicle rearrangement following ischemia.
V. Sources of Neurons for Structural Studies Many structural investigations of neurons have utilized intact tissue. The obvious advantages of such methods are the nativelike state of the neurons and their associated synapses, as well as of the surrounding cells, primarily glial. One of the disadvantages is the processing time required between a given treatment and sample preservation by either rapid freezing or chemical fixation; there is also diYculty from the thickness of the samples and poor reproducibility of the sample treatments. For these reasons, we and others have focused on the use of primary cultured hippocampus neurons for studying the architecture of synaptic plasticity. The ability to reliably culture neurons in vitro (Banker and Goslin, 1991) has opened up a number of areas in neuronal studies. It has permitted the observation of neural patterning (Fromherz and Schaden, 1994), growth-cone targeting (Ming et al., 2002), and synaptogenesis (Ahmari et al., 2000), to name just a few of the areas that have benefited from this methodology. One of the most important aspects of culture methods for hippocampus neurons is that the process can be done at relatively low cell density. Such culture conditions open the possibility of collecting cryo-electron tomographic data on individual synapses that have been grown under defined conditions and subject to defined stimulation protocols. For structural studies, neurons can be cultured on a variety of substrates, including glass, carbon-coated Formvar or collodion (nitrocellulose), as well as filter disks. Our laboratory has also investigated a number of substrates that are compatible with lithographic manufacturing procedures and that allow thin electrontransparent windows to be fabricated. These include silicon, silicon nitride, and anodized alumina oxide (AAO) (Fig. 2). While various substrates will support primary neuronal culture, several hurdles must typically be overcome; these vary with the substrate. In the case of silicon and silicon nitride substrates, pretreatment of the substrates with oxygen plasma greatly improves the reproducibility of the cultures. In addition, a variety of more standard surface treatments are needed to help the substrate to support neurons. These include the application of MatrigelÔ, polylysine, and/or polylysine/laminin. Our most reproducible and successful results have been achieved with polylysine/ laminin. Our typical procedure starts with brains from P0 to P2 neonatal mice, dissected in calcium-magnesium-free Hank’s balanced salt solution (HBSS). After the brain was separated into the two hemispheres, the C-shaped structure in the
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Fig. 2 Light micrographs of low-density, primary-cultured murine hippocampus neurons grown on silicon oxide/nitride windows showing a high degree of diVerentiation and synapse formation. (A) Field view, (B) Close-up of the boxed area in (A) (500 500 mm2), showing the degree of branching and connectivity. (C) Primary neurons grown on AAO substrates.
posterior half of both hemispheres, hippocampi, were removed under a dissection microscope at 10 to 15 magnification. The meninges and choroid plexus were next removed, and the tissue treated with papain for 1½ h. The tissue was then gently triturated with a fire-polished Pasteur pipet. This material was layered on top of a Percoll gradient (30–60%) and centrifuged for 30 min at 3000 rpm without brake. The first band from the bottom of the tube was collected and concentrated by centrifugation. The resulting pellet was washed with HBSS medium several times and resuspended in neurobasal medium without serum, supplemented with B27, glutaMAX (Invitrogen) streptomycin/penicillin, and glutamate. Generally, 5.0 106 cells/ml were plated on polylysine/laminin-treated substrates immobilized on glass coverslips and allowed to stand for 40 min. Then, 2.5 ml of medium was added to the plate and the neurons were incubated at 37 C for 2 days. Half of the medium was then replaced with fresh medium without glutamate. Subsequently, the medium was replaced every week. The number of plated cells varies slightly depending on the substrate, as not all substrates provide good initial cell adhesion. This simple procedure, modified from that of Banker, provides reproducible cultures on a wide range of substrates treated with laminin/polylysine.
VI. Methods for Assessing Neuron and Synapse Integrity Although primary cultured hippocampus neurons are regarded as the ‘‘gold standard’’ of neuronal systems, it is important to use various quantitative techniques to assess the proper formation of functional synapses prior to structural studies. This can be achieved in several ways. One method is to employ a battery of known synaptic markers for an analysis by fluorescence microscopy. A second method is to use quantitative polymerase chain reaction (QPCR) to measure the
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Postsynaptic markers
Bassoon Piccolo Munc 13 Cask RIM 1 RIM 2 Snap 25 Syntaxin Synaptobrevin
Gephyrin Grip Neuroligin NMDA receptor PSD95 SAP 102 AMPA receptor CaM kinase Syntenin
level of expression of known synaptic marker proteins during synaptogenesis. Table I lists a series of presynaptic and postsynaptic markers that can be used to quantitatively assess the formation of synapses from neurons cultured on various substrates. The advantage of QPCR is that one can assay a much larger number of markers in a single experiment. Previous work has utilized a battery of 500 gene markers, of which approximately half are involved in synaptogenesis (Saito et al., 2005). Thus, QPCR allows one to develop a quantitative although general measure of the biological relevance of neurons cultured on various substrates relative to traditionally cultured neurons; with the resulting information one can rapidly home in on optimal conditions. The important fundamental diVerence between these methods is that QPCR provides a global readout of all the neurons whereas fluorescence methods allow one to examine individual neurons and synapses within a culture. One obvious question to address using such methods is the diVerence between the amounts of various synaptic markers in cells grown on various substrates. To date, we have not observed any substrate-dependent diVerences. The main application of synaptic marker quantification is to assess the diVerentiation of neurons, which in turn enables a study of synaptogenesis, and to monitor the viability and reproducibility of our cultures.
VII. Rapid Freezing and HPF of Neurons Neurons grown on various substrates can either be plunge frozen or highpressure frozen to maintain their structural integrity. Both of these methods are adequately covered in other sections of this book. Here it suYces to say that HPF has the advantage that larger sample masses can be well preserved. We have cultured neurons on silicon substrates patterned by photolithography to display 50–100 mm SU8 epoxy-based polymer surface features and obtained good sample
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Fig. 3 Thin sections cut from samples of 10-day-old hippocampus neurons grown on a silicon substrate patterned with SU8 and plunge frozen in liquid ethane. (A) Overview (2 2 mm2) of two neuron wells with an SU8 picket fence and outgrowth track. (B) More detailed view [boxed area in (A) 200 200 mm2] showing neuronal cell bodies within the ‘‘picket fences’’ (dark circles). (C) A single slice (1 1 mm2) from a tomogram taken from the sections in (B), showing well-preserved synaptic vesicles.
preservation by either plunge freezing in liquid ethane or HPF, followed by freezesubstituting with standard procedures. The results of such studies are shown in Fig. 3. Following plunge freezing and freeze-substitution, we cut thin sections that demonstrated good overall preservation of the sample and localization of neurons within the designed surface structures. The importance of this result is that it demonstrates good sample preservation for structural studies on substrates that are suitable for functional analysis using field potential electrophysiological methods as discussed below.
VIII. Modeling and Analysis of Synaptic Structures Once suitable tomograms have been collected, data processing and analysis proceeds in a similar fashion to other samples. Other chapters of this book deal extensively with modeling, so here we have only described the aspects that are important for understanding synaptic structure and function. A. Size and Organization of the Vesicle Pool The size and organization of the pool of synaptic vesicles is one of the first elements to be investigated. As discussed above, the size of this pool can vary greatly, depending on the type of neuron. In addition, stimulation-dependent changes can occur, and dramatic changes can occur when important traYcking protein are either removed or inhibited. A classic example of this phenomenon is
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Fig. 4 (A) Images from neonate mice of wild-type (WT) and kif1a-knockout (KO) genotypes. The knockout mice show a distinct loss of the clustered synaptic vesicles seen in the WT. (B) Twentyone-day-old primary cell culture of murine hippocampus pyramidal neurons grown on a treated silicon substrate with clearly observed synaptic vesicles and microtubules. (C) Three-dimensional (3D) model of the tomographic data in (B) created using the IMOD suite of programs.
the loss of typical asymmetric synapses in kif1a-knockout mice. Kif1A is a kinesinlike motor protein responsible for traYcking to the presynaptic active zone. In the kif1a-knockout mice there is a pronounced accumulation of synaptic-like vesicle at the Golgi and the loss of a well-organized synaptic pool (Fig. 4A). Another important study was that by Betz and colleagues (Rizzoli and Betz, 2004) in which they combined ET with photoconversion of dye-labeled vesicles to study the structural organization of the vesicle pool at the neuromuscular junction.
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B. Organization of the Postsynaptic Density Proteins The postsynaptic density (PSD) and the organization of the various channels and channels associated complexes are of particular interest to those studying synaptic plasticity. Dramatic PSD reorganization is known to occur following stimulations such as siezure (Martone et al., 2000). Recently, an intriguing study that combined ET, electrophysiology, and modeling, investigated the role of acetylcholine isoforms in the chick ciliary ganglion synapse (Coggan et al., 2005). This study provided compelling evidence that ectopic neurotransmission is functionally relevant, thereby expanding the modes of biologically important neurotransmission. This study also highlights the need to determine the localization of neurotransmitter receptors in order to fully understand synaptic transmission and its modulation.
C. Synaptic Cleft Complexes The organization of the cell adhesion proteins that interact across the synaptic cleft is important not only for understanding synaptogenesis but for its important consequences with regard to receptor localization and arrangement. A recent cryo-ET study has given some insight into this organization and discovered that there are substantial lateral connections within the synaptic cleft as well as the expected intercellular connections. (Lucic et al., 2005).
D. Endosomal Compartments The vesicle cycle of the small synapse of the brain is an intriguing and controversial area of study. At least two modes of vesicle fusion and recycling have been observed: full fusion and ‘‘kiss and run’’ (Fernandez-Peruchena et al., 2005). Understanding how these diVerent types of fusion, and subsequent endocytosis, are regulated is critical to understanding presynaptic modes of plasticity. While studies at the neuromuscular junction (Rizzoli and Betz, 2004) have implicated a random distribution of ‘‘kiss and run’’ vesicles, such studies have not been performed on the small synapse of the CNS. It will be important to understand how these diVerent mechanisms of vesicle fusion are regulated and how this regulation aVects the activity of a given synapse.
E. Microtubule and Neurofilament Arrangements The organization of the microtubule network associated with both the axon and the dendrite is of particular interest when trying to understand both synaptogenesis and long-term plasticity. Although little has been done to investigate the detailed organization of microtubules, it is well established that they are critical for cargo transport to both the pre- and postsynaptic zones. Disruption of microtubules
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prevents the formation of new synaptic boutons (Ruiz-Canada et al., 2004), and as discussed above, microtubule motor proteins are essential for proper formation of a synaptic pool. The proposed roles of neurofilaments have included a ‘‘girder’’-like function for the organization of acetylcholine receptors as well as a role in defining microtubule organization for material transport (Paysan et al., 2000).
IX. Stimulation-Dependent Changes in Synaptic Structure One of the most intriguing aspects of synaptic structure is the structural change that occurs during specific stimulations. While very few studies of this nature have yet been performed, this is certainly an area that will be moving forward with great speed and anticipation. We mentioned above the studies of the frog neuromuscular junction vesicle cycle, as well as PSD reorganization following seizure in hippocampal neurons. Another important study was the investigation of the auditory ribbon synapse of the Xenopus saccular hair cells, before and after stimulation (Lenzi et al., 2002). Figure 5 shows the results from this study, which demonstrated a dramatic reorganization of the ribbon synapse vesicle pool following stimulation. The ability to perform defined stimulation protocols and then to collect structural data provides the possibility of important insights into the organization and mechanistic functioning of synapses. Such data cannot be obtained with any other techniques.
X. The Future of EM Tomography and Synapses One of the most fascinating areas for synaptic structure studies by ET is the ultrastructural analysis of synapses that have recently undergone electrical stimulation with recording. While this kind of study can in principle be performed using traditional electrophysiological methods, as were used for studies on the frog neuromuscular junction, such methods are impractical when studying early stages of plasticity. Only one or a few neurons per culture could be properly stimulated and high-pressure frozen for subsequent ET analysis. The solution to such a problem is to utilize multielectrode arrays. Current technologies for studying neurons in arrays have been optimized for signal generation and detection, using either cultured neurons or brain slices. Several commercial systems are presently available that employ large-format arrays and are optimized mainly for slice work and perfusion. However, these devices are not suitable for EM-tomographic analysis because the samples cannot be rapidly frozen or even high-pressure frozen. Both the size and the thermal mass of these samples are too great to allow rapid cooling, a requirement for adequate sample preservation. To achieve cooling rates suitable for HPF, while retaining the capability to stimulate and pattern the neurons, we have designed and fabricated new devices specifically for this purpose (Fig. 6). With such tools, researchers will be able to recapitulate a variety
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Fig. 5 Stimulation-dependent changes in the saccular ribbon synapse, reproduced from Lenzi et al. (2002). Electron tomographic reconstructions of hair cell synapses exposed to 0-Ca2þ saline (A and C) or high-Kþ saline (B and D), which stimulates the synapse. (A and B) Single x–y planes through the reconstructed volumes are shown. The hair cell cytoplasm contains the synaptic body (SB) surrounded by synaptic vesicles, outlying cytoplasmic vesicles [arrows in (A)], coated vesicles [arrows in (B)], membrane-bounded cistern (c), mitochondria (m), and postsynaptic cell (p). (C and D) 3D structure of presynaptic organelles from the same reconstructions shown in (A) and (B). The SB (blue) was rendered as hollow and semitransparent and lies adjacent to the hair cell’s plasma membrane (red). SB-associated vesicles (yellow) surround the SB; outlying vesicles (green, thick long arrows) lie further out in the cytoplasm. Also visible are coated vesicles (gold, thin long arrows), cisternae (purple, thick short arrows), and mitochondria (blue, thin short arrows). (D) Regions of presynaptic density (pink) lie beneath the SB on the plasma membrane, which formed a tubular invagination (arrowhead). The flat surfaces of the mitochondria and the open face of the SBs denote the edges of the reconstruction. Synapses 1 and 11; scale bars ¼ 200 nm.
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Fig. 6 (A) Silicon chip for use in a high-pressure freezer. The silicon chip is designed so that the inner core of electrodes can be snapped oV and introduced directly into the cap of an HPF device. Left: overview of the chip designed for the HPF; CD, central disk for culturing neurons; IC, interconnect for direct hookup to both a multichannel systems or Panasonic MED-64 stimulation and recording amplifier system; SC, snapoV connections between the CD and the main chip body. Right: detail of overview; NW, neuron well; SNW, silicon nitride window for synaptic observation. (B) SEM image of a silicon EM chip with SU8 features for culturing neurons. (C) Detail of (B) showing the picket fence cell body containment posts.
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of synaptic phenomena that have previously been characterized by fluorescent light microscopy and electrophysiology, and then go on to observe the identical synapse using ET.
XI. Conclusions The use of ET to study synaptic structure and function holds tremendous promise, yet there are substantial hurdles that need to be overcome. The ultimate goal of such studies is to identify and ascertain the organization of membranes and supramolecular complexes in a resting synapse or in a synapse after treatment with well-defined stimulation protocols. Many of the relevant molecular complexes are of suYcient mass (>200 kDa) that their appearance will cause clear contrast in conventional EM. These include such important assemblies as the SNARE complex, NSF, and NMDA-PSD-95, the bassoon and piccolo complexes that order synaptic vesicles, polysomes, Ca2þ-release channels, microtubules, and associated transport vesicles. While the study of several systems, such as the neuromuscular junction and ribbon synapses following stimulation, has been achieved and important insights gained, such studies cannot currently be performed with primary cultured hippocampus neurons. This limitation can be overcome by employing microfabrication methods and multielectrode stimulation technologies, so that well-defined stimulation protocols can be achieved and their eVects rapidly analyzed by EM and ET. In the end, a knowledge of the distribution and organization of relevant macromolecular complexes, including membranes, will give us important new insights into the workings of the synapse. Acknowledgments We would like to thank our colleagues at the University of Colorado for helpful comments and advice, J. Richard McIntosh for patients and insight, and the Beckman Foundation for financial support.
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Electron Tomographic Methods for Studying the Chemical Synapse Christopher P. Arthur, David B. Serrell, Maria Pagratis, David L. Potter, Dudley S. Finch, and Michael H. B. Stowell Department of MCD Biology University of Colorado Boulder, Colorado 80309
I. II. III. IV.
V. VI. VII. VIII.
IX. X. XI.
Introduction and Rationale Neurons, Neural Networks, and the Synapse ET and the Synapse Types of Neurons and Synapses A. Neuromuscular Junction Synapse B. Ribbon Synapses C. CNS Synapses Sources of Neurons for Structural Studies Methods for Assessing Neuron and Synapse Integrity Rapid Freezing and HPF of Neurons Modeling and Analysis of Synaptic Structures A. Size and Organization of the Vesicle Pool B. Organization of the Postsynaptic Density Proteins C. Synaptic Cleft Complexes D. Endosomal Compartments E. Microtubule and Neurofilament Arrangements Stimulation-Dependent Changes in Synaptic Structure The Future of EM Tomography and Synapses Conclusions References
This chapter focuses on the use of electron tomography (ET) to study the chemical synapse. It discusses the diVerent types of chemical synapses and their properties as well as the biological questions that are addressable using ET. We present METHODS IN CELL BIOLOGY, VOL. 79 Copyright 2007, Elsevier Inc. All rights reserved.
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methods for culturing neurons on a variety of substrates suitable for ET and discuss the development of novel tools that combine electrophysiological methods with ET methods for studying active synapses.
I. Introduction and Rationale Our current understanding of synapse formation and organization comes from a variety of studies that used immunofluorescence (or GFP-tagged proteins), fluorescent dyes, and/or electron microscopy (EM) of fixed embedded samples (Garner et al., 2002). All these methods are limited in their resolution and their inability to distinguish a functioning synapse from one that is inactive or undergoing formation or decay. In addition, both detailed structural information about the molecular and supramolecular organization of the synapse and knowledge about the dynamic events associated with synaptic transmission are lacking. Such information is important for understanding phenomena such as synaptogenesis, synaptic transmission and plasticity, and synaptic regeneration following neurological damage. During the last few years, neuroscientists have begun to dig more deeply into the structures and process of synaptic behavior by using electron tomography (ET). In this chapter, we will review these methods, recent studies of synapses, their limitations, and consider the ways in which future developments may allow for the systematic structural investigation of synaptic architecture under controlled conditions.
II. Neurons, Neural Networks, and the Synapse Living neural networks (LNN) are one of Nature’s greatest technological achievements. They can be massively parallel and dynamic computational devices with built-in plasticity and repair mechanisms. The most complex LNN, a human brain, comprises over 1014 connections, more connections than all the computing power manufactured between 1981 and 2001 (Stowell, 2003). The chemical synapse is at the heart of a functioning LNN. During brain development, chemical synapses are constructed following the establishment of contact between a growth-cone at the end of a growing axon and the cell that will receive its signal. Synaptogenesis is the process by which this complex, intercellular connection is formed. Once formed, a functional synapse has a dynamic life. Of course it transmits signals between neurons, but it is also subject to slower modulations that aVect its behavior and function. Individual synaptic signals undergo modulations, such as long-term potentiation (LTP) and long-term depression (LTD) (Bliss et al., 2003; Gilbert, 1998; Lisman, 2003), and in neurodegenerative diseases they can disintegrate. These adaptive changes in synapse function, which can occur on timescales ranging from seconds to minutes, are the foundation for more permanent changes in LNN wiring that may involve the formation of new synapses (Knott et al., 2002;
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Trachtenberg et al., 2002), giving rise to biologically indelible memory. All these phenomena are central to our understanding of memory and learning, and they also serve as a foundation for understanding biological computation in general and the development of neuromorphic technologies in the future. Naturally, a great deal of eVort has been spent on trying to understand the genetic and molecular bases of neuronal function and in particular memory and learning.
III. ET and the Synapse The last decade has seen a strong resurgence of interest in biological EM. While this trend has been led by dramatic progress in EM of macromolecules, the analysis of cellular ultrastructure has not lagged far behind (McIntosh, 2001). Several technological advances have facilitated this renaissance. On the instrumentation front, field emission guns (FEGs), which provide electron beams with good coherence and brightness, have improved the imaging of frozen-hydrated samples. Computer-controlled specimen stages have improved the regulation of sample position and automated data collection. Greatly improved CCD cameras have significantly increased the speed of image acquisition and analysis. Finally, as in almost every area of research, the dramatic decrease in price for computational speed and data storage has also played an important role. However, as in most areas of research, perhaps the most dramatic improvements have been the result of greatly improved methods for sample preparation; in particular, rapid freezing of biological specimens has opened the door for higher resolution studies of macromolecules and cells in their native states (Dubochet et al., 1983; Lepault et al., 1983). As a result, the use of ET for ultrastructural analysis has been applied extensively to a variety of cells and organisms such as Caenorhabditis elegans (Mu¨ller-Reichert et al., 2003), Saccharomyces cerevisiae kineticore distribution (O’Toole et al., 2002), and cytoplasmic membrane structures (Duman et al., 2002). The emergence and interest in cryo-ET has produced some exceptional results for small cells (Grimm et al., 1998; Nickell et al., 2003) and isolated organelles or cell components (Medalia et al., 2002; Nicastro et al., 2000). EM tomographic studies of neurons have focused primarily on the neuromuscular junction (Harlow et al., 2001; Rizzoli and Betz, 2004) and the ribbon synapse of frog saccular hair cells (Lenzi et al., 1999); for a review of ultrastructural studies of the synapse see Torrealba and Carrasco (2004). In addition, ultrastructural studies of stimulation-dependent changes in the synapse have been carried out for the ribbon synapse of saccular hair cells (Lenzi et al., 2002) as well as the frog neuromuscular junction (Rizzoli and Betz, 2004). These studies have all used chemical fixation, a method that greatly reduces the quality and reliability of the biological representation derived from ET. A major eVort of current studies is to develop tools that will allow the study of stimulation-dependent phenomena by ET, using techniques such as rapid-freezing and freeze-substitution (RF-FS),
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high-pressure freezing and freeze-substitution (HPF-FS), and potentially, cryo-ET. At present, such studies are not possible because there is no technology that can combine neuronal patterning, axon stimulation, and optical observation with ET for a single platform. This chapter focuses on the use of ET to study the chemical synapse and in particular the small synapse of the brain. The best characterized of these are the glutamatergic synapses between primary cultured hippocampus neurons. Although mention is made of other neuronal systems and types of synapses, our focus will be on the hippocampus synapses, which have functionally well-characterized plasticity events and can be regarded as the ‘‘gold standard’’ sample for studying memory and learning.
IV. Types of Neurons and Synapses While all metazoans (save for sponges) are a potential source of neurons for structural investigation, a few model organisms have been primarily utilized. These range from the invertebrate neurons of Aplysia to vertebrate hippocampus neurons. While each type of neuron and neuron source may have particular experimental advantages or disadvantages, the biological question is the main driver in neuron choice. Here we will give a brief description of three diVerent types of neurons that have been used to investigate synaptic function with structural methods. At this stage, it is important not only to distinguish between chemical and electrical synapses but also between inhibitory and excitatory synapses, as well as the type of neuronal connection that is made. Chemical synapses involve the release of neurotransmitters and require the packaging of neurotransmitter in ‘‘quanta,’’ the term used to refer to the amount of neurotransmitter in a single synaptic vesicle. Electrical or fast synapses are mediated by connexons or gap junctions; they do not involve transmitter release and are not known to exhibit plasticity in the traditional sense; however, recent studies of Mauthner cells suggest that such connections can undergo a form of plasticity (Pereda et al., 2004). In the following sections, we discuss the three main synapses that have been studied using EM and ET: neuromuscular junction synapses, ribbon synapses, and hippocampal neuron synapses. While all of these synapses have common features, they diVer in a number of important ways. For example, the number and arrangement of vesicles at the synapse show considerable variation. Neuromuscular synapses contain hundreds to thousands of vesicles; they are thought to be stable and not to undergo plasticity events, such as LTP and LTD, but they do undergo more subtle remodeling during postnatal growth (Wilson and Deschenes, 2005). Hippocampal neurons and synapses are undoubtedly the most studied because they display plasticity phenomena with great clarity. Hippocampus neurons form synapses that can undergo a variety of plasticity events, and it is for this reason that much of our knowledge about synaptic plasticity derives from data obtained on hippocampus neurons.
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A. Neuromuscular Junction Synapse The neuromuscular junction is an archetypal synapse, and many of the early structural studies of synapses used the frog neuromuscular junction (Fig. 1A). Seminal studies by Heuser and colleagues (Heuser and Reese, 1981; Heuser et al., 1971) provided much of our fundamental structural understanding of the neuromuscular chemical synapse in both the resting and active states. Recent EM tomographic studies of this system have described a series of structural features that link synaptic vesicles to the ‘‘active zone,’’ from which neurotransmitter will subsequently be secreted (Harlow et al., 2001). More recent studies by Betz and colleagues have investigated vesicle recycling at the frog neuromuscular junction using fixed samples and ET (Rizzoli and Betz, 2004). B. Ribbon Synapses Structural studies on the ribbon synapse found in auditory, vestibular, and bipolar photoreceptor cells of vertebrates (Sterling and Matthews, 2005) have demonstrated that these synapses have a remarkably regular arrangement of synaptic vesicles (Fig. 1B). Although the precise role of this regular arrangement is not fully understood, it has been conjecturally viewed as a ‘‘conveyer belt’’ for vesicle delivery; plasticity phenomena in these synapses appear to revolve around variations in the size and number of vesicles on the conveyer belt (Vollrath and Spiwoks-Becker, 1996). C. CNS Synapses The synapses of the brain can occur at a wide range of places on the neurons. The interactions can be axo-axonic, somato-axonic, somato-dendritic, dendroaxonic, and even dendro-dendritic. The types of activity at these various sites can
Fig. 1 Electron micrographs of the three main types of synapse being studied by ET. (A) Neuromuscular junction, (B) ribbon synapse, from Sterling and Matthews (2005), and (C) a small synapse of the brain, Stowell et al. (unpublished data). The main structural diVerences between these types of synapses are the number and organization of synaptic vesicles.
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be either excitatory (glutamatergic) or inhibitory (GABAergic). The earliest ultrastructural investigation of the brain described two types of synapses, type 1 or asymmetric and type 2 or symmetric (Colonnier, 1968; Gray, 1959). The vast majority of synapses of the brain are asymmetric (Fig. 1C). Ellisman and colleagues have investigated structural changes that occur in the rat hippocampus following ischemia (Martone et al., 2000) and found an intriguing vesicle rearrangement following ischemia.
V. Sources of Neurons for Structural Studies Many structural investigations of neurons have utilized intact tissue. The obvious advantages of such methods are the nativelike state of the neurons and their associated synapses, as well as of the surrounding cells, primarily glial. One of the disadvantages is the processing time required between a given treatment and sample preservation by either rapid freezing or chemical fixation; there is also diYculty from the thickness of the samples and poor reproducibility of the sample treatments. For these reasons, we and others have focused on the use of primary cultured hippocampus neurons for studying the architecture of synaptic plasticity. The ability to reliably culture neurons in vitro (Banker and Goslin, 1991) has opened up a number of areas in neuronal studies. It has permitted the observation of neural patterning (Fromherz and Schaden, 1994), growth-cone targeting (Ming et al., 2002), and synaptogenesis (Ahmari et al., 2000), to name just a few of the areas that have benefited from this methodology. One of the most important aspects of culture methods for hippocampus neurons is that the process can be done at relatively low cell density. Such culture conditions open the possibility of collecting cryo-electron tomographic data on individual synapses that have been grown under defined conditions and subject to defined stimulation protocols. For structural studies, neurons can be cultured on a variety of substrates, including glass, carbon-coated Formvar or collodion (nitrocellulose), as well as filter disks. Our laboratory has also investigated a number of substrates that are compatible with lithographic manufacturing procedures and that allow thin electrontransparent windows to be fabricated. These include silicon, silicon nitride, and anodized alumina oxide (AAO) (Fig. 2). While various substrates will support primary neuronal culture, several hurdles must typically be overcome; these vary with the substrate. In the case of silicon and silicon nitride substrates, pretreatment of the substrates with oxygen plasma greatly improves the reproducibility of the cultures. In addition, a variety of more standard surface treatments are needed to help the substrate to support neurons. These include the application of MatrigelÔ, polylysine, and/or polylysine/laminin. Our most reproducible and successful results have been achieved with polylysine/ laminin. Our typical procedure starts with brains from P0 to P2 neonatal mice, dissected in calcium-magnesium-free Hank’s balanced salt solution (HBSS). After the brain was separated into the two hemispheres, the C-shaped structure in the
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Fig. 2 Light micrographs of low-density, primary-cultured murine hippocampus neurons grown on silicon oxide/nitride windows showing a high degree of diVerentiation and synapse formation. (A) Field view, (B) Close-up of the boxed area in (A) (500 500 mm2), showing the degree of branching and connectivity. (C) Primary neurons grown on AAO substrates.
posterior half of both hemispheres, hippocampi, were removed under a dissection microscope at 10 to 15 magnification. The meninges and choroid plexus were next removed, and the tissue treated with papain for 1½ h. The tissue was then gently triturated with a fire-polished Pasteur pipet. This material was layered on top of a Percoll gradient (30–60%) and centrifuged for 30 min at 3000 rpm without brake. The first band from the bottom of the tube was collected and concentrated by centrifugation. The resulting pellet was washed with HBSS medium several times and resuspended in neurobasal medium without serum, supplemented with B27, glutaMAX (Invitrogen) streptomycin/penicillin, and glutamate. Generally, 5.0 106 cells/ml were plated on polylysine/laminin-treated substrates immobilized on glass coverslips and allowed to stand for 40 min. Then, 2.5 ml of medium was added to the plate and the neurons were incubated at 37 C for 2 days. Half of the medium was then replaced with fresh medium without glutamate. Subsequently, the medium was replaced every week. The number of plated cells varies slightly depending on the substrate, as not all substrates provide good initial cell adhesion. This simple procedure, modified from that of Banker, provides reproducible cultures on a wide range of substrates treated with laminin/polylysine.
VI. Methods for Assessing Neuron and Synapse Integrity Although primary cultured hippocampus neurons are regarded as the ‘‘gold standard’’ of neuronal systems, it is important to use various quantitative techniques to assess the proper formation of functional synapses prior to structural studies. This can be achieved in several ways. One method is to employ a battery of known synaptic markers for an analysis by fluorescence microscopy. A second method is to use quantitative polymerase chain reaction (QPCR) to measure the
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248 Table I Synaptic Markers for QPCR Presynaptic markers
Postsynaptic markers
Bassoon Piccolo Munc 13 Cask RIM 1 RIM 2 Snap 25 Syntaxin Synaptobrevin
Gephyrin Grip Neuroligin NMDA receptor PSD95 SAP 102 AMPA receptor CaM kinase Syntenin
level of expression of known synaptic marker proteins during synaptogenesis. Table I lists a series of presynaptic and postsynaptic markers that can be used to quantitatively assess the formation of synapses from neurons cultured on various substrates. The advantage of QPCR is that one can assay a much larger number of markers in a single experiment. Previous work has utilized a battery of 500 gene markers, of which approximately half are involved in synaptogenesis (Saito et al., 2005). Thus, QPCR allows one to develop a quantitative although general measure of the biological relevance of neurons cultured on various substrates relative to traditionally cultured neurons; with the resulting information one can rapidly home in on optimal conditions. The important fundamental diVerence between these methods is that QPCR provides a global readout of all the neurons whereas fluorescence methods allow one to examine individual neurons and synapses within a culture. One obvious question to address using such methods is the diVerence between the amounts of various synaptic markers in cells grown on various substrates. To date, we have not observed any substrate-dependent diVerences. The main application of synaptic marker quantification is to assess the diVerentiation of neurons, which in turn enables a study of synaptogenesis, and to monitor the viability and reproducibility of our cultures.
VII. Rapid Freezing and HPF of Neurons Neurons grown on various substrates can either be plunge frozen or highpressure frozen to maintain their structural integrity. Both of these methods are adequately covered in other sections of this book. Here it suYces to say that HPF has the advantage that larger sample masses can be well preserved. We have cultured neurons on silicon substrates patterned by photolithography to display 50–100 mm SU8 epoxy-based polymer surface features and obtained good sample
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Fig. 3 Thin sections cut from samples of 10-day-old hippocampus neurons grown on a silicon substrate patterned with SU8 and plunge frozen in liquid ethane. (A) Overview (2 2 mm2) of two neuron wells with an SU8 picket fence and outgrowth track. (B) More detailed view [boxed area in (A) 200 200 mm2] showing neuronal cell bodies within the ‘‘picket fences’’ (dark circles). (C) A single slice (1 1 mm2) from a tomogram taken from the sections in (B), showing well-preserved synaptic vesicles.
preservation by either plunge freezing in liquid ethane or HPF, followed by freezesubstituting with standard procedures. The results of such studies are shown in Fig. 3. Following plunge freezing and freeze-substitution, we cut thin sections that demonstrated good overall preservation of the sample and localization of neurons within the designed surface structures. The importance of this result is that it demonstrates good sample preservation for structural studies on substrates that are suitable for functional analysis using field potential electrophysiological methods as discussed below.
VIII. Modeling and Analysis of Synaptic Structures Once suitable tomograms have been collected, data processing and analysis proceeds in a similar fashion to other samples. Other chapters of this book deal extensively with modeling, so here we have only described the aspects that are important for understanding synaptic structure and function. A. Size and Organization of the Vesicle Pool The size and organization of the pool of synaptic vesicles is one of the first elements to be investigated. As discussed above, the size of this pool can vary greatly, depending on the type of neuron. In addition, stimulation-dependent changes can occur, and dramatic changes can occur when important traYcking protein are either removed or inhibited. A classic example of this phenomenon is
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Fig. 4 (A) Images from neonate mice of wild-type (WT) and kif1a-knockout (KO) genotypes. The knockout mice show a distinct loss of the clustered synaptic vesicles seen in the WT. (B) Twentyone-day-old primary cell culture of murine hippocampus pyramidal neurons grown on a treated silicon substrate with clearly observed synaptic vesicles and microtubules. (C) Three-dimensional (3D) model of the tomographic data in (B) created using the IMOD suite of programs.
the loss of typical asymmetric synapses in kif1a-knockout mice. Kif1A is a kinesinlike motor protein responsible for traYcking to the presynaptic active zone. In the kif1a-knockout mice there is a pronounced accumulation of synaptic-like vesicle at the Golgi and the loss of a well-organized synaptic pool (Fig. 4A). Another important study was that by Betz and colleagues (Rizzoli and Betz, 2004) in which they combined ET with photoconversion of dye-labeled vesicles to study the structural organization of the vesicle pool at the neuromuscular junction.
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B. Organization of the Postsynaptic Density Proteins The postsynaptic density (PSD) and the organization of the various channels and channels associated complexes are of particular interest to those studying synaptic plasticity. Dramatic PSD reorganization is known to occur following stimulations such as siezure (Martone et al., 2000). Recently, an intriguing study that combined ET, electrophysiology, and modeling, investigated the role of acetylcholine isoforms in the chick ciliary ganglion synapse (Coggan et al., 2005). This study provided compelling evidence that ectopic neurotransmission is functionally relevant, thereby expanding the modes of biologically important neurotransmission. This study also highlights the need to determine the localization of neurotransmitter receptors in order to fully understand synaptic transmission and its modulation.
C. Synaptic Cleft Complexes The organization of the cell adhesion proteins that interact across the synaptic cleft is important not only for understanding synaptogenesis but for its important consequences with regard to receptor localization and arrangement. A recent cryo-ET study has given some insight into this organization and discovered that there are substantial lateral connections within the synaptic cleft as well as the expected intercellular connections. (Lucic et al., 2005).
D. Endosomal Compartments The vesicle cycle of the small synapse of the brain is an intriguing and controversial area of study. At least two modes of vesicle fusion and recycling have been observed: full fusion and ‘‘kiss and run’’ (Fernandez-Peruchena et al., 2005). Understanding how these diVerent types of fusion, and subsequent endocytosis, are regulated is critical to understanding presynaptic modes of plasticity. While studies at the neuromuscular junction (Rizzoli and Betz, 2004) have implicated a random distribution of ‘‘kiss and run’’ vesicles, such studies have not been performed on the small synapse of the CNS. It will be important to understand how these diVerent mechanisms of vesicle fusion are regulated and how this regulation aVects the activity of a given synapse.
E. Microtubule and Neurofilament Arrangements The organization of the microtubule network associated with both the axon and the dendrite is of particular interest when trying to understand both synaptogenesis and long-term plasticity. Although little has been done to investigate the detailed organization of microtubules, it is well established that they are critical for cargo transport to both the pre- and postsynaptic zones. Disruption of microtubules
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prevents the formation of new synaptic boutons (Ruiz-Canada et al., 2004), and as discussed above, microtubule motor proteins are essential for proper formation of a synaptic pool. The proposed roles of neurofilaments have included a ‘‘girder’’-like function for the organization of acetylcholine receptors as well as a role in defining microtubule organization for material transport (Paysan et al., 2000).
IX. Stimulation-Dependent Changes in Synaptic Structure One of the most intriguing aspects of synaptic structure is the structural change that occurs during specific stimulations. While very few studies of this nature have yet been performed, this is certainly an area that will be moving forward with great speed and anticipation. We mentioned above the studies of the frog neuromuscular junction vesicle cycle, as well as PSD reorganization following seizure in hippocampal neurons. Another important study was the investigation of the auditory ribbon synapse of the Xenopus saccular hair cells, before and after stimulation (Lenzi et al., 2002). Figure 5 shows the results from this study, which demonstrated a dramatic reorganization of the ribbon synapse vesicle pool following stimulation. The ability to perform defined stimulation protocols and then to collect structural data provides the possibility of important insights into the organization and mechanistic functioning of synapses. Such data cannot be obtained with any other techniques.
X. The Future of EM Tomography and Synapses One of the most fascinating areas for synaptic structure studies by ET is the ultrastructural analysis of synapses that have recently undergone electrical stimulation with recording. While this kind of study can in principle be performed using traditional electrophysiological methods, as were used for studies on the frog neuromuscular junction, such methods are impractical when studying early stages of plasticity. Only one or a few neurons per culture could be properly stimulated and high-pressure frozen for subsequent ET analysis. The solution to such a problem is to utilize multielectrode arrays. Current technologies for studying neurons in arrays have been optimized for signal generation and detection, using either cultured neurons or brain slices. Several commercial systems are presently available that employ large-format arrays and are optimized mainly for slice work and perfusion. However, these devices are not suitable for EM-tomographic analysis because the samples cannot be rapidly frozen or even high-pressure frozen. Both the size and the thermal mass of these samples are too great to allow rapid cooling, a requirement for adequate sample preservation. To achieve cooling rates suitable for HPF, while retaining the capability to stimulate and pattern the neurons, we have designed and fabricated new devices specifically for this purpose (Fig. 6). With such tools, researchers will be able to recapitulate a variety
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Fig. 5 Stimulation-dependent changes in the saccular ribbon synapse, reproduced from Lenzi et al. (2002). Electron tomographic reconstructions of hair cell synapses exposed to 0-Ca2þ saline (A and C) or high-Kþ saline (B and D), which stimulates the synapse. (A and B) Single x–y planes through the reconstructed volumes are shown. The hair cell cytoplasm contains the synaptic body (SB) surrounded by synaptic vesicles, outlying cytoplasmic vesicles [arrows in (A)], coated vesicles [arrows in (B)], membrane-bounded cistern (c), mitochondria (m), and postsynaptic cell (p). (C and D) 3D structure of presynaptic organelles from the same reconstructions shown in (A) and (B). The SB (blue) was rendered as hollow and semitransparent and lies adjacent to the hair cell’s plasma membrane (red). SB-associated vesicles (yellow) surround the SB; outlying vesicles (green, thick long arrows) lie further out in the cytoplasm. Also visible are coated vesicles (gold, thin long arrows), cisternae (purple, thick short arrows), and mitochondria (blue, thin short arrows). (D) Regions of presynaptic density (pink) lie beneath the SB on the plasma membrane, which formed a tubular invagination (arrowhead). The flat surfaces of the mitochondria and the open face of the SBs denote the edges of the reconstruction. Synapses 1 and 11; scale bars ¼ 200 nm.
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Fig. 6 (A) Silicon chip for use in a high-pressure freezer. The silicon chip is designed so that the inner core of electrodes can be snapped oV and introduced directly into the cap of an HPF device. Left: overview of the chip designed for the HPF; CD, central disk for culturing neurons; IC, interconnect for direct hookup to both a multichannel systems or Panasonic MED-64 stimulation and recording amplifier system; SC, snapoV connections between the CD and the main chip body. Right: detail of overview; NW, neuron well; SNW, silicon nitride window for synaptic observation. (B) SEM image of a silicon EM chip with SU8 features for culturing neurons. (C) Detail of (B) showing the picket fence cell body containment posts.
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of synaptic phenomena that have previously been characterized by fluorescent light microscopy and electrophysiology, and then go on to observe the identical synapse using ET.
XI. Conclusions The use of ET to study synaptic structure and function holds tremendous promise, yet there are substantial hurdles that need to be overcome. The ultimate goal of such studies is to identify and ascertain the organization of membranes and supramolecular complexes in a resting synapse or in a synapse after treatment with well-defined stimulation protocols. Many of the relevant molecular complexes are of suYcient mass (>200 kDa) that their appearance will cause clear contrast in conventional EM. These include such important assemblies as the SNARE complex, NSF, and NMDA-PSD-95, the bassoon and piccolo complexes that order synaptic vesicles, polysomes, Ca2þ-release channels, microtubules, and associated transport vesicles. While the study of several systems, such as the neuromuscular junction and ribbon synapses following stimulation, has been achieved and important insights gained, such studies cannot currently be performed with primary cultured hippocampus neurons. This limitation can be overcome by employing microfabrication methods and multielectrode stimulation technologies, so that well-defined stimulation protocols can be achieved and their eVects rapidly analyzed by EM and ET. In the end, a knowledge of the distribution and organization of relevant macromolecular complexes, including membranes, will give us important new insights into the workings of the synapse. Acknowledgments We would like to thank our colleagues at the University of Colorado for helpful comments and advice, J. Richard McIntosh for patients and insight, and the Beckman Foundation for financial support.
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