Name: preparation of newborn rat brain tissue for ultrastructural morphometric analysis of synaptic vesicle distribution at nerve terminals
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This manuscript was supported by NIH/NIGMS K08 123321 (to N.L.) and by funds from the Department of Anesthesiology at the University of Virginia. The Authors wish to thank Alev Erisir (Department of Biology, University of Virginia, Charlottesville, VA) for excellent training and technical assistance with TEM, and for her invaluable manuscript criticism. The Authors also thank the Advanced Electron Microscopy facility at the University of Virginia for technical assistance with specimen sectioning and post-staining.

All studies were approved by the Institutional Animal Care and Use Committee at the University of Virginia (Charlottesville, VA) and conducted in accordance with the National Institutes of Health guidelines.
1. Fixation by Vascular Perfusion
NOTE: A general description of the method for rat brain vascular perfusion has been already detailed in this journal13 and is beyond the scope of this protocol. However, the following steps...

<ol>
	<li>Lunardi, N., Oklopcic, A., Prillaman, M., Erisir, A., Jevtovic-Todorovic, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+exposure+to+general+anesthesia+disrupts+spatial+organization+of+presynaptic+vesicles+in+nerve+terminals+of+the+developing+subiculum.">Early exposure to general anesthesia disrupts spatial organization of presynaptic vesicles in nerve terminals of the developing subiculum.</a> Molecular Neurobiology. 52 (2), 942-951 (2015).</li><li>Lunardi, N., Ori, C., Erisir, A., Jevtovic-Todorovic, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=General+anesthesia+causes+long-lasting+disturbances+in+the+ultrastructural+properties+of+developing+synapses+in+young+rats.">General anesthesia causes long-lasting disturbances in the ultrastructural properties of developing synapses in young rats.</a> Neurotoxicity Research. 17 (2), 179-188 (2010).</li><li>Sanchez, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=General+anesthesia+causes+long-term+impairment+of+mitochondrial+morphogenesis+and+synaptic+transmission+in+developing+rat+brain.">General anesthesia causes long-term impairment of mitochondrial morphogenesis and synaptic transmission in developing rat brain.</a> Anesthesiology. 115 (5), 992-1002 (2011).</li><li>Boscolo, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+abolishment+of+anesthesia-induced+cognitive+impairment+by+timely+protection+of+mitochondria+in+the+developing+rat+brain:+the+importance+of+free+oxygen+radicals+and+mitochondria+integrity.">The abolishment of anesthesia-induced cognitive impairment by timely protection of mitochondria in the developing rat brain: the importance of free oxygen radicals and mitochondria integrity.</a> Neurobiology of Disease. 45 (3), 1031-1041 (2012).</li><li>Aghajanian, G. 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Handling of tissue sections during specimen preparation for TEM requires a considerable degree of finesse, concentration and patience. When using a micropipette to add and remove solutions, specimens can be sucked into the pipette tip by surface tension, so great care should be taken to avoid tissue damage by the pipette. Also, certain steps of the dehydration sequence can be as quick as 1 min, hence the operator needs to work swiftly to ensure that the next dehydration step is started on time and the specimen does not d...

We describe methods for the preparation of newborn rat brain tissue to obtain high-quality electron micrographs for in-depth morphometric analysis of synaptic vesicle spatial distribution at nerve terminals1,2. The high-contrast micrographs that can be obtained by processing specimens following these methods can also be used to study the detailed morphology of a number of cellular components and their dimensional structural relations3,4.
The transmission electron microscope (TEM) is a powerful tool to study the morphology of organelles and other cellular structures quantitatively. As of this decade, there are no other methods of investigation that can provide the same degree of resolution of lipid membranes and organelles without immuno-tagging, with the exception of cryofixation by high pressure freezing. However, freeze substitution techniques are not widely used, and normally require expensive equipment and long preparation times.
In order to take advantage of TEM&#39;s high resolving power, optimal specimen preparation is of paramount importance. The main goals of specimen preparation are to preserve tissue structure with minimum alteration from the living state, enhance specimen contrast, and stabilize the tissue against extraction of cellular components during processing and exposure to the electron beam. Numerous protocols for TEM tissue preparation have been introduced and perfected by several laboratories over the years. Many of them have focused on methods for optimal visualization of synaptic vesicles5,6,7,8,9,10,11. Among a number of well-established, gold standard methods currently in use, we chose procedures for chemical fixation, post fixation, en bloc staining, sequential dehydration, resin embedding and post staining that aim to preserve optimal tissue structure and achieve excellent image contrast. Of note, preservation of fine ultrastructure can be particularly challenging when working with newborn rat brain tissue. In fact, the central nervous system of very young animals is characterized by a higher water content than the adult brain, more prominent enlargement of extracellular spaces, and looser connections between cells12. This makes newborn rat brain tissue profoundly sensitive to changes in osmolarity, and exquisitely prone to artifactual shrinkage and/or swelling when processed through sequential solutions of different tonicity12. Therefore, our methods employed solutions for specimen processing that are of osmolarity as close as possible to that of rat newborn brain. Our goal was to obtain high-quality, high-resolution electron microscopy images for quantitative assessment of synaptic vesicle spatial distribution at nerve terminals. Specifically, we sought to measure the number of vesicles within the nerve terminal, the distance of synaptic vesicles from the pre-synaptic plasma membrane, the number of vesicles docked at the pre-synaptic membrane, the size of synaptic vesicles and the inter-vesicle distances1.
Satisfactory chemical fixation is a prerequisite for obtaining high-quality electron micrographs that can provide the morphological detail necessary to study synaptic vesicle morphology and spatial organization. Although several modes of fixation exist, fixation of brain tissue by vascular perfusion is decidedly superior to other methods. Since fixation via vascular perfusion begins immediately after the arrest of systemic circulation, it shortens the interval between deoxygenation of the brain tissue and cross-linking of proteins with fixatives, resulting in minimum alterations in cell structure. Furthermore, it accomplishes fast and uniform penetration, because of the rapid flow of the fixative from the vascular bed to the extracellular and cellular compartments12,13,14. Primary fixation with glutaraldehyde, followed by secondary fixation (post-fixation) with osmium tetroxide, yields excellent preservation of the fine structure15,16,17. A mixture of glutaraldehyde and paraformaldehyde has the additional advantage of more rapid penetration into the tissue12.
Since biological tissues are not sufficiently rigid to be cut into thin sections without the support of a resin matrix, they need to be embedded in a medium before thin sectioning. Water-immiscible epoxy resins are commonly used as an embedding medium in TEM. When this type of matrix is used, all specimen&#39;s free water must be replaced with an organic solvent before resin infiltration. Water is removed by passing the specimen through a series of solutions of ascending concentrations of ethanol and/or acetone12. In this protocol, specimens are first flat embedded between flexible aclar sheets, then embedded in a capsule. The final result is tissue situated at the tip of a cylindrical resin block, which has the ideal geometry to be least affected by vibrations arising during microtome sectioning.
Staining with heavy metals to enhance endogenous tissue contrast is another important aspect of specimen preparation. Image contrast in TEM is due to electron scattering by the atoms in the tissue. However, biological materials consist largely of low atomic weight molecules (i.e., carbon, hydrogen, oxygen and nitrogen). Therefore, the generation of sufficient scattering contrast requires the incorporation of high atomic weight atoms into the cellular components of the tissue. This is achieved through staining of the specimen with heavy metals12,18,19. Osmium tetroxide, uranium and lead, which bind strongly to lipids, are the most common heavy metals used as electron stains.
Osmium (atomic number 76) is one of the densest metals in existence. It is both a fixative and a stain, although its primary role in TEM is as a reliable fixative12. Among various fixation protocols in use, the method of double fixation with glutaraldehyde followed by osmium is the most effective in reducing the extraction of cell constituents during specimen preparation. These two fixatives are used to stabilize the maximum number of different types of molecules, especially proteins and lipids, and result in superior preservation of tissue ultrastructure12,14,15,16,17.
Uranium (atomic number 92) is the heaviest metal used as electron stain, most typically in the form of uranyl acetate. Similarly to osmium, it acts as a stain and fixative, although its primary role in TEM is as a stain20,21. Nucleic acid-containing and membranous structures are strongly and preferentially stained with uranyl salts in aldehyde-fixed tissues22,23. Treatment of tissues with uranyl acetate after osmication and before dehydration results in stabilization of membranous and nucleic acid-containing structures, as well as enhanced contrast, and permits identification of some structural details that would not be easily detected in specimens stained with osmium alone12,24,25. It is thought that uranyl acetate may stabilize the fine structure by combining with reduced osmium that has been deposited on lipid membranes during osmication24. Maximum contrast is achieved when uranyl acetate is applied before embedding and as a post-stain in thin sections12.
Lead (atomic number 82) is the most common stain used for TEM and is mainly employed for post-staining of thin sections. Lead salts have high electron opacity and show affinity for a wide range of cellular structures, including membranes, nuclear and cytoplasmic proteins, nucleic acids and glycogen26,27. When the double staining method is employed (i.e., staining with uranyl acetate is followed by treatment with lead), the latter acts as a developer of uranyl acetate staining. For instance, lead post-staining of chromatin fixed with glutaraldehyde increases uranyl acetate uptake by a factor of three28,29,30,31,32. Lead also enhances the staining imparted by other metals such as osmium. It is thought that lead salts stain the membranes of osmium-fixed tissues by attaching to the polar group of phosphatides in the presence of reduced osmium33. A potential disadvantage of staining with both uranyl acetate and lead, especially for prolonged durations, is that many different structural elements are stained equally and non-specifically, and thus may not be easily distinguished from one another12.
The recent introduction of alternative light sources, such as in optical super-resolution photo-activated localization microscopy, has significantly improved light microscopy resolution34. However, because light microscopy relies on histochemical and immune-cytochemical methods to visualize individually-labelled proteins or enzymes, the power of TEM to display all structural elements at once remains unsurpassed for in-depth study of the morphology and dimensional relationships of tissue structures. In particular, no other technique can provide the morphological detail necessary to perform morphometric analysis of synaptic vesicle distribution at pre-synaptic nerve boutons. Nevertheless, it is important to note that electron micrographs capture the structure of the tissue after the organism dies, and therefore they cannot provide information regarding the dynamics of pre-synaptic vesicle trafficking and exocytosis. Hence, other tools, such as FM dye-live imaging and patch-clamp electrophysiology, should be considered when the main objective is to study dynamic and/or functional aspects of synaptic vesicle trafficking and exocytosis.

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			<td>4% Osmium tetroxide</td>
			<td>Electron Microscopy Sciences</td>
			<td>19170</td>
			<td>acqueous</td>
		</tr>
		<tr>
			<td>50% Glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16310</td>
			<td>EM grade, acqueous</td>
		</tr>
		<tr>
			<td>Aclar 33 C embedding film</td>
			<td>Electron Microscopy Sciences</td>
			<td>50425-25</td>
			<td>7.8 mil thickness, size 8&#34;x10&#34;</td>
		</tr>
		<tr>
			<td>BEEM capsule holder</td>
			<td>Electron Microscopy Sciences</td>
			<td>69916</td>
			<td>holds size &#34;00&#34; capsules</td>
		</tr>
		<tr>
			<td>BEEM embedding capsules</td>
			<td>Electron Microscopy Sciences</td>
			<td>70021</td>
			<td>size &#34;00&#34;, flat</td>
		</tr>
		<tr>
			<td>Butler block trimmer</td>
			<td>Electron Microscopy Sciences</td>
			<td>69945-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Camel hair paint brush</td>
			<td>Electron Microscopy Sciences</td>
			<td>65576-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Disc punch</td>
			<td>Electron Microscopy Sciences</td>
			<td>77850-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Embed 812 kit</td>
			<td>Electron Microscopy Sciences</td>
			<td>14120</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>17600</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead citrate</td>
			<td>Electron Microscopy Sciences</td>
			<td>17800</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead nitrate</td>
			<td>Electron microscopy Sciences</td>
			<td>17900</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica UC7 ultracut microtome</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micro scale</td>
			<td>Electron Microscopy Sciences</td>
			<td>62091-23</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>19208</td>
			<td>EM grade, granular</td>
		</tr>
		<tr>
			<td>Precision Thelco laboratory oven</td>
			<td>Thelco</td>
			<td>51221159</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Sigma-Aldricht</td>
			<td>S2002</td>
			<td></td>
		</tr>
		<tr>
			<td>StatMark pen</td>
			<td>Electron Microscopy Sciences</td>
			<td>72109-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Tyrode solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>11760-05</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>22400</td>
			<td>powder</td>
		</tr>
	</tbody>
</table>

preparation of newborn rat brain tissue for ultrastructural morphometric analysis of synaptic vesicle distribution at nerve terminals

Our laboratory and many others have exploited the high resolving power of transmission electron microscopy to study the morphology and spatial organization of synaptic vesicles. In order to obtain high-quality electron micrographs that can yield the degree of morphological detail necessary for quantitative analysis of pre-synaptic vesicle distribution, optimal specimen preparation is critical. Chemical fixation is the first step in the process of specimen preparation, and of utmost importance to preserve fine ultrastructure. Vascular fixation with a glutaraldehyde-formaldehyde solution, followed by treatment of vibratome-sectioned specimens with osmium tetroxide, stabilizes the maximum number of molecules, especially proteins and lipids, and results in superior conservation of ultrastructure. Tissue is then processed with counterstaining, sequential dehydration and resin-embedding. En bloc staining with uranyl acetate (i.e., staining of vibratome-sectioned tissue before resin embedding) enhances endogenous contrast and stabilizes cell components against extraction during specimen processing. Contrast can be further increased by applying uranyl acetate as a post-stain on ultrathin sections. Double-staining of ultrathin sections with lead citrate after uranyl acetate treatment also improves image resolution, by intensifying electron-opacity of nucleic acid-containing structures through selective binding of lead to uranyl acetate. Transmission electron microscopy is a powerful tool for characterization of the morphological details of synaptic vesicles and quantification of their size and spatial organization in the terminal bouton. However, because it uses fixed tissue, transmission electron microscopy can only provide indirect information regarding living or evolving processes. Therefore, other techniques should be considered when the main objective is to study dynamic or functional aspects of synaptic vesicle trafficking and exocytosis.

General criteria that are mostly accepted as indicative of satisfactory or defective preservation of specimen for TEM have been established. These criteria are exemplified in four selected electron micrographs (two examples of optimal preparation, two examples of defective preparation) that were obtained by treating young rat brain tissue following the methods described in this protocol.
In general, a good-quality electron micrograph appears as an orderly, distinct and overall grayish image. I...

Watch this Scientific Journal Video about Preparation of Newborn Rat Brain Tissue for Ultrastructural Morphometric Analysis of Synaptic Vesicle Distribution at Nerve Terminals at JoVE.com

Preparation of Newborn Rat Brain Tissue for Ultrastructural Morphometric Analysis of Synaptic Vesicle Distribution at Nerve Terminals

We describe procedures for processing newborn rat brain tissue to obtain high-resolution electron micrographs for morphometric analysis of synaptic vesicle distribution at nerve terminals. The micrographs obtained with these methods can also be used to study the morphology of a number of other cellular components and their dimensional structural relationships.

Our laboratory and many others have exploited the high resolving power of transmission electron microscopy to study the morphology and spatial ...

Among several existing protocols for tissue preparation for transmission electron microscopy, we selected methods that yield optimal structural preservation and high resolution micrographs to perform morphometric analysis of synaptic vesicles distribution at presynaptic terminals. The high contrast images obtained with these methods allow to quantify parameters such as distance of synaptic vesicles from the presynaptic membrane number of vesicles docked at the presynaptic membrane and inter vesicle distances. All of these parameters are indicative of synaptic function.Since alterations in the spatial organization of synaptic vesicles may be associated with the disruption of synaptic function, these methods have been used to examine the effects of general anesthetics on synaptic development. These methods could also be used to examine the effects of any drug or treatment on the detailed morphology of synapses to provide insight into their function. Begin by preparing osmium tetroxide for post fixation of rat brain sections previously fixed with paraformaldehyde and glutaraldehyde via vascular perfusion.Sections become rigid after treatment with osmium tetroxide. Therefore tissue handling requires attention from then on. Also, before adding osmium, make sure that your sections are flattened as any fold will likely result in tissue fracture.Open one five milliliter glass ampule of 4%osmium tetroxide and water and drop it inside a brown glass bottle. Add five milliliters of 0.2 molar phosphate buffer and then add an additional 10 milliliters of 0.1 molar phosphate buffer to achieve a 1%final solution. Use a 20 milliliter syringe fitted with a long needle to draw up the 1%osmium tetroxide and then add the osmium tetroxide to a brown glass jar covered with aluminum foil.After making sure that the specimen has been unfolded and flattened, add 1%osmium tetroxide and 0.1 molar PB to the specimen vial. Incubate the specimen in osmium tetroxide for one hour at room temperature. Extract the osmium tetroxide with a micropipet after incubation is finished and rinse the sections as described in the protocol.To prepare uranyl acetate, place a 200 milliliter volumetric glass flask containing 100 milliliters of 70%ethanol on a stirring plate. Add four grams of uranyl acetate to the flask, wrap an aluminum foil to prevent precipitation by light and stir continuously. Dehydrate the sections in 50%ethanol.Stain with the prepared the 4%uranyl acetate for one hour and follow with serial dehydration and rinsing as described in the manuscript. Remove the syringe plunger from a 60 milliliter gavage syringe and cap it with a safety needle. Position the syringe with the open side up and add each ingredient of the resin mix one by one.Finish by adding 525 microliters of DMP30 with a pipet. Moving the plunger of the pipet very slowly as DMP is very viscous. Put the plunger back in the syringe.Invert the syringe a few times to mix the contents. The color will change from yellow to amber. Evacuate the air inside the syringe.Then place it on a rocker with continuous shaking for at least 30 minutes. Add one volume of epoxy resin and one volume of acetone to a scintillation vial and shake to mix. Apply this one to one resin to acetone mixture on the tissue section after the last acetone rinse keeping in covered to avoid acetone evaporation.Remove the one to one mixture of resin and acetone after two to four hours and replace it with full resin and incubate for four hours. Put all resin waste in a collection container under the hood to polymerize and to be disposed of later. Cut two rectangular pieces of clear polychlorotrifluoroethylene film and wipe them with 70%ethanol.Trim them so that there is at least 1.5 centimeter of tissue-free plastic on every side of the section, making sure that they have the same width but the height of the sheet on top is approximately 2/3 of the bottom sheet. Slowly tilt the vial and use a fine camel paintbrush and a flexible micro-spatula to very gently lift the specimen from the bottom. Then carefully move the section along the vial walls and transfer onto the bottom PCTFE sheet.Remove excess resin with a micro-spatula and gently cover the specimen with the top sheet. Gently push out any trapped air bubbles without applying direct pressure onto the tissue section and wipe out the excess resin. Use a solvent resistant pen to label the sheets and place in an oven at 60 degrees celsius for two to three days to polymerize.Gently pull apart the sandwiched PCTFE sheets to open them. Use a solvent resistant pen to mark the side of the sheet that contains the adhering section marking it near the tissue. Use a disc punch to obtain a circle sample of the section making sure that the pen mark is part of the punched out tissue.The pen mark will gleam when light is shined on the specimen. Place the cap of an embedding capsule side up on a capsule holder. Place a drop of resin on the cap and insert the punched disk inside the cap with the tissue section facing up.Insert a capsule into the cap using a gentle corkscrew movement. Use fine tweezers to roll and lower a printed two centimeter long piece of paper label into the capsule, making sure it fits to the curvature of the side walls of the capsule. Pour the embedding resin inside the capsule, fill up to edge of the capsule and use a pointed wooden stick to push the tissue specimen down to the bottom.Place the capsule in the oven at 60 degree celsius for two to three days for polymerization. And then continue with sectioning and staining as described in the protocol. Electron micrograph of satisfactory preservation of neuronal cell structure shows layered cell membranes without breaks.The cytoplasm is finally granular and without empty spaces. Mitochondria are neither swollen nor shrunken with a conserved outer double membrane and intact internal cristae. Optimal preservation of neuronal ultrastructure and fine morphological detail is essential in order to visualize synaptic vesicles and measure their distance from the presynaptic membrane.Synaptic vesicles should be distinct and lined by an unbroken single membrane. In order to facilitate morphometric analysis of synaptic vesicle distribution, presynaptic and postsynaptic membranes should be parallel in their continuity preserved. Electron micrograph of defective preservation of tissue structure shows the distortion and breakage of neuronal cell membranes and the presence of markedly enlarged extracellular spaces.Mitochondria appear distended and have swollen cristae. In another example of defective tissue structure preservation, there were large white empty spaces within the cytoplasm in place of finely granular cytoplasmic substance with enlarged extracellular spaces. An artificial membrane as forall likely resulted from mobilization of lipids during fixation with glutaraldehyde.When working young brain tissue use 1 molar phosphate buffer as a solvent for all your solution so that you keep tonicity as close as possible to that of newborn rat brain and minimize artefactual tissue shrinkage and or swelling. The micrographs obtained with these methods can be used to study the detailed morphology and dimensional structural relations of a number of cellular components other than synaptic vesicles. For example, mitochondria, golgi apparatus and nuclear chromatin.Several steps in this protocol require chemicals that can be toxic. Always work under a fume hood and wear personal protective equipment when handling aldehydes, osmium, uranium and lead compounds.

preparation-newborn-rat-brain-tissue-for-ultrastructural-morphometric

Research

JoVE Journal

Neuroscience

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Preparation of Oligomeric &beta;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). &beta;-amyloid (A&beta;), a peptide produced in high amounts in AD, is known to reduce Long-Term Potentiation (LTP), a cellular correlate of learning and memory. Indeed, LTP impairment caused by A&beta; is a useful experimental paradigm for studying synaptic dysfunctions in AD models and for screening drugs capable of mitigating or reverting such synaptic impairments. Studies have shown that A&beta; produces the LTP disruption preferentially via its oligomeric form. Here we provide a detailed protocol for impairing LTP by perfusion of oligomerized synthetic A&beta;1-42 peptide onto acute hippocampal slices. In this video, we outline a step-by-step procedure for the preparation of oligomeric A&beta;1-42. Then, we follow an individual experiment in which LTP is reduced in hippocampal slices exposed to oligomerized A&beta;1-42 compared to slices in a control experiment where no A&beta;1-42 exposure had occurred.

Preparation of Oligomeric &amp;#946;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

One feature of Alzheimer's Disease is the elevation of A&beta;1-42 peptide. Here we provide a protocol for preparing synthetic A&beta;1-42 oligomers, which impairs hippocampal Long-Term Potentiation, a cellular correlate of memory. This procedure is useful for investigating mechanisms of A&beta;-induced pathology and drug screening.

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Morphometric Analyses of Retinal Sections

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the retinal ganglion cell layer (RGCL) in rat models with N-methyl-D-aspartate (NMDA)&#x2013;induced excitotoxicity1, retinal ischemia-reperfusion injury2 and glaucoma3. Reduction of INL and inner plexiform layer (IPL) thicknesses were reversed with citicoline treatment in rats' eyes subjected to kainic acid-mediated glutamate excitotoxicity4. Alteration of RGC density and soma sizes were observed with different drug treatments in eyes with elevated intraocular pressure3,5,6. Therefore, having objective methods of analyzing the retinal morphometries may be of great significance in evaluating retinal pathologies and the effectiveness of therapeutic strategies.
The retinal structure is multi-layers and several different kinds of neurons exist in the retina. The morphometric parameters of retina such as cell number, cell size and thickness of different layers are more complex than the cell culture system. Early on, these parameters can be detected using other commercial imaging software. The values are normally of relative value, and changing to the precise value may need further accurate calculation. Also, the tracing of the cell size and morphology may not be accurate and sensitive enough for statistic analysis, especially in the chronic glaucoma model. The measurements used in this protocol provided a more precise and easy way. And the absolute length of the line and size of the cell can be reported directly and easy to be copied to other files. For example, we traced the margin of the inner and outer most nuclei in the INL and formed a line then using the software to draw a 90 degree angle to measure the thickness. While without the help of the software, the line maybe oblique and the changing of retinal thickness may not be repeatable among individual observers. In addition, the number and density of RGCs can also be quantified. This protocol successfully decreases the variability in quantitating features of the retina, increases the sensitivity in detecting minimal changes.
 	This video will demonstrate three types of morphometric analyses of the retinal sections. They include measuring the INL thickness, quantifying the number of RGCs and measuring the sizes of RGCs in absolute value. These three analyses are carried out with Stereo Investigator (MBF Bioscience &mdash; MicroBrightField, Inc.). The technique can offer a simple but scientific platform for morphometric analyses.

This video demonstrates three types of morphometric analyses of the retina, which include measuring the inner nuclear layer thickness, quantifying the number of retinal ganglion cells (RGCs) and measuring the sizes of RGCs. The technique can offer a simple but scientific platform for morphometric analyses.

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the ...

High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs). Physiological data have indicated that glutamate receptors at RGCs are expressed not only in postsynaptic but also in perisynaptic or extrasynaptic membrane compartments. However, precise anatomical locations for glutamate receptors at RGC synapses have not been determined. Although a high-resolution quantitative analysis of glutamate receptors at central synapses is widely employed, this approach has had only limited success in the retina. We developed a postembedding immunogold method for analysis of membrane receptors, making it possible to estimate the number, density and variability of these receptors at retinal ribbon synapses. Here we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of retinal fixation, 2) freeze-substitution, 3) postembedding immunogold electron microscope (EM) immunocytochemistry and, 4) quantitative visualization of glutamate receptors at ribbon synapses.

The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA ...

Peering at Brain Polysomes with Atomic Force Microscopy

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, formed by ribosomes, mRNAs, several proteins and non-coding RNAs, represent integrated platforms where translational controls take place. However, while the ribosome has been widely studied, the organization of polysomes is still lacking comprehensive understanding. Thus much effort is required in order to elucidate polysome organization and any novel mechanism of translational control that may be embedded. Atomic force microscopy (AFM) is a type of scanning probe microscopy that allows the acquisition of 3D images at nanoscale resolution. Compared to electron microscopy (EM) techniques, one of the main advantages of AFM is that it can acquire thousands of images both in air and in solution, enabling the sample to be maintained under near physiological conditions without any need for staining and fixing procedures. Here, a detailed protocol for the accurate purification of polysomes from mouse brain and their deposition on mica substrates is described. This protocol enables polysome imaging in air and liquid with AFM and their reconstruction as three-dimensional objects. Complementary to cryo-electron microscopy (cryo-EM), the proposed method can be conveniently used for systematically analyzing polysomes and studying their organization.

While ribosome structure has been extensively characterized, the organization of polysomes is still understudied. To overcome this lack of knowledge, we present here a detailed preparation protocol for accurate imaging of mammalian polysomes by atomic force microscopy (AFM) in air and liquid.

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, ...

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at the synaptic level. The Active Zone (AZ) is the region at the presynaptic membrane that mediates neurotransmitter release and is composed of a dense collection of scaffold proteins. AZs of Drosophila melanogaster (Drosophila) photoreceptors undergo molecular remodeling after prolonged exposure to natural ambient light. Thus the level of neuronal activity can rearrange the molecular composition of the AZ and contribute to the regulation of the functional output.
Starting from the light exposure set-up preparation to the immunohistochemistry, this protocol details how to quantify the number, the spatial distribution, and the delocalization level of synaptic molecules at AZs in Drosophila photoreceptors. Using image analysis software, clusters of the GFP-fused AZ component Bruchpilot were identified for each R8 photoreceptor (R8) axon terminal. Detected Bruchpilot spots were automatically assigned to individual R8 axons. To calculate the distribution of spot frequency along the axon, we implemented a customized software plugin. Each axon&#39;s start-point and end-point were manually defined and the position of each Bruchpilot spot was projected onto the connecting line between start and end-point. Besides the number of Bruchpilot clusters, we also quantified the delocalization level of Bruchpilot-GFP within the clusters. These measurements reflect in detail the spatially resolved synaptic dynamics in a single neuron under different environmental conditions to stimuli.

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

Here we show how to quantify the number and spatial distribution of synaptic active zones in Drosophila melanogaster&#160;photoreceptors, highlighted with a genetically encoded molecular marker, and their modulation after prolonged exposure to light.

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.