Name: an optical assay for synaptic vesicle recycling in cultured neurons overexpressing presynaptic proteins
Uploaded: 2018-06-26T15:00:00
Description: Watch this Scientific Journal Video about An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins at JoVE.com

We thank Irmgard Weiss for expert technical assistance. This work was supported by the DFG via the Cluster of excellence for microscopy at the nanometer range and molecular physiology of the brain (CNMPB, B1-7, to T.D.).

No studies with live animals were conducted. Experiments involving euthanized animals to obtain cell cultures were approved by the local animal protection authorities (Tierschutzkommission der Universit&#228;tsmedizin G&#246;ttingen) under the approval number T10/30. The experiments were conducted with the approved protocols.
1. Primary Hippocampal Cell Culture
<ol>
	<li>Prepare the dissociated cell culture of the rat hippocampus on embryonic day 1916,<sup ...

<ol>
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There are three assays routinely used to study synaptic vesicle (SV) recycling. The first two include the use of a) fluorescent styryl dyes such as FM1-43 (which incorporate into membranes, are taken up into organelles during endocytosis, and are released after exocytosis); and b) fluorescently tagged recombinant SV proteins (which, upon overexpression, incorporate into the proteinaceous recycling machinery). If the attached fluorophores change their fluorescence depending on the pH, they can be used to monitor changes b...

Studying synaptic vesicle recycling is important in determining how presynaptic properties change, either during synaptic plasticity or in response to perturbation of synaptic function. Studying Synaptotagmin-1 (Syt1) antibody uptake provides one method of measuring the amount of SV recycling. Syt1 is a SV-associated protein that acts as a Ca2+ sensor and is necessary for exocytotic release of the neurotransmitter1,2. It is a transmembrane protein with a C-terminal cytoplasmic domain outside the SV and an N-terminal luminal domain inside the SV3. During exocytosis, the luminal domain of Syt1 becomes exposed to the external medium. To this external medium, we add antibodies directed against the cytoplasmic domain, which becomes internalized during endocytosis. These antibodies can be either pre-conjugated with fluorophores or immunostained with secondary antibodies4,5,6,7. The fluorescence intensity of the resulting immunosignal is proportional to the amount of SV recycling. This approach can be used to determine both constitutive and depolarization-induced SV recycling6,8.
Syt1 uptake assays can be performed after virus-mediated gene transfer to virtually all cells in the dish or after sparse transfection of a small number of cells. Our method combines the assay with sparse double transfection of primary hippocampal neurons using calcium phosphate9. We use a recombinant marker protein known to accumulate at presynapses, fluorescently tagged Synaptophysin, to locate presynaptic terminals and overexpress our protein of interest, Rogdi. This allows us to test whether or not Rogdi targets functional synapses and affects SV recycling. The gene encoding Rogdi was originally identified in a screen for Drosophila mutants characterized by impaired memory10. In humans, mutations in the Rogdi gene cause a rare and devastating disease called Kohlsch&#252;tter-T&#246;nz syndrome. Patients suffer from dental enamel malformations, pharmacoresistant epilepsy, and psychomotor delays; however, the subcellular localization of the gene product remained elusive11. Thus, the Syt1 uptake assay provided key evidence for the localization of GFP-tagged Rogdi at functional synapses9.
This uptake technique has several benefits. First, SV recycling can be observed both in real time by performing live imaging7,12, and after fixation6,9 by measuring the fluorescence intensity of the Syt1 fluorescence label. Additionally, several Syt1 antibody variants have been developed. There are untagged variants that can be labeled with a secondary antibody following a standard immunostaining protocol after fixation, and pre-conjugated variants with a fluorescence label already attached. Finally, antibody-based fluorescence is advantageous due to the large selection of commercially available secondary or conjugated dyes that can be used.
When fixing and immunostaining the neurons, it is also possible to stain for additional proteins and perform colocalization analysis. This can help determine where they are located in relation to recycling SVs. The intensity of the fluorescence label is the direct measure of the amount of SV recycling. In addition, the antibodies selectively label Syt1-containing structures, resulting in high specificity and little background fluorescence4. Different stimulation protocols can also be used, such as depolarization buffers or electric stimulation protocols9,12,13,14. However, basal SV recycling can be measured without stimulating the neuronal cultures15.
Our method specifically addresses Syt1 antibody uptake in double-transfected neurons with secondary antibody immunolabeling after fixation. However, we refer to all routinely used variants of the assay in our discussion to give viewers an opportunity to adapt the protocol to specific needs.

<table>
	<tbody>
		<tr>
			<td>B27</td>
			<td>Gibco</td>
			<td>17504-044</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A7030-50g</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma-Aldrich</td>
			<td>C3306-100g</td>
			<td></td>
		</tr>
		<tr>
			<td>CoolSNAP HQ2</td>
			<td>Photometrics</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>dH2O</td>
			<td>Invitrogen</td>
			<td>15230</td>
			<td></td>
		</tr>
		<tr>
			<td>DABCO</td>
			<td>Merck</td>
			<td>8.03456.0100</td>
			<td></td>
		</tr>
		<tr>
			<td>donkey anti mouse Alexa 647</td>
			<td>Jackson-Immunoresearch</td>
			<td>715605151</td>
			<td>antibody</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Invitrogen</td>
			<td>41966</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>Gibco</td>
			<td>14190</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf tubes</td>
			<td>Eppendorf</td>
			<td>30120094</td>
			<td></td>
		</tr>
		<tr>
			<td>multiwell 24 well</td>
			<td>Fisher Scientific</td>
			<td>087721H</td>
			<td></td>
		</tr>
		<tr>
			<td>tube (50 mL)</td>
			<td>Greiner Bio-One</td>
			<td>227261</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS superior</td>
			<td>BiochromAG</td>
			<td>S0615</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Merck</td>
			<td>1,083,421,000</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Invitrogen</td>
			<td>14170</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H4034-500g</td>
			<td></td>
		</tr>
		<tr>
			<td>Hera Cell 150 (Inkubator)</td>
			<td colspan="2">ThermoElectron Corporation</td>
			<td></td>
		</tr>
		<tr>
			<td>KCL</td>
			<td>Sigma-Aldrich</td>
			<td>P9541-500g</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamin</td>
			<td>Gibco</td>
			<td>25030</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Honeywell</td>
			<td>M0250-500g</td>
			<td></td>
		</tr>
		<tr>
			<td>microscope slides</td>
			<td>Fisher Scientific</td>
			<td>10144633CF</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsoft Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mowiol4-88</td>
			<td>Calbiochem</td>
			<td>475904</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>BioFroxx</td>
			<td>1394KG001</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>BioFroxx</td>
			<td>5155KG001</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurobasal</td>
			<td>Invitrogen</td>
			<td>21103049</td>
			<td></td>
		</tr>
		<tr>
			<td>OpenView Experiment Analysis Application</td>
			<td>Free software, see comments</td>
			<td></td>
			<td>written by Noam E. Ziv, Technion &#8211; Israel Institute of Technology, Haifa, Israel</td>
		</tr>
		<tr>
			<td>PBS (10x)</td>
			<td>Roche</td>
			<td>11666789001</td>
			<td></td>
		</tr>
		<tr>
			<td>Optimem</td>
			<td>Invitrogen</td>
			<td>31985</td>
			<td></td>
		</tr>
		<tr>
			<td>Penstrep</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>PFA</td>
			<td>Sigma</td>
			<td>P6148-1kg</td>
			<td></td>
		</tr>
		<tr>
			<td>safety hood</td>
			<td>ThermoElectron</td>
			<td></td>
			<td>Serial No. 40649111</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>neoFroxx</td>
			<td>1104kg001</td>
			<td></td>
		</tr>
		<tr>
			<td>Synaptotagmin1</td>
			<td>Synaptic Systems</td>
			<td>105311</td>
			<td>mouse monoclonal; clone 604.2</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Merck</td>
			<td>1,086,031,000</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex Genius 3&#160;</td>
			<td>IKA</td>
			<td>3340001</td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>GFL</td>
			<td>1004</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss Observer. Z1&#160;</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

an optical assay for synaptic vesicle recycling in cultured neurons overexpressing presynaptic proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

An expected result of this approach is locating approximately 50 double-transfected neurons per coverslip at a density of 50,000 neurons per well. The axon of each neuron is expected to show multiple hotspots of fluorescently-tagged Synaptophysin accumulation, indicating clusters ofSVs. At functional presynaptic sites, the recombinant Synaptophysin signal colocalizes with punctate Syt1 fluorescence. Using double transfection, either GFP-Rogdi as the protein of interest (<strong class="xfi...

Watch this Scientific Journal Video about An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins at JoVE.com

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...

The main advantage of this technique is that it is very simple, requires only standard equipment, and benefits from the primary advantage of antibodies, which is their specificity. The protocol was originally introduced by Michela Matteoli and colleagues, and has proven highly effective. I've used it to test whether a certain protein cordocti is localized to active synapses.Begin this procedure by culturing primary hippocampal cells on coverslips in a 24-well plate for three days, as described in the text protocol. Prewarm the reduced-serum medium, cell-culture medium, and distilled water to 37 degrees celsius in the water bath. Working under the laminar flow hood to ensure sterile working conditions, prepare the transfection mix in a 1.5-milliliter microcentrifuge tube.First, mix 7.5 microliters of two-molar calcium chloride with four micrograms of each endotoxin-free DNA. Add water to reach a total volume of 60 microliters in a 1.5-milliliter microcentrifuge tube. Now, add 60 microliters of transfection buffer to the mix.To obtain the best results, add the transfection buffer dropwise while shaking the DNA mixture gently on the vortex. Incubate the resulting transfection mix at room temperature for 20 minutes. Avoid shaking the tube during the incubation time.Under the laminar flow hood, remove the cell-culture medium from the wells using a 1, 000-microliter pipette, store it in a separate container, and immediately add 500 microliters of reduced-serum medium to each well. Store the conditioned medium in the incubator. Incubate the cells at 37 degrees celsius and 5%CO2 until the 20-minute incubation period for the transfection mix is over.Then, add 30 microliters of transfection mix to each well by pipetting several drops. Discard the residue at the bottom of the tube. After all the wells have been supplied with transfection mix gently shake the 24-well plate to ensure distribution of the transfection mix in the medium.Now incubate the wells for 60 minutes at 37 degrees celsius and 5%CO2. Following incubation, remove and discard the transfection mix, and wash the cells three times with cell-culture medium. The washing step is critical.Minimize the time that each well has no medium by removing and replacing well-by-well. Also, make sure to add the medium gently. To wash the cells, add one milliliter of cell-culture medium to each well, and incubate them for 30 seconds at room temperature.Then, remove 750 microliters of medium, and add the same amount of fresh medium. After repeating this process three times, remove and discard the cell-culture medium, and add 450 microliters of the conditioned medium well-by-well. Now let the neurons mature in the incubator at 37 degrees celsius and 5%CO2 to DIV 10.Prewarm 600 microliters of 1X depolarization buffer and 10 milliliters of cell-culture medium to 37 degrees celsius in the water bath. Add one microliter of mouse Anti-Syt1 antibody to the 1X depolarization buffer, and vortex for 10 seconds. After removing and discarding the cell-culture medium from the cells, add 200 microliters of the depolarization antibody mix to each well.Incubate the plate for five minutes at 37 degrees celsius and 5%CO2 in the incubator. Remove and discard the depolarization antibody mix, and wash the cells three times with cell-culture medium. Add one milliliter of cell-culture medium to each well, and incubate for 30 seconds at room temperature.Then, remove 750 microliters of medium, and add the same amount of fresh medium. Repeat this process three times. Next, remove and discard the cell-culture medium, and add 300 microliters of 4%PFA in 1X PBS.Finally, incubate for 20 minutes at four degrees celsius before washing three times for five minutes each with 1X PBS. Dilute the secondary fluorophore-coupled antibody in 200 microliters of blocking buffer for each well at a dilution of one to 1, 000. Remove and discard the 1X PBS from each well containing coverslips with the cultured neurons.Add 200 microliters of blocking buffer antibody mix to each well, and incubate for 60 minutes at room temperature. After incubation, wash the cells three times for five minutes with one milliliter of 1X PBS. Next, add a seven-microliter drop of embedding medium onto the microscope slide.Remove the coverslip from the 24-well plate by lifting it with a canula and grabbing it with forceps. Dip the coverslip into distilled water to remove the PBS, and dry it carefully by touching one edge to a soft tissue. Flip the coverslip onto the embedding-medium droplet so that the surface carrying the cells faces the microscope slide, thereby embedding the cells into the embedding medium.Leave the slides to dry under the hood for one to two hours, covering them to avoid light exposure. Store the dried slides in a microscope slide box at four degrees celsius. After immunolabeling the Syt1 antibody with a fluorescently-labeled secondary antibody, the extent of Syt1 uptake is finally quantified using the immunosignal.Punctate immunofluorescence indicates sites of Syt1 uptake in the field of view. These sites thus contain synaptic vesicles that recycled during the experiment. Synaptophysin-mOrange fluorescence indicates accumulations of synaptic vesicles in the axon of a transfected neuron.GFP-Rogdi fluorescence indicates accumulations of Rogdi in the axon of the transfected neuron. This part of the experiment reveals extensive colocalization of GFP-Rogdi and Synaptophysin-mOrange, suggesting that GFP-Rogdi accumulates at presynaptic sites within the axon. The overlay reveals which of the coaccumulations of GFP-Rogdi and Synaptophysin-mOrange colocalize with sites of Syt1 antibody uptake.The sites marked by arrows represent accumulations of recombinant Rogdi and recombinant synaptophysin at functional synapses. The protocol depicts a technically straightforward way to determine the extent of synaptic residue recycling. This method can help answer key questions in molecular and cellular science, such as where presynaptic nerve terminals allocated in an axon of a certain neuron and whether a recombinant or endogenous protein of interest accumulates at these terminals.After its development, this technique has been used in many ways to monitor synaptic vesicle recycling in presynaptic terminals. For example, it can be used to determine synaptic vesicle recycling in neurons after overexpression on adaptive proteins as well as in neurons obtained from animals or from stem-cell-derived neurons.

an-optical-assay-for-synaptic-vesicle-recycling-cultured-neurons

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 ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

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 ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

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 ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...