Name: measuring synaptic vesicle endocytosis in cultured hippocampal neurons
Uploaded: 2017-09-04T12:30:00
Description: Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

Nous remercions m. Zhu Yong-Ling prévoyant la fourniture VAMP2-phluorin de construction synaptophysine-pHluorin2x et Dr. James E. Rothman. Nous remercions le Dr Susan Cheng et Virginia Crocker de NINDS Electron Microscopy Facility pour leur soutien technique et aide. Ce travail a été soutenu par le National Institute of Neurological Disorders et programme de recherche intra-muros accident vasculaire cérébral aux États-Unis et une subvention de la KRIBB Research Initiative (programme coréen biomédicale scientifique Fellowship), Korea Research Institute of Bioscience et biotechnologie, République de Corée.

Remarque : le protocole suivant décrit les méthodes d’imagerie pHluorin et les méthodes de EM utilisés dans des neurones hippocampal cultivés. pHluorin moniteurs Vésicule synaptique protéine absorption dans les cellules vivantes et EM détecte l’absorption des vésicules synaptiques et modifications ultrastructurales.
 soins aux animaux et la procédure suivie NIH guidelines et ont été approuvés par le Comité de l’utilisation et le NIH animalerie.
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Nous démontrons ici deux méthodes pour surveiller l’endocytose de la vésicule synaptique. Dans la première méthode, nous avons suivi pHluorin fondu avec une protéine de la vésicule synaptique dans les neurones transfectées et stimulé par la suite électriquement. Deuxièmement, nous avons utilisé l’imagerie EM d’absorption HRP car induite par le KCl. Nous avons utilisé différents stimuli pour deux raisons. Tout d’abord, application de potassium élevé induit une dépolarisation de tous les neurones d...

Neurotransmetteurs sont stockées dans des vésicules synaptiques et libérées par exocytose. La membrane des vésicules synaptiques et la protéine sont internalisés par endocytose, puis réutilisés dans la prochaine série d’exocytose. L’endocytose des vésicules synaptiques est important pour le maintien des piscines de la vésicule synaptique et supprime la saillie vésicules de la membrane plasmique. La protéine fluorescente verte sensibles au pH pHluorin, qui est trempé dans des conditions acides et dequenched à pH neutre, a été utilisé pour mesurer le temps d’endocytose cours à vivent les cellules1,2,3. La protéine pHluorin est généralement attachée au côté lumière des protéines des vésicules synaptiques, comme synaptophysine ou VAMP2/synaptobrévine. Au repos, pHluorin est trempé dans le lumen de pH 5,5 des vésicules synaptiques. La fusion des vésicules de la membrane plasmique expose la lumière vésiculaire à la solution extracellulaire où le pH est ~ 7,3, résultant en une augmentation pHluorin de fluorescence. Après exocytose, l’augmentation de la fluorescence décroît, en raison de l’endocytose des protéines Vésicule synaptique, suivie par une vésicule ré-acidification dans ces vésicules récupérés. Bien que la désintégration reflète l’endocytose et vésiculaire ré-acidification, il reflète surtout endocytose, parce que la réacidification est plus rapide que l’endocytose dans la plupart des conditions1,4. La constante de temps de ré-acidification est 3-4 s ou moins5,6, qui est généralement plus rapide que les 10 s ou plus requis pour la vésicule endocytose4,5. Si les expériences sont nécessaires pour distinguer l’endocytose de ré-acidification, les expériences trempe acides avec un pH de 5.5 à l’aide de la solution de (MES) acide 4-Morpholineethanesulfonic (25 mM) peuvent être utilisés pour déterminer si les protéines des vésicules synaptiques sont récupérés de la membrane plasmique par endocytose1,3,4. Ainsi, l’augmentation d’intensité de fluorescence pHluorin fait apparaître un solde d’exo - et l’endocytose et la diminution après une stimulation nerveuse reflète précisément l’endocytose.
pHluorin imagerie peut être utilisé non seulement pour mesurer l’évolution temporelle de l’endocytose, mais aussi la taille des vésicules synaptiques piscines7,8, et la probabilité de libération évoquée et spontanée de sortie9. Plusieurs facteurs et les protéines impliquées dans la régulation de l’endocytose, tels que le calcium, les protéines solubles NSF-attachment protein receptor (SNARE), factor(BDNF) neurotrophique dérivé du cerveau et de la calcineurine sont identifiés au moyen d’imagerie pHluorin1 , 2 , 10 , 11 , 12 , 13 , 14 , 15 , 16. en outre, la libération du neurotransmetteur pouvait être détectée dans les neurones pas primaires mais dans les cellules de neuroblastome avec TIRFM17. Récemment, pHTomato, dsRed, mOrange et variantes de pHluorin ont été développés pour le suivi des enregistrements simultanés de facteurs multiples dans une seule synapse18,19. Par exemple, pHTomato a été fusionné avec synaptophysine et utilisé avec un indicateur de calcium génétiquement encodé (GCaMP5K) pour surveiller la fusion des vésicules présynaptiques et l’influx de Ca2 + dans le compartiment post-synaptique20. Par conséquent, pHluorin attaché aux protéines synaptiques fournit une méthode utile pour analyser la relation entre l’endocytose et l’exocytose.
EM est une autre méthode couramment utilisée pour étudier l’endocytose, en raison de la haute résolution spatiale qui montre les changements ultrastructuraux au cours de l’endocytose. Deux domaines généraux sont la possibilité de visualiser des changements pathologiques dans les cellules neuronales21 et de suivre les vésicules protéines22. En particulier, l’observation d’absorption de la vésicule synaptique, courbure de membrane enduite de clathrine dans la zone de periactive, et les structures endosomale sont possibles avec EM3,23,24,25 ,26,27,28. Tandis qu’EM implique des artefacts potentiels, tels que des malformations induites par le fixateur, qui peuvent affecter l’endocytose, et analyse des données est de main-d'oeuvre, que la résolution offre la possibilité de visualiser la structure cellulaire attrayante. Les problèmes potentiels de fixateur et la limitation en résolution temporelle EM peuvent être surmontés en haute pression, congélation, fournissant une méthode rapide et non chimiques de stabiliser les structures délicates présents au cours de l’endocytose27.

<table>
	<tbody>
		<tr>
			<td>Lipofectamine LTX with Plus</td>
			<td>Thermo Fisher</td>
			<td>15338-100</td>
			<td>Transfection of plasmid DNA including synaptophysin or VAMP2-pHluorin</td>
		</tr>
		<tr>
			<td>neurobasal medium</td>
			<td>Thermo Fisher</td>
			<td>21103-049</td>
			<td>Growth medium for neuron, Warm up to 37&#176;C before use</td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Thermo Fisher</td>
			<td>17504-044</td>
			<td>Gradient for neuronal differentiation</td>
		</tr>
		<tr>
			<td>Glutamax</td>
			<td>Thermo Fisher</td>
			<td>35050-061</td>
			<td>Gradient for neuronal culture</td>
		</tr>
		<tr>
			<td>Poly-D-Lysine coated coverslip</td>
			<td>Neuvitro</td>
			<td>GG-25-pdl</td>
			<td>Substrate for neuronal growth and imaging of pHluorin</td>
		</tr>
		<tr>
			<td>Trypsin XI from bovine pancrease</td>
			<td>Sigma</td>
			<td>T1005</td>
			<td>Neuronal culture-digest hippocampal tissues</td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancreas</td>
			<td>Sigma</td>
			<td>D5025</td>
			<td>Neuronal culture-inhibits viscous cell suspension</td>
		</tr>
		<tr>
			<td>pulse stimulator</td>
			<td>A-M systems</td>
			<td>model 2100</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>Slotted bath with field stimulation</td>
			<td>Warner Instruments</td>
			<td>RC-21BRFS</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>stimulus isolation unit</td>
			<td>Warner Instruments</td>
			<td>SIU102</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>lubricant</td>
			<td>Dow corning</td>
			<td>111</td>
			<td>pHluorin imaging-seal with coverslip and imaging chamber, avoid leak from chamber</td>
		</tr>
		<tr>
			<td>AP5</td>
			<td>Tocris</td>
			<td>3693</td>
			<td>Gradient for normal saline, selective NMDA receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td>
		</tr>
		<tr>
			<td>CNQX</td>
			<td>Tocris</td>
			<td>190</td>
			<td>Gradient for normal saline, competitive AMPA/kainate receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td>
		</tr>
		<tr>
			<td>Illuminator</td>
			<td>Nikon</td>
			<td>C-HGFI</td>
			<td>Metal halide light source for pHluorin</td>
		</tr>
		<tr>
			<td>EMCCD camera</td>
			<td>Andor</td>
			<td>iXon3</td>
			<td>pHluorin imaging, detect pHluorin fluorescence intensity</td>
		</tr>
		<tr>
			<td>Inverted microscopy</td>
			<td>Nikon</td>
			<td>Ti-E</td>
			<td>Imaging for synaptophysin or VAMP2 pHluorin transfected cells</td>
		</tr>
		<tr>
			<td>NIS-Elements AR</td>
			<td>Nikon</td>
			<td>NIS-Elements Advanced Research</td>
			<td>Software for imaging acquisition and analysis</td>
		</tr>
		<tr>
			<td>Igor Pro</td>
			<td>WaveMetrics</td>
			<td>Igor pro</td>
			<td>Software for imaging analysis and data presentation</td>
		</tr>
		<tr>
			<td>imaging chamber</td>
			<td>Warner Instruments</td>
			<td>RC21B</td>
			<td>pHluorin imaging, apply field stimulation on living cells</td>
		</tr>
		<tr>
			<td>poly-l-lysine</td>
			<td>Sigma</td>
			<td>P4832</td>
			<td>Electron microscopy, substrate for neuronal growth, apply on multiwell plate for 1 h at room temperature then wash with sterilized water 3 times</td>
		</tr>
		<tr>
			<td>Horseradish peroxidase(HRP)</td>
			<td>Sigma</td>
			<td>P6782</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles by catalyzing DAB in presence hydrogen peroxide, final concentration is 5 mg/mL in normal saline, make fresh before use</td>
		</tr>
		<tr>
			<td>Na cacodylate</td>
			<td>Electron Microscopy Sciences</td>
			<td>12300</td>
			<td>Electron microscopy, buffer for fixatives and washing, final concentration is 0.1 N</td>
		</tr>
		<tr>
			<td>3,3&#8242;-Diaminobenzidine(DAB)</td>
			<td>Sigma</td>
			<td>D8001</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles, substrate for HRP, final concentration is 0.5 mg/mL in DDW and filtered, make fresh before use</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide solution</td>
			<td>Sigma</td>
			<td>H1009</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles by inducing HRP-DAB reaction, final concentration is 0.3% in DDW, make fresh before use</td>
		</tr>
		<tr>
			<td>glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16365</td>
			<td>Electron microscopy, fixatives, final concentration is 4% in Na-cacodylate buffer, make fresh before use, shake well before to use</td>
		</tr>
		<tr>
			<td>TEM</td>
			<td>JEOL</td>
			<td>200CX</td>
			<td>Electron microscopy, imaging of endocytosed vesicles and ultrastructural changes</td>
		</tr>
		<tr>
			<td>CCD digital camera</td>
			<td>AMT</td>
			<td>XR-100</td>
			<td>Electron microscopy, capturing images</td>
		</tr>
		<tr>
			<td>Lead citrate</td>
			<td>Leica microsystems</td>
			<td>16707235</td>
			<td>Electron microscopy, grid staining</td>
		</tr>
	</tbody>
</table>

measuring synaptic vesicle endocytosis in cultured hippocampal neurons

Au cours de l’endocytose, fusion des vésicules synaptiques sont récupérées dans les terminaisons nerveuses, permettant de recyclage de la vésicule et donc l’entretien de la transmission synaptique au moment du tir nerf répétitives. Avec facultés affaiblies endocytose dans certaines pathologies conduit à une diminution des fonctions synaptiques de force et le cerveau. Nous décrivons ici les méthodes utilisées pour mesurer l’endocytose de la vésicule synaptique synaptique hippocampal mammifères en culture neuronale. Nous avons suivi l’endocytose de protéine Vésicule synaptique en fusionnant une protéine de la membrane de vésicules synaptiques, y compris synaptophysine et VAMP2/synaptobrévine, sur le côté lumière vésiculaire, avec pHluorin, une protéine fluorescente verte sensibles au pH qui augmente sa intensité de la fluorescence lorsque le pH augmente. Au cours de l’exocytose, pH lumen vésiculaire augmente, alors qu’au cours de la lumière vésiculaire endocytose pH est re-acidifié. Ainsi, une augmentation de l’intensité de fluorescence pHluorin indique fusion, alors qu’une diminution indique l’endocytose de la protéine marquée des vésicules synaptiques. Outre l’utilisation de la méthode d’imagerie pHluorin pour enregistrer l’endocytose, nous avons suivi endocytose de la membrane vésiculaire par microscopie électronique (me) mesures d’absorption de raifort peroxydase de raifort (HRP) par des vésicules. Enfin, nous avons suivi la formation de fosses terminal membrane nerveuse à des moments différents après forte dépolarisation induite par le potassium. L’évolution temporelle de la HRP absorption et membrane fosse formation indique l’évolution temporelle de l’endocytose.

En utilisant la méthode de transporteur de lipides, SpH a été exprimé dans les neurones de l’hippocampe, permettant l’identification des boutons (Figure 1 a). La stimulation électrique des cellules induites par exocytose et une augmentation correspondante dans l’intensité de la fluorescence. L’augmentation de fluorescence (ΔF) a été arrêtée en mettant fin à la stimulation (Figure 1 b). L’augmentation de la f...

Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Endocytose Vésicule synaptique est détectée par microscopie de pHluorin fusionnée avec des protéines de la vésicule synaptique et par microscopie électronique de l’absorption de la vésicule.

Mesurer l’endocytose de la vésicule synaptique dans des neurones Hippocampal cultivés

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

The overall goal of this labeling methodology is to measure synaptic vesicle endocytosis. This method can help answer key questions in the neuroscience field such as the dispersion of formed endocytic vesicles. The main advantage of this technique is the resolution during imaging and the visualization of changes by examining the complete structure of a cell.Generally, individuals new to this method will struggle because they fail to remove tiny air bubbles while preparing the epoxy or dry out the sample during the ethanol washes. To begin this procedure, prepare a poly-d-lysine coated six-well plate by applying 1.5 milliliters of 0.01%sterile filtered poly-d-lysine solution to each well for one hour at room temperature, then washing it three times with sterilized water. Next, prepare a high potassium stimulation solution with horse radish peroxidase and adjust the solution to pH 7.4 with five molar sodium hydroxide.After that, stimulate the hippocampal neuron culture by adding 1.5 milliliters of high potassium stimulation solution to each well at room temperature for 90 seconds. In this step, prepare a 0.1 molar sodium cacodylate buffer at pH 7.4. Fix the cells with 4%glutaraldehyde in 0.1 molar sodium cacodylate buffer for at least one hour at room temperature, then wash the cells three times with 0.1 molar sodium cacodylate buffer each time for seven minutes.Following that, prepare DAB solution and filter it with a 0.22 micrometer filter. Next, apply 1.5 milliliters of DAB solution to the cells for 30 minutes at 37 degrees Celsius, then wash the cells three times with 0.1 molar sodium cacodylate buffer each time for seven minutes. As post-fixation, incubate the neurons with 1.5 milliliters of 1%osmium tetroxide in 0.1 molar sodium cacodylate buffer for one hour at four degrees Celsius.Afterward, wash the neurons three times with 1.5 milliliters of 0.1 molar sodium cacodylate buffer each time for seven minutes. Subsequently, prepare 0.1 molar sodium acetate buffer with sodium acetate and glacial acetic acid. Wash the cells three times with 1.5 milliliters of 0.1 molar acetate buffer each time for seven minutes.After that, incubate the cells with 1.5 milliliters of 1%uranyl acetate in 0.1 molar acetate buffer for one hour at four degrees Celsius, then wash the cells three times with 1.5 milliliters of 0.1 molar acetate buffer for seven minutes each time. In a fume hood, dehydrate the neuronal culture with single 1.5 milliliter washes with 50%70%and 90%ethanol for seven minutes each wash, followed by three washes with 1.5 milliliters of 100%ethanol for seven minutes each time. Next, create the epoxy resin and mix thoroughly and store it under vacuum to remove air bubbles.Infiltrate the sample by replacing the ethanol with 50%epoxy resin in ethanol on a shaker for 30 minutes at room temperature, then 70%epoxy resin in ethanol for 30 minutes at room temperature on a shaker. Switch the epoxy resin solution with 100%epoxy resin and incubate for 10 minutes at 50 degrees Celsius. Perform two exchanges of fresh 100%epoxy resin, each exchange is incubated for one hour at 50 degrees Celsius.Then add fresh 100%epoxy resin and allow it to harden at 50 degrees Celsius overnight and then at 60 degrees Celsius for over 36 hours. Afterward, remove each sample from the multi-well plate with a driller's handsaw. Using an inverted light microscope, select the regions of interest that have dense concentrations of cells, then cut the blocks of four by five by eight millimeters cube for sectioning.Afterward, counterstain the sections by submersion with 1%aqueous uranyl acetate for 15 minutes and then 3%aqueous lead citrate for five minutes to improve the contrast of the samples. In this study, electron microscopy was performed in cultured neuron and the clathrin-coated vesicles were examined in comparison to the control samples which were incubated with five milligrams per milliliter of HRP for 90 seconds in the absence of stimulus. The clathrin-coated pits were observed at a probability of 0.05 per bouton in the absence of stimulus.In these images, stimulation with high potassium induced bulk endosome and synaptic vesicles uptake were identified by HRP labeling. Synaptic vesicles were defined as vesicles with a diameter less than 50 nanometers and bulk endosomes were defined as having a diameter over 80 nanometers or with a cross-sectional area of more than an 80-nanometer vesicle. Induced bulk endosome uptake was decreased by recovery with normal saline.Once mastered, this technique can be done in 8-1/2 hours with three additional days to harden the epoxy if it is performed properly. Following this procedure, other methods such as immunogold labeling can be performed in order to answer additional questions like distribution of a target protein at various points in stimulation recovery. After watching this video, you should have a good understanding of how to stabilize and stain samples in preparation for imaging by electron microscopy.

measuring-synaptic-vesicle-endocytosis-in-cultured-hippocampal-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.

Etudier Piscines vésicules synaptiques utilisant des colorants styryle photoconversion

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

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Neurosciences

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.

Récupération de fluorescence après photoblanchiment (FRAP) de fluorescence protéines marquées dans les épines dendritiques des neurones de l&#39;hippocampe de culture

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 ...

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.

L&#39;analyse de la morphologie des épines dendritiques dans les neurones du SNC de culture

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.

Analyse quantitative des vésicules reconstitution Piscine synaptique dans des cultures de neurones granulaires du cervelet utilisant des colorants FM

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.

Induire la croissance dendritique dans des cultures de neurones sympathiques

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.

Diffusion latérale et l&#39;exocytose des protéines membranaires dans des neurones en culture évaluée à l&#39;aide de récupération de fluorescence et de fluorescence de perte de photoblanchiment

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

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Détection de la palmitoylation des protéines dans les neurones de l&#39;hippocampe en culture par immunoprécipitation et Acyl-biotine Exchange (ABE)

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

Le dosage de dommages de l&#39;ADN dans les neurones hippocampiques Utilisation de l&#39;essai de comète

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Étiquetage immunologique post-enrobage des protéines synaptiques dans l&#39;hippocampe cultures Slice

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

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.

Examen de la vésicule synaptique recyclage utilisant des colorants FM Pendant évoqués, spontanée, et les activités synaptiques miniatures

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