We thank Yehui Qin and Mallory Myers for technical support. We also thank Dr. Sigismund Huck for providing us the anti-&#945;4-ECD antibody. This work is supported by National Institutes of Health grant NS22061 to L. W. R.

Toutes les expériences sur les animaux ont été effectuées en conformité avec les National Institutes of Health Guide pour le soin et l&#39;utilisation des animaux de laboratoire (NIH Publications n ° 80-23, révisé 2012) et des études ont été approuvés par Institutional Animal Care et à utiliser pour les comités de recherche à Stony Brook University (# 1618 et # 1792). 1. vHipp ANCC synaptiques Co-cultures <ol><li> souris Sacrifice (jour postnatal 0 - 3, de type sauvage (WT) ou nAChRs α7 de ligne de souris transgénique) avec CO 2. Décapitez chaque chiot et enregistrer la queue dans un tube de 1,5 ml de centrifugeuse pour le génotypage post facto. Effectuez les étapes suivantes dans une hotte stérile. </li><li> Retirez le sommet du crâne avec des ciseaux et des pinces pour exposer l&#39;ensemble du cerveau. À partir de la jonction où les cortex cérébraux partie caudale de l&#39;autre, d&#39;obtenir une section coronale qui contient les cortex postérieur dont la vHipp 5,21 sous un microscope à dissection. </li><li> Disséquer vBandes de tissu Hipp contenant la région CA1-subiculum selon un cerveau de souris en développement atlas 22 (figure 1A). Transfert à une boîte de culture de 35 mm contenant des milieux de culture froid (4 ° C), les couper en petites tranches minces (environ 100 um x 100 um de microslices). </li><li> Pré-incuber 12 mm de poly-D-lysine / laminine lamelles de verre revêtu à l&#39;intérieur 24 des plaques de culture de tissu avec un milieu de culture (~ 500 ul) pendant au moins 30 min à l&#39;étuve à 37 ° C. Retirez le support avant l&#39;étalement microslices. </li><li> Plaque avec une pipette Pasteur polie au feu au centre de la lamelle avec une quantité minimale de médias (~ 50 pi, contenant ~ 20 morceaux de microslices) pour faciliter la fixation des explants. Plate microslices vHipp provenant d&#39;un seul animal (soit le + / + ou - / - génotype de la ligne de α7 transgénique) sur chaque lamelle. </li><li> Après les explants se déposent sur la lamelle, ajouter doucement les médias supplémentaires (~ 100 pi) par ee mur de sorte que le niveau des médias est suffisamment élevé pour couvrir complètement les explants sur la périphérie, mais pas ceux qui sont au centre de la lamelle. </li><li> Après un O / N incubation dans 37 ° C, 5% de CO 2 incubateur, disperser neurones embryonnaires CNAC de jours 18 en jour 1 souris WT postnatales et ajouter à des explants. <ol><li> Disséquer les tissus CNAC selon un développement du cerveau de la souris atlas 20 (figure 1B). </li><li> Coupez-les en petits morceaux (environ 500 um x 500 um microslices) et transférer dans un tube de 15 ml. </li><li> Traiter avec 0,25% de trypsine (2 ml) pendant 15 min à 37 ° C. Wash Les morceaux de tissus à trois reprises avec les médias de lavage à froid (5 ml, 4 ° C), suivie par un lavage dans des milieux de culture froide (5 ml, 4 ° C). Laisser la suspension reposer pendant 5 min dans chaque étape de lavage, puis versez le surnageant avec précaution. </li><li> Dissocier les cellules par trituration ménagée dans 2 ml de milieux de culture avec une pipette Pasteur légèrement poli-le-feu. </li><li> Transfer suspension cellulaire à un autre tube de 15 ml et on centrifuge à 2000 x rpm pendant 5 min. Retirer le surnageant et remettre en suspension les cellules dans des milieux de culture à 1 ml par 6 chiots. Disperser les cellules par pipetage dans les médias à plusieurs reprises avec une pipette Pasteur polie au feu. </li><li> Ajouter 0,25 ml de cellules CNAC dispersées à chaque lamelle avec des explants vHipp. </li><li> Maintenir les cultures dans une humidifié à 37 ° C, 5% de CO 2 incubateur. Placez des plaques de culture sur des tampons de gaze stérile humide pour stabiliser mécaniquement et maintenir l&#39;humidité, faciliter la formation des synapses. </li></ol></li></ol> 2. Immunocytochemistry <ol><li> Fixer cultures (5 - 7 jours in vitro) en paraformaldéhyde 4% / 4% de saccharose / PBS (~ 500 pi par lamelle, 20 min, RT). </li><li> Perméabiliser les cultures avec 0,25% de Triton X-100 / PBS (~ 500 ul par lamelle couvre-objet, 5 min, RT). </li><li> cultures de bloc avec 10% de sérum âne normale dans PBS (~ 500 pi par lamelle, 30 min, RT) avant Antibody coloration. Incuber avec des anticorps primaires O / N à 4 ° C. Utiliser les anticorps primaires suivants: anti-nAChR (α 4 -ECD 1: 500; anti-sous-unités, α 5 1: 500), anti-glutamate transporteur vésiculaire 1 (1: 250), anti-GAD65 (1: 100). </li><li> Laver les anticorps primaires avec du PBS (3 fois / ~ 500 ul par lamelle couvre-objet, 5 min) et ensuite incuber les cultures dans des anticorps secondaires conjugués à de l&#39;Alexa 488 (~ 500 ul par une lamelle couvre-objet,: 500) ou Alexa 594 (~ 500 ul par lamelle couvre- , 1: 500) pendant 1 heure à 37 ° C. </li><li> Incuber les cultures à αBgTx conjugué à Alexa 594 (~ 500 ul par une lamelle couvre-objet,: 1000 dans du milieu de culture) pendant 45 min à 37 ° C avant la fixation pour le marquage surface α7 nAChR *. </li><li> Montage et de phoques lamelles. </li><li> Capturez des images avec un microscope à fluorescence équipé d&#39;objectifs Plan-Apochromat (20X avec 0,8 NA ou 63X huile avec 1,4 NA) et une caméra CCD. </li></ol> 3. FM1-43 Based vésiculeuse Fusion <ol><li> Maintenir cultures (5 - 7 jours in vitro) dans une chambre de l&#39;imagerie, monter la chambre sur disque rotatif microscope confocal. Perfuser en continu (1 ml / min) avec HBS cocktail au RT. </li><li> Arrêtez perfusion et de charge cultures avec 10 uM FM1-43 dans 56 mM K + ACSF (~ 500 pi) pendant 90 secondes (coloration), laver colorant externe distance pendant 15 min avec Ca 2+ HBS gratuit contenant ADVASEP-7 (0,1 mM) pour piéger FM1-43 liée à la membrane. </li><li> Stimuler les cultures avec 56 mM K + ACSF (~ 500 pi) sans FM1-43 pendant 120 secondes (décoloration), rechargez cultures avec FM1-43 en utilisant les mêmes conditions, et lavez à nouveau avec Ca 2+ HBS gratuit contenant ADVASEP-7 pour 15 min. </li><li> Acquérir FM1-43 images de fluorescence avec disque rotatif microscope confocal équipé d&#39;un objectif Plan-Apochromat (60X eau avec 1,4 NA, excitation 488 nm, émission 530 nm) et de capturer des images avec une caméra CCD. </li><li> Recueillir base d&#39;images de fluorescence de FM1-43 (tous les 2 s pendant 1 min) Que le contrôle pré-nicotine; appliquer la nicotine (1 uM) par perfusion rapide (2 ml / min) pendant 1 min, puis laver la nicotine avec HBS cocktail. Gardez capture des images en accéléré avant, pendant, et après l&#39;application de la nicotine toutes les 2 secondes pendant 5 min. </li><li> Quantifier FM1-43 intensité de fluorescence avant (F de coloration) et après (F décoloration) l&#39;application de la nicotine à chaque bouton synaptique. </li><li> Calculer la quantité totale de fluorescence de FM1-43 amovible le long des axones vHipp en quantifiant la différence de FM1-43 intensité de fluorescence avant et après l&#39;application de nicotine (AF = F -F coloration de décoloration). Analyser fraction de la diminution de l&#39;intensité de fluorescence après la nicotine avec l&#39;équation suivante: F% = diminution AF / F coloration. </li></ol> 4. Imagerie de calcium <ol><li> Rincer cultures (5 - 7 jours in vitro) rapidement avec HBS normale, puis charger cultures avec 5 uM Fluo-4 Ca 2+ indicateur (ester AM) et 0,02% de Pluronic F-127 dans du HBS (2 ml) pendant 30 min à 37 ° C et 5% de CO 2. </li><li> Laver Fluo-4 solution avec HBS (3 fois / 5 min). Retour cultures dans l&#39;incubateur (37 ° C / 5% CO 2) pendant au moins 30 min. </li><li> Maintenir les cultures dans la même chambre de l&#39;imagerie et les mêmes conditions que celles décrites dans FM1-43 section de fusion vésiculaire à base (voir l&#39;étape 3.1). </li><li> Recueillir images Fluo-4 fluorescence des projections axonales des micro-tranches vHipp avec un objectif Plan-Apochromat (60X eau avec 1,4 NA, excitation 488 nm, émission 530 nm) et une caméra CCD. </li><li> Recueillir base Fluo-4 images de fluorescence (toutes les 10 sec pendant 2 min) que le contrôle pré-nicotine, appliquer la nicotine (1 uM) par perfusion rapide (2 ml / min) pendant 1 min, puis laver la nicotine avec HBS cocktail. Gardez capture des images en accéléré avant, pendant, et après l&#39;application de la nicotine toutes les 10 secondes pendant 30 min. </li><li> Enregistrer tous les cadres des premières fluo-4 images de fluorescence, l&#39;exportation comme une série deLes images au format TIFF pour pousser l&#39;analyse. </li><li> Recueillir l&#39;intensité intégrée de fluo-4 fluorescence le long des axones vHipp avant et après l&#39;application de la nicotine. </li><li> Normaliser l&#39;intensité de fluorescence intégrée avec l&#39;équation suivante: AF / F 0 = (FF 0) / F 0, où F 0 est la pré-nicotine intensité de fluorescence intégrée de fond corrigé et F est l&#39;intensité de fluorescence intégrée le long des axones vHipp à chaque point de temps après l&#39;application de la nicotine. Analyser et tracer l&#39;intensité de fluorescence intégrée normalisée. </li></ol>

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La préparation de co-culture décrit re-capitule circuits de l&#39;hippocampe-accumbens ventrales in vitro. Cet examen permet de préparation relativement simple et fiable des profils spatiaux et temporels qui activation des nAChR pré-synaptiques susciter améliorée transmission glutamatergique 5, 21. Co-cultures sont définis comme la croissance de différents types de cellules spécifiques dans un plat qui peut fournir des conditions physiologiques in vitro ...

Modulation cholinergique du circuit excitabilité contribue à des aspects fondamentaux de la cognition, et la modulation cholinergique modifié est une caractéristique de neurodégénératives et neuropsychiatriques troubles y compris la maladie d&#39;Alzheimer, la maladie de Parkinson, la schizophrénie et la toxicomanie 1-4. Un mécanisme établi de la facilitation de la transmission synaptique cholinergique dans le SNC est via l&#39;activation directe de nAChRs localisées sur des sites pré-synaptiques. L&#39;activation de ces récepteurs pré-synaptiques entraîne une augmentation de Ca 2+ intracellulaire ([Ca2 +] i) dans les terminaux présynaptiques - à la fois directement, en raison de la conductance de calcium relativement élevé de certains sous-types de nAChR, et indirectement, par l&#39;intermédiaire de cascades de signalisation intracellulaires 5, améliorant ainsi la libération de neurotransmetteurs. En effet, l&#39;activation des nAChR pré-synaptiques a été liée à des changements dans la libération d&#39;une grande variété de neurotransmetteurs, y compris le glutamate, GABA, acétylcholine, une dopamine 6-10. Bien que ce processus a été étudié indirectement en utilisant des méthodes électrophysiologiques à divers synapses, les journalistes optiques de [Ca 2+] i et vésicule synaptique recyclage permettent de mesurer plus directe et temporellement précise des phénomènes pré-synaptiques. Localisation pré-synaptique de nAChR a été démontré de façon convaincante direct étiquetage immuno-or de nAChRs au microscope électronique (EM) de niveau 11,12. Plusieurs autres techniques ont également été utilisés pour traiter la localisation nAChR indirectement, y compris la détection de l&#39;emplacement des chimères protéiques fluorescents de nAChR dans des neurones en culture 13,14, enregistrement électrophysiologique des courants nAChR dans les terminaux synaptiques 15,16, la surveillance des changements induits par la nicotine en [Ca 2+] i dans les terminaisons nerveuses synaptiques par imagerie de cellules vivantes 17, et la surveillance indirecte de la libération des neurotransmetteurs à la terminaison synaptique partechniques d&#39;imagerie cellulaire en direct avec des indicateurs fluorescents, y compris l&#39;exocytose des vésicules synaptiques vus par les colorants de styryle amphipathiques FM (FM1-43 et FM4-64) et / ou synapto-pHluorin et par des journalistes de neurotransmetteurs fluorescentes spécifiques, tels que CNiFERs pour ACh et iGluSnFr pour le glutamate 18-20. Globalement, ces approches actuelles pour identifier la localisation pré-synaptique de nAChRs sont complexes, et nécessitent des systèmes et des techniques spéciales pour permettre une identification fiable et surveillance physiologique de l&#39;activité pré-synaptique. Nous décrivons ici les protocoles et l&#39;équipement pour un système de co-culture in vitro d&#39;un hippocampe ventral (vHipp) - noyau accumbens (NACC) circuit qui offre un accès direct à identifier et analyser les deux composants pré et post-synaptiques de la transmission synaptique. Nous montrons des exemples de localisation pré-synaptique de nAChRs et l&#39;imagerie des cellules vivantes de nAChR médiation Ca 2+ signalisation et de la libération de neurotransmetteurs aloaxones ng vHipp. Une extension naturelle (et simple) du protocole présenté ici est la préparation des contacts pré- et post-synaptiques constitués de neurones à partir de différents génotypes. De cette manière, la contribution d&#39;un produit de gène particulier de la pré- et / ou post-synaptiques des mécanismes de modulation peut être évaluée directement.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>1, Culture Media (50 ml)</td>
		</tr>
		<tr>
			<td>Neurobasal&#160;</td>
			<td>GIBCO</td>
			<td>10888022</td>
			<td>48 ml</td>
		</tr>
		<tr>
			<td>B-27 Supplements</td>
			<td>GIBCO</td>
			<td>0080085-SA</td>
			<td>1 ml</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>GIBCO</td>
			<td>10908-010</td>
			<td>0.5 ml</td>
		</tr>
		<tr>
			<td>GlutaMAX Supplement</td>
			<td>GIBCO</td>
			<td>35050-061</td>
			<td>0.5 ml</td>
		</tr>
		<tr>
			<td>Brain-derived neurotrophic factor (BDNF)</td>
			<td>GIBCO</td>
			<td>15140-122</td>
			<td>20 ng/ml</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>2, washing media (HBSS, 100 ml)</td>
		</tr>
		<tr>
			<td>HBSS, no calcium, no magnesium, no phenol red&#160;</td>
			<td>GIBCO</td>
			<td>14175-095</td>
			<td>99 ml</td>
		</tr>
		<tr>
			<td>HEPES ( 1M)</td>
			<td>GIBCO</td>
			<td>15630-130</td>
			<td>1 ml</td>
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		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>3, HEPES buffered saline&#160; (HBS)&#160;&#160; pH=7.3</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888&#160;</td>
			<td>135 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333&#160;</td>
			<td>5 mM</td>
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		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266&#160;</td>
			<td>1 mM</td>
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		<tr>
			<td>CaCl2,</td>
			<td>Sigma</td>
			<td>C1016&#160;</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>4, HBS Cocktail for live imaging pH=7.3</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888&#160;</td>
			<td>135 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333&#160;</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266&#160;</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>CaCl2,</td>
			<td>Sigma</td>
			<td>C1016&#160;</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>tetrodotoxin&#160;</td>
			<td>Tocris</td>
			<td>1078</td>
			<td>2 &#181;M</td>
		</tr>
		<tr>
			<td>bicuculline</td>
			<td>Tocris</td>
			<td>131</td>
			<td>10 &#181;M</td>
		</tr>
		<tr>
			<td>D-AP-5</td>
			<td>Tocris</td>
			<td>105</td>
			<td>50 &#181;M</td>
		</tr>
		<tr>
			<td>CNQX</td>
			<td>Tocris</td>
			<td>1045</td>
			<td>20 &#181;M</td>
		</tr>
		<tr>
			<td>LY341495</td>
			<td>Tocris</td>
			<td>1209</td>
			<td>10 &#181;M</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>5, Calcium-free&#160; HBS&#160;&#160; pH=7.3</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888&#160;</td>
			<td>135 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333&#160;</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266&#160;</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>6, 56 mM Potassium ACSF pH=7.4</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888</td>
			<td>119 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333</td>
			<td>56 mM</td>
		</tr>
		<tr>
			<td>MgSO4.7H</td>
			<td>Sigma</td>
			<td>M1880</td>
			<td>1.3 mM</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma</td>
			<td>C1016</td>
			<td>2.5 mM</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Sigma</td>
			<td>S8282</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma</td>
			<td>S5761</td>
			<td>26.2 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
	</tbody>
</table>

live imaging of nicotine induced calcium signaling and neurotransmitter release along ventral hippocampal axons

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors (nAChRs) is an important mechanism for neuromodulation by acetylcholine (ACh). The difficulty with access to probing the signaling mechanisms within intact axons and at nerve terminals both in vitro and in vivo has limited progress in the study of the pre-synaptic components of synaptic plasticity. Here we introduce a gene-chimeric preparation of ventral hippocampal (vHipp)&#8211;accumbens (nAcc) circuit in vitro that allows direct live imaging to analyze both the pre- and post-synaptic components of transmission while selectively varying the genetic profile of the pre- vs post-synaptic neurons. We demonstrate that projections from vHipp microslices, as pre-synaptic axonal input, form multiple, reliable glutamatergic synapses with post-synaptic targets, the dispersed neurons from nAcc. The pre-synaptic localization of various subtypes of nAChRs are detected and the pre-synaptic nicotinic signaling mediated synaptic transmission are monitored by concurrent electrophysiological recording and live cell imaging. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of glutamatergic synaptic plasticity in vitro.

La préparation employée se compose de gènes chimère co-cultures de circuits vHipp ANCC in vitro. Projections émanant microslices vHipp, comme entrée axonale pré-synaptique, peuvent faire des contacts synaptiques avec les objectifs post-synaptiques, les neurones dispersés du CNAC. La nicotine induit une (≥ 30 min) la facilitation de la transmission glutamatergique soutenue à partir de neurones CNAC innervés par vHipp axones 21 et prolongée calcium signalisation le long des axones vHipp <s...

Watch this Scientific Journal Video about Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons at JoVE.com

Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons

We developed a gene-chimeric preparation of ventral hippocampal &#8211; accumbens circuit in vitro that allows direct live imaging to analyze presynaptic mechanisms of nicotinic acetylcholine receptors (nAChRs) mediated synaptic transmission. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of synaptic plasticity.

Imagerie en direct de la nicotine induit la signalisation du calcium et de la libération de neurotransmetteurs long ventrales hippocampe axones

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors ...

live-imaging-nicotine-induced-calcium-signaling-neurotransmitter

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Drugs Acting on Autonomic Nervous System: Adrenergic Agonists and Antagonists Agents

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8.9K vues.  Stony Brook University. We developed a gene-chimeric preparation of ventral hippocampal – accumbens circuit in vitro that allows direct live imaging to analyze presynaptic mechanisms of nicotinic acetylcholine receptors (nAChRs) mediated synaptic transmission. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of synaptic plasticity.

Vidéo: Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

Neuromodulation et mitochondriale de transport: Live in Imaging neurones de l&#39;hippocampe sur des durées longues

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

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Neurosciences

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

Imagerie en temps réel des axones de la racine dorsale après Rhizotomie

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Analyzing Murine Schwann Cell Development Along Growing Axons

The development of peripheral nerves is an intriguing process. Neurons send out axons to innervate specific targets, which in humans are often more than 100 cm away from the soma of the neuron. Neuronal survival during development depends on target-derived growth factors but also on the support of Schwann cells (SCs). To this end SC ensheath axons from the region of the neuronal soma (or the transition from central to peripheral nervous system) to the synapse or neuromuscular junction. Schwann cells are derivatives of the neural crest and migrate as precursors along emerging axons until the entire axon is covered with SCs. This shows the importance of SC migration for the development of the peripheral nervous system and underlines the necessity to investigate this process. In order to analyze SC development, a setup is needed which next to the SCs also includes their physiological substrate for migration, the axon. Due to intrauterine development in vivo time-lapse imaging, however, is not feasible in placental vertebrates like mouse (mus musculus). To circumvent this, we adapted the superior cervical ganglion (SCG) explant technique. Upon treatment with nerve growth factor (NGF) SCG explants extend axons, followed by SC precursors migrating along the axons from the ganglion to the periphery. The beauty of this system is that the SC are derived from a pool of endogenous SC and that they migrate along their own physiological axons which are growing at the same time. This system is especially intriguing, because the SC development along axons can be analyzed by time-lapse imaging, opening further possibilities to gain insights into SC migration.

Here we describe a Schwann cell (SC) migration assay in which SCs are able to develop along extending axons.

Analyser murin développement des cellules de Schwann le long des axones en croissance

The  development of peripheral nerves is an intriguing process. Neurons send out  axons to innervate specific targets, which in humans are often more than ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Phosphate de calcium transfection de primaire neurones de l&#39;hippocampe

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Imagerie en direct de<em&gt; Drosophila</em&gt; larves Neuroblastes

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Dorsal Root Ganglia Isolation and Primary Culture to Study Neurotransmitter Release

Dorsal root ganglia (DRG) contain cell bodies of sensory neurons. This type of neuron is pseudo-unipolar, with two axons that innervate peripheral tissues, such as skin, muscle and visceral organs, as well as the spinal dorsal horn of the central nervous system. Sensory neurons transmit somatic sensation, including touch, pain, thermal, and proprioceptive sensations. Therefore, DRG primary cultures are widely used to study the cellular mechanisms of nociception, physiological functions of sensory neurons, and neural development. The cultured neurons can be applied in studies involving electrophysiology, signal transduction, neurotransmitter release, or calcium imaging. With DRG primary cultures, scientists may culture dissociated DRG neurons to monitor biochemical changes in single or multiple cells, overcoming many of the limitations associated with in vivo experiments. Compared to commercially available DRG-hybridoma cell lines or immortalized DRG neuronal cell lines, the composition and properties of the primary cells are much more similar to sensory neurons in tissue. However, due to the limited number of cultured DRG primary cells that can be isolated from a single animal, it is difficult to perform high-throughput screens for drug targeting studies. In the current article, procedures for DRG collection and culture are described. In addition, we demonstrate the treatment of cultured DRG cells with an agonist of neuropeptide FF receptor type 2 (NPFFR2) to induce the release of peptide neurotransmitters (calcitonin gene-related peptide (CRGP) and substance P (SP)).

Dorsal root ganglia (DRG) primary cultures are frequently used to study physiological functions or pathology-related events in sensory neurons. Here, we demonstrate the use of lumbar DRG cultures to detect the release of neurotransmitters after neuropeptide FF receptor type 2 stimulation with a selective agonist.

Isolement des ganglions de la racine dorsale et la Culture primaire afin d’étudier la libération des neurotransmetteurs

Dorsal root ganglia (DRG) contain cell bodies of sensory neurons. This type of neuron is pseudo-unipolar, with two axons that innervate peripheral ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Ex Vivo Imagerie calcique pour visualiser le cerveau réponses à signalisation endocrinienne chez la drosophile

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its highly-organized subcellular architecture, we recently developed a new imaging modality, named spectral reflectometry (SpeRe), which enables unprecedented label-free nanoscale imaging of the live myelinated axons in situ. The underlying principle is to obtain nanostructural information by analyzing the reflectance spectrum of the multilayered subcellular structure. In this article, we describe a detailed step-by-step protocol for performing a basic SpeRe imaging of the nervous tissues using a commercial confocal microscopic system, equipped with a white-light laser and a tunable filter. We cover the procedures of sample preparation, acquisition of spectral data, and image processing for obtaining nanostructural information.

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

Microscopie spectrale réflectométrique sur les axones myélinisés In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its ...

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of individual cells. Since an increase of intracellular calcium (Ca2+) in the cytosol often occurs because of the electrical activity, or in response to a myriad of other stimuli, this process can be monitored by the imaging of cells loaded with fluorescent calcium-sensitive dyes.&#160; However, it is difficult to image this response in an individual cell type within whole tissue because these dyes are taken up by all cell types within the tissue. In contrast, genetically encoded calcium indicators (GECIs) can be expressed by an individual cell type and fluoresce in response to an increase of intracellular Ca2+, thus permitting the imaging of Ca2+ signaling in entire populations of individual cell types. Here, we apply the use of the GECIs GCaMP3/6 to the mouse neuromuscular junction, a tripartite synapse between motor neurons, skeletal muscle, and terminal/perisynaptic Schwann cells. We demonstrate the utility of this technique in classic ex vivo tissue preparations. Using an optical splitter, we perform dual-wavelength imaging of dynamic Ca2+ signals and a static label of the neuromuscular junction (NMJ) in an approach that could be easily adapted to monitor two cell-specific GECI or genetically encoded voltage indicators (GEVI) simultaneously. Finally, we discuss the routines used to capture spatial maps of fluorescence intensity. Together, these optical, transgenic, and analytic techniques can be employed to study the biological activity of distinct cell subpopulations at the NMJ in a wide variety of contexts.

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

Here we present a protocol to image calcium signaling in populations of individual cell types at the murine neuromuscular junction.

Ex Vivo Imagerie du Calcium cellulaire spécifique de signalisation à la Synapse Tripartite du diaphragme souris

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of ...

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster In Vivo