Financement: Ce travail a été soutenu par une subvention de la Fondation Bertarelli (DJL), un MED-EL subvention (DJL) et les Instituts nationaux de la santé Subventions DC01089 (RCM).

NOTE: Toutes les procédures expérimentales sont effectuées en conformité avec le soin et l&#39;utilisation Comité pour les Animaux du Massachusetts Eye and Ear Infirmary et Harvard Medical School, qui suivent les lignes directrices de soins aux animaux nationaux, y compris la politique de santé publique service sur Humane soin et l&#39;utilisation des animaux de laboratoire, le Guide ILAR, et la Loi sur la protection des animaux. Les procédures expérimentales énumérés ci-dessous l&#39;exposition détaillée de la DCN gauche. Utilisez des instruments stériles tout en effectuant la chirurgie de survie. 1. Craniotomie primaire et Dorsale Cochlear Nucleus exposition <ol><li> Anesthésie <ol><li> Anesthésier une souris, âgées de 8 à 12 semaines, pesant 18 à 24 g, de xylazine 20 mg / kg de kétamine et 100 mg / kg par une administration intrapéritonéale. Confirmez anesthésie appropriée par le taux de surveillance cardiaque, fréquence respiratoire, ainsi que pincement de l&#39;orteil réflexe de retrait. Lieu vétérinaire onguent sur les yeux pour prévenir la sécheresse sous anesthésie. Une fois correctement sous anesthésie, cheveux rasage recouvrant til cuir chevelu pour fournir un accès libre sur le site chirurgical. </li></ol></li><li> Positionnement chirurgical <ol><li> Placez la souris en toute sécurité dans un petit porte-stéréotaxique animal, maintenu en place par une pince de museau. </li><li> Assurez-vous que la pince de museau est assez lâche pour permettre une respiration adéquate, mais assez serré pour immobiliser complètement la tête de la souris. Si la tête de la souris est lâche, l&#39;animal est susceptible de tomber sur le support de tête pendant la partie craniotomie de la procédure. </li><li> Placez électrodes implant du tronc cérébral à la mode standard, ce qui permet un suivi de la fréquence cardiaque. La fréquence respiratoire doit être surveillée par la visualisation. Notez que la fréquence cardiaque normale et la fréquence respiratoire varient en fonction de l&#39;âge de la souris. </li><li> Un thermomètre est placé et euthermia est assurée par le placement sur une couverture chauffante homéotherme. </li></ol></li><li> incision de la peau et l&#39;exposition des os interpariétales et occipital du crâne <ol><li> Sous un Microscope, faire une incision verticale à travers la peau à partir de la ligne médiane, directement entre le pavillon de l&#39;oreille, et se étendant jusqu&#39;à la partie caudale de l&#39;occiput. Après incision médiane de la peau, la peau déplacer latéralement. Visualiser les muscles couvrant la gauche de la face postérieure du crâne. </li><li> Retirer muscle recouvrant la pariétale gauche, interpariétal, et les os occipital du crâne par désarticulation des attaches musculaires à leurs os respectifs avec un scalpel ou de l&#39;iris ciseaux. Observer un petit degré de saignement le long des bords de coupe de muscle. Réduire cette par une légère pression avec un applicateur coton-tige pour 10-15 sec. </li></ol></li><li> Recouvrant Craniotomie Cochlear Nucleus <ol><li> Identifier les lignes de suture pertinents, y compris des sutures sagittale et lambda lignes (Figure 1, panneau de gauche). </li><li> Utilisation de rongeurs, faire une craniotomie sur l&#39;os interpariétal, à gauche de la ligne médiane, ~ 2 mm caudale à la ligne lambda de suture. Cette région recouvre la DCN. </li><li> Following craniotomie, observer une mince couche de dure qui recouvre le cervelet. En utilisant une lame de scalpel, enlever la dure-mère. (Figure 1, panneau moyen). REMARQUE: Enlèvement de dure peut causer un faible degré de saignement. Ce processus d&#39;utilisation de la lame de scalpel pour enlever la dure-mère est destiné à réduire la quantité de perte de sang pendant l&#39;aspiration de cervelet, qui mettra en commun sur la surface du tronc cérébral et l&#39;identification obscur de la DCN. Le volume de sang total de souris est d&#39;environ 1,5 ml devraient pas dépasser&gt; à 15%. Si le saignement dépasse ce montant, la souris doit être muni d&#39;une supplémentation fluide. </li><li> Retirer sang coagulé recouvrant le cervelet avec la pointe du coton applicateur. Eventuellement, égoutter doucement solution saline en utilisant un point dentaires sur le cervelet pour effacer le sang loin sang. </li></ol></li><li> Cérébelleuse aspiration <ol><li> L&#39;utilisation d&#39;un 5 aspiration français, aspirer la partie latérale la plus du cervelet gauche recouvrant la DCN jusqu&#39;à ce que la DCN est visUAL (Figure 1, panneau de droite). Retirer environ 1/4 à 1/3 du cervelet gauche. Le principal point de repère à côté de la DCN est l&#39;ampoule du canal semi-circulaire supérieur. NOTE: L&#39;aspiration du cervelet est l&#39;étape clé du protocole. Réalisé aspiration du cervelet est effectué le plus de succès si elle se produit en une seule tentative et non avec de multiples passes de l&#39;aspiration car cela pourrait causer des saignements. Pour aider à l&#39;aspiration, régler le plan focal du microscope à la profondeur prévue de la NC, qui sera légèrement distale par rapport à la surface du cervelet. Réglage du plan focal du CN améliorer la visualisation et d&#39;assurer une image nette. </li><li> Après aspiration, le saignement, liquide céphalo-rachidien (LCR) accumulation, et le déplacement du cervelet sur la DCN sont attendus. Instiller 0,5 ml de solution saline stérile rapidement dans la craniotomie pour empêcher la coagulation du sang. Une combinaison de tamponnant doucement avec un point dentaire et aspiration peut alors êtreutilisé pour déblayer un chemin pour la visualisation directe de la DCN. Ne pas directement contact avec la surface de la DCN. </li></ol></li></ol> 2. microinjection à pression à médiation virale Gene Transfer and Recovery chirurgical <ol><li> Après la DCN est libre de sang recouvrant et le LCR et est clairement visible, assurez microinjections de pression dans la DCN en utilisant une seringue Hamilton de 10 ul sur une période de 2 min. </li><li> Introduire l&#39;aiguille avec un micromanipulateur jusqu&#39;au bout ne est plus visible sous la surface de la DCN. </li><li> Pour des performances optimales, utilisez une aiguille de calibre 33 ou 34 (utiliser une seringue étanche aux gaz avec l&#39;aiguille 34 gauge) avec un biseau peu profonde, comme 45 °, afin de minimiser traumatisme contondant et localiser le volume d&#39;injection dans le DCN, qui est un mince et la structure du tronc cérébral peu profondes (&lt;300 um d&#39;épaisseur). NOTE: D&#39;autres instruments de micro-injection peuvent être fondées utilisé. Idéalement, l&#39;instrument d&#39;injection doit être souple pour permettre une légère bend, si nécessaire pour cibler directement la DCN. En outre, comme la souris se déplace légèrement tout au long de la procédure en raison de la respiration, l&#39;instrument doit être suffisamment robuste pour supporter des quantités mineures de mouvement. </li></ol> 3. Récupération chirurgicale <ol><li> Immédiatement après l&#39;injection, re-rapprochant la peau et laissez la souris pour récupérer par la procédure de récupération standard. Restant tissu cicatriciel cervelet et remplira la cavité provoquée par l&#39;aspiration du cervelet. Surveiller constamment animaux jusqu&#39;à ce que la conscience suffisante est retrouvé à maintenir décubitus sternal. Surveiller le rythme cardiaque, fréquence respiratoire, ainsi que la capacité à nourrir. REMARQUE: Aucun animal qui a subi une intervention chirurgicale est retourné dans une cage avec les autres animaux jusqu&#39;à guérison complète. </li><li> Si une souris commence à tourner en rond après l&#39;opération (&lt;5% des cas), généralement 2-4 heures après la fermeture de l&#39;incision de la peau, immédiatement sacrifier la souris, comme il aurait probablement un temps à se nourrir difficile. Tson effet secondaire est probablement due à l&#39;aspiration du cervelet. L&#39;animal doit être surveillé pour la douleur post-opératoire, et les stupéfiants appropriées devrait être accordée pour assurer le confort des animaux basée sur des normes institutionnelles. Les antibiotiques peuvent être nécessaires, si la preuve de l&#39;infection. </li></ol> 4. secondaire exposition Craniotomie et Cochlear Nucleus <ol><li> Après 2-4 semaines de cicatrisation et virus transfert génique à médiation par incubation, re-anesthésier une souris et répétez les étapes 1.1 à 1.5 temps d&#39;incubation optimal précédemment injectée varie en fonction du type de gène transféré. </li><li> Retirer le tissu de cicatrice sur le site de craniotomie par une combinaison de pinces, scalpel, et rongeurs. </li><li> Après la visualisation de la craniotomie, identifier une combinaison de cervelet et tissu cicatriciel recouvrant la DCN reste. Utilisez un 5 aspiration française pour aspirer cervelet recouvrant / tissu cicatriciel (similaire à l&#39;aspiration cérébelleuse précédente). </li><li> Après aspiration, attendre le saignement, CSF benforcer-up, et les bits restants du cervelet. Introduire rapidement une solution saline dans la craniotomie pour empêcher la coagulation du sang. Utilisez une combinaison de douceur dentaires tamponnant point et d&#39;aspiration pour dégager le chemin pour la visualisation directe de la DCN. </li><li> Observer la surface de la DCN. Effectuer des expériences de physiologie base optogénétique-DCN est maintenant accessible. Présentez un stimulus lumineux, par exemple avec une fibre optique (Figure 2), à conduire une expérience basée optogénétique. </li></ol> 5. histologie <ol><li> Après la conclusion d&#39;expériences, euthanasier la souris avec une surdose de kétamine. Perfuser la souris avec une solution saline normale suivie par 4% de paraformaldéhyde. </li><li> Extraire le tronc cérébral du crâne et de post-correctif pour 2 heures. Cryoprotect le tronc cérébral dans 30% de saccharose pendant 24-48 h. Section du tronc cérébral au moyen d&#39;un cryostat standard en utilisant 60 sections um. </li></ol>

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Les auteurs ne ont rien à divulguer.

Cet article décrit la technique de visualisation directe de la DCN dans le modèle murin pour la manipulation du système auditif central. L&#39;approche décrite de la visualisation directe offre des avantages significatifs par rapport à la principale alternative, qui sont des approches stéréotaxiques. Principalement, la visualisation directe de la DCN permet de confirmation immédiate du site du tronc cérébral, alors que les approches stéréotaxiques ne offrent pas la visualisation directe. Dans les expérience...

Virus-mediated gene transfer to reverse hearing loss has largely been focused on the peripheral auditory system.1 Targeting the cochlea, investigators have examined a host of delivery routes, including osmotic minipump infusion2, vector-transgene complex-soaked Gelfoam&#174;2 or gelatin sponge3, direct microinjection4; numerous gene transfer vectors, including adeno-associated viral vectors5,6, lentiviral vectors7, and cationic liposomes2; and the dissemination of gene transfer vectors beyond the target tissue2. Most recently, adeno-associated virus (AAV)-1 has been introduced in the cochlea in order to treat deafness in mice due to loss of vesicular glutamate transporter-3.8 Further, the application of optogenetics in peripheral auditory system has recently been described.9
Few studies, however, have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem contain second order neurons of the auditory pathway. While gene transfer techniques in the cochlear nucleus (CN) may be utilized for a host of investigations, gene transfer of opsins, light-sensitive proteins, to the DCN may also be utilized to enable optogenetics-based experimental techniques. Following virus-mediated gene transfer delivery of an opsin, such as channelrhodopsin-2 (ChR2), the neurons of the DCN becomes sensitive to light stimuli. Optogenetic gene transfer has been previously attempted in several brainstem regions, including the rat retrotrapezoid nucleus, mouse locus coeruleus, monkey superior colliculus, and mouse ventral tegmental area.10-14
Recently, investigators have examined the use of optogenetics in the DCN.15,16 The DCN is the location of placement of auditory brainstem implants in humans, making it an attractive part of the auditory system to study for translational studies on auditory neuroprostheses. However, given the location of the DCN, surgical exposure is challenging. The technique described herein provides a protocol for maximal exposure of the DCN via posterior fossa approach to enable viral vector gene transfer and optogenetics-based experiments in a murine model. Previous studies used stereotactic microinjection into the DCN with channelrhodopsin-2.16 Stereotaxic injections, however, are potentially less accurate than injections made by direct visualization, especially in a nucleus as small and deep along the brainstem as the DCN. Transgenic mice expressing tissue specific proteins in the CN are also an attractive option and would obviate the need for gene transfer. Our protocol for exposure of the DCN would also work in transgenic mice as the DCN would need to be directly exposed for optical stimulation. This technique for surgical exposure of the DCN is adapted from previous protocols involving recordings from the auditory nerve and cochlear nucleus in mice and rat models.15,17-20
The overall goal of the protocol is to provide direct exposure to the CN to allow for gene transfer techniques. More specifically, the approach is compatible with both acute and survival surgery and the preparation can be repeated in the same animal for subsequent neurophysiological testing. The direct exposure of the DCN protocol has implications for optogenetics- and virus-mediated gene transfer-based experimentation in other nuclei of the brainstem.

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			<td height="21" style="height:21px;width:197px;">Stereotaxic holder</td>
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			<td height="21" style="height:21px;width:197px;">Scalpel blade #11</td>
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			<td>10011-00</td>
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			<td height="21" style="height:21px;width:197px;">5 French suction</td>
			<td style="width:227px;">Symmetry Surgical</td>
			<td>2777914</td>
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			<td height="21" style="height:21px;width:197px;">Dental Points</td>
			<td style="width:227px;">Henry Schein</td>
			<td>100-8170</td>
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		<tr height="25">
			<td height="46" rowspan="2" style="height:46px;width:197px;">Bone rongeur</td>
			<td rowspan="2" style="width:227px;">Fine Surgical Tools</td>
			<td rowspan="2">16020-14</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="30">
			<td height="51" rowspan="2" style="height:52px;width:197px;">10 &#181;l Hamilton syringe</td>
			<td rowspan="2" style="width:227px;">Hamilton&#160;</td>
			<td rowspan="2">7633-01</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="21">
			<td height="42" rowspan="2" style="height:42px;width:197px;">34 gauge, needle</td>
			<td rowspan="2" style="width:227px;">Hamilton&#160;</td>
			<td rowspan="2">207434</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rongeurs</td>
			<td>Fine Surgical Tools</td>
			<td>16021-14</td>
		</tr>
	</tbody>
</table>

direct visualization of the murine dorsal cochlear nucleus for optogenetic stimulation of the auditory pathway

Enquête sur l&#39;utilisation du transfert de gène à médiation virale pour arrêter ou inverser la perte auditive a largement été reléguée au système auditif périphérique. Peu d&#39;études ont examiné le transfert de gènes au système auditif central. Le noyau cochléaire dorsal (DCN) du tronc cérébral, qui contient des neurones de second ordre de la voie auditive, est un site potentiel de transfert de gènes. Dans ce protocole, une technique pour une exposition directe et maximale de la DCN murin par une approche de la fosse postérieure est démontrée. Cette approche permet une chirurgie aiguë ou la survie. Suite à la visualisation directe de la DCN, une multitude d&#39;expériences sont possibles, y compris l&#39;injection de opsins dans le noyau cochléaire et une stimulation ultérieure par une fibre optique couplée à un laser à lumière bleue. D&#39;autres expériences de neurophysiologie, telles que la stimulation électrique et tracés d&#39;injection de neurones sont également possibles. Le niveau de visualisation et la durée de la stimulation font réalisables cette approche applicable à un large éventail d&#39;expériences.

Partielle cérébelleuse aspiration démontre avoir accès à noyau cochléaire Après la peau et le muscle recouvrant le crâne, les sites d&#39;intérêt sont éliminés de la surface du crâne, comme les lignes de suture coronale et lamda, démontrer la localisation approximative de la craniotomie. Après craniotomie avec rongeurs, le cervelet est visualisée. Une aspiration soigneuse de la petite partie du cervelet montre la visualisation de la NC, qui peut alors être injecté ...

Watch this Scientific Journal Video about Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway at JoVE.com

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Visualisation directe de la Dorsale murin Cochlear Nucleus pour optogénétiques stimulation de la voie auditive

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

direct-visualization-murine-dorsal-cochlear-nucleus-for-optogenetic

Research

JoVE Journal

Neuroscience

9.1K vues.  Harvard Medical School. The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model. 

Vidéo: Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

TMS: Using the Theta-Burst Protocol to Explore Mechanism of Plasticity in Individuals with Fragile X Syndrome and Autism

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS display abnormalities in the expression of FMR1 - a protein required for typical, healthy neural development.3 Recent data has suggested that the loss of this protein can cause the cortex to be hyperexcitable thereby affecting overall patterns of neural plasticity.4,5
In addition, Fragile X shows a strong comorbidity with autism: in fact, 30% of children with FXS are diagnosed with autism, and 2 - 5% of autistic children suffer from FXS.6
Transcranial Magnetic Stimulation (a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability via the application of localized magnetic field pulses 7,8) represents a unique method of exploring plasticity and the manifestations of FXS within affected individuals. More specifically, Theta-Burst Stimulation (TBS), a specific stimulatory protocol shown to modulate cortical plasticity for a duration up to 30 minutes after stimulation cessation in healthy populations, has already proven an efficacious tool in the exploration of abnormal plasticity.9,10 
Recent studies have shown the effects of TBS last considerably longer in individuals on the autistic spectrum - up to 90 minutes.11 This extended effect-duration suggests an underlying abnormality in the brain's natural plasticity state in autistic individuals - similar to the hyperexcitability induced by Fragile X Syndrome.
In this experiment, utilizing single-pulse motor-evoked potentials (MEPs) as our benchmark, we will explore the effects of both intermittent and continuous TBS on cortical plasticity in individuals suffering from FXS and individuals on the Autistic Spectrum.

In this article, we examine the effects of Theta-Burst TMS stimulation on cortical plasticity in individuals suffering from Fragile X syndrome and individuals on the autistic spectrum.

TMS: Utilisation du protocole Theta-Burst à explorer Mechasnism de la plasticité chez les individus atteints du syndrome du X fragile et autisme

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS ...

Recherche

Neurosciences

The NeuroStar TMS Device: Conducting the FDA Approved Protocol for Treatment of Depression

The Neuronetics NeuroStar Transcranial Magnetic Stimulation (TMS) System is a class II medical device that produces brief duration, pulsed magnetic fields. These rapidly alternating fields induce electrical currents within localized, targeted regions of the cortex which are associated with various physiological and functional brain changes.1,2,3
In 2007, O'Reardon et al., utilizing the NeuroStar device, published the results of an industry-sponsored, multisite, randomized, sham-stimulation controlled clinical trial in which 301 patients with major depression, who had previously failed to respond to at least one adequate antidepressant treatment trial, underwent either active or sham TMS over the left dorsolateral prefrontal cortex (DLPFC). The patients, who were medication-free at the time of the study, received TMS five times per week over 4-6 weeks.4
The results demonstrated that a sub-population of patients (those who were relatively less resistant to medication, having failed not more than two good pharmacologic trials) showed a statistically significant improvement on the Montgomery-Asberg Depression Scale (MADRS), the Hamilton Depression Rating Scale (HAMD), and various other outcome measures. In October 2008, supported by these and other similar results5,6,7, Neuronetics obtained the first and only Food and Drug Administration (FDA) approval for the clinical treatment of a specific form of medication-refractory depression using a TMS Therapy device (FDA approval K061053).
In this paper, we will explore the specified FDA approved NeuroStar depression treatment protocol (to be administered only under prescription and by a licensed medical profession in either an in- or outpatient setting).

In this article, we examine the methodology and considerations relevant to the FDA approved depression treatment protocol using the Neuronetics NeuroStar TMS device.

Le dispositif NeuroStar TMS: Mener la FDA a approuvé protocole pour le traitement de la dépression

The Neuronetics NeuroStar Transcranial Magnetic Stimulation (TMS) System is a class II medical device that produces brief duration, pulsed magnetic ...

Electrode Positioning and Montage in Transcranial Direct Current Stimulation

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a non-invasive and safe alternative to change cortical excitability2. The effects of one session of tDCS can last for several minutes, and its effects depend on polarity of stimulation, such as that cathodal stimulation induces a decrease in cortical excitability, and anodal stimulation induces an increase in cortical excitability that may last beyond the duration of stimulation6. These effects have been explored in cognitive neuroscience and also clinically in a variety of neuropsychiatric disorders &#x2013; especially when applied over several consecutive sessions4. One area that has been attracting attention of neuroscientists and clinicians is the use of tDCS for modulation of pain-related neural networks3,5. Modulation of two main cortical areas in pain research has been explored: primary motor cortex and dorsolateral prefrontal cortex7. Due to the critical role of electrode montage, in this article, we show different alternatives for electrode placement for tDCS clinical trials on pain; discussing advantages and disadvantages of each method of stimulation.

Transcranial direct current stimulation (tDCS) is an established technique to modulate cortical excitability1,2. It has been used as an investigative tool in neuroscience due to its effects on cortical plasticity, easy operation, and safe profile. One area that tDCS has been showing encouraging results is pain alleviation 3-5.

Positionnement des électrodes et de montage de stimulation transcrânienne Direct Current

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Visualisation des propriocepteurs dans<em&gt; Drosophila</emLes larves&gt; et pupes

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Fibre optique d&#39;implantation pour la stimulation chronique optogenetic de tissu cérébral

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

Stimulation optogenetic de comportement de fuite en<em&gt; Drosophila melanogaster</em

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

In vivo optogénétiques stimulation du système nerveux central pour rongeurs

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Stimulation optogenetic du nerf auditif

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective treatment for several movement disorders that have failed to respond to medication. Recent progress in the field of DBS surgery has begun to extend the application of this surgical technique to other conditions as diverse as morbid obesity, depression and obsessive compulsive disorder. Despite these expanding indications, little is known about the underlying physiological mechanisms that facilitate the beneficial effects of DBS surgery. One approach to this question is to perform gene expression analysis in neurons that receive the electrical stimulation. Previous studies have shown that neurogenesis in the rat dentate gyrus is elicited in DBS targeting of the anterior nucleus of the thalamus1. DBS surgery targeting the ATN is used widely for treatment refractory epilepsy. It is thus of much interest for us to explore the transcriptional changes induced by electrically stimulating the ATN. In this manuscript, we describe our methodologies for stereotactically-guided DBS surgery targeting the ATN in adult male Wistar rats. We also discuss the subsequent steps for tissue dissection, RNA isolation, cDNA preparation and quantitative RT-PCR for measuring gene expression changes. This method could be applied and modified for stimulating the basal ganglia and other regions of the brain commonly clinically targeted. The gene expression study described here assumes a candidate target gene approach for discovering molecular players that could be directing the mechanism for DBS.

The mechanism underlying the therapeutic effects of Deep Brain Stimulation (DBS) surgery needs investigation. The methods presented in this manuscript describe an experimental approach to examine the cellular events triggered by DBS by analyzing the gene expression profile of candidate genes that can facilitate neurogenesis post DBS surgery.

Analyse des modifications d&#39;expression génique dans l&#39;hippocampe de rat après stimulation cérébrale profonde de l&#39;antérieur thalamique Nucleus

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective ...

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These neurotropic cancer cells have a unique ability to migrate unidirectionally along nerves towards the central nervous system (CNS). The dorsal root ganglia (DRG)/cancer cell model is a three dimensional (3D) in vitro model frequently used for studying the interaction between neural stroma and cancer cells. In this model, mouse or human cancer cell lines are grown in ECM adjacent to preparations of freshly dissociated cultured DRG. In this article, the DRG isolation protocol from mice, and implantation in petri dishes for co-culturing with pancreatic cancer cells are demonstrated. Five days after implantation, the cancer cells made contact with the DRG neurites. Later, these cells formed bridgeheads to facilitate more extensive polarized, neurotropic migration of cancer cells.

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

This video article shows the use of the dorsal root ganglia (DRG)/cancer cell model in pancreatic ductal adenocarcinoma.

In Vitro Modélisation de Cancerous Neural Invasion: Le ganglion de racine dorsale Modèle

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These ...