Finanziamento: Questo lavoro è stato sostenuto da una sovvenzione della Fondazione Bertarelli (DJL), una borsa di MED-EL (DJL), e un National Institutes of Health sovvenzioni DC01089 (MCB).

NOTA: Tutte le procedure sperimentali sono eseguite in conformità con la cura e l&#39;uso degli animali Comitato del Massachusetts Eye and Ear Infirmary e Harvard Medical School, che seguono le linee guida nazionali per la cura degli animali, tra cui la politica di sanità pubblica Service Humane cura e l&#39;uso di animali da laboratorio, la ILAR Guida, e la legge sulla protezione degli animali. Procedure sperimentali elencate di seguito dettagliato esposizione della DCN di sinistra. Usare strumenti sterili durante l&#39;esecuzione di un intervento chirurgico di sopravvivenza. 1. primario Craniotomia e dorsale cocleare Nucleus Esposizione <ol><li> Anestesia <ol><li> Anestetizzare un mouse, età 8-12 settimane, del peso di 18-24 g, con xilazina 20 mg / kg e 100 mg ketamina / kg tramite somministrazione intraperitoneale. Verificare il corretto anestesia per il monitoraggio della frequenza cardiaca, frequenza respiratoria, nonché punta ritiro pizzico reflex. Luogo veterinario pomata sugli occhi per prevenire la secchezza mentre sotto anestesia. Una volta adeguatamente sotto anestesia, capelli barba sovrastante tegli cuoio capelluto per fornire libero accesso al sito chirurgico. </li></ol></li><li> Posizionamento chirurgico <ol><li> Posizionare il mouse saldamente in un piccolo supporto stereotassico animali, tenuto in posizione da un morsetto muso. </li><li> Assicurarsi che il morsetto muso è abbastanza largo per permettere un&#39;adeguata respirazione, ma abbastanza stretto per immobilizzare completamente la testa del mouse. Se la testa del mouse è allentato, l&#39;animale rischia di cadere dal supporto testa durante la parte craniotomia di procedura. </li><li> Posizionare uditive elettrodi impianto tronco in modo standard, che consente il monitoraggio della frequenza cardiaca. La frequenza respiratoria deve essere monitorata da visualizzazione. Si noti che la frequenza cardiaca normale e la frequenza respiratoria variano a seconda dell&#39;età del mouse. </li><li> Un termometro è posto e eutermia è assicurata tramite il posizionamento su una coperta riscaldamento omeotermi. </li></ol></li><li> Incisione cutanea e l&#39;esposizione delle ossa interparietale e occipitale del cranio <ol><li> Sotto un microscOPE, fare una incisione verticale attraverso la pelle a partire dalla linea mediana, direttamente tra il padiglione auricolare, e si estende alla porzione caudale dell&#39;occipite. Dopo mediana pelle incisione, spostare la pelle lateralmente. Visualizza i muscoli che coprono la sinistra la faccia posteriore del cranio. </li><li> Rimuovere sovrastante parietale sinistra, interparietale muscolare, e le ossa occipitali del cranio per disarticolazione di inserzioni muscolari ai rispettivi ossa con un bisturi o iris forbice. Osservare un piccolo grado di sanguinamento lungo i bordi muscolari taglio. Minimizzare questo con una leggera pressione con un applicatore con punta di cotone per 10-15 sec. </li></ol></li><li> Sovrastante Craniotomy Cochlear Nucleus <ol><li> Identificare linee di sutura pertinenti, tra cui suture sagittale e lambda linee (Figura 1, pannello di sinistra). </li><li> Utilizzando rongeurs, fare una craniotomia sopra l&#39;osso interparietale, a sinistra di linea mediana, ~ 2 mm caudale alla linea lambda sutura. Questa regione si sovrappone il DCN. </li><li> Following craniotomia, osservare un sottile strato di dura che sovrasta il cervelletto. Usando un bisturi, rimuovere la dura. (Figura 1, pannello centrale). NOTA: La rimozione di dura può causare un piccolo grado di sanguinamento. Questo processo di utilizzare il bisturi per rimuovere la dura lo scopo di ridurre la quantità di perdita di sangue durante l&#39;aspirazione del cervelletto, che riunirà in superficie tronco encefalico e oscura l&#39;identificazione del DCN. Volume di sangue totale di mouse è ~ 1,5 ml non deve superare i&gt; del 15%. Se l&#39;emorragia supera tale importo, il mouse deve essere provvisto di integrazione con il fluido. </li><li> Rimuovere sangue coagulato sovrapponendo il cervelletto con il cotton-fioc. Facoltativamente, gocciola delicatamente salina con un punto dentale sul cervelletto per eliminare il sangue via il sangue. </li></ol></li><li> Cerebellare Aspirazione <ol><li> Utilizzando una aspirazione 5 French, aspirare la porzione laterale più a sinistra del cervelletto sovrastante il DCN finché il DCN è visUAL (Figura 1, pannello di destra). Rimuovere circa 1/4 a 1/3 del cervelletto sinistro. Il punto di riferimento principale adiacente al DCN è l&#39;ampolla del canale semicircolare superiore. NOTA: L&#39;aspirazione del cervelletto è il passo fondamentale del protocollo. Diretto aspirazione del cervelletto viene eseguita con maggior successo se si verifica in un unico tentativo, e non con più passaggi di aspirazione ciò causerebbe sanguinamento. Per facilitare l&#39;aspirazione, impostare il piano focale del microscopio alla profondità prevista della NC, che sarà leggermente distale alla superficie del cervelletto. Impostazione del piano focale per il CN migliorare la visualizzazione e di garantire un&#39;immagine nitida. </li><li> Dopo l&#39;aspirazione, ulteriori emorragie, liquido cerebrospinale (CSF) accumulo, e lo spostamento del cervelletto sulla DCN sono attesi. Instillare 0,5 cc di soluzione fisiologica sterile rapidamente nel craniotomia per evitare la coagulazione del sangue. Una combinazione di dolce tamponando con un punto dentale e di aspirazione può quindi essereutilizzata per eliminare un percorso per la visualizzazione diretta del DCN. Non contattare direttamente la superficie del DCN. </li></ol></li></ol> 2. Pressione microiniezione di trasferimento genico Virus-mediata e ripristino chirurgico <ol><li> Dopo l&#39;DCN è libera di sangue sovrastante e CSF ed è chiaramente visibile, fare microiniezioni pressione nel DCN utilizzando una siringa da 10 microlitri Hamilton un periodo di 2 min. </li><li> Introdurre l&#39;ago con un micromanipolatore finché la punta non è più visibile sotto la superficie della DCN. </li><li> Per ottenere prestazioni ottimali, utilizzare un ago calibro 33 o 34 (utilizzare una siringa a tenuta di gas con l&#39;ago 34 gauge) con uno smusso superficiale, come ad esempio 45 °, per ridurre al minimo il trauma smussato e localizzare il volume di iniezione ai DCN, che è un sottile e struttura tronco superficiale (&lt;300 micron di spessore). NOTA: Altri strumenti microiniezione possono basarsi utilizzato. Idealmente, lo strumento di iniezione deve essere flessibile per consentire qualche lieve bend, se necessario di indirizzare direttamente il DCN. Inoltre, come il mouse si sposta un po &#39;tutta la procedura a causa di respirazione, lo strumento dovrebbe essere abbastanza robusta per sopportare piccole quantità di movimento. </li></ol> 3. Recupero Surgical <ol><li> Subito dopo l&#39;iniezione, ri-approssimare la pelle e lasciare il mouse per recuperare per procedura di recupero standard. Rimanendo cervelletto e tessuto cicatriziale riempirà nella cavità causata dalla aspirazione del cervelletto. Monitorare costantemente animali fino coscienza sufficiente riguadagnato a mantenere decubito sternale. Monitorare la frequenza cardiaca, frequenza respiratoria, nonché la capacità di nutrire. NOTA: Nessun animale che ha subito un intervento chirurgico viene restituita una gabbia con altri animali fino alla completa guarigione. </li><li> Se un mouse comincia a camminare in cerchio post-operatorio (&lt;5% dei casi), tipicamente 2-4 ore dopo la chiusura della incisione cutanea, sacrificare immediatamente il mouse, come sarebbe probabilmente un&#39;alimentazione momento difficile. Til suo effetto collaterale è probabilmente dovuto all&#39;aspirazione del cervelletto. L&#39;animale deve essere monitorata per il dolore postoperatorio, e narcotici appropriate deve essere somministrato a garantire il comfort degli animali sulla base di norme istituzionali. Gli antibiotici possono essere necessarie, se le prove di infezione. </li></ol> 4. secondaria Craniotomia e Cochlear Nucleus Esposizione <ol><li> Dopo 2-4 settimane di guarigione e virus-mediata gene trasferimento incubazione, ri-anestetizzare un precedentemente iniettato mouse e ripetere il passo 1,1-1,5 tempo di incubazione ottimale varia con il tipo di gene trasferito. </li><li> Rimuovere il tessuto cicatriziale sul sito craniotomia da una combinazione di pinzette, bisturi, e rongeurs. </li><li> Dopo visualizzare la craniotomia, individuare una combinazione di rimanere cervelletto e tessuto cicatriziale sovrastante il DCN. Utilizzare un 5 di aspirazione francese per aspirare cervelletto sovrastante / tessuto cicatriziale (simile al precedente aspirazione cerebellare). </li><li> Dopo l&#39;aspirazione, si aspettano ulteriori emorragie, CSF build-up, e rimanenti bit di cervelletto. Introdurre rapidamente salina nel craniotomia per prevenire la coagulazione del sangue. Utilizzare una combinazione di dolce punto tamponando e di aspirazione dentale per spianare la strada per la visualizzazione diretta del DCN. </li><li> Osservare la superficie della DCN. Effettuare esperimenti di fisiologia basato optogenetics-come DCN è ora accessibile. Introdurre un stimolo luminoso, ad esempio con una fibra ottica (Figura 2), per guidare un esperimento basato optogenetics-. </li></ol> 5. Istologia <ol><li> Dopo la conclusione degli esperimenti, eutanasia il mouse con una overdose di ketamina. Profumato il mouse con soluzione fisiologica seguita da 4% paraformaldeide. </li><li> Estrarre il tronco cerebrale dal cranio e post-fix per 2 ore. Cryoprotect tronco encefalico nel 30% di saccarosio per 24-48 ore. Sezione tronco cerebrale utilizzando un criostato standard utilizzando 60 sezioni micron. </li></ol>

<ol>
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Questo documento descrive la tecnica di visualizzazione diretta del DCN nel modello murino per la manipolazione del sistema uditivo centrale. L&#39;approccio delineato di visualizzazione diretta offre notevoli vantaggi rispetto l&#39;alternativa principale, che sono approcci stereotassica. In primo luogo, la visualizzazione diretta della DCN consente conferma immediata del sito del tronco encefalico, considerando approcci stereotassica non danno visualizzazione diretta. In esperimenti che necessitano prolungati periodi ...

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>
			<td style="width:227px;">Fine Surgical Tools</td>
			<td>10011-00</td>
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			<td height="21" style="height:21px;width:197px;">Iris scissor</td>
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			<td>14084-08</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>
		</tr>
		<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

Indagine sull&#39;uso di trasferimento genico virus-mediata di arrestare o invertire la perdita dell&#39;udito è stata in gran parte relegato al sistema uditivo periferico. Pochi studi hanno esaminato il trasferimento di geni al sistema uditivo centrale. Il nucleo dorsale cocleare (DCN) del tronco encefalico, che contiene neuroni secondo ordine del percorso uditivo, è un sito di capacità di trasferimento genico. In questo protocollo, una tecnica per l&#39;esposizione diretta e massima della murino DCN mediante un approccio posteriore fossa è dimostrata. Questo approccio permette sia per la chirurgia acuta o sopravvivenza. Dopo la visualizzazione diretta del DCN, una serie di esperimenti sono possibili, compresa l&#39;iniezione di opsine nel nucleo cocleare e la stimolazione successiva da una fibra ottica accoppiata ad un laser a luce blu. Altri esperimenti di neurofisiologia, come la stimolazione elettrica e tracciati iniettori neurali sono anche possibili. Il livello di Visualizazione e la durata della stimolazione rendono realizzabile questo approccio applicabile ad una vasta gamma di esperimenti.

Parziale cerebellare aspirazione possa accedere alla Cochlear Nucleus Dopo la pelle ei muscoli sovrastante il cranio vengono rimossi, monumenti superficie del cranio, come le linee di sutura coronale e Lamda, dimostrano la localizzazione approssimativa della craniotomia. A seguito di craniotomia con rongeurs, il cervelletto è visualizzato. Accurata aspirazione della piccola porzione del cervelletto dimostra visualizzazione della NC, che possono poi essere iniettato (Figura ...

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.

Visualizzazione diretta del Murine Dorsale Cochlear Nucleus per optogenetic stimolazione del percorso uditivo

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 Views.  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. 

Video: 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: Utilizzo del Theta Burst protocollo Esplora Mechasnism di plasticità in individui con Sindrome X Fragile e autismo

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 ...

Ricerca

Neuroscienze

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.

Il dispositivo TMS NeuroStar: Condurre la FDA ha approvato protocollo per il trattamento della depressione

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.

Posizionamento elettrodo e Montage stimolazione transcranica diretta in corrente

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.

Visualizzazione dei Propriorecettori in<em&gt; Drosophila</em&gt; Larve e pupe

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.

Fibra ottica Impianto per la Stimolazione Optogenetic cronica del tessuto cerebrale

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.

Stimolazione Optogenetic del comportamento in Fuga<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 optogenetic stimolazione del sistema nervoso centrale Roditore

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.

Stimolazione optogenetic del nervo uditivo

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.

Analisi delle variazioni di espressione genica nel Rat Hippocampus Dopo stimolazione cerebrale profonda del nucleo anteriore talamiche

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 Modellazione di cancerose Neural Invasion: La dorsale della radice Ganglio Modello

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 ...