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>

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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.

<table><tbody><tr><td>Stereotaxic holder</td><td>Stoelting</td><td>51500</td><td></td></tr><tr><td>Homeothermic blanket</td><td>Harvard</td><td>507214</td><td></td></tr><tr><td>Scalpel blade #11</td><td>Fine Surgical Tools</td><td>10011-00</td><td></td></tr><tr><td>Iris scissor</td><td>Fine Surgical Tools</td><td>14084-08</td><td></td></tr><tr><td>5 French suction</td><td>Symmetry Surgical</td><td>2777914</td><td></td></tr><tr><td>Dental Points</td><td>Henry Schein</td><td>100-8170</td><td></td></tr><tr><td>Bone rongeur</td><td>Fine Surgical Tools</td><td>16020-14</td><td></td></tr><tr><td>10 &amp;micro;l Hamilton syringe</td><td>Hamilton&amp;nbsp;</td><td>7633-01</td><td></td></tr><tr><td>34 gauge, needle</td><td>Hamilton&amp;nbsp;</td><td>207434</td><td></td></tr><tr><td>Rongeurs</td><td>Fine Surgical Tools</td><td>16021-14</td><td></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

9.2K 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

Canalostomy As a Surgical Approach to Local Drug Delivery into the Inner Ears of Adult and Neonatal Mice

Local delivery of therapeutic drugs into the inner ear is a promising therapy for inner ear diseases. Injection through semicircular canals (canalostomy) has been shown to be a useful approach to local drug delivery into the inner ear. The goal of this article is to describe, in detail, the surgical techniques involved in canalostomy in both adult and neonatal mice. As indicated by fast-green dye and adeno-associated virus serotype 8 with the green fluorescent protein gene, the canalostomy facilitated broad distribution of injected reagents in the cochlea and vestibular end-organs with minimal damage to hearing and vestibular function. The surgery was successfully implemented in both adult and neonatal mice; indeed, multiple surgeries could be performed if required. In conclusion, canalostomy is an effective and safe approach to drug delivery into the inner ears of adult and neonatal mice and may be used to treat human inner ear diseases in the future.

Here we describe canalostomy procedure which allows local drug delivery into the inner ears of adult and neonatal mice through the semicircular canal with minimum damage to hearing and vestibular function. This method can be used to inoculate viral vectors, pharmaceuticals, and small molecules into the mouse inner ear.

Canalostomy come un approccio chirurgico alla consegna di droga locale nelle orecchie interno dei topi adulti e neonati

Local delivery of therapeutic drugs into the inner ear is a promising therapy for inner ear diseases. Injection through semicircular canals (canalostomy) ...

Ricerca

Genetica

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Selettiva trasduzione virale di neuroni olfattivi per adulti nati per cronica In vivo Optogenetic Stimolazione

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

Neuroscienze

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

Laser-scanning Fotostimolazione di circuiti prosencefalo Optogenetically mirati

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

Cultura primaria e elettroporazione plasmidi dell&#39;Organo murini di Corti.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

Biologia

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 ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

Una guida per<em&gt; In vivo</em&gt; Singola unità di registrazione da Optogenetically Identificato corticale inibitorio interneuroni

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

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 ...

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a condition that typically involves damage to or loss of the delicate mechanosensory structures of the inner ear. Sophisticated in vitro and ex vivo assays have emerged in recent years, enabling the screening of an increasing number of potentially therapeutic compounds while minimizing resources and accelerating efforts to develop cures for SNHL. Though homogenous cultures of certain cell types continue to play an important role in current research, many scientists now rely on more complex organotypic cultures of murine inner ears, also known as cochlear explants. The preservation of organized cellular structures within the inner ear facilitates the in situ evaluation of various components of the cochlear infrastructure, including inner and outer hair cells, spiral ganglion neurons, neurites, and supporting cells. Here we present the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The careful preparation of these explants facilitates the identification of mechanisms that contribute to SNHL and constitutes a valuable tool for the hearing research community.

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

The goal of this protocol is to demonstrate the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The technique can be utilized as an in vitro screening tool in hearing research.

Neonatal Murine Cochlear Explant Technique come un<em&gt; In vitro</em&gt; Strumento di screening nella ricerca acustica

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a ...

Medicina

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

Preparazione della fetta di cuneo in vitro per imitare la connettività del circuito neuronale in vivo

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...

Preparing Wedge Brain Slices to Visualize Auditory Neurons

Source: Fischl, M. J. et al., <a href="https://app.jove.com/v/61664">In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity.</a>&#160;J. Vis. Exp.&#160;(2020)
This video demonstrates the process of preparing wedge-shaped brain slices in a laboratory setting. Using a vibratome, a wedge-shaped slice from a mouse&#39;s brain is crafted with varying thickness. This slice contains the auditory nerve root positioned on the thicker end, while the hearing-related neurons are located on the thinner end.

Preparazione di fette di cervello a cuneo per visualizzare i neuroni uditivi

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Brain ...

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

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