Name: direct visualization of the murine dorsal cochlear nucleus for optogenetic stimulation of the auditory pathway
Uploaded: 2015-01-20T15:00:00.000+00:00
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Description: Watch this Scientific Journal Video about Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway at JoVE.com

Financiación: Este trabajo fue apoyado por una beca de la Fundación Bertarelli (DJL), un MED-EL subvención (DJL), y los Institutos Nacionales de Salud subvenciones DC01089 (MCB).

NOTA: Todos los procedimientos experimentales se realizan de acuerdo con el Comité de Cuidado de Animales y el empleo de la Massachusetts Eye and Ear Infirmary y la Escuela Médica de Harvard, que siguen las directrices nacionales de cuidado de animales, incluyendo la Política Pública de Salud Servicio de Cuidado Humano y Uso de Animales de laboratorio, la Guía ILAR, y la Ley de Protección de los Animales. Los procedimientos experimentales indican a continuación la exposición detallada de la DCN izquierda. Utilice instrumentos estériles mientras se realiza la cirugía de supervivencia. 1. Craneotomía Primaria y Dorsal coclear Núcleo exposición <ol><li> Anestesia <ol><li> Anestesiar a un ratón, a la edad de 8-12 semanas, con un peso de 18-24 g, con xilazina 20 mg / kg de ketamina y 100 mg / kg a través de una administración intraperitoneal. Confirmar la anestesia adecuada por la tasa de monitorización cardiaca, frecuencia respiratoria, así como reflejo de retirada pizca dedo del pie. Lugar veterinario ungüento en los ojos para evitar la sequedad mientras que bajo anestesia. Una vez adecuadamente bajo anestesia, el pelo afeitado recubre tque el cuero cabelludo para proporcionar un acceso sin obstrucciones a la zona quirúrgica. </li></ol></li><li> Posicionamiento quirúrgico <ol><li> Coloca el ratón de forma segura en un pequeño titular estereotáxica animal, sujeto por una abrazadera hocico. </li><li> Asegúrese de que la abrazadera hocico es lo suficientemente floja para permitir la respiración adecuada, pero lo suficientemente apretado para inmovilizar por completo la cabeza del ratón. Si la cabeza del ratón está suelto, el animal es probable que caiga fuera del soporte de cabeza durante la parte craneotomía de procedimiento. </li><li> Coloque los electrodos del implante auditivo de tronco cerebral en forma estándar, lo que permite la monitorización de la frecuencia cardíaca. La frecuencia respiratoria debe ser supervisada por la visualización. Tenga en cuenta que la frecuencia cardíaca normal y la frecuencia respiratoria variarán dependiendo de la edad del ratón. </li><li> Un termómetro se coloca y euthermia está asegurada a través de la colocación de una manta de calefacción homeotérmico. </li></ol></li><li> Incisión de la piel y la exposición de los huesos interparietales y occipital del cráneo <ol><li> Bajo un Microscopa, hacer una incisión vertical a través de la piel a partir de la línea media, directamente entre el pabellón de la oreja, y se extiende hasta la porción caudal del occipucio. Después de incisión en la piel en la línea media, desplazar lateralmente la piel. Visualizar los músculos que cubren la izquierda de la cara posterior del cráneo. </li><li> Retire el músculo que recubre el parietal izquierdo, interparietal, y los huesos occipitales del cráneo por la desarticulación de las inserciones musculares a sus respectivos huesos con un bisturí o tijera iris. Observar un pequeño grado de sangrado a lo largo de los bordes musculares corte. Minimizar esta por presión suave con un aplicador con punta de algodón para 10-15 seg. </li></ol></li><li> Suprayacente craneotomía coclear Núcleo <ol><li> Identificar las líneas de sutura pertinentes, incluidas las líneas de suturas sagital y lambda (Figura 1, panel izquierdo). </li><li> Usando unas pinzas, hacer una craneotomía sobre el hueso interparietal, izquierda de la línea media, ~ 2 mm caudal a la línea de sutura lambda. Esta región se superpone a la DCN. </li><li> Following craneotomía, observar una fina capa de duramadre que recubre el cerebelo. Usando una hoja de bisturí, retire la duramadre. (Figura 1, el panel medio). NOTA: La eliminación de la duramadre puede causar un pequeño grado de sangrado. Este proceso de usar la hoja de bisturí para eliminar la duramadre está destinado a reducir la cantidad de pérdida de sangre durante la aspiración cerebelo, que será común en la superficie del tronco cerebral y la identificación oscura de la DCN. El volumen de sangre total de ratón es ~ 1,5 ml no deben exceder&gt; al 15%. Si el sangrado es superior a esta cantidad, el ratón debe estar provista con suplementos de fluido. </li><li> Retire la sangre coagulada superponer el cerebelo con el aplicador con punta de algodón. Opcionalmente, escurrir suavemente salina utilizando una punta dental en el cerebelo para limpiar la sangre fuera sangre. </li></ol></li><li> Cerebelosa Aspiración <ol><li> Usando una succión Francés 5, aspirar la porción lateral más del cerebelo izquierdo que recubre el DCN hasta que el DCN es visual (Figura 1, panel derecho). Retire aproximadamente 1/4 a 1/3 del cerebelo izquierdo. El hito principal adyacente a la DCN es la ampolla del canal semicircular superior. NOTA: La aspiración del cerebelo es el paso clave del protocolo. Dirigida aspiración del cerebelo se lleva a cabo con mayor éxito si se produce en un solo intento y no con varias pasadas de la succión, ya que esto causar sangrado. Para ayudar en la aspiración, definir el plano focal del microscopio a la profundidad prevista de la NC, que será ligeramente distal a la superficie del cerebelo. Ajuste del plano focal a la CN mejorará la visualización y asegurar una imagen nítida. </li><li> Se espera que después de la aspiración, más sangrado, el líquido cefalorraquídeo (LCR) la acumulación y el desplazamiento cerebelo en la DCN. Inculcar 0,5 cc de solución salina estéril rápidamente en la craneotomía para prevenir la coagulación de la sangre. Una combinación de dabbing suave con un punto dental y la succión puede ser entoncesutilizado para despejar un camino para la visualización directa de la DCN. No entre en contacto directo con la superficie de la DCN. </li></ol></li></ol> 2. La microinyección de presión para la transferencia génica mediada por virus y la recuperación quirúrgica <ol><li> Después de la DCN es libre de sangre suprayacente y CSF y es claramente visible, hacer microinyecciones de presión en el DCN utilizando una jeringa Hamilton de 10 l durante un período de 2 min. </li><li> Introducir la aguja con un micromanipulador hasta que la punta ya no es visible bajo la superficie de la DCN. </li><li> Para un rendimiento óptimo, utilice una aguja de calibre 33 o 34 (utilizar una jeringa estanca a los gases con la aguja de calibre 34) con un bisel de poca profundidad, tales como 45 °, para reducir al mínimo el traumatismo cerrado y localizar el volumen de inyección dentro de la DCN, que es un delgado y estructura del tronco cerebral profunda (&lt;300 micras de espesor). NOTA: Otros instrumentos de microinyección pueden basarse utilizado. Idealmente, el instrumento de inyección debe ser flexible para permitir una ligera bend, si es necesario apuntar directamente a la DCN. Además, como el ratón se mueve ligeramente durante todo el procedimiento debido a la respiración, el instrumento debe ser lo suficientemente robusta como para soportar pequeñas cantidades de movimiento. </li></ol> 3. Recuperación Quirúrgica <ol><li> Inmediatamente después de la inyección, vuelva a la aproximación de la piel y permitir el ratón para recuperar por procedimiento de recuperación estándar. Restante cerebelo y tejido cicatrizal se llenará en la cavidad causada por la aspiración del cerebelo. Constantemente observar a los animales hasta que la conciencia suficiente se recuperó para mantener decúbito esternal. Monitorear la frecuencia cardiaca, la frecuencia respiratoria, así como la capacidad para alimentarse. NOTA: No animal que ha sido sometido a cirugía es devuelto a una jaula con otros animales hasta que se recupere totalmente. </li><li> Si un ratón comienza a caminar en círculos después de la intervención (&lt;5% de los casos), típicamente 2-4 horas después del cierre de la incisión de la piel, sacrificar inmediatamente el ratón, ya que probablemente tendría una alimentación momento difícil. Tsu efecto secundario es probable debido a la aspiración del cerebelo. El animal debe ser supervisada para el dolor después de la operación, y los narcóticos también se debe prestar para asegurar el confort de los animales sobre la base de las normas institucionales. Los antibióticos pueden ser necesarios, si la evidencia de infección. </li></ol> 4. Secundaria exposición craneotomía y coclear Núcleo <ol><li> Después de 2-4 semanas de curación y el gen de incubación transferencia mediada por virus, volver a anestesiar un ratón y repita el paso 1.1 a 1.5 tiempo de incubación óptimo previamente inyectado varía con el tipo de gen transferido. </li><li> Quitar el tejido de la cicatriz sobre el sitio de la craneotomía por una combinación de pinzas, bisturí, y rongeurs. </li><li> Después de visualizar la craneotomía, identificar una combinación de permanecer cerebelo y tejido cicatrizal que recubre la DCN. Utilice una succión Francés 5 para aspirar cerebelo suprayacente / tejido de la cicatriz (similar a la aspiración cerebelar anterior). </li><li> Después de la aspiración, le espera un sangrado posterior, CSF build-up, y el resto de bits de cerebelo. Introducir rápidamente salina en la craneotomía para evitar la coagulación de la sangre. Use una combinación de suave dabbing punto dental y de succión para despejar el camino para la visualización directa de la DCN. </li><li> Observar la superficie de la DCN. Realizar experimentos de fisiología con sede en la optogenética como DCN es ahora accesible. Introducir un estímulo de luz, por ejemplo, con una fibra óptica (Figura 2), para conducir un experimento basado en la optogenética. </li></ol> 5. Histología <ol><li> Tras la conclusión de los experimentos, sacrificar al ratón con una sobredosis de ketamina. Perfundir el ratón con solución salina normal seguido de paraformaldehído al 4%. </li><li> Extraer el tronco cerebral del cráneo y post-fix durante 2 horas. Cryoprotect el tronco cerebral en el 30% de sacarosa durante 24 a 48 horas. Sección del tronco cerebral usando un criostato estándar con 60 micras secciones. </li></ol>

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En este trabajo se describe la técnica de la visualización directa de la DCN en el modelo murino para la manipulación del sistema auditivo central. El enfoque esbozado de visualización directa ofrece ventajas significativas sobre la principal alternativa, que son enfoques estereotáxica. Principalmente, la visualización directa de la DCN permite confirmación inmediata del sitio del tronco cerebral, mientras que los enfoques estereotáxicas no proporcionan visualización directa. En experimentos que requieren prolo...

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

Investigación sobre el uso de la transferencia génica mediada por virus para detener o revertir la pérdida de la audición en gran medida ha sido relegado al sistema auditivo periférico. Pocos estudios han examinado la transferencia de genes al sistema auditivo central. El núcleo coclear dorsal (DCN) del tronco cerebral, que contiene las neuronas de segundo orden de la vía auditiva, es un sitio potencial para la transferencia de genes. En este protocolo, una técnica para la exposición directa y máxima de la murino DCN a través de un enfoque de la fosa posterior se demuestra. Este enfoque permite ya sea la cirugía aguda o supervivencia. Después de la visualización directa de la DCN, una serie de experimentos son posibles, incluyendo la inyección de opsinas en el núcleo coclear y la estimulación posterior por una fibra óptica acoplada a un láser de luz azul. Otros experimentos neurofisiología, como la estimulación eléctrica y trazados inyectores neuronales también son factibles. El nivel de Visualización y la duración de la estimulación alcanzable hacen de este enfoque aplicable a una amplia gama de experimentos.

Parcial cerebelosa Aspiración demuestre tener acceso a la coclear Núcleo Después de la piel y el músculo que recubre el cráneo se retiran, superficie cráneo puntos de referencia, tales como las líneas de sutura coronal y Lamda, demostrar la localización aproximada de la craneotomía. Siguiendo craneotomía con gubia, el cerebelo se visualiza. La aspiración cuidadosa de la pequeña porción del cerebelo demuestra la visualización de la CN, que luego se puede inyectar <stron...

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.

La visualización directa de la murino Dorsal coclear Núcleo de optogenética La estimulación de la vía auditiva

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

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

9.2K vistas.  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 como un abordaje quirúrgico a la entrega del Local de la droga en los oídos interiores de ratones adultos y Neonatal

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

Investigación

Genética

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.

Transducción viral selectiva de adultos nacidos en las neuronas olfativas de las enfermedades crónicas En vivo Estimulación Optogenetic

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

Neurociencias

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.

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 y electroporación plásmido del órgano de Corti murino.

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 ...

Biología

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 optogenético la estimulación del sistema nervioso central de roedores

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 guía para<em&gt; En vivo</em&gt; Single-unidad de grabación de Optogenetically Identificado cortical inhibitorio interneuronas

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.

Estimulación Optogenética del nervio auditivo

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.

Técnica de Explante Coclear Murino Neonatal como<em&gt; In Vitro</em&gt; Herramienta de detección en la investigación de la audición

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.

In Vitro Wedge Slice Preparación para imitar la conectividad del circuito neuronal 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://appjove.unicartagenaproxy.elogim.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.

Preparación de cortes de cerebro en cuña para visualizar neuronas auditivas

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

La visualización directa de la murino Dorsal coclear Núcleo de optogenética La estimulación de la vía auditiva

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