<p class="jove_content"&gt; Este trabajo fue apoyado por el programa intramural de los Institutos Nacionales de Trastornos Neurológicos y Accidentes Cerebrovasculares de los Institutos Nacionales de Salud. También nos gustaría agradecer al Dr. George Mentis por sus contribuciones anteriores al método y los datos de la Figura 3.</p

Dextranos fluorescentes se han utilizado como herramientas anatómicas y para la actividad neuronal imagen 1-4. Campos et al, (2009) 4 publicó un protocolo para la aplicación de iones y sensibles al voltaje tintes de vías axonales con un enfoque en la médula espinal como un sistema modelo. Aquí se describe un procedimiento más detallado para la aplicación de tinte fluorescente y / o de iones sensibles a las raíces cortadas ventrales, raíces dorsales o cualquier tracto neuronal de la médula espinal para los estudios in vitro optophysiological y morfológicas. 1. Tipo I y Tipo II Pipetas Para empezar, tirando de dos tramos cortos de tubo de polietileno (PE90, Clay Adams Marca) sobre la llama de una lámpara de alcohol 7 para producir pipetas con puntas afiladas. Una pipeta (Tipo I) debe ser corto (3-7 mm) con un pequeño consejo que bien puede contener la raíz de destino o en el tracto (Diámetro exterior: 0.2-0.4 mm; Diámetro interior: 0,1-0,3 mm). A continuación, tire de una pipeta segundo (Tipo-II) con un tiempo (8-12 cm) más delgada, el eje y una punta muy fina (diámetro exterior: 0.2-0.3 mm, diámetro interior: 0.1-0.2 mm) que se puede insertar en el Tipo I pipetear la medida de lo la punta. La pipeta segundo diluyente será utilizada para aspirar e introducir fluido cerebroespinal artificial (aCSF) y la solución colorante, respectivamente, de la pipeta Tipo-I. 2. Posicionamiento de tipo I Pipetear Disecar y aislar a la médula espinal 8,9 y colocarlo en un baño, con frío superfused aCSF (~ 16 ° C) durante todo el proceso de carga. Coloque el tipo I de la pipeta en un soporte de electrodo (H1/12 porta-electrodos, Narishige) de manera que su apertura posterior puede ser fácilmente conectado a una jeringa (1 ml jeringa U-100 a la insulina, Becton Dickenson o similar) a través de un tubo flexible (PharMed BPT, Cole-Parmer, # AY242002; 10 cm) (Fig. 1A-B). 3. Dye aplicación <ol><li> Con la jeringa, primero trazaaCSF en la pipeta Tipo-I (Fig. 1B; 2A), seguido por el tracto axonal o raíz para ser llenado (Fig. 2B). El tubo flexible debe entonces ser retirado de la abertura trasera. La pipeta de tipo II se une a otra jeringuilla y se inserta en el Tipo I pipeta de sujeción del tracto axonal o raíz. Uso de la jeringa con la pipeta de tipo II, aspirado aCSF para que el aCSF residuales sólo cubre el tracto axonal (Fig. 1c; 2C-D). Luego retirar la pipeta de tipo II a partir de la pipeta de tipo I. Controlar el nivel de aCSF durante unos minutos para verificar un buen sello en el tracto axonal o los nervios (ver Sección 3 Solución de problemas de &quot;buen cierre&quot;). </li><li> Disolver el colorante en aCSF o el 0,2% Triton X-100 en agua destilada dos veces y dibujarlo en la pipeta de tipo II, mientras que prestar atención de no incluir burbujas de aire. Insertar la pipeta de tipo II que contiene el colorante en la pipeta Tipo-I y en la solución residual aCSF (Fig. 2E). Suelte lentamenteel medio de contraste en el uso de una presión suave aCSF positivo (Fig. 2F). Tenga cuidado de no introducir burbujas de aire o para provocar el desplazamiento del tejido diana desde el interior de la punta. Retirar la pipeta de tipo II, después de una cantidad suficiente de medio de contraste ha sido liberado (3-5 l de solución de tinte). </li></ol> 4. Incubación Incubar el tejido en la oscuridad entre 6 a 20 horas, dependiendo del tipo de experimento. Una vez que el tiempo suficiente se ha permitido para el llenado, tire suavemente lejos del electrodo del tejido dejando el tejido listo para formación de imágenes o histología. Solución de problemas <ol><li> Si el tejido diana no entra fácilmente en la punta de la pipeta Tipo-I o está suelto una vez dentro, entonces el diámetro de la punta es el tamaño incorrecto. Si esto ocurre, tirar y usar una pipeta diferente. </li><li> Antes de añadir el colorante a la aCSF en la pipeta de tipo I, asegurarse de que aCSF queda suficiente para que pueda ser alcanzado con elTipo II punta de la pipeta (Fig. 2D). De lo contrario, un espacio de aire existirá una vez que el colorante se añade. Si esto sucede, no agregue el colorante y eliminar el tracto axonal de la pipeta y luego dibujarla de nuevo con aCSF suficiente para ser alcanzado por la pipeta de tipo II. </li><li> Si el nivel de aCSF en los aumentos de pipeta I Tipo-después de que se aspiró con la pipeta de tipo II, significa que el sello con el tejido no es buena. Utilizar un electrodo diferente, de lo contrario el colorante se diluyen durante el procedimiento de etiquetado. </li><li> Si una burbuja de aire se introduce en la pipeta de tipo I, mientras que la inyección del colorante, drenar la burbuja por el avance del cierre de tipo II punta de pipeta a la burbuja y suavemente aplicando una presión negativa débil con la jeringa adjunta. </li></ol> 5. Los resultados representativos Motoneuronas Para ilustrar una aplicación del método, al mismo tiempo cargadas, las fibras aferentes sensoriales y interneuronas de la médula con tres diferentes DEXTcorrió y conjugado con los tintes fluorescentes (Fig. 3). Como se ilustra en la figura. 3B, tres clases de neuronas han sido etiquetados, las fibras aferentes sensoriales (color rojo), las interneuronas se proyectan en el ventral, funículo (verde) y las neuronas motoras (azul). <img alt="Figura 1" src="/files/ftp_upload/4008/4008fig1.jpg" /> Figura 1. Esquema que muestra con una pipeta de tipo I y tipo II Asamblea. (A) Un tipo I pipeta (luz azul) se coloca en un soporte de electrodo de modo que la espalda de apertura puede ser fácilmente conectado a la tubería de elastómero flexible (B) y en el que una pipeta de tipo II se puede insertar (C). <img alt="Figura 2" src="/files/ftp_upload/4008/4008fig2.jpg" /> Figura 2. Esquema que muestra el proceso de carga de las motoneuronas con los tintes fluorescentes. (A) Una pipeta de tipo I se coloca cerca de la raíz ventral a ser llenado. (B) de succión se aplica a la pipeta para dibujar aCSF en la pipeta seguido por la raíz. (C-D) Desconectar el tubo de elastómero flexible desde el servidor de tipo I pipeta y reducir la cantidad de aCSF mediante la aplicación de succión a la pipeta de tipo II. (E) Introducir la punta de la pipeta de tipo II que contiene el colorante disuelto en el aCSF restante en la pipeta Tipo-I. (EF) Poco a poco llenar la pipeta de tipo I con el tinte y luego extraiga la pipeta de tipo II. <img alt="Figura 3" src="/files/ftp_upload/4008/4008fig3.jpg" /> Figura 3. La aplicación del procedimiento de llenado de la médula espinal de ratón aislado. (A) Esquema que muestra los tres de tipo I pipetas usadas para llenar diferentes clases de neuronas en la médula espinal. Motoneuronas (azul, Blue Cascade dextrano) y sacros aferentes sensoriales (color rojo; de Texas-Red dextrano) se rellenaron a través de las raíces ventral y dorsal, respectivamente. Fluoresceína dextrano se utiliza para cargar las interneuronas espinales (verde) cuyos axones ascienden a través del funículo ventral (FV). 1 mg de cada tinte dextrano (10.000 MW, Molecular Probes)se disolvió en 6 l de agua destilada que contenía 0,2% de Triton X-100 (B). La imagen confocal de una sección de la médula sacra de la médula espinal en la que las neuronas estaban de vuelta marcado, como se describe en la Nota A. que los aferentes sensoriales son etiquetados principalmente ipsilateral con una proyección contralateral pocos. Las motoneuronas son etiquetados ipsilateral a la raíz ventral llena mientras que las interneuronas etiquetados son contralateral al lado del relleno (cabeza de flecha). Calibración de barra de 200 m.

<ol>
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Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+functional+impairment+of+sensory-motor+connectivity+in+a+mouse+model+of+spinal+muscular+atrophy.">Early functional impairment of sensory-motor connectivity in a mouse model of spinal muscular atrophy.</a> Neuron. 69, 453-467 (2011).</li><li>Blivis, D., Mentis, Z., O'Donovan, J. M. G., Lev-Tov, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+of+sacral+neurons+involved+in+activation+of+the+lumbar+central+pattern+generator+for+locomotion+in+the+neonatal+rodent+spinal+cord.">Studies of sacral neurons involved in activation of the lumbar central pattern generator for locomotion in the neonatal rodent spinal cord.</a> Soc. Neurosci. Abstr. 564.8, (2009).</li><li>O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+the+spatiotemporal+organization+of+neural+activity+in+the+developing+spinal+cord.">Imaging the spatiotemporal organization of neural activity in the developing spinal cord.</a> Dev. Neurobiol. 68, 788-803 (2008).</li><li>Kasumacic, N., Glover, J. C., Perreault, M. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segmental+patterns+of+vestibular-mediated+synaptic+inputs+to+axial+and+limb+motoneurons+in+the+neonatal+mouse+assessed+by+optical+recording.">Segmental patterns of vestibular-mediated synaptic inputs to axial and limb motoneurons in the neonatal mouse assessed by optical recording.</a> J. Physiol. (Lond). 588, 4905-4925 (2010).</li><li>Szokol, K., Glover, J. C., Perreault, M. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+of+functional+synaptic+connections+between+medullary+reticulospinal+neurons+and+lumbar+descending+commissural+interneurons+in+the+neonatal+mouse.">Organization of functional synaptic connections between medullary reticulospinal neurons and lumbar descending commissural interneurons in the neonatal mouse.</a> J. Neurosci. 31, 4731-4742 (2011).</li></ol>

<p class="jove_content"&gt; No tenemos nada que revelar.</p

<p class="jove_content"&gt; Se describe aquí un protocolo simple y rentable para la introducción de colorantes en las neuronas, los nervios y tractos espinales. Este método consiste en exponer identificados vías anatómicas a una solución de tinte altamente concentrada para la duración del proceso de carga. Esto se traduce en el etiquetado retrógrada del sitio de destino con poca experiencia en comparación con microinyecciones, la aplicación de baños y técnicas de electroporación. Sin embargo, el método está limitado a los sitios ...

<table border="1"&gt;<tbody&gt;<tr&gt;<td colspan="2"<strong&gt; Nombre del Material</strong</td&gt;<td<strong&gt; Empresa</strong</td&gt;<td<strong&gt; Número de catálogo</strong</td&gt;<td<strong&gt; Comentarios</strong</td&gt;</tr&gt;<tr&gt;<td rowspan="2"&gt; Tubos</td&gt;<td&gt; PE90 (IDxOD: 0.034 &quot;x0.050&quot;, Espesor de pared: 0.008 &quot;)</td&gt;<td&gt; Clay Adams Marca Intermedic</td&gt;<td&gt; 427421</td&gt;<td&gt; Por tipo I y tipo II pipetas</td&gt;</tr&gt;<tr&gt;<td&gt; NSF-51 (IDxOD: 1/16x1/8; Espesor de pared: 1/32)</td&gt;<td&gt; PharMed BPT, Cole-Parmer</td&gt;<td&gt; AY242002</td&gt;<td&gt; Para los tubos flexibles</td&gt;</tr&gt;<tr&gt;<td&gt; Jeringa</td&gt;<td&gt; 1 ml jeringa de insulina U-100</td&gt;<td&gt; Becton Dickenson</td&gt;<td&gt; 329650</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Aguja</td&gt;<td&gt; 19G x1-1/2 &quot;Metal cubo de la aguja</td&gt;<td&gt; MONOJECT</td&gt;<td&gt; 200136</td&gt;<td&gt; Conexión de Tipo II pipeta a la jeringa</td&gt;</tr&gt;<tr&gt;<td&gt;</td&gt;<td&gt; El alcohol de la lámpara</td&gt;<td&gt;</td&gt;<td&gt;</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Llave de paso</td&gt;<td&gt; Llave de tres vías con adaptador Luer Slip Mujer</td&gt;<td&gt; Baxter Healthcare Corp.</td&gt;<td&gt; 2C6241</td&gt;<td&gt; Utilice con la jeringa</td&gt;</tr&gt;<tr&gt;<td rowspan="3"&gt; Soporte</td&gt;<td&gt; Titular de la H-1 electrodo</td&gt;<td&gt; Narishige</td&gt;<td&gt; H-1/12</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Imán soporte</td&gt;<td&gt; Narishige</td&gt;<td&gt; GJ-8</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Micromanipulador</td&gt;<td&gt; Narishige</td&gt;<td&gt; M-3333</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Pinzas</td&gt;<td&gt; S &amp; T DUMONT suiza 00632-11</td&gt;<td&gt; DUMONT</td&gt;<td&gt; JF-5 TC</td&gt;<td&gt;</td&gt;</tr&gt;</tbody&gt;</table&gt;<p class="jove_content"&gt; * Estos son los materiales sugeridos. Puede ser reemplazado por cualquiera de los productos compatibles</p

retrograde loading of nerves, tracts, and spinal roots with fluorescent dyes

Etiquetado retrógrada de las neuronas es un 1,2 anatómica método estándar que se ha utilizado también para cargar calcio y tintes sensibles al voltaje en neuronas 3-6. En general, los colorantes se aplican en forma de cristales sólidos o por inyección a presión local utilizando pipetas de vidrio. Sin embargo, esto puede resultar en la dilución de la intensidad colorante y etiquetado reducida, particularmente cuando se requieren varias horas para la difusión del colorante. Aquí se demuestra una técnica sencilla y de bajo costo para la introducción de colorantes fluorescentes y de iones sensibles a las neuronas con una pipeta de succión de polietileno llena de la solución de colorante. Este método ofrece una forma fiable para mantener una alta concentración del colorante en contacto con los axones en todo el procedimiento de carga.

Watch this Scientific Journal Video about Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes at JoVE.com

Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Se describe una técnica sencilla y de bajo costo para la introducción de alta concentración de colorantes fluorescentes y sensible de calcio en las neuronas o cualquier aparato neuronal con una pipeta de succión de polietileno.

Cargando retrógrada de los nervios, panfletos, y las raíces espinales con tintes fluorescentes

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. ...

retrograde-loading-nerves-tracts-spinal-roots-with-fluorescent

Research

JoVE Journal

Neuroscience

12.2K vistas.  National Institute of Neurological Disorders and Stroke, National Institutes of Health. Se describe una técnica sencilla y de bajo costo para la introducción de alta concentración de colorantes fluorescentes y sensible de calcio en las neuronas o cualquier aparato neuronal con una pipeta de succión de polietileno.

Video: Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

El estudio de Piscinas vesícula sináptica con fotoconversión de tintes estirilo

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Investigación

Neurociencias

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Imagen óptica de neuronas en el ganglio Cangrejo Stomatogastric con tintes sensibles al voltaje

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

Marcaje fluorescente retrógrada Permite dirigida extracelular sola unidad de grabación de neuronas identificadas<em&gt; En vivo</em

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Promotion of Survival and Differentiation of Neural Stem Cells with Fibrin and Growth Factor Cocktails after Severe Spinal Cord Injury

Neural stem cells (NSCs) can self-renew and differentiate into neurons and glia. Transplanted NSCs can replace lost neurons and glia after spinal cord injury (SCI), and can form functional relays to re-connect spinal cord segments above and below a lesion. Previous studies grafting neural stem cells have been limited by incomplete graft survival within the spinal cord lesion cavity. Further, tracking of graft cell survival, differentiation, and process extension had not been optimized. Finally, in previous studies, cultured rat NSCs were typically reported to differentiate into glia when grafted to the injured spinal cord, rather than neurons, unless fate was driven to a specific cell type. To address these issues, we developed new methods to improve the survival, integration and differentiation of NSCs to sites of even severe SCI. NSCs were freshly isolated from embryonic day 14 spinal cord (E14) from a stable transgenic Fischer 344 rat line expressing green fluorescent protein (GFP) and were embedded into a fibrin matrix containing growth factors; this formulation aimed to retain grafted cells in the lesion cavity and support cell survival. NSCs in the fibrin/growth factor cocktail were implanted two weeks after thoracic level-3 (T3) complete spinal cord transections, thereby avoiding peak periods of inflammation. Resulting grafts completely filled the lesion cavity and differentiated into both neurons, which extended axons into the host spinal cord over remarkably long distances, and glia. Grafts of cultured human NSCs expressing GFP resulted in similar findings. Thus, methods are defined for improving neural stem cell grafting, survival and analysis of in vivo findings.

Fibrin matrices containing growth factors were used to retain grafted neural stem cells into sites of complete spinal cord transection. Grafted cells completely filled the lesion cavity and differentiated into multiple neural cell types, including neurons that extended axons into host spinal cord over long distances.

Promoción de la supervivencia y la diferenciación de células madre neurales con fibrina y factor de crecimiento cócteles después de la médula espinal severa lesión

Neural stem cells (NSCs) can self-renew and differentiate into neurons and glia. Transplanted NSCs can replace lost neurons and glia after spinal cord ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

Etiquetado retrógrada de células ganglionares retinianas en adultos de pez cebra con tintes fluorescentes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

En Vivo electrofisiológicos mediciones sobre los nervios ciáticos de ratones

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Membrana de imagen potencial con dos tipos de sensores de tensión fluorescentes genéticamente codificados

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Seguimiento Neuroanatomical retrógrada frénico de neuronas de Motor en ratones

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

Activity-based Training on a Treadmill with Spinal Cord Injured Wistar Rats

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) on a treadmill is widely used as a rehabilitation tool in the SCI population with many benefits and improvements to daily life. We utilize this method of activity-based task-specific training (ABT) in rodents after SCI to both elucidate the mechanisms behind such improvements and to enhance and improve upon existing clinical rehabilitation protocols. Our current goal is to determine the mechanisms underlying ABT-induced improvements in urinary, bowel, and sexual function in SCI rats after a moderate to severe level of contusion. After securing each individual animal in a custom-made adjustable vest, they are secured to a versatile body weight support mechanism, lowered to a modified three-lane treadmill and assisted in step-training for 58 minutes, once a day for 10 weeks. This setup allows for the training of both quadrupedal and forelimb-only animals, alongside two different non-trained groups. Quadrupedal-trained animals with body weight support are aided by a technician present to assist in stepping with proper hind limb placement as necessary, while forelimb-only trained animals are raised at the caudal end to ensure no hind limb contact with the treadmill and no weight-bearing. One non-trained SCI group of animals is placed in a harness and rests next to the treadmill, while the other control SCI group remains in its home cage in the training room nearby. This paradigm allows for the training of multiple SCI animals at once, thus making it more time-efficient in addition to ensuring that our pre-clinical animal model mimics the clinical representation as close as possible, particularly with respect to the body weight support with manual assistance.

This protocol demonstrates our model of activity-based locomotor treadmill training for rats with spinal cord injury (SCI). Included is both quadrupedal and forelimb-only groups, in addition to two distinct types of non-trained control groups. Investigators are able to assess training effects on SCI rats using this protocol.

Formación basada en una cinta rodante con la médula espinal lesionado ratas Wistar

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) ...

Dissection of Pelvic Autonomic Ganglia and Associated Nerves in Male and Female Rats

The bilateral major pelvic ganglia (MPG; synonym, pelvic ganglia) are the primary source of postganglionic sympathetic and parasympathetic neurons innervating pelvic organs of rodents; the functionally equivalent structure in humans is the inferior hypogastric plexus. The major pelvic ganglia also provide the route by which lumbar and sacral sensory axons reach the pelvic organs. These complex, mixed ganglia can prove challenging to identify and dissect for further experimental study of normal autonomic mechanisms or to establish preclinical models of disease, injury or visceral pain. Here we describe a protocol to access and visualize these ganglia and their associated nerve tracts. We provide this protocol with schematics for both male and female rats, as the ganglion size and landmarks for identification differ between sexes. The protocol describes removal of the ganglion for in vitro studies, but this method can be integrated into a surgical recovery protocol for experimental interventions (e.g., nerve crush, nerve resection) or for mapping neuronal circuits (e.g., by microinjection of neural tracers). We also demonstrate the primary structures of the ganglion and its associated nerves immediately following dissection and following immunohistochemical staining.

The major pelvic ganglia contain parasympathetic and sympathetic neurons that innervate pelvic organs. Here we describe a dissection method and provide schematics for identification of these ganglia and their associated nerves. These methods can be applied to experimental manipulation of these ganglia in vivo or removal post-mortem for further study.