The project was supported by Award AAES 461Hatch/Multistate Grants ALA08-045 and ALA015-1-10026 to N.L.

1. Preparación de Instrumentos, Soluciones de estímulos, y los insectos de cama <ol><li> Preparar una solución de 50% KNO 2 (w / v) en una botella de 20 ml. </li><li> Afilar dos microelectrodos de tungsteno en solución KNO 2 a 5 V por inmersión en repetidas ocasiones los electrodos de tungsteno dentro y fuera de la solución. <ol><li> Aproximadamente afilar el alambre de tungsteno por inmersión de aproximadamente 10 mm del alambre de tungsteno dentro y fuera de la solución de KNO 2 a la velocidad de 2 inmersiones / seg durante aproximadamente 5 min, lo que puede consumir en gran medida el extremo frontal del alambre de tungsteno. </li><li> Delicadamente afilar el electrodo mediante la inmersión de aproximadamente 1 mm de la punta del alambre dentro y fuera de la solución a la velocidad de 2 inmersiones / seg durante al menos 1 min con el fin de hacer un punto del electrodo fino y agudo. Compruebe el diámetro de la punta del electrodo con el microscopio con frecuencia hasta que llega a 0,2 a 0,5 micras, que debe ser lo suficientemente fina como para perforar la cutícula del insecto de la cama sensillum olfativo. Nota: Si bien afilada manualmenteEning el electrodo, la velocidad de inmersión del alambre de tungsteno en la solución de KNO 2 no es constante todo el tiempo. Con más práctica, es mucho más fácil mantener una velocidad relativamente constante en el afilado del electrodo. La hora de afilar también es incierto en función de cómo bien debe ser el electrodo. Aquí, una punta de electrodo con el diámetro de ~ 0,2 micras es suficiente para perforar a través de la sensillum olfativo. </li></ol></li><li> Diluir cada uno de los estímulos químicos en sulfóxido de dimetilo (DMSO) a partir del compuesto ordenado a una concentración inicial de 01:10 v / v como una solución madre. Crear una serie de diluciones decimal dependiendo del número de dosis requeridas en el experimento, de nuevo con DMSO, a partir de cada una de las soluciones madre para cada producto químico. Aquí, utilizar 10% (+) - β-pineno y eucaliptol. </li><li> Coloca los días sin alimentar o siete después de la alimentación de las chinches adultos (ya sea hombre o mujer) de la Ft. Colonia Dix (un regalo del doctor Haynes en la Universidad de Kentucky) que se utilizará en tque experimentar en una placa de Petri. Nota: No hay un número exacto de chinches colocados en la placa de Petri. Puede ser un poco o mucho. </li></ol> 2. Cama Bug Preparación Antenas <ol><li> Anestesiar las chinches en el hielo (2-3 min). </li><li> Fijar tanto las antenas y el cuerpo del insecto en un cubreobjetos de microscopio con cinta de doble cara y retire las piernas con unas tijeras finas. </li><li> Use una aguja pequeña para tocar suavemente las antenas a fin de que se peguen en la cinta de manera constante. </li><li> Descansa el cubreobjetos contra una pequeña bola (~ 1 cm de diámetro) de cera dental para facilitar la manipulación y ajustarlo a un ángulo apropiado (~ 90 °) para el electrodo de registro (Figura 1). </li><li> Una vez asegurado, coloque la chinche de cama bajo un microscopio estéreo, encienda la fuente de luz fría y ajustar la intensidad de la iluminación hasta que la antena se presenta claramente, y el enfoque del microscopio en el segundo flagelo de la antena de chinches a gran aumento (720X) . Nota: La intensidad de la iluminación utilizada en el experimento no se cuantifica, lo que realmente depende de cómo los ojos del experimentador sentir la intensidad de la iluminación. </li></ol> 3. Individual sensillum grabación <ol><li> Conectar el preamplificador (10X) con el controlador de adquisición de la señal, que está conectado con el ordenador para la grabación de la señal y visualización. Encienda el ordenador e inicie el software, por ejemplo, AutoSpike32 y haga clic en el modo de &quot;Grabar&quot; en la barra de menú. A continuación, elija la &quot;onda&quot; con el fin de iniciar la grabación de las señales de onda. Nota: Una línea plana que va desde la izquierda a la derecha de la pantalla en varias ocasiones ahora debe ser visible. Aquí, la ventana de grabación tiene una duración de 40 seg. Grabación Max onda es 10 seg. Frecuencia de muestreo seleccionada es 96.000 y la tasa de muestreo digital es 240. No es la posición 0% y sin filtrar, sin rectificación de las señales de grabación. Todos estos ajustes de los parámetros en el software pueden ser modificados según sea necesario. </li> &lt;li&gt; Activar el altavoz conectado al preamplificador, que se utiliza para presentar el modo de tono para las respuestas neuronales a partir de sensillum antenal. </li><li> Inserte el electrodo de referencia en el abdomen del insecto de la cama estabilizado. Nota: El electrodo de referencia se llevó a cabo por un soporte del metal se adhiere magnéticamente a la mesa de aire. </li><li> Después de que el electrodo de referencia se ha conectado al abdomen del insecto de la cama, mover el electrodo de registro, que está conectado al preamplificador y manipulada por un micromanipulador, hacia el extremo posterior de la antena del insecto de la cama. </li><li> Cuando el electrodo de registro está en contacto con la punta derecha de la antena, encienda el microscopio y localizar el electrodo a bajo aumento. </li><li> Ajuste el electrodo de registro al tiempo que aumenta gradualmente la magnificación hasta que tanto el electrodo y la sensillum antenal están en el mismo plano y claramente visible bajo el microscopio. Nota: En ese momento, el microscopio es habitualmente con el máximo aumento. </li><li> Inserte el electrodo de registro en el eje del sensillum usando el micromanipulador e ir un poco más si el ruido de fondo es alto en comparación con el potencial de acción. </li><li> Una vez que los potenciales de acción claras se observan desde el sensillum grabada, llenar una micropipeta con 10% (+) - β-pineno. Utilice la micropipeta para depositar 10 l alícuota de 10% (+) - β-pineno sobre una tira de papel de filtro (~ 3 x 15 mm) colocado dentro de una pipeta Pasteur de vidrio. <ol><li> Conectar la pipeta cargada a la salida del tubo de flujo de impulsos del controlador de estímulo y coloque la punta de la pipeta en el pequeño orificio en el tubo orientado hacia la antena. </li></ol></li><li> Cuando todas estas conexiones se han estabilizado, presione el pedal del controlador de estímulo para entregar un soplo 0,5 seg de estímulo (0,5 L / min) en la corriente de aire humidificado continua. El registro de los potenciales de acción se iniciará simultáneamente cuando el footswitch es deprimido. El proceso de grabación será el último durante 10 segundos a partir del 1 seg antes de la estimulación. </li><li> Cuente los potenciales de acción fuera de línea durante dos períodos de 500 ms, una antes y otra después de la estimulación. Restar cualquier cambio en la tasa de pico durante el ms post-estimulación 500 de la actividad espontánea registrado durante los 500 ms precedentes y convertir los conteos en la escala convencional de espigas / s multiplicándolos por 2. </li></ol> Reemplazo 4. Estímulo en el SSR <ol><li> Una vez que el interruptor de pedal ha sido activado, entregar el 10% de (+) - β-pineno en la pipeta sobre la antenas bug cama y registrar la respuesta a este odorizante específico para 10 seg, después de lo cual se retira la pipeta. </li><li> Etiquetar otra nueva pipeta con el eucaliptol 0,001% a ensayar. Coloque un pequeño trozo de papel de filtro sobre el que se ha aplicado 10 l de estímulo, en la nueva pipeta. </li><li> Espere 2-5 minutos hasta que el estímulo es completamente vaporized en la pipeta de vidrio. Coloque la pipeta sobre la salida del tubo de flujo de pulso. </li><li> Inserte la punta de la pipeta en el pequeño orificio del tubo orientada hacia la antena. Pise el pedal y empezar la grabación de 10 segundos. </li><li> Desconecte la pipeta y preparar otra pipeta con 0,01% eucaliptol. </li><li> Prueba de todo el resto de las dosis de eucaliptol (de 0,001% a 10%) en la sensilla antenal para observar las respuestas dependientes de la dosis. Prueba de la más diluida, a las dosis menos diluidas. </li></ol>

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The authors have nothing to disclose.

La técnica individual sensillum grabación ha sido ampliamente utilizado en la prueba de las respuestas neurales de insectos como las moscas de la fruta, mosquitos y chinches a diferentes estímulos químicos en el medio ambiente. Estos estímulos químicos a menudo se disuelven y se diluyen en un disolvente común a fin de preparar diferentes dosis de los tratamientos. Sin embargo, diferentes disolventes pueden producir muy diferentes tasas de liberación de los estímulos. Estudios anteriores sobre algunos insectos e...

La chinche de cama común Cimex lectularius L (Hemiptera: Cimicidae), como un ectoparásito temporal, es un insecto hematófago obligado, lo que significa su supervivencia, desarrollo y reproducción requieren fuentes de sangre de los ejércitos, incluidos los seres humanos y los animales 1,2. Aunque la transmisión del virus rara vez ha sido reportado por C. lectularius, las molestias morder generada por una infestación afecta gravemente a los ejércitos, tanto física como psicológicamente 3. La introducción y el uso generalizado de insecticidas químicos, especialmente DDT, bajaron el riesgo de plagas y para el final de las infestaciones de 1950 se encontraban en un nivel tan bajo que ya no eran una grave preocupación pública. Sin embargo, una serie de posibles factores han llevado a resurgimiento en las poblaciones de chinches en todo el mundo, como la reducción del uso de insecticidas, una disminución de la conciencia pública, el aumento de la actividad de los viajes, y el desarrollo de resistencia a los insecticidas 4-9. &lt;/ p&gt; Señales químicas en el medio ambiente son detectados y reconocidos por los insectos a través de órganos olfativos, como antenas y palpos maxilares. El olfativo sensilla en las antenas de insectos juegan un papel crucial en la detección de estas señales químicas. Las moléculas químicas entran en la cutícula antenal través de los poros en la superficie de la cutícula. Odorante proteínas de unión en el enlace linfático antenal a estas moléculas químicas y transportarlos a los receptores de olor 10. Los receptores de olor y su co-receptor desde el canal iónico de cationes no selectivo en la membrana neuronal, que se despolariza una vez que estas moléculas químicas son reconocidos por los receptores de olor 11. Grabación sensillum Individual (SSR) fue desarrollado para detectar el cambio extracelular en el potencial de acción causada por la aplicación de cualquiera de los estímulos químicos o no químicos. Mediante la inserción de un electrodo de registro en la linfa sensillum y un electrodo de referenciaen alguna otra parte del cuerpo del insecto (por lo general, ya sea los ojos compuestos o el abdomen), la tasa de disparo de las neuronas en respuesta a estímulos se puede grabar 12. Los cambios en el número de espigas representan la sensibilidad del insecto a estímulos específicos. Estímulos químicos de diferentes identidades y concentración provocarán diferentes respuestas neuronales, con diferentes tasas de disparo y las estructuras temporales, y por lo tanto pueden ser utilizados para investigar el proceso de codificación del insecto a los productos químicos específicos. Para la chinche común, ambas formas sexuales comparten el mismo patrón de sensilla olfativo en las antenas: de nueve ranurado sensilla clavija C, 29-pelo como E (E1 y E2) sensilla, y un par cada uno de Dα, Dβ, Dγ peg suave sensilla 13,14. Como múltiples neuronas han sido identificados en cada tipo de sensillum, no es fácil distinguir los potenciales de acción de diferentes neuronas alojados en la misma sensillum, por lo que para este experimento la totanúmeros l de potenciales de acción se contaron fuera de línea durante un período mseg 500 antes y después de la estimulación. A continuación, el número de potenciales de acción después de la estimulación se restará de la cantidad de potenciales de acción antes de la estimulación y se multiplica por dos con el fin de cuantificar los cambios en la velocidad de disparo en cada sensillum individual en espigas por segundo 15.

<table border="0" cellpadding="0" cellspacing="0" width="1064">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:299px;">Tungsten wire</td>
			<td style="width:139px;">A-M SYSTEMS</td>
			<td style="width:117px;">#716500</td>
			<td style="width:511px;">Used for preparing the electrode</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">KNO2</td>
			<td>Sigma</td>
			<td>#310484</td>
			<td>Used for sharpening the tungsten wire</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">AC Power Supply</td>
			<td>BK Precision</td>
			<td>1653A</td>
			<td>Providing the voltage in sharpening the tungsten wire</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Leica Z6 APO Microscope</td>
			<td>Leica</td>
			<td>10447424</td>
			<td>Used for observing the sensilla on antennae</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Simulus controller</td>
			<td>Syntech</td>
			<td>CS-55</td>
			<td>Used for controlling the stimulus application</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">4-Channel USB Acquisition Controller</td>
			<td>Syntech</td>
			<td>IDAC-4</td>
			<td>Real-time on screen display of all signals before and during recording</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Light Source</td>
			<td>SCHOTT</td>
			<td>A20500</td>
			<td>Providing light sources for observation</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Micromanupulator</td>
			<td>Leica</td>
			<td>115378</td>
			<td>Used for minor movement of electrode</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Speaker</td>
			<td>Juster</td>
			<td>95a</td>
			<td>Connected with Acquisition Controller IDAC-4 and providing sound for the signal</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Magnetic stand</td>
			<td>Narishige</td>
			<td>GJ-1</td>
			<td>Used to hold the reference electrode, stablized bed bug and stimulus delivery tube</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">TMC Vibration Isolation Table</td>
			<td>TMC</td>
			<td>63-500</td>
			<td>Used for isolating the vibration from the equipments</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Coverslip</td>
			<td>Tedpella</td>
			<td>2225-1</td>
			<td>Used for holding the bed bug</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Double-sided Tape</td>
			<td>3M</td>
			<td>XT6110</td>
			<td>Used for stablizing the bed bug on the coverclip</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dental Wax</td>
			<td>Dentakit</td>
			<td>DK-R012</td>
			<td>Used for supporting the coverclip where bed bug is stablized&#160;</td>
		</tr>
	</tbody>
</table>

using single sensillum recording to detect olfactory neuron responses of bed bugs to semiochemicals

El sistema olfativo insecto desempeña un papel importante en la detección de semioquímicos en el ambiente. En particular, la sensilla antenal que alojan las neuronas individuales o múltiples en el interior, se consideran para hacer la mayor contribución en la respuesta a los estímulos químicos. Al registrar directamente potencial de acción en el sensillum olfatoria después de la exposición a los estímulos, grabación sensillum sola técnica (SSR) ofrece un enfoque poderoso para investigar las respuestas neurales de los insectos a los estímulos químicos. Para el insecto de la cama, que es un conocido parásito humano, se han caracterizado múltiples tipos de sensillum olfativo. En este estudio, hemos demostrado respuestas neurales de chinches sensilla olfativa a dos estímulos químicos y las respuestas dependientes de la dosis a una de ellas por el método de la RSS. Este enfoque permite a los investigadores para llevar a cabo la detección temprana de los estímulos químicos individuales en el olfativo sensilla chinches, que proporcionarían información valiosa para el desarrollo de nuevos atrayentes o repelentes de chinches y los beneficios de los esfuerzos de control de chinches.

Grabación individual sensillum es una poderosa técnica de investigación utilizada en los estudios de ecología química de insectos y fisiología neuronal. La investigación de las respuestas neurales de los insectos a los diferentes compuestos volátiles, especialmente los que piensa que es ecológico relacionado con la supervivencia y el desarrollo de los insectos, no sólo nos da una visión muy valiosa en el proceso de olfato de insectos, pero también abre nuevas vías prometedor...

Watch this Scientific Journal Video about Using Single Sensillum Recording to Detect Olfactory Neuron Responses of Bed Bugs to Semiochemicals at JoVE.com

Using Single Sensillum Recording to Detect Olfactory Neuron Responses of Bed Bugs to Semiochemicals

Bed bugs rely on olfactory receptor neurons housed in their antennal olfactory sensilla to detect semiochemicals in the environment. Utilizing single sensillum recording, we demonstrate a method to evaluate bed bug response to semiochemicals and explore the coding process involved.

Uso Individual sensillum grabación para detectar respuestas olfativo neurona de chinches de cama a Semioquímicos

The insect olfactory system plays an important role in detecting semiochemicals in the environment. In particular, the antennal sensilla which house ...

using-single-sensillum-recording-to-detect-olfactory-neuron-responses

Research

JoVE Journal

Neuroscience

9.8K vistas.  Auburn University. Bed bugs rely on olfactory receptor neurons housed in their antennal olfactory sensilla to detect semiochemicals in the environment. Utilizing single sensillum recording, we demonstrate a method to evaluate bed bug response to semiochemicals and explore the coding process involved.

Video: Using Single Sensillum Recording to Detect Olfactory Neuron Responses of Bed Bugs to Semiochemicals

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

Grabaciones solo Sensillum en los insectos Drosophila melanogaster Y Anopheles gambiae</em

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Investigación

Neurociencias

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Las respuestas de los análisis de las neuronas sensoriales olfativas ratón Utilizando la grabación Electroolfactogram Aire fase

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Recording Behavioral Responses to Reflection in Crayfish

Social behavior depends on sensory input from the visual, mechanical and olfactory systems. One important issue concerns the relative roles of each sensory modality in guiding behavior. The role of visual inputs has been examined by isolating visual stimuli from mechanical and chemosensory stimuli. In some studies (Bruski &amp; Dunham, 1987: Delgado-Morales et al., 2004) visual inputs have been removed with blindfolds or low light intensity, and effects of remaining sensory modalities have been elucidated. An alternative approach is to study the effects of visual inputs in the absence of any appropriate mechanical and chemosensory cues. This approach aims to identify the exclusive role of visual inputs. 
We have used two methods to provide visual stimuli to crayfish without providing chemical and mechanical cues. In one method, crayfish are videotaped in an aquarium where half of the walls are covered in mirrors to provide a reflective environment, and the other half are covered in a non-reflective (matte finish) plastic. This gives the crayfish a choice between reflective and non-reflective environments. The reflective environment provides visual cues in the form of reflected images of the crayfish as it moves throughout half of the tank; these visual cues are missing from the non-reflective half of the tank. An alternative method is to videotape the behavior of crayfish in an aquarium separated by a smaller chamber at each end, with a crayfish in one small chamber providing visual cues and an inert object in the opposite small chamber providing visual input from a non-moving, non-crayfish source. 
Our published results indicate that responses of crayfish to the reflective environment depend on socialization and dominance rank. Socialized crayfish spent more time in the reflective environment and exhibited certain behaviors more frequently there than in the non-reflective environment; isolated crayfish showed no such differences. Crayfish that were housed in same-sex pairs developed a social rank of either dominant or subordinate. Responses to reflection differed between dominant and subordinate crayfish (May &amp; Mercier, 2006; May &amp; Mercier, 2007). Dominant crayfish spent more time on the reflective side, entered reflective corners more frequently and spent more time in reflective corners compared to the non-reflective side. Subordinate crayfish walked in reverse more often on the reflective side than on the non-reflective side. Preliminary data suggest similar effects from visual cues provided by a crayfish in a small adjoining chamber (May et al., 2008).

We have developed two methods for studying effects of visual cues on behavior in the absence of tactile and chemical cues. One method involves videotaping responses of crayfish to reflective walls in an aquarium; the other examines effects of visual inputs provided by a live crayfish behind a transparent partition.

Grabación de las respuestas de comportamiento a la reflexión en el cangrejo de río

Social behavior depends on sensory input from the visual, mechanical and olfactory systems.  One important issue concerns the relative roles of each ...

Using a Comparative Species Approach to Investigate the Neurobiology of Paternal Responses

A goal of behavioral neuroscience is to identify underlying neurobiological factors that regulate specific behaviors. Using animal models to accomplish this goal, many methodological strategies require invasive techniques to manipulate the intensity of the behavior of interest (e.g., lesion methods, pharmacological manipulations, microdialysis techniques, genetically-engineered animal models). The utilization of a comparative species approach allows researchers to take advantage of naturally occurring differences in response strategies existing in closely related species. In our lab, we use two species of the Peromyscus genus that differ in paternal responses. The male California deer mouse (Peromyscus californicus) exhibits the same parental responses as the female whereas its cousin, the common deer mouse (Peromyscus maniculatus) exhibits virtually no nurturing/parental responses in the presence of pups. Of specific interest in this article is an exploration of the neurobiological factors associated with the affiliative social responses exhibited by the paternal California deer mouse. Because the behavioral neuroscience approach is multifaceted, the following key components of the study will be briefly addressed: the identification of appropriate species for this type of research; data collection for behavioral analysis; preparation and sectioning of the brains; basic steps involved in immunocytochemistry for the quantification of vasopressin-immunoreactivity; the use of neuroimaging software to quantify the brain tissue; the use of a microsequencing video analysis to score behavior and, finally, the appropriate statistical analyses to provide the most informed interpretations of the research findings.

The comparative species approach allows behavioral neuroscientists to explore various neurobiological factors associated with specific behaviors viewed as characteristic of a specific animal model. Taking advantage of naturally occurring differences in behavior between closely related species, this technique doesn&#x2019;t require invasive techniques to manipulate the expression of the behavior.

Con un enfoque comparativo de especies para la Investigación de Neurobiología de las respuestas Paternal

A goal of behavioral neuroscience is to identify underlying neurobiological factors that regulate specific behaviors.  Using animal models to accomplish ...

Odorant-induced Responses Recorded from Olfactory Receptor Neurons using the Suction Pipette Technique

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex behaviors such as food choice, predator, conspecific and mate recognition and other socially relevant cues. Olfactory receptor neurons (ORNs) are located in the dorsal part of the nasal cavity embedded in the olfactory epithelium. These bipolar neurons send an axon to the olfactory bulb (see Fig. 1, Reisert &amp; Zhao1, originally published in the Journal of General Physiology) and extend a single dendrite to the epithelial border from where cilia radiate into the mucus that covers the olfactory epithelium. The cilia contain the signal transduction machinery that ultimately leads to excitatory current influx through the ciliary transduction channels, a cyclic nucleotide-gated (CNG) channel and a Ca2+-activated Cl- channel (Fig. 1). The ensuing depolarization triggers action potential generation at the cell body2-4.
In this video we describe the use of the "suction pipette technique" to record odorant-induced responses from ORNs. This method was originally developed to record from rod photoreceptors5 and a variant of this method can be found at jove.com modified to record from mouse cone photoreceptors6. The suction pipette technique was later adapted to also record from ORNs7,8. Briefly, following dissociation of the olfactory epithelium and cell isolation, the entire cell body of an ORN is sucked into the tip of a recording pipette. The dendrite and the cilia remain exposed to the bath solution and thus accessible to solution changes to enable e.g. odorant or pharmacological blocker application. In this configuration, no access to the intracellular environment is gained (no whole-cell voltage clamp) and the intracellular voltage remains free to vary. This allows the simultaneous recording of the slow receptor current that originates at the cilia and fast action potentials fired by the cell body9. The difference in kinetics between these two signals allows them to be separated using different filter settings. This technique can be used on any wild type or knockout mouse or to record selectively from ORNs that also express GFP to label specific subsets of ORNs, e.g. expressing a given odorant receptor or ion channel.

Olfactory receptor neurons (ORNs) convert odor signals first into a receptor current that in turn triggers action potentials that are conveyed to second order neurons in the olfactory bulb. Here we describe the suction pipette technique to record simultaneously the odorant-induced receptor current and action potentials from mouse ORNs.

Inducidas Odorante-respuestas registradas de neuronas olfativas del receptor utilizando la técnica de aspiración con una pipeta

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Varias unidades Métodos de grabación para caracterizar la actividad neuronal en el Locust (<em&gt; Schistocerca americana</em&gt;) Circuitos olfativas

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

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 ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Método electrofisiológico para registrar las respuestas de voltaje intracelular<em&gt; Drosophila</em&gt; Los fotorreceptores y interneuronas a la luz Estímulos<em&gt; En Vivo</em

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

A Free-breathing fMRI Method to Study Human Olfactory Function

The study of human olfaction is a highly complex and valuable field with applications ranging from biomedical research to clinical evaluation. Currently, evaluation of the functions of the human central olfactory system with functional magnetic resonance imaging (fMRI) is still a challenge because of several technical difficulties. There are some significant variables to take into account when considering an effective method for mapping the function of the central olfactory system using fMRI, including proper odorant selection, the interaction between odor presentation and respiration, and potential anticipation of or habituation to odorants. An event-related, respiration-triggered olfactory fMRI technique can accurately administer odorants to stimulate the olfactory system while minimizing potential interference. It can effectively capture the precise onsets of fMRI signals in the primary olfactory cortex using our data post-processing method. The technique presented here provides an efficient and practical means for generating reliable olfactory fMRI results. Such a technique can ultimately be applied in the clinical realm as a diagnostic tool for diseases associated with olfactory degeneration, including Alzheimer&#39;s and Parkinson&#39;s disease, as we begin to further understand the complexities of the human olfactory system.

We present the technical challenges and solutions for obtaining reliable functional magnetic resonance imaging (fMRI) data from the human central olfactory system. This includes special considerations in olfactory fMRI paradigm design, descriptions of fMRI data acquisition with an MRI-compatible olfactometer, odorant selection, and a special software tool for data post-processing.

Un método fMRI de respiración libre para estudiar la función olfativa humana

The study of human olfaction is a highly complex and valuable field with applications ranging from biomedical research to clinical evaluation. Currently, ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.