Este estudio fue apoyado por la Juvenile Diabetes Research Foundation conceder 5-2007-97 (a VEP), por el Instituto Nacional de Diabetes y Enfermedades Digestivas y Renales subvenciones R01DK064973 (a VEP), R01DK48494 (a LHP), T32DK07074 (apoyo SCE), K01DK083552 (a MDK), y por P60DK20595 a la Universidad de Chicago Investigación de la Diabetes y el Centro de Formación. El contenido es responsabilidad exclusiva de sus autores y no representan necesariamente la opinión oficial de la NIDDK oa los NIH.

1. Los preparativos pre-inyección <ol><li> Rápidamente el pescado por lo menos 24 horas antes de la inyección. Esto vaciará el bulbo intestinal (estómago) contenido. El protocolo de ayuno básica es la transferencia de los peces, a su densidad normal, a un tanque limpio, y luego abstenerse de ingerir alimentos. A más largo plazo que requiere el ayuno condiciones más rigurosas (por ejemplo, para los estudios de glucosa en la sangre), ver las consideraciones adicionales en la discusión. </li><li> Prepare una solución de sal de Cortland (Perry et al., 1984). Para un volumen de 100 ml, disolver el siguiente en agua destilada: 725 mg de NaCl (124,1 mM) 38 mg de KCl (5,1 mM) 41 mg Na 2 HPO 4 (2,9 mM) 24 mg MgSO4 ∙ 7H 2 O (1,9 mM) 16 mg de CaCl 2 ∙ 2 H 2 O (1,4 mM) 100 mg NaHCO3 (11,9 mm) 4 g polivinilpirrolidona (PVP) (4%) 1,000 unidades USP de heparina Filtro, esterilizar y almacenar a 4 ° C. </li><li> Preparar el microscopio. <ul><li> Cubra la base del microscopio con una envoltura de plástico para su protección en caso de derrames. </li><li> Coloque una toalla de papel encima de la envoltura de plástico. La mesa de operaciones se sentará en la parte superior de la toalla de papel. </li><li> Pre-ajustar el enfoque al ver la mesa de operaciones y centrarse en la esponja. </li></ul> Consejo: Ponga el dedo en la parte superior de la esponja y se centran en eso. Así se eliminarán o minimizar aún más el ajuste focal, una vez que el pescado esté en la mesa de operaciones. </li><li> Que pesar los peces. <ul><li> Llene un vaso de 500 ml aproximadamente 1 / lleno de peces de agua instalación 3. </li><li> Tarar la balanza. </li><li> Recoger los peces con una red. Mecha exceso de agua lejos de la red y los peces con una breve taponando la red en toallas de papel. La transferencia de los peces en el vaso. </li><li> Que pesar los peces. </li><li> La transferencia de los peces a un tanque limpio. </li><li> La transferencia de cada pez pesado a su propio tanque de la etiqueta. </li><li> Calcular el volumen de inyección para cada pez basado en el peso de pescado. </li></ul></li><li> Prepare la jeringa y equipos relacionados con la inyección. Para la inyección, se recomienda una aguja 35G de acero biselado y una microjeringa de 10 l NanoFil. Prepare la jeringa y el tubo NanoFil Silflex siguiendo las instrucciones del fabricante. Es importante eliminar las burbujas de la jeringa y el tubo. Después de llenar la jeringa y el tubo, coloque la jeringa de la bomba, y el programa del volumen de inyección para el primer pez. </li><li> Prepare la mesa de operaciones. <ul><li> Cortar una esponja suave (como # L800-D, Jaece Industrias), de modo que es de aproximadamente 20 mm de altura. En la cara plana, hacer un corte que es de 10-15 mm de profundidad. Este corte es el canal que llevará a cabo la pesca de la inyección. </li><li> Establecer la esponja en un 60 mm placa de Petri. </li><li> Ajuste la placa de Petri con una esponja en una tapa de dimensiones adecuadas pipeta caja de punta. La tapa tiene que ser lo suficientemente grande como para contener el agua para ayudar a mantener la temperatura de la esponja, pero deben ser poco profundos como para no ponerse en el camino. Nosotros utilizamos una tapa de una caja de punta P200 que es de 11,4 cm de largo x 7,7 cm x 1,5 cm W D. </li></ul> Estos tres elementos reunidos (esponja en placa de Petri en la tapa de la caja) constituyen la mesa de operaciones. </li><li> Prepare la anestesia. <ul><li> Hacer uso de cubos de hielo picado a partir de peces de agua las instalaciones. </li></ul> Consejo: Con típicas bandejas de cubitos de hielo, que se llevará a tres bandejas para anestesiar a los peces 10-12. <ul><li> Llenar un cubo de hielo limpio con el hielo picado. </li><li> Ponga la mesa de operaciones en un recipiente más grande, como un contenedor de 2,4 litros de almacenamiento Rubbermaid alimentos. </li><li> Vierta un poco de agua instalación (caliente) en el recipiente exterior y la mesa de operaciones. Mantenga una reserva de agua caliente instalaciones cercanas. </li><li> Ponga un termómetro en el recipiente exterior. </li></ul></li></ol> 2. Anestesia, de inyección y recuperación <ol><li> Coloque el recipiente de anestesia exterior y una mesa quirúrgica junto al microscopio. Tiene el cubo de cubitos de hielo cerca. </li><li> Llevar la temperatura de agua a 17 ° C mediante la adición de trozos de hielo. Importante: No se vaya por debajo de 17 ° C para este paso. </li><li> Utilice una red para la transferencia de los peces en el recipiente exterior. </li><li> Poco a poco agregue cubos de hielo en el contenedor para bajar la temperatura a 12 ° C, en el transcurso de varios minutos. </li><li> Monitorear el comportamiento de los peces: A los 17 ° C o ligeramente más baja, los peces normalmente se extenderá sus aletas pectorales en posición horizontal, jadear y tener movimientos rápidos opérculo. A medida que la temperatura baja, los peces nadan más lentamente y, finalmente, dejar de nadar. A medida que el plano quirúrgico de anestesia se acercó, jadeando se detendrá y los movimientos lentos opérculo. El pescado está listo para la inyección cuando no reacciona a los que se manejan. Para la mayoría de los peces, 12 ° C es suficiente. Los peces más grandes pueden requerir más frío water. </li><li> A medida que la temperatura deseada (~ 12 ° C o inferior), pulse sobre la esponja para saturar. </li><li> Mantenga sus dedos en el agua fría suficiente para que no se caliente el pescado y llevarlo a cabo de la anestesia durante la manipulación. </li><li> Con los dedos fríos, con cuidado de transferencia de los peces al canal de la esponja. Coloque el pescado en el abdomen y las agallas en el comedero. </li><li> Rápidamente la transferencia de la mesa de operaciones de la platina del microscopio. </li><li> Trabajando rápidamente, cuidadosamente inserte la aguja en la línea media entre las aletas pélvicas. La aguja debe apuntar hacia craneal y se inserta cerca de la cintura pélvica que en el ano. Usted debe ser capaz de sentir cuando la aguja se encuentra profundamente a la pared del cuerpo. Inyectar el volumen adecuado y retirar la aguja. </li><li> Después de la inyección, inmediatamente la transferencia de los peces de nuevo a su agua caliente (~ 28,5 ° C) el tanque de recuperación por la liberación de los peces de la esponja en el agua del tanque. Consejo: Si los peces no comienzan a nadar inmediatamente, contribuir a su recuperación girando suavemente el agua hacia las branquias. </li><li> Compruebe la aguja. En ocasiones una escala puede estar unido y debe ser removido antes de la próxima inyección. </li><li> Las inyecciones siguientes, el uso de agua caliente para instalaciones de llevar la cámara de anestesia temperatura del agua de nuevo hasta 17 ° C antes de introducir el pescado que viene. </li></ol> 3. Los resultados representativos: <img src="/files/ftp_upload/2126/2126fig1.jpg" alt="Figura 1" /> Figura 1. Los resultados representativos después de la inyección intraperitoneal de 0,5 mg / g de glucosa o de un vehículo. Peces se mantuvieron en ayunas durante 72 horas antes de la inyección. El eje X muestra el tiempo, después de la inyección. Media ± SEM.

<ol>
	<li>Perry, S. F., Davie, P. S., Daxboeck, C., Ellis, A. G., Smith, D. G., Hoar, W. S., Randall, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+methods+for+the+study+of+gill+physiology.">Perfusion methods for the study of gill physiology.</a> Fish Physiology Volume X: Gills, Part B: Ion and Water. , 325-388 (1984).</li><li>Brown, L. A., Stoskopf, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthesia+and+restraint.">Anesthesia and restraint.</a> Fish Medicine. , 79-90 (1993).</li><li>Eames, S. C., Philipson, L., Prince, V. E., Kinkel, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+sugar+measurement+in+zebrafish+reveals+dynamics+of+glucose+homeostastis.">Blood sugar measurement in zebrafish reveals dynamics of glucose homeostastis.</a> Zebrafish. 7, 205-213 (2010).</li><li>Chavin, W., Young, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factors+in+the+determination+of+normal+serum+glucose+levels+of+goldfish+Carassius+auratus+L.">Factors in the determination of normal serum glucose levels of goldfish Carassius auratus L.</a> Comp Biochem Physiol. 33, 629-653 (1970).</li><li>Groff, J. M., Zinkl, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hematology+and+clinical+chemistry+of+cyprinid+fish.+Common+carp+and+goldfish.">Hematology and clinical chemistry of cyprinid fish. Common carp and goldfish.</a> Vet Clin North Am Exot Anim Pract. 2, 741-776 (1999).</li><li>Brand, M., Granato, M., Nusslein-Volhard, C., Nusslein-Volhard, C., Dahm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Keeping+and+raising+zebrafish.">Keeping and raising zebrafish.</a> Zebrafish: A Practical Approach. , 7-37 (2002).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book. , (1995).</li><li>Iwama, G. K., Ackerman, P. A., Hochachka, P. W., Mommsen, T. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anaesthetics.">Anaesthetics.</a> Biochemistry and Molecular Biology of Fishes, Volume 3: Analytical Techniques. , 1-15 (1994).</li><li>Reavill, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Common+diagnostic+and+clinical+techniques+for+fish.">Common diagnostic and clinical techniques for fish.</a> Vet Clin North Am Exot Anim Pract. 9, 223-235 (2006).</li><li>Stoskopf, M. K., Stoskopf, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surgery.">Surgery.</a> Fish Medicine. , 91-97 (1993).</li></ol>

Inyección intraperitoneal de cinco pasos: en ayunas, con un peso, anestesia, la inyección, y la recuperación. Para cada etapa existen las mejores prácticas que pueden garantizar el éxito. El éxito incluye un paciente saludable de peces, así como un buen resultado del experimento. Ayuno: Un ayuno de 24 horas debe vaciar el bulbo intestinal. Esta práctica se ha tomado de la literatura de pescado veterinaria (por ejemplo, Brown 1993). Consideraciones adicionales en ayunas se discuten a continuación. A largo plazo en ayunas: Hemos descubierto que un ayuno de 72 horas es necesario para disminuir la glucosa en sangre a un nivel de referencia antes de la inyección (Eames et al, 2010).. También hemos encontrado que para los estudios de la glucosa son varios los procedimientos que se requieren para garantizar que el pescado en ayuno correctamente. Comience con un tanque limpio (sin restos en la parte inferior). Los tanques deben estar fuera de línea, claramente etiquetados como &quot;ayuno&quot;, y en un lugar donde entusiasta personal de atención de los peces no les dará de comer. Evaluar el ambiente externo de la cisterna y tomar medidas para evitar que los peces se hizo hincapié en las perturbaciones, como el estrés se sabe que aumentan la glucosa en sangre (Chavín y Young, 1970;. Groff et al, 1999). Por ejemplo, tuvimos un experimento de ayuno en la que fue operado de una radio a diario en el banco que llevaba los tanques de peces. Se encontró que la glucosa en sangre fue inusualmente alta y llegó a la conclusión de que el pescado se destacó por las vibraciones. Otro factor de estrés es el hacinamiento. El pescado debe mantenerse a una densidad que cumpla con las buenas prácticas de cría de peces. Para recomendaciones, consulte Brand et al. (2002) y Westerfield (1995). Hemos tenido buenos resultados en ayunas nuestros peces a una densidad de 10 a 12 peces en un tanque de 9 litros (con 3 capas de mármoles de tomar parte de ese volumen). La separación de los sexos puede causar estrés, por lo que se recomienda mantener una población de ambos sexos durante el ayuno. Esto significa que los huevos pueden ser puestos, y los huevos deben ser secuestrado para que no se comen. Una manera sencilla de capturar los huevos es para cubrir el fondo del tanque con 2-3 capas de canicas. La calidad del agua debe mantenerse mediante la eliminación de los huevos y los desechos y mediante la sustitución de alrededor del 10-15% del agua del tanque, todos los días. Para la eliminación de los huevos y los residuos, sifón funciona bien. Pesaje: Al pesar los peces que no están anestesiados, se debe tener cuidado para reducir al mínimo la transferencia de agua de la red en el vaso, para garantizar la exactitud de pesaje. Si la red (con pescado) se transfirieron a las toallas de papel, la mayor parte del exceso de agua se puede quitar, y el peso se puede medir con precisión. Puede ser más fácil para anestesiar a los peces antes del pesaje, pero no hemos probado los posibles efectos de la anestesia un pescado dos veces en un día. Hemos probado nuestra técnica pesando los primeros peces con la red / secante método y volver a pesar el pescado después de haber sido anestesiado, y suavemente borrado en seco. No se encontraron diferencias significativas en el peso entre los métodos (p = 0,7927, t-test). Además, hemos probado si esta red / secante de glucosa en la sangre método afectadas, en comparación con la simple transferencia de los peces en el vaso tan pronto como se anotó (sin borrar). No se encontraron diferencias significativas en el nivel de glucosa en sangre entre los dos métodos de transferencia (P = 0,2241, t-test). Anestesia: La anestesia química puede ser adecuado para muchos estudios. Aquí hemos demostrado anestesia agua fría como una alternativa, ya que anestésicos muchos (incluyendo tricaine/MS-222 (Brown, 1993)), elevar la glucosa en la sangre. En estudios anteriores, hemos determinado que el agua fría no elevan la glucosa en la sangre en el pez cebra (Eames et al., 2010). Para la anestesia de agua fría, la temperatura debe reducirse lentamente. La tasa de disminución parece depender del tamaño de los peces, con peces más pequeños va en más rápido que los peces más grandes. Después de la inyección, se puede observar que el pescado se está recuperando muy lentamente de la anestesia (ver más abajo). Esto puede dar lugar, cuando sea la temperatura inicial es demasiado bajo, o cuando la temperatura se reduce demasiado rápido. La temperatura inicial es muy baja si la curva de pescado lateralmente al entrar en el agua. Si la temperatura inicial es correcta, los peces mantener su equilibrio inicial. Girará sus aletas pectorales a una posición horizontal, jadear y tener movimientos rápidos opérculo. Por lo general, se va a nadar. A medida que disminuye la temperatura, los movimientos se reducirá y el pescado pierda el equilibrio. Un plano quirúrgico de anestesia se alcanza cuando los peces se pueden manejar sin reaccionar. Para mantener a los peces bajo anestesia quirúrgica, los dedos deben estar fríos, por lo que mantenerlos en el agua antes de manipular el pescado. La esponja también deben mantenerse en frío a la misma temperatura que el agua utilizada para anestesiar a los peces. Es importante para saturar la esponja con agua que se sufficiently fría para mantener la anestesia una vez que el pescado se coloca en la misma. Inyección: Antes de las inyecciones de empresa, es posible que desee analizar por lo menos un pescado para tener una idea del espesor de pared del cuerpo. Esto puede ayudarle a juzgar hasta qué punto la aguja tiene que insertar a entrar en la cavidad abdominal. Además, al insertar la aguja, se puede sentir la pared del cuerpo &quot;dar&quot; cuando la aguja entra en la cavidad abdominal. Durante la inyección, tomar medidas para mantener al paciente feliz. Asegúrese de que la esponja se satura con la temperatura del agua fría correcta para evitar que el pescado de reactivación durante la inyección. Una esponja muy suave y saturado es importante para minimizar los daños a las escalas y el moco que cubre de la piel. Una esponja bien saturada es también importante para mantener las branquias aireado. Le recomendamos la esponja de espuma se enumeran a continuación en Materiales. Finalmente, una vez que el pescado esté anestesiado, trabajar con rapidez para minimizar el tiempo que el pescado es bajo. Recuperación: El pescado debe recuperarse de la anestesia prácticamente al entrar en el tanque de agua caliente. Si el pescado no comienzan a nadar inmediatamente, agitar suavemente el agua hacia sus branquias para acelerar la recuperación. Si la recuperación es lenta, entonces los peces se hundió muy rápido y se debe ajustar el procedimiento de anestesia adecuada. Las posibles causas de la lenta recuperación se discuten en La anestesia.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Foam Sponge</td>
<td>&nbsp;</td>
<td>Jaece Industries</td>
<td>L800-D</td>
<td>&nbsp;</td>
<tr>
<td>60 mm Petri dish</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Pipet tip box lid</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>not too deep, e.g. 1.5 cm</td>
</tr>
<tr>
<td>Plastic storage container</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>deep, e.g. 7 cm </td>
</tr>
<tr>
<td>Thermometer</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Crushed ice</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>made from facility water</td>
</tr>
<tr>
<td>Warm facility water</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>1 liter or more</td>
</tr>
<tr>
<td>500 ml beaker</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>for weighing</td>
</tr>
<tr>
<td>NanoFil syringe</td>
<td>&nbsp;</td>
<td>World Precision Instruments (WPI)</td>
<td>NANOFIL</td>
<td>or Hamilton syringe</td>
</tr>
<tr>
<td>35 gauge needle</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>NF35BV-2</td>
<td>beveled</td>
</tr>
<tr>
<td>Silflex tubing</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>SILFLEX-2</td>
<td>&nbsp;</td>
<tr>
<td>UltraMicroPump III and Micro4 controller</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>UMPS-1</td>
<td>&nbsp;</td>
<tr>
<td>Foot switch</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>15867</td>
<td>&nbsp;</td>
<tr>
<td>Dissecting microscope</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Plastic wrap</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Paper towels</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Cortland salt solution</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

intraperitoneal injection into adult zebrafish

Un método conveniente para tratar químicamente el pez cebra es introducir el reactivo en el agua del tanque, donde será absorbido por los peces. Sin embargo, este método hace que sea difícil saber la cantidad de reactivo es absorbido o tomado por los peces. Algunas de las preguntas experimentales, en particular los relacionados con los estudios metabólicos, pueden ser mejor atendidos por la entrega de una cantidad determinada a cada uno de los peces, en función del peso. Aquí presentamos un método para intraperitoneal (IP) la inyección en adultos de pez cebra. La inyección es en la cavidad abdominal, por detrás de la cintura pélvica. Este procedimiento es una adaptación de los métodos de uso veterinario para peces más grandes. Es seguro, como hemos observado la mortalidad cero. Además, hemos visto el sangrado en el sitio de la inyección en sólo 5 de cada 127 inyecciones, y en cada uno de esos casos, el sangrado fue breves segundos, durando varios meses, y la cantidad de sangre perdida era pequeño. Éxito con este procedimiento requiere de un manejo suave de los peces a través de varias medidas incluyendo el ayuno, con un peso, anestesia, la inyección, y la recuperación. Las precauciones son necesarias para reducir al mínimo el estrés durante el procedimiento. Nuestras precauciones incluyen el uso de un volumen de inyección de pequeñas y una aguja 35G. Usamos solución Cortland sal como el vehículo, que es equilibrada por ósmosis de peces de agua dulce. Aireación de las branquias se mantiene durante el procedimiento de inyección, en primer lugar con lo que el pescado en un plano quirúrgico de anestesia, que permite movimientos lentos opérculo, y en segundo lugar, mediante la celebración de los peces en un canal dentro de una esponja saturada de agua durante la inyección en sí. Demostrar la utilidad de la inyección de IP mediante la inyección de glucosa y el control de la elevación del nivel de glucosa en la sangre y su posterior regreso a la normalidad. Ya que el estrés se sabe que aumenta la glucosa en sangre en los peces teleósteos, se comparan los niveles de glucosa en sangre en adultos inyectados con vehículo y no inyecta-y demostrar que el procedimiento no causa un aumento significativo de la glucosa en sangre.

Watch this Scientific Journal Video about Intraperitoneal Injection into Adult Zebrafish at JoVE.com

Intraperitoneal Injection into Adult Zebrafish

Demostramos inyección intraperitoneal en adultos de pez cebra. Nosotros usamos un 10 l microjeringa NanoFil controlado por un controlador Micro4 y III UltraMicroPump. Esta demostración incluye el uso de agua fría como un anestésico.

Inyección intraperitoneal en adultos de pez cebra

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, ...

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57.1K vistas.  The University of Chicago. Demostramos inyección intraperitoneal en adultos de pez cebra. Nosotros usamos un 10 l microjeringa NanoFil controlado por un controlador Micro4 y III UltraMicroPump. Esta demostración incluye el uso de agua fría como un anestésico.

Video: Intraperitoneal Injection into Adult Zebrafish

Transplantation of Whole Kidney Marrow in Adult Zebrafish

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an organism. The functional definition of a HSC is a transplanted cell that has the ability to reconstitute all the blood lineages of an irradiated recipient long term. This designation was established by decades of seminal work in mammalian systems. Using hematopoietic cell transplantation (HCT) and reverse genetic manipulations in the mouse, the underlying regulatory factors of HSC biology are beginning to be unveiled, but are still largely under-explored. Recently, the zebrafish has emerged as a powerful genetic model to study vertebrate hematopoiesis. Establishing HCT in zebrafish will allow scientists to utilize the large-scale genetic and chemical screening methodologies available in zebrafish to reveal novel mechanisms underlying HSC regulation. In this article, we demonstrate a method to perform HCT in adult zebrafish. We show the dissection and preparation of zebrafish whole kidney marrow, the site of adult hematopoiesis in the zebrafish, and the introduction of these donor cells into the circulation of irradiated recipient fish via intracardiac injection. Additionally, we describe the post-transplant care of fish in an "ICU" to increase their long-term health. In general, gentle care of the fish before, during, and after the transplant is critical to increase the number of fish that will survive more than one month following the procedure, which is essential for assessment of long term (&lt;3 month) engraftment. The experimental data used to establish this protocol will be published elsewhere. The establishment of this protocol will allow for the merger of large-scale zebrafish genetics and transplant biology.

In this article, we demonstrate a method to perform HCT in adult zebrafish.

El trasplante de médula del riñón entero en adultos de pez cebra

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an ...

Investigación

Biología

Zebrafish Brain Ventricle Injection

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved cavities within the brain that are filled with cerebrospinal fluid.  In zebrafish, after neural tube formation, the neuroepithelium undergoes a series of constrictions and folds while it fills with fluid resulting in brain ventricle formation.  In order to understand the process of ventricle formation, and the neuroepithelial shape changes that occur at the same time, we needed a way to visualize the ventricle space in comparison to the brain tissue.  However, the nature of transparent zebrafish embryos makes it difficult to differentiate the tissue from the ventricle space.  Therefore, we developed a brain ventricle injection technique where the ventricle space is filled with a fluorescent dye and imaged by brightfield and fluorescent microscopy.  The brightfield and the fluorescent images are then processed and superimposed in Photoshop.  This technique allows for visualization of the ventricle space with the fluorescent dye, in comparison to the shape of the neuroepithelium in the brightfield image.  Brain ventricle injection in zebrafish can be employed from 18 hours post fertilization through early larval stages.  We have used this technique extensively in our studies of brain ventricle formation and morphogenesis as well as in characterizing brain morphogenesis mutants (1-3).

After neural tube formation, the neuroepithelium constricts and folds while the tube fills with embryonic cerebrospinal fluid (eCSF) to form the embryonic brain ventricles. We developed this ventricle injection technique to better visualize the fluid filled space in contrast to the neuroepithelial shape in a live embryo.

Cerebro del pez cebra ventrículo inyección

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved ...

Retro-orbital Injection in Adult Zebrafish

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is often the most effective and noninvasive form of delivery of an agent of interest. In the zebrafish (Danio rerio), IV injection of drugs has long been a challenge because of the small vessel diameter. This has also proved a significant hurdle for the injection of cells during hematopoeitic stem cell transplantation. Historically, injections into the bloodstream were done directly through the heart. However, this intra-cardiac procedure has a very high mortality rate as the heart is often punctured during injection leaving the fish prone to infection, massive blood loss or fatal organ damage. Drawing on our experience with the mouse, we have developed a new injection procedure in the zebrafish in which the injection site is behind the eye and into the retro-orbital venous sinus. This retro-orbital (RO) injection technique has been successfully employed in both the injection of drugs in the adult fish as well as transplantation of whole kidney marrow cells. RO injection has a much lower mortality rate than traditional intra-cardiac injection. Fish that are injected retro-orbitally tend to bleed less following injection and are at a much lower risk of injury to a major organ like the heart. Further, when performed properly, injected cells and/or drugs quickly enter the bloodstream allowing compounds to exert their effect on the whole fish and kidney cells to easily home to their niche. Thus, this new injection technique minimizes mortality while allowing efficient delivery of material into the bloodstream of adult fish. Here we exemplify this technique by retro-orbital injection of Tg(globin:GFP) cells into adult casper fish as well as injection of a red fluorescent dye (dextran, Texas Red ) into adult casper fish. We then visualize successful injections by whole animal fluorescence microscopy.

Here we show how to do retro-orbital injection in adult zebrafish.

Retro-orbital de inyección en adultos de pez cebra

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is ...

Dissection of Organs from the Adult Zebrafish

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental analysis of the embryo and larva is extensive and the morphology has been well documented, descriptions of adult zebrafish anatomy and studies of development of the adult structures and organs, together with techniques for working with adults are lacking. The organs of the larva undergo significant changes in their overall structure, morphology, and anatomical location during the larval to adult transition.  Externally, the transparent larva develops its characteristic adult striped pigment pattern and paired pelvic fins, while internally, the organs undergo massive growth and remodeling.  In addition, the bipotential gonad primordium develops into either testis or ovary.  This protocol identifies many of the organs of the adult and demonstrates methods for dissection of the brain, gonads, gastrointestinal system, heart, and kidney of the adult zebrafish. The dissected organs can be used for in situ hybridization, immunohistochemistry, histology, RNA extraction, protein analysis, and other molecular techniques.  This protocol will assist in the broadening of studies in the zebrafish to include the remodeling of larval organs, the morphogenesis of organs specific to the adult and other investigations of the adult organ systems.

This protocol describes a procedure for identifying and dissecting organs from the adult zebrafish.

La disección de los órganos del pez cebra adultos

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental ...

Direct Delivery of MIF Morpholinos Into the Zebrafish Otocyst by Injection and Electroporation Affects Inner Ear Development

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the eye, brain, and somites of zebrafish. The advantage of electroporation over other methods of genetic manipulation is that specific tissues can be targeted, both spatially and temporally, for the introduction of macromolecules by the application of electrical current. Here we describe the use of electroporation for transfecting mif and mif-like morpholinos into the tissues of the developing inner ear of the zebrafish. In past studies, mif morpholino injected into embryos at the 1- to 8-cell stage resulted in widespread morphological changes in the nervous system and eye, as well as the ear. By targeting the tissues of the inner ear at later stages in development, we can determine the primary effects of MIF in the developing inner ear, as opposed to secondary effects that may result from the influence of other tissues. By using phalloidin and acetylated tubulin staining to study the morphology of neurons, neuronal processes, and hair cells associated with the posterior macula, we were able to assess the efficacy of electroporation as a method for targeted transfection in the zebrafish inner ear. The otic vesicles of 24hpf embryos were injected with morpholinos and electroporated and were then compared to embryos that had received no treatment or had been only injected or electroporated. Embryos that were injected and electroporated showed a decrease in hair cell numbers, decreased innervation by the statoacoustic ganglion (SAG) and fewer SAG neurons compared with control groups. Our results showed that direct delivery of morpholinos into otocysts at later stages avoids the non-specific nervous system and neural crest effects of morpholinos delivered at the 1-8 cell stage. It also allows examination of effects that are directed to the inner ear and not secondary effects on the ear from primary effects on the brain, neural crest or periotic mesenchyme.

A method to deliver morpholinos directly into the zebrafish otocyst at 24hpf has been developed. Using microinjection of morpholinos into the lumen of otic vesicle and electroporation to effect penetration, we were able to bypass the effect of morpholinos on the brain and obtain effects specific to the inner ear.

Entrega directa de Morpholinos FOMIN Hacia el otocyst pez cebra por inyección y electroporación afecta el desarrollo del oído interno

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the ...

RNAi Interference by dsRNA Injection into Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements and tools for genetic manipulation have become available in Drosophila 2. Soon after the initial discovery by Frie and Mello 3 that double stranded RNA can be used to knockdown the activity of individual genes in Caenorhabditis elegans, RNA interference (RNAi) was shown to provide a powerful reverse genetic approach to analyze gene functions in Drosophila organ development 4, 5.
Many organs, including lung, kidney, liver, and vascular system, are composed of branched tubular networks that transport vital fluids or gases 6, 7. The analysis of Drosophila tracheal formation provides an excellent model system to study the morphogenesis of other tubular organs 8. The Berkeley Drosophila genome project has revealed hundreds of genes that are expressed in the tracheal system. To study the molecular and cellular mechanism of tube formation, the challenge is to understand the roles of these genes in tracheal development. Here, we described a detailed method of dsRNA injection into Drosophila embryo to knockdown individual gene expression. We successfully knocked down endogenous dysfusion(dys) gene expression by dsRNA injection. Dys is a bHLH-PAS protein expressed in tracheal fusion cells, and it is required for tracheal branch fusion 9, 10. dys-RNAi completely eliminated dys expression and resulted in tracheal fusion defect. This relatively simple method provides a tool to identify genes requried for tissure and organ development in Drosophila.

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNA interference has been proven very effective to analyze gene function in Drosophila tracheal development. A detailed protocol used by Jiang lab to inject dsRNA into fly embryos to knockdown gene expression is illustrated. This technique has the potential for screening genes required for tissue and organ development in Drosophila.

RNAi interferencia por inyección de dsRNA en Drosophila Los embriones

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements ...

Transplantation of Cells Directly into the Kidney of Adult Zebrafish

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic diseases1. However, most organs are not easily accessible, necessitating the need to develop surgical methods to gain access to these structures. In this video article, we describe a method for transplanting cells directly into the kidney of adult zebrafish, a popular model to study regeneration and disease2. Recipient fish are pre-conditioned by irradiation to suppress the immune rejection of the injected cells3. We demonstrate how the head kidney can be exposed by a lateral incision in the flank of the fish, followed by the injection of cells directly in to the organ. Using fluorescently labeled whole kidney marrow cells comprising a mixed population of renal and hematopoietic precursors, we show that nephron progenitors can engraft and differentiate into new renal tissue - the gold standard of any cell-based regenerative therapy. This technique can be adapted to deliver purified stem or progenitor cells and/or small molecules to the kidney as well as other internal organs and further enhances the zebrafish as a versatile model to study regenerative medicine.

Cell transplantation is an essential technique for studying tissue regeneration and for developing cell-based therapies of disease. We demonstrate here a microsurgical technique that permits the transplantation of genetically labeled cells directly into the kidney of adult zebrafish fish.

El trasplante de células directamente en el riñón de adultos de pez cebra

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic ...

Dissection of the Adult Zebrafish Kidney

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem cells during normal homeostasis and disease states. The understanding of adult stem cells has broad reaching implications for the future of regenerative medicine1. For example, better knowledge about adult stem cell biology can facilitate the design of therapeutic strategies in which organs are triggered to heal themselves or even the creation of methods for growing organs in vitro that can be transplanted into humans1. The zebrafish has become a powerful animal model for the study of vertebrate cell biology2. There has been extensive documentation and analysis of embryonic development in the zebrafish3. Only recently have scientists sought to document adult anatomy and surgical dissection techniques4, as there has been a progressive movement within the zebrafish community to broaden the applications of this research organism to adult studies. For example, there are expanding interests in using zebrafish to investigate the biology of adult stem cell populations and make sophisticated adult models of diseases such as cancer5. Historically, isolation of the zebrafish adult kidney has been instrumental for studying hematopoiesis, as the kidney is the anatomical location of blood cell production in fish6,7. The kidney is composed of nephron functional units found in arborized arrangements, surrounded by hematopoietic tissue that is dispersed throughout the intervening spaces. The hematopoietic component consists of hematopoietic stem cells (HSCs) and their progeny that inhabit the kidney until they terminally differentiate8. In addition, it is now appreciated that a group of renal stem/progenitor cells (RPCs) also inhabit the zebrafish kidney organ and enable both kidney regeneration and growth, as observed in other fish species9-11. In light of this new discovery, the zebrafish kidney is one organ that houses the location of two exciting opportunities for adult stem cell biology studies. It is clear that many outstanding questions could be well served with this experimental system. To encourage expansion of this field, it is beneficial to document detailed methods of visualizing and then isolating the adult zebrafish kidney organ. This protocol details our procedure for dissection of the adult kidney from both unfixed and fixed animals.
Dissection of the kidney organ can be used to isolate and characterize hematopoietic and renal stem cells and their offspring using established techniques such as histology, fluorescence activated cell sorting (FACS)11,12, expression profiling13,14, and transplantation11,15. 
We hope that dissemination of this protocol will provide researchers with the knowledge to implement broader use of zebrafish studies that ultimately can be translated for human application.

The zebrafish kidney is home to both renal and hematopoietic adult stem/progenitor cells, and represents an outstanding opportunity to study these cell types and their progeny in a vertebrate model organism. Here, we demonstrate a detailed dissection procedure that enables the researcher to identify and surgically remove the adult zebrafish kidney, which can be used for applications such as cell isolation, transplantation, and expression studies of kidney and/or blood cell populations.

La disección del riñón adultos de pez cebra

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

En electroporación in vivo de Morpholinos en la retina de pez cebra adultos

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

En electroporación in vivo de Morpholinos en la regeneración de adultos de pez cebra Aleta de cola

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...