This work was supported by AI101263 from the National Institutes of Health (T.V.B), and by a grant from The Hartwell Foundation (M.L.M). M.L.M. is a member of the Duke Comprehensive Cancer Center.

NOTA: Todos los procedimientos experimentales sobre la utilización de las ratas se realizaron de acuerdo con los protocolos aprobados por el Comité de Cuidado y Uso de Animales de la Universidad de Duke. 1. Preparación de la intubación traqueal Cánula <ol><li> Corte el extremo de la aguja de un 2 pulgadas 14 G aguja angiocathether con alicates de corte. </li><li> Apriete la luz de la aguja cerrada con unos alicates de punta fina. </li><li> Cortar la sección pellizcado hasta el borde de la porción lumen abierto con unos alicates de corte, y luego cortar los bordes en ambos lados de el extremo restante a 30 - 45 ° ángulo con el borde de la aguja. </li><li> Coloque la parte inferior 3 - 4 mm de la cánula en la zona abierta de alicates de junta deslizante recta y tire hacia arriba hasta una curva ligeramente hacia arriba se ha formado en el extremo. NOTA: Esto facilitará dirigir el catéter en la tráquea (Figura 1). </li><li> Utilice papel de lija fino para suavizar los bordes en el extremo de la cánula. </li><li&gt; Envíe el instrumento romo en el tubo angiocatéter. Asegúrese de que el tubo es ligeramente más corta que la aguja roma. </li></ol> Procedimientos 2. Pre-quirúrgicos <ol><li> Configurar el sitio para el procedimiento mediante la colocación de un microscopio de 10X de aumento sobre el área de operación. </li><li> Coloque una almohadilla térmica en el área de operaciones y cubrir con una almohadilla absorbente limpio. </li><li> Establecer un monitor de frecuencia cardíaca y oxigenación de la sangre cerca del campo quirúrgico. </li><li> Pese la rata para la dosificación basada en el peso de medicamentos (por ejemplo, la analgesia postoperatoria, antibióticos, o anticuerpo linfocito ozono). NOTA: Típico ratas adultas pesan entre 350 a 450 g. </li><li> Sedar al rata con vaporizado 3% de isoflurano-O 2 (3 L / min) usando una cámara de inducción ventila a un sistema de recogida de gases anestésicos residuales que contienen carbón activado. Permita 5 min para la inducción de anestesia para proporcionar anestesia profunda necesaria para la intubación endotraqueal. </li><li> Administrar carprofeno o Meloxicane (4,4 mg / kg) por vía subcutánea a la rata antes de proceder con la intubación y la cirugía. </li><li> Utilice una maquinilla eléctrica de afeitar el cuello y el pecho de la rata anestesiada antes de la intubación. </li></ol> 3. La intubación <ol><li> Preparar la configuración de la intubación por el roce de una pequeña cantidad de lubricación (por ejemplo, KY gel) en el extremo de la cánula de intubación. Encienda el ventilador para comenzar el flujo de gas isoflurano a través del tubo conector. </li><li> Transferencia de la rata en el aparato de intubación y suspender la rata en la barra de metal por su dientes incisivo superior (Figura 2A). NOTA: Algunos aparatos de intubación pueden ser comprados en línea. El modelo que se muestra aquí es hecho en casa utilizando una hoja de acrílico moldeado alrededor de las secciones de una pulgada pieza de 2 x 4 lumbar utilizando un soplete de butano, y los refuerzos están unidos con cloroformo. La barra (o alambre de metal) se une por los agujeros de perforación más cinta adhesiva colocadas en el side del acrílico para evitar que se astille o formación de grietas. </li><li> Trans-iluminar el cuello de la rata mediante la colocación de una fuente de luz de alta intensidad flexible de 1 - 2 cm de la superficie ventral del cuello (Figura 2B). </li><li> Use un par de pinzas de estudiantes patrón estándar para tirar de la lengua suavemente hacia arriba y hacia el lado de los dientes inferiores. Grip la lengüeta entre el pulgar y el dedo índice de una mano mientras que la colocación de la superficie interior plana de un lado de la pinza contra el extremo inferior de la lengua. Pulse ventralmente para exponer la epiglotis y la abertura de la laringe (Figura 2C). </li><li> Visualizar las cuerdas vocales y guiar el extremo vuelto hacia arriba, romo de la cánula de intubación anterior a través de la glotis abierta en la tráquea hasta el centro de la angiocatheter toca los incisivos. </li><li> Retire el estilete de metal, y conecte el tubo de anestesia para la apertura angiocatéter para iniciar la ventilación con isoflurano. NOTA: El flo típica de oxígenotasa de w para el ventilador es de 3 L / min con 3% de isoflurano. <ol><li> Ajuste el ventilador a una velocidad de 60 respiraciones / min en un modo de ventilador de volumen controlado que alcanza una presión de ~ 12 - 14 mmHg. Use una presión positiva espiratoria final (PEEP) de 3 cm H 2 O. </li></ol></li><li> Observar la expansión de la pared torácica bilateral en sincronía con el ventilador para asegurar la colocación correcta del tubo endotraqueal. </li><li> Aplicar veterinario recomendado ungüento para los ojos a los ojos de la rata para evitar la sequedad mientras que bajo anestesia. </li><li> Aplique un ungüento veterinario en los ojos de la rata para evitar la sequedad mientras que bajo anestesia. </li><li> Asegure el tubo endotraqueal a la cabeza de la rata con una tira de cinta de tela. Asegurar una conexión firme entre el tubo endotraqueal y el tubo de entrada de anestésico. </li><li> Coloque la oxigenación de la sangre y el monitor de ritmo cardíaco para el pie de la rata y comenzar la vigilancia. Confirme anestesia adecuada al confirmar que no hay reacción al ae-pellizco. </li></ol> 4. La toracotomía y timectomía <ol><li> Siga técnica aséptica estándar para la totalidad del procedimiento. <ol><li> Limpie la tabla del área de trabajo y de funcionamiento y desinfectar con una solución de etanol al 70%. </li><li> Utilizar guantes quirúrgicos estériles durante el procedimiento, y el autoclave todos los instrumentos y materiales utilizados durante el procedimiento. </li><li> Aplicar providona-yodo para toda la zona del pecho y deje que se seque. A continuación, limpiar la superficie de la piel con etanol al 70% en gasa. Cubra la rata con una envoltura de plástico transparente cortar un agujero para exponer el campo operatorio estéril. </li></ol></li><li> Identificar la horquilla esternal en la región torácica superior. Hacer un 2 cm incisión en la línea media a través de la piel a partir de 2 - 3 mm por encima de la horquilla esternal y se extiende a distancia de la línea media a lo largo del esternón con unas tijeras de punta roma Shea. </li><li> Realice una esternotomía 1,5 cm mediana de la horquilla esternal de nuevo con unas tijeras de punta roma Shea. KEEP el borde inferior de las tijeras justo debajo del esternón y avanzar lentamente. </li><li> Inserte un pequeño retractor Alm justo debajo del esternón se separó y se abren para revelar el músculos correa pre-traqueal (esternohioideo y músculos esternotiroideo). Separar los músculos de la correa pre-traqueales utilizando fórceps Graefe contundentes. NOTA: En este punto, la tráquea se puede ver, y el tubo de intubación se debe visualizar el interior de la tráquea. </li><li> Coloque los dientes del pequeño retractor Alm debajo de los músculos de la correa separadas y el esternón. Abra el retractor para exponer la cara superior del timo. </li><li> Utilice unas pinzas finas Dumont para liberar los bordes laterales del tejido del timo y exponer los lóbulos tímicos inferiores. </li><li> Tire el timo suavemente superiormente en el sitio de la incisión abierta, teniendo cuidado de evitar el contacto con los vasos de la vena cava, subclavia y carótida superiores, y para minimizar la interrupción del delicado revestimiento pleural entre el timo y los pulmones. </li><li> Como tél tímica vasos se disecan y reveló, utilice micro-tijeras para dividirlos. Use un hisopo de algodón para mantener la presión sobre los vasos de la hemostasia si es necesario. </li><li> Entregar los lóbulos tímicos más bajos dentro de la incisión y luego bruscamente lisar los archivos adjuntos posteriores. Retire el timo intacta e inspeccionar cuidadosamente el timo retirado por cualquiera de las secciones que faltan. NOTA: Los pequeños ganglios linfáticos torácicos pueden ser visualizados rodea el timo y son a menudo difíciles de diferenciar de tejido del timo. Estos serán los nodos redondos discretos con un aspecto similar al tejido del timo, pero no estarán en continuidad con el timo. </li><li> Retire el retractor Alm y cerrar los músculos esternohioideo y sternothyoid con dos suturas interrumpidas 5-0 Maxon. </li><li> Aplicar 2 gotas de grado quirúrgico adhesivo tisular de cianoacrilato a través de los puntos de sutura para cerrar el mediastino bajo ventilación con presión positiva. NOTA: Esto disminuirá la incidencia de neumotórax y hematoma. </li><li> Cierre el sternum con dos interrumpido suturas de seda 4-0 en una aguja de corte. Insertar la aguja a través de huecos entre las costillas, teniendo cuidado para guiar la aguja justo debajo del esternón para evitar la capa muscular subyacente. </li><li> Cierre la capa de la piel con un corredor de nylon 4-0 sutura. Suspenda el isoflurano en este punto para acortar el período de recuperación de la anestesia después de la operación. </li><li> Limpie el sitio de la incisión y la piel circundante con una gasa húmeda con salina. </li><li> Aplicar varias gotas de bupivicaína (0,25%) para la anestesia local, seguido de 1 - 2 gotas de pegamento de cianoacrilato sobre la incisión para sellar la incisión. </li><li> Continuar para ventilar hasta que la rata muestra signos de esfuerzo respiratorio independiente y comienza a mover sus extremidades. Retirar la intubación a la rata y deje que se recupere bajo observación en una jaula colocada sobre una almohadilla térmica. </li><li> Administrar buprenorfina (0,01-0,05 mg / kg) por vía subcutánea a la rata inmediatamente después de la operación y repita cada 8-12 h durante 48 h post-operativamente (normalmente 5 dosis en total). A las 24 y 48 horas después de la operación, administrar Carprofeno o Meloxicane (4,4 mg / kg) por vía subcutánea (3 dosis totales incluyendo la dosis preoperatoria). </li><li> Seguir vigilando hasta que la rata se mueve alrededor de la jaula. En cualquier signo de dificultad respiratoria, realizar re-intubación y re-exploración del pecho. <ol><li> No deje a un animal sin vigilancia hasta que se haya recuperado el conocimiento suficiente para mantener decúbito esternal. </li><li> No devuelva un animal que ha sido sometido a cirugía para la compañía de otros animales hasta que se recupere completamente. </li></ol></li><li> Retire las suturas de la piel de la capa 1 semana después de la operación. </li></ol>

<ol>
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The authors have nothing to disclose.

El presente protocolo para la timectomía completa ofrece un abordaje mínimamente invasivo, con un cierre de la incisión de tres capas con aplicación de pegamento quirúrgico diseñado para minimizar las complicaciones. La eliminación completa del timo se demostró por la pérdida de células T vírgenes y por tinción IHC de tejido linfoide mediastinal para citoqueratina. El procedimiento de la timectomía rata adulta se ha complicado por las tasas de mortalidad que van desde 1,5 hasta ...

Desde principios de la década de 1960, el timo ha sido reconocido por su papel crítico en el desarrollo de la tolerancia inmunológica central. Timectomía roedores ha demostrado ser un procedimiento esencial en la definición de la función del timo en la diferenciación de linfocitos, la auto-tolerancia y la inmunotolerancia en el contexto de trasplante de aloinjerto y la metástasis tumoral. La eliminación del timo de rata permite estudios con depleción de células T o la transferencia adoptiva de las poblaciones de células T definidas sin la reaparición de células T vírgenes nativos. Timectomías en roedores neonatales pueden llevarse a cabo usando una técnica de aspiración con resultados fiables 1. En ratas adultas, esta técnica está asociado con una tasa de mortalidad del 20% aproximada y con frecuencia resulta en una timectomía incompleta 2. Para lograr consistentemente timectomía completa en ratas adultas, se requiere la exposición pública del mediastino a través de una esternotomía media. Sin embargo, esteprocedimiento se asocia con complicaciones que incluyen lesión traqueal, hemorragia y neumotórax que conduce a una tasa de mortalidad global que oscila 1,5-6% 2- 4. En las últimas dos décadas las mejoras en las técnicas de timectomía han disminuido las complicaciones perioperatorias y han mejorado las tasas de supervivencia. La intubación endotraqueal permitiendo la ventilación con presión positiva ha disminuido las tasas de neumotórax 5. Métodos de intubación descritos anteriormente rango de la exposición abierta a la tráquea a los métodos menos invasivos que utilizan la visualización directa de las cuerdas vocales. Las complicaciones asociadas con el procedimiento de intubación incluyen lesión traqueal, la rotura de las cuerdas vocales, la intubación esofágica no intencional, y la hemorragia resultante de punción cardíaca o laceración de la vena cava superior. Además, la proximidad de los lóbulos tímicos inferiores a el revestimiento pleural puede resultar en neumotórax. Aquí se describe una técnica detimectomía a través de una incisión en la piel mínimamente invasiva 2 cm tras un método simple de la intubación endotraqueal usando un angiocatéter de extremo romo y la iluminación traqueal transcutánea. La timectomía implica una esternotomía 1,5 cm y un cierre de tres capas con la aplicación de pegamento quirúrgico para sellar el mediastino y minimizar la incidencia de hemorragia y complicaciones respiratorias. Este método da como resultado de forma fiable en la timectomía completa como lo demuestra la desaparición de las células T ingenuas CD4 + y CD8 + después de la timectomía y la ausencia de tejido tímico en la tinción IHC. Los tiempos operatorios y la mortalidad peri-procedimiento se reducen al mínimo.

<table border="0" cellpadding="0" cellspacing="0" width="718">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Name of Reagent/ Equipment</td>
			<td style="width:117px;">Company</td>
			<td style="width:152px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">2&#34; 14-gauge angiocatheter</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Operating microscope</td>
			<td style="width:117px;">Zeiss</td>
			<td style="width:152px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Warming pad</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:283px;">Heart rate and blood oxygenation monitor for rodents with foot sensors</td>
			<td style="width:117px;">Harvard Apparatus</td>
			<td style="width:152px;">ST1 72-8010, ST1 72-8098 (Rat foot sensor)</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Intubation apparatus (plastic with metal bar at the top)</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td style="width:167px;">See Figure 2&#160;</td>
		</tr>
		<tr height="67">
			<td height="67" style="height:67px;width:283px;">Small animal anesthesia system with induction box, isoflurane tank and O2 tank</td>
			<td style="width:117px;">Harvard Apparatus</td>
			<td style="width:152px;">ST1 72-6420</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:283px;">Small animal ventilator with tubing</td>
			<td style="width:117px;">CWE</td>
			<td style="width:152px;">12-02000 (ventilator)and 12-04000 (external valve assembly for mice/rats)</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">High-intensity Fiber-Optic Illuminator</td>
			<td style="width:117px;">Dolan Jenner</td>
			<td style="width:152px;">EEG 2823M</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Student standard pattern forceps</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">91100-16</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Fine straight scissors</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">14060-09</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Blunt-tipped Shea scissors</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">14105-12</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Small Alm retractor (for sternum)</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">17008-07</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Blunt Graefe forceps</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">11050-10</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Fine Dumont forceps</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">11254-20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">5-0 Maxon sutures</td>
			<td style="width:117px;">Ethicon</td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">4-0 Silk sutures (with cutting needle)</td>
			<td style="width:117px;">Ethicon</td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">6-0 Nylon suture</td>
			<td style="width:117px;">Ethicon</td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Cyanoacrylate glue (Endermil)</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Lubrication gel</td>
			<td style="width:117px;">Akorn Animal Health</td>
			<td style="width:152px;">NDC 17478-162-35</td>
			<td></td>
		</tr>
	</tbody>
</table>

complete thymectomy in adult rats with non-invasive endotracheal intubation

La timectomía en roedores neonatales es un procedimiento establecido y fiable para estudios inmunológicos. Sin embargo, en ratas adultas, las complicaciones de hemorragia y neumotórax de interrupción pleural pueden resultar en una tasa de mortalidad significativa. Este protocolo es un método simple de la timectomía rata que utiliza un mini-intubación endotraqueal y esternotomía. La intubación se logra con un método no invasivo y fácilmente reproducible y permite la ventilación con presión positiva para prevenir el neumotórax y una vía aérea controlada que permite el tiempo suficiente para la disección timo cuidado para minimizar la interrupción pleural. A 1,5 cm incisión esternal disminuye el contacto con los vasos mediastínicos y pleura, mientras que todavía proporciona visualización completa del timo. Después de la exposición del mediastino, el timo se retira mediante disección roma con una lupa. El espacio pleural se sella entonces por el cierre de la sutura de los músculos pre-traqueales seguido por la aplicación de pegamento quirúrgico. Latórax se cierra entonces por el cierre de sutura del esternón, seguido por el cierre de la sutura de la piel. Todos timectomías estaban completas como se evidencia por inmunohistoquímica (IHC) tinción del tejido mediastinal, y la ausencia de células T ingenuas por citometría de flujo, y el procedimiento tuvo una tasa de supervivencia del 96%. Este método es adecuado cuando se desea timectomía completa con mínimas complicaciones de realizar más estudios inmunológicos en ratas adultas sin timo.

Este procedimiento se realizó en ratas Lewis adultas (n = 26). El tiempo de operación promedio fue de 15 ± 3 min. No hubo mortalidad intraoperatoria. El tiempo de intubación media - a partir de la colocación de la rata en el aparato de intubación para el establecimiento de la ventilación - fue de 45 ± 5 seg. 24 ratas tuvieron una recuperación completa de la operación, sin evidencia de dificultades respiratorias o hemorragia a través día postoperatorio (POD) 14. Una rata desarrollado dificultad respiratoria e...

Watch this Scientific Journal Video about Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation at JoVE.com

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described.

La timectomía completo en ratas adultas con no invasiva La intubación endotraqueal

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and ...

complete-thymectomy-adult-rats-with-non-invasive-endotracheal

Research

JoVE Journal

Immunology and Infection

16.4K vistas.  Duke University Medical Center. Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described. 

Video: Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists. The value of 2P microscopy is that it provides rich spatiotemporal information regarding cell behaviors within intact tissues and in live mice. Leukocyte recruitment plays a significant role in host defense against infection and when unchecked, can contribute to inflammatory and autoimmune diseases. Studying leukocyte recruitment in vivo is technically challenging since cells are moving rapidly within vessels located deep within light scattering tissues. To date, most intravital imaging studies require surgical preparation to expose the blood vessels and tissues. To avoid the tissue damage and inflammation induced by surgery itself, here, we describe a non-invasive single-cell imaging approach that can be used to study leukocyte trafficking in the mouse footpad and phalanges. We discuss the technical aspects of our 2P imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Here, we describe a non-invasive two-photon (2P) microscopy approach to study leukocyte homing in the mouse footpad. We discuss the technical aspects of our tissue imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

De imagen no invasivas de Homing leucocitos y Migración In vivo</em

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists.  The value of 2P microscopy is that it ...

Investigación

Inmunología e Infección

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 to 40% attributable mortality6. Systemic candidiasis is normally controlled by innate immunity, and individuals with genetic defects in innate immune cell components such as phagocyte NADPH oxidase are more susceptible to candidemia7-9. Very little is known about the dynamics of C. albicans interaction with innate immune cells in vivo. Extensive in vitro studies have established that outside of the host C. albicans germinates inside of macrophages, and is quickly destroyed by neutrophils10-14. In vitro studies, though useful, cannot recapitulate the complex in vivo environment, which includes time-dependent dynamics of cytokine levels, extracellular matrix attachments, and intercellular contacts10, 15-18. To probe the contribution of these factors in host-pathogen interaction, it is critical to find a model organism to visualize these aspects of infection non-invasively in a live intact host. 
The zebrafish larva offers a unique and versatile vertebrate host for the study of infection. For the first 30 days of development zebrafish larvae have only innate immune defenses2, 19-21, simplifying the study of diseases such as disseminated candidiasis that are highly dependent on innate immunity. The small size and transparency of zebrafish larvae enable imaging of infection dynamics at the cellular level for both host and pathogen. Transgenic larvae with fluorescing innate immune cells can be used to identify specific cells types involved in infection22-24. Modified anti-sense oligonucleotides (Morpholinos) can be used to knock down various immune components such as phagocyte NADPH oxidase and study the changes in response to fungal infection5. In addition to the ethical and practical advantages of using a small lower vertebrate, the zebrafish larvae offers the unique possibility to image the pitched battle between pathogen and host both intravitally and in color. 
The zebrafish has been used to model infection for a number of human pathogenic bacteria, and has been instrumental in major advances in our understanding of mycobacterial infection3, 25. However, only recently have much larger pathogens such as fungi been used to infect larva5, 23, 26, and to date there has not been a detailed visual description of the infection methodology. Here we present our techniques for hindbrain ventricle microinjection of prim25 zebrafish, including our modifications to previous protocols. Our findings using the larval zebrafish model for fungal infection diverge from in vitro studies and reinforce the need to examine the host-pathogen interaction in the complex environment of the host rather than the simplified system of the Petri dish5.

The rapid development, small size and transparency of zebrafish are tremendous advantages for the study of innate immune control of infection1-4. Here we demonstrate techniques for infecting zebrafish larvae using the fungal pathogen Candida albicans by microinjection, methodology recently used to implicate phagocyte NADPH oxidase activity in control of fungal dimorphism5.

La imagen no invasiva de la candidiasis diseminada en larvas de pez cebra

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

No invasiva imágenes ópticas de la vasculatura linfática de un ratón

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Culturing Microglia from the Neonatal and Adult Central Nervous System

Microglia are the resident macrophage-like cells of the central nervous system (CNS) and, as such, have critically important roles in physiological and pathological processes such as CNS maturation in development, multiple sclerosis, and spinal cord injury. Microglia can be activated and recruited to action by neuronal injury or stimulation, such as axonal damage seen in MS or ischemic brain trauma resulting from stroke. These immunocompetent members of the CNS are also thought to have roles in synaptic plasticity under non-pathological conditions. We employ protocols for culturing microglia from the neonatal and adult tissues that are aimed to maximize the viable cell numbers while minimizing confounding variables, such as the presence of other CNS cell types and cell culture debris. We utilize large and easily discernable CNS components (e.g. cortex, spinal cord segments), which makes the entire process feasible and reproducible. The use of adult cells is a suitable alternative to the use of neonatal brain microglia, as many pathologies studied mainly affect the postnatal spinal cord. These culture systems are also useful for directly testing the effect of compounds that may either inhibit or promote microglial activation. Since microglial activation can shape the outcomes of disease in the adult CNS, there is a need for in vitro systems in which neonatal and adult microglia can be cultured and studied.

We outline methods for the efficient and quick isolation/culture of viable microglia from the neonatal cerebral cortex and adult spinal cord. The dissection and plating of cortical microglia can be accomplished within 90 minutes, with the subsequent microglial harvest taking place ~ 10 days following the initial dissection.

Microglia cultivo del Sistema Nervioso Central Neonatal y Adultos

Microglia are the resident macrophage-like cells of the central nervous system (CNS) and, as such, have critically important roles in physiological and ...

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal features of the natural infection. This protocol describes procedures for the establishment and evaluation of systemic infection due to neuropathogenic Escherichia coli K1 in the neonatal rat. Colonization of the gastrointestinal tract leads to dissemination of the pathogen along the gut-lymph-blood-brain course of infection and the model displays strong age dependency. A strain of E. coli O18:K1 with enhanced virulence for the neonatal rat produces exceptionally high rates of colonization, translocation to the blood compartment and invasion of the meninges following transit through the choroid plexus. As in the human host, penetration of the central nervous system is accompanied by local inflammation and an invariably lethal outcome. The model is of proven utility for studies of the mechanism of pathogenesis, for evaluation of therapeutic interventions and for assessment of bacterial virulence.

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Here, a procedure is described for the establishment of systemic infection in the neonatal rat with cultures of Escherichia coli K1. This non-invasive procedure permits colonization of the gastrointestinal tract, translocation of the pathogen to the systemic circulation, and invasion of the central nervous system at the choroid plexus.

Modelo no invasiva de neuropatogénico<em&gt; Escherichia coli</em&gt; La infección en Neonatología Rata

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal ...

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of causing systemic disease in both fish and humans. A better understanding of S. iniae disease pathogenesis requires an appropriate model system. The genetic tractability and the optical transparency of the early developmental stages of zebrafish allow for the generation and non-invasive imaging of transgenic lines with fluorescently tagged immune cells. The adaptive immune system is not fully functional until several weeks post fertilization, but zebrafish larvae have a conserved vertebrate innate immune system with both neutrophils and macrophages. Thus, the generation of a larval infection model allows the study of the specific contribution of innate immunity in controlling S. iniae infection.
The site of microinjection will determine whether an infection is systemic or initially localized. Here, we present our protocols for otic vesicle injection of zebrafish aged 2-3 days post fertilization as well as our techniques for fluorescent confocal imaging of infection. A localized infection site allows observation of initial microbe invasion, recruitment of host cells and dissemination of infection. Our findings using the zebrafish larval model of S. iniae infection indicate that zebrafish can be used to examine the differing contributions of host neutrophils and macrophages in localized bacterial infections. In addition, we describe how photolabeling of immune cells can be used to track individual host cell fate during the course of infection.

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Here, we present a protocol for the generation and imaging of a localized bacterial infection in the zebrafish otic vesicle.

Imagen no invasiva de la respuesta inmune innata en un modelo de larvas de pez cebra de<em&gt; Streptococcus iniae</em&gt; Infección

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of ...

A Non-invasive and Technically Non-intensive Method for Induction and Phenotyping of Experimental Bacterial Pneumonia in Mice

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant preclinical advances in our understanding of the underlying cellular and molecular pathogenesis. In vivo mouse models capture the integrated physiology and resilience of the host defense response in a manner not revealed by alternative, simplified ex vivo approaches. Several methods have been described in the literature for intrapulmonary inoculation of bacteria in mice, including aerosolization, intranasal delivery, peroral endotracheal cannulation under &#39;blind&#39; and visualized conditions, and transcutaneous endotracheal cannulation. All methods have relative merits and limitations. Herein, we describe in detail a non-invasive, technically non-intensive, inexpensive, and rapid method for intratracheal delivery of bacteria that involves aspiration (i.e., inhalation) by the mouse of an infectious inoculum pipetted into the oropharynx while under general anesthesia. This method can be used for pulmonary delivery of a wide variety of non-caustic biological and chemical agents, and is relatively easy to learn, even for laboratories with minimal prior experience with pulmonary procedures. In addition to describing the aspiration pneumonia method, we also provide step-by-step procedures for assaying the subsequent in vivo pulmonary innate immune response of the mouse, in particular, methods for quantifying bacterial clearance and the cellular immune response of the infected airway. This integrated and simple approach to pneumonia assessment allows for rapid and robust evaluation of the effect of genetic and environmental manipulations upon pulmonary innate immunity.

Several methods have been described in the literature for modeling bacterial pneumonia in mice. Herein, we describe a non-invasive, inexpensive, rapid method for inducing pneumonia via aspiration (i.e., inhalation) of a bacterial inoculum pipetted into the oropharynx. Downstream methods for assessment of the pulmonary innate immune response are also detailed.

Un método no invasivo y técnicamente no intensivos para la inducción y Fenotipificación Experimental de neumonía bacteriana en ratones

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant ...

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo fluorescence optical imaging (OI), in two different mouse models of inflammation: a rheumatoid arthritis (RA) and a contact hypersensitivity reaction (CHR) model. Light with a wavelength in the near infrared (NIR) window (650 - 950 nm) allows a deeper tissue penetration and minimal signal absorption compared to wavelengths below 650 nm. The major advantages using fluorescence OI is that it is cheap, fast and easy to implement in different animal models.
Activatable fluorescent probes are optically silent in their inactivated states, but become highly fluorescent when activated by a protease. Activated MMPs lead to tissue destruction and play an important role for disease progression in delayed-type hypersensitivity reactions (DTHRs) such as RA and CHR. Furthermore, MMPs are the key proteases for cartilage and bone degradation and are induced by macrophages, fibroblasts and chondrocytes in response to pro-inflammatory cytokines. Here we use a probe that is activated by the key MMPs like MMP-2, -3, -9 and -13 and describe an imaging protocol for near infrared fluorescence OI of MMP activity in RA and control mice 6 days after disease induction as well as in mice with acute (1x challenge) and chronic (5x challenge) CHR on the right ear compared to healthy ears.

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper explains the application of fluorescent imaging using an activatable optical imaging probe to visualize the in vivo activity of key matrix metalloproteinases in two different experimental models of inflammation.

No invasiva<em&gt; En Vivo</emHome Idiomas Ingresar a Epistemonikos Búsqueda avanzada Fluorescencia de imágenes ópticas de la actividad MMP inflamatoria utilizando un agente de imagen fluorescente activable

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo ...

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte extravasation might result in pathophysiological changes in the CNS. Thus, investigation of lymphocyte migration into the CNS is important to understand inflammatory CNS diseases and to develop new therapy approaches. Here we present an in vitro model of the human blood-brain barrier to study lymphocyte extravasation. Human brain microvascular endothelial cells (HBMEC) are confluently grown on a porous polyethylene terephthalate transwell insert to mimic the endothelium of the blood-brain barrier. Barrier function is validated by zonula occludens immunohistochemistry, transendothelial electrical resistance (TEER) measurements as well as analysis of evans blue permeation. This model allows investigation of the diapedesis of rare lymphocyte subsets such as CD56brightCD16dim/- NK cells. Furthermore, the effects of other cells, cytokines and chemokines, disease-related alterations, and distinct treatment regimens on the migratory capacity of lymphocytes can be studied. Finally, the impact of inflammatory stimuli as well as different treatment regimens on the endothelial barrier can be analyzed.

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Here, we describe a human blood-brain barrier model enabling to investigate lymphocyte transmigration into the central nervous system in vitro.

Análisis de linfocitos extravasación El uso de una<em&gt; In Vitro</em&gt; Modelo de la barrera sangre-cerebro humano

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte ...

A Non-invasive Way to Isolate and Phenotype Cells from the Conjunctiva

Traditionally, ocular surface cytology is studied with techniques such as spatula technology and brush technology. The problem with these techniques is that they may induce traumatic lesions on the surface of the eye, which can progress to scarring, eyelid deformity, limbal stem cell deficiency and in some cases, cause great discomfort to the subject. To avoid these clinical problems, impression cytology (IC) was developed to diagnose dry eye disease and later neoplasia, atopic disease, vernal keratoconjunctivitis and keratoconjunctivitis sicca. Typically, clinicians manually cut filter papers into required shapes and apply these to the ocular surface. Here, we describe how to perform IC using a commercially available medical device. This technique is explained here followed by immunophenotyping by flow cytometry. This technique requires less manual handling and causes less injury to the ocular surface.

The exposed normal ocular surface consists of cornea and conjunctiva. Epithelial cells, goblet cells and immune cells are present in the conjunctiva. Here, a non-invasive, technique of impression cytology is described using an impression cytology device and flow cytometry to analyze immune cells in the conjunctiva.

Una forma no invasiva de aislar y células fenotípicas de la conjuntiva

Traditionally, ocular surface cytology is studied with techniques such as spatula technology and brush technology. The problem with these techniques is ...