The authors would like to acknowledge Jonathan Chou, MD for his assistance on preparation and editing of the final manuscript and Wenzhong Liu for the OCT data. We would also like to acknowledge support from the Macula Society Research Grant (AAF), support from an unrestricted grant to Northwestern University from Research to Prevent Blindness, Inc., New York, NY, USA, and support from NIH-EY019951.

Todos los animales son tratados de acuerdo con la Guía del Cuidado y Uso de Animales de Laboratorio 2013 Edition, la Asociación para la Investigación en Visión y Oftalmología (ARVO) Declaración para el uso de animales en Oftálmica y Vision Research, y aprobada por el Animal Institucional Cuidado y el empleo Comisión de la Universidad Northwestern. Nota: El siguiente procedimiento se puede realizar en su totalidad con un operador; sin embargo, se llevó a cabo mucho más eficiente con dos operadores con las tareas se dividen en consecuencia. 1. Prepare la estación láser y Pre-láser <ol><li> Coloque el láser y lámpara de hendidura donde puede ser fácilmente accesible. A su vez en láser y ajustado a los parámetros predeterminados (por ejemplo, 75 micras de tamaño punto, 100 mW de potencia, 100 ms de duración). PRECAUCIÓN: Asegúrese de que el operador lleva todas las señales de seguridad láser animales y equipo de protección personal y de seguridad láser pertinente se muestran fuera de la sala de procedimientos. <ol><li> Antes de usar cualquier animales de experimentación, encontrar parámetros ideales utilizando una calibración, el ratón no experimental. Los parámetros del láser dependerá del láser utilizado. Ideal parámetros son definidos por el ajuste de la potencia del láser más bajo que hace que constantemente la formación de &quot;burbuja&quot; cuando se enfoca correctamente. Ver video de ejemplo de lesión con la formación adecuada &quot;burbuja&quot;. </li></ol></li><li> Prepare la estación de pre-láser de modo que la anestesia, todas las gotas para los ojos (tropicamida, tetracaína, y lágrimas artificiales) animales más cálido, toallitas de tejidos, y son fácilmente a su alcance para el operador. <ol><li> Asegúrese de que los animales más cálido se precalienta para corregir la temperatura (37 ° C) antes de la inyección de anestésico en primer ratón con el fin de evitar la hipotermia inducida por anestesia. </li></ol></li><li> Coloque etapa del ratón en más cálido por lo que puede retener el calor y permanecer caliente una vez que comience el procedimiento láser. </li></ol> 2. Ratón Anestesia y Preparación Pre-láser <ol><li> Antes de inyectaring anestesia, inspeccione el ojo macroscópicamente para asegurarse de que no tiene deformaciones o anomalías que disminuyen la claridad de la córnea (por ejemplo, cataratas). </li><li> Pesar ratón. </li><li> Apunte el peso, el sexo y el número de identificación de los animales. </li><li> Usando peso, calcular la cantidad adecuada de anestesia que se utilizará en base a las directrices dadas por la institución (por ejemplo, 100 mg / kg de clorhidrato de ketamina, 10 mg / kg de xilazina O tribromoetanol 250 mg / kg; para un ratón de 20 g inyectar 0,20 ml de xilazina / cóctel de ketamina O 0,25 ml de tribromoetanol). Tener una tabla de dosis previamente calculado por peso en incrementos de 1 g con el fin de reducir los errores matemáticos. </li><li> Scruff ratón e inyectar anestesia intraperitoneal basan en cálculos del paso 2.4. </li><li> Coloca el ratón sobre los animales más cálido y espere hasta que el ratón está completamente anestesiado marcando el reflejo pedal a través pizca dedo del pie (aproximadamente 3-5 minutos). </li><li> Enrolle un tejido limpiar y proteger la zona nasal del ratón para evitar que aspiratión de líquido roll-off. Ratón rollo en su lado y colocar una gota (aproximadamente 30 l) de clorhidrato de tetracaína en cada ojo para la anestesia tópica. Espere 2 minutos para la solución tenga efecto. </li><li> Repita el paso 2.7 con una gota de tropicamida tópico para la dilatación pupilar. Como alternativa, utilice clorhidrato de fenilefrina (2,5%) para la dilatación. </li><li> Espere 2 minutos para las soluciones que se apliquen; mantener a los animales en el más cálido durante este tiempo. </li><li> Una vez transcurrido el tiempo adecuado, colocar rápidamente el ratón en el escenario del ratón y colocar la etapa en la barbilla resto de lámpara de hendidura. </li><li> Encienda la lámpara de hendidura para el brillo de la luz más baja y comprobar el grado de dilatación pupilar. Si alumno no se dilata de manera adecuada (aproximadamente 2,5 a 3 mm), devuelva ratón para animales más caliente y esperar. Alternativamente, administrar otra gota de tropicamida. Una vez que el ojo es suficientemente dilatada, continúe con el procedimiento con láser. </li></ol> Procedimiento 3. Láser Nota: Asegúrese de otra personas en la sala de llevar gafas de protección cuando están lejos de láser protegido lámpara de hendidura ocular <ol><li> Ajuste la colocación del ratón en la etapa del ratón, por lo que está en una posición ideal para la visualización de nervio óptico (ver 3.1.2). <ol><li> Oriente el ratón sobre su soporte de manera que quede en posición horizontal, perpendicular a la hendidura del haz de la lámpara, con la cabeza en un lado y la cola en el otro. Colocación Ideal ratón hará que la visualización del nervio óptico mucho más fácil una vez que se aplica hoja de la cubierta. </li><li> Ligeramente gire el ratón por lo que es en un ángulo de 170 ° ~ con la cabeza más cerca de operador de láser. </li><li> Asegúrese de que el ratón es tan cerca como sea posible de la lámpara de hendidura, sin embargo, todavía en una posición en la que es estable y donde la mano del operador tendrá espacio suficiente para la manipulación fina. </li></ol></li><li> Después de que el ratón está en una posición ideal, coloque una gota de solución de lágrimas artificiales en un 25 mm x 25 mm cubreobjetos de vidrio. <ol><li> Coloque una gota de solución de lágrima artificial en del opp del ratónojo osite - este ojo se asegurará se hidrata y la formación ayuda retardo de cataratas. </li></ol></li><li> Mantenga esquina de cubreobjetos entre el pulgar y los dedos de puntero; posición de modo que el vidrio se comprime entre las puntas de los dedos de ambas. </li><li> Envolver suavemente los tres dedos restantes en todo el cuerpo del animal para el apoyo y la estabilización mano. Posición de la mano de modo que el cubreobjetos de vidrio se puede colocar fácilmente en el ojo del ratón. <ol><li> Asegúrese de que la muñeca se estabiliza en una superficie firme con el fin de reducir el temblor de las manos. </li></ol></li><li> Una vez que se obtiene la posición estable, presione con cuidado cubreobjetos de vidrio (con la gota de lágrima artificial todavía adherido) en el ojo del ratón. <ol><li> Asegúrese de que el cubreobjetos se coloca lo más perpendicular posible al rayo láser con el fin de prevenir la dispersión de haz de láser o la reflexión. El cubreobjetos actúa como una lente de contacto para aplanar la córnea. </li></ol></li><li> Mira a través de la lámpara de hendidura y con la mano libre de palanca centrarse untretina il se pueden visualizar. La retina tendrá un color claro-amarillo / rojo dependiendo de la ubicación visualizada, vasos, rojas definidas serán visibles. </li><li> Poco a poco y con cuidado manipular cabeza de ratón y / o cubreobjetos hasta visualizar el nervio óptico. El nervio óptico será de color amarillo con múltiples vasos que irradian de él. </li><li> Una vez que el operador ha confirmado la visualización del nervio óptico, encienda el láser de enfoque del haz. </li><li> Una vez que haz de láser se ha encendido, maniobrar enfoque del láser de haz a la posición deseada (aproximadamente 1 diámetro del disco del nervio óptico). </li><li> Enfoque rayo láser en la RPE del fondo de ojo. El enfoque correcto se logra haciendo que el rayo láser más nítida y más clara. Si el objetivo del haz se ve oval o fuera de foco, alternar el foco de la lámpara de hendidura o colocar cubreobjetos de vidrio. </li><li> Una vez que el haz de encuadre se centra en RPE, iniciar la administración láser utilizando gatillo pie del láser. <ol><li> Asegúrese de evitar los vasos de la retina para prevenir intraocular quemorrhage. </li></ol></li><li> Atento a la aparición de una burbuja inmediatamente después de la administración láser. El contorno de la disparo de láser debe ser clara y no turbia de ninguna manera. <ol><li> Si el disparo de láser no da lugar a la formación de burbujas, o área de impacto se ve nebuloso (aspecto turbio con borde circular mal definido vs. clara frontera claramente definida de un impacto con éxito), o si la hemorragia es visto después de la administración de láser, NO incluir estas lesiones para el análisis futuro. </li></ol></li><li> Repita los pasos 03.10 a 03.12 para todas las posiciones deseadas de la CNV (por lo general a las 3, 6, 9, y 12 en punto alrededor del nervio óptico). Si se aplican las inducciones láser en aproximadamente misma distancia del nervio óptico, el nuevo enfoque no debería ser necesario. Sin embargo, debido a la fuerte curvatura del ojo del ratón y pequeñas variaciones que puedan existir en la retina, la reorientación de la viga puede ser necesario entre las administraciones de láser consecutivos. </li><li> Grabar en un cuaderno la ubicación y el resultado de correoadministración disparo de láser ada y resultado (éxito, nebuloso, hemorragia, etc.) de cada disparo se administra para el ojo. Asegúrese de colocar el láser en modo de espera cuando no esté en uso. </li><li> Repita 03/01 a 03/14 para el ratón del otro ojo, si es necesario, usando la mano opuesta para la estabilización y un nuevo cubreobjetos. </li><li> Después se administran todos los disparos de láser deseados, apague láser y con lámpara de hendidura. </li><li> Deseche cubreobjetos y el lugar del ratón en más cálido para la recuperación de la anestesia. Macroscópicamente inspeccionar ojo por cualquier lesión y colocar una gota de solución de lágrimas artificiales para mantener el ojo hidratado y potencialmente prevenir futuro desarrollo de cataratas. Una vez que el ratón se recupere de la anestesia, volver a la jaula. </li></ol><table fo:keep-together.within-page="1"><tbody><tr><td></td><td> AVERTIN </td><td> AVERTIN </td><td> AVERTIN </td><td> XYL / KET </td><td> XYL / KET </td></tr><tr><td> Ratón Peso (g) </td><td> Dosis (mg / kg) </td><td&gt; concentración de la solución (mg / ml) </td><td> Dosis anestésico (ml) </td><td> Dosis (mg / kg) </td><td> Dosis anestésico (ml) </td></tr><tr><td> 15 </td><td> 250 </td><td> 20 </td><td> 0.1875 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.15 </td></tr><tr><td> 16 </td><td> 250 </td><td> 20 </td><td> 0.2 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.16 </td></tr><tr><td> 17 </td><td> 250 </td><td> 20 </td><td> 0.2125 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.17 </td></tr><tr><td> 18 </td><td> 250 </td><td> 20 </td><td> 0,225 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.18 </td></tr><tr><td> 19 </td><td> 250 </td><td> 20 </td><td> 0.2375 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.19 </td></tr><tr><td> 20 </td><td> 250 </td><td> 20 </td><td> 0.25</td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.2 </td></tr><tr><td> 21 </td><td> 250 </td><td> 20 </td><td> 0.2625 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.21 </td></tr><tr><td> 22 </td><td> 250 </td><td> 20 </td><td> 0,275 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.22 </td></tr><tr><td> 23 </td><td> 250 </td><td> 20 </td><td> 0.2875 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.23 </td></tr><tr><td> 24 </td><td> 250 </td><td> 20 </td><td> 0.3 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.24 </td></tr><tr><td> 25 </td><td> 250 </td><td> 20 </td><td> 0.3125 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.25 </td></tr><tr><td> 26 </td><td> 250 </td><td> 20 </td><td> 0,325 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.26 </td></tr><tr><td> 27 </td><td> 250 </td><td&gt; 20 </td><td> 0.3375 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.27 </td></tr><tr><td> 28 </td><td> 250 </td><td> 20 </td><td> 0.35 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.28 </td></tr><tr><td> 29 </td><td> 250 </td><td> 20 </td><td> 0.3625 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.29 </td></tr><tr><td> 30 </td><td> 250 </td><td> 20 </td><td> 0,375 </td><td> 100 mg / kg de ketamina; 10 mg / kg de xilazina </td><td> 0.3 </td></tr></tbody></table> Tabla 1: Dosis XyIKet Chart.

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The authors declare they have no competing financial interests.

Existen múltiples factores que pueden afectar la entrega de láser y la consiguiente desarrollo de la lesión NVC tras éxito láser inducción. Estos factores deben ser controlados por y estandarizados con el fin de tener los resultados más confiables. El más pertinente de estos factores son la selección del ratón (genotipo, edad y sexo), la selección de anestésico, y la configuración de láser. El modelo específico de ratón utilizada puede tener un efecto significativo en el curs...

Edad relacionados con la degeneración macular (AMD) es una de las principales causas de ceguera en personas mayores de 50 1.3. AMD se puede clasificar en dos formas: atrófica (&quot;secar&quot;) y AMD neovascular (&quot;húmeda&quot;) de AMD. El primero se caracteriza por la atrofia geográfica del epitelio pigmentario de la retina (RPE), coriocapilar, y los fotorreceptores, mientras que el último se caracteriza por la invasión de vasos anormales de la coroides en las capas externas de la retina causando fugas, hemorragia y fibrosis, y en última instancia que conduce a la ceguera 1,2. De las dos formas, AMD neovascular da cuenta de la mayoría de la pérdida de visión 1. Afortunadamente, esta forma tiene numerosas opciones de gestión farmacológicos eficaces, mientras que su contraparte atrófica Actualmente no ha probado los tratamientos médicos 3. Además, debido a la forma neovascular ha sido re-capitulado fácilmente en un modelo animal, ha sido más ampliamente accesible a A básicaInvestigación MD explorar los mecanismos patológicos subyacentes a fin de desarrollar nuevas terapias 4. El primer modelo animal experimental de la neovascularización coroidea (CNV) fue desarrollado por Ryan et al. en primates no humanos 5. Esta ruptura modelo inducido de la membrana de Bruch a través de la fotocoagulación con láser, lo que provocó una respuesta inflamatoria local dando como resultado en la angiogénesis similar a la observada en neovascular AMD. Se encontró que la progresión histopatológico de la angiogénesis inducción post-láser para imitar la DMAE neovascular, que confirmó la validez del modelo de 6. Los primates no humanos ofrecer la anatomía más similares a los humanos, pero desafortunadamente, son caros de mantener, no puede ser manipulado fácilmente genéticamente, y tienen un curso lento tiempo de progresión de la enfermedad 7. En contraste, los modelos de roedores son mucho más rentable para mantener, pueden ser manipulados genéticamente con relativa facilidad, y tienen un mucho más rápido course de progresión de la enfermedad (experimentos pueden llevarse a cabo en una escala de tiempo de semana frente a meses). Estos experimentos solamente deben llevarse a cabo en roedores pigmentadas ya que es muy difícil de visualizar en animales albinos. El modelo CNV inducida por láser del ratón, primero desarrollado por el grupo Campochiaro a finales del 90 de 10, se ha convertido en el modelo animal dominante en la mayoría de los estudios recientes 11-16. Debido a la patogénesis complejo y todavía no está claro de CNV, el modelo de láser se ha aplicado en todos los aspectos de la investigación AMD húmeda que van desde el estudio de los mecanismos moleculares de conducción de la angiogénesis para la evaluación de nuevas modalidades de tratamiento para el futuro uso humano. Por ejemplo, Sakurai et al. y Espinosa-Heidmann et al. utilizado el modelo de láser para investigar el efecto de los macrófagos en el desarrollo de la CNV utilizando ratones transgénicos y los tratamientos farmacológicos de agotamiento 15, 16. Giani et al. y Hoerster et al. usado tomografía óptica de coherencia-(OCT) para la imagen de la CNV en un esfuerzo por caracterizar la progresión de la CNV y comparar los resultados con los hallazgos histopatológicos observados en imágenes de OCT 12,17 inducida por láser. Finalmente, los estudios que implican la inyección intravítrea de agentes anti-angiogénicos se han utilizado como pre-requisitos para los ensayos en humanos y fueron vitales en el desarrollo de la primera generación de agentes anti-VEGF utilizadas en la gestión de AMD neovascular hoy 10,18,19. Modelos alternativos para la CNV experimental utilizan métodos quirúrgicos para inducir la CNV. Este procedimiento implica la inyección de sustancias pro-angiogénicos (por ejemplo, vectores virales recombinantes que sobreexpresan VEGF, inyección subretiniana de células del EPR y / o perlas de poliestireno) para imitar el aumento de la expresión del VEGF se ve en neovascular AMD, con el objetivo de causar la angiogénesis 8,20. Sin embargo, este método produce una incidencia drásticamente inferior de la neovascularización; estos estudios mostraron que en la CNVRatones C57 / BL6 se produce en 31% de las inyecciones frente al ~ 70% de éxito visto en el método de la fotocoagulación con láser en la misma cepa de ratones 8,14. Por estas razones, y dadas las ventajas del uso de roedores en comparación con primates no humanos, el modelo de ratón de la CNV inducida por láser se ha convertido en el modelo animal estándar para la mayoría de CNV neovasculares experimentos de estudio de AMD 8. El ojo es un ratón, delicado tejido minúsculo para trabajar. Maniobras del ojo para visualizar la retina es difícil y requiere mucha práctica hasta que se alcanza la maestría. Esta tarea se complica por el hecho de que debe ser aprendido con la mano dominante y no dominante. Por otra parte, después de que los finos movimientos necesarios para visualizar la retina se han aprendido, la coordinación entre ambas manos y el pedal de funcionamiento del láser son importantes. En este trabajo, hemos tratado de destilar los desafíos de aprender todas las manipulaciones físicas involucradas en la CNV proc inducida por láseredure en una guía que ayudará a los operadores a lograr un rápido éxito con este modelo.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>532 nm (green) argon ophthalmic laser</td>
			<td>IRIDEX</td>
			<td>GLx</td>
			<td>any ophthalmic 532 nm (green) argon laser can be used</td>
		</tr>
		<tr>
			<td>slit lamp</td>
			<td>Carl Zeiss</td>
			<td>30SL-M</td>
			<td>any slit lamp can be used as long as it is compatible with the laser</td>
		</tr>
		<tr>
			<td>tribromoethanol</td>
			<td>Sigma</td>
			<td>T48402-25G</td>
			<td>used to make anesthetic</td>
		</tr>
		<tr>
			<td>tert-amyl alcohol</td>
			<td>Sigma</td>
			<td>152463-1L</td>
			<td>used to make anesthetic</td>
		</tr>
		<tr>
			<td>amber glass vials + septa</td>
			<td>Wheaton</td>
			<td>WH-223696</td>
			<td>tribromoethanol storage</td>
		</tr>
		<tr>
			<td>tissue wipes</td>
			<td>VWR</td>
			<td>82003-820</td>
			<td>miscellaneous&#160;</td>
		</tr>
		<tr>
			<td>1% tropicamide</td>
			<td>Falcon Pharmaceuticals</td>
			<td>RXD2974251</td>
			<td>pupillary dilation</td>
		</tr>
		<tr>
			<td>0.5% tetracaine hydrochloride</td>
			<td>Alcon</td>
			<td>&#160;0065-0741-12</td>
			<td>topical anesthesia</td>
		</tr>
		<tr>
			<td>artificial tears</td>
			<td>Alcon</td>
			<td>&#160;58768-788-25</td>
			<td>hydration</td>
		</tr>
		<tr>
			<td>heat therapy pump (for animal warming)</td>
			<td>Kent Scientific</td>
			<td>HTP-1500</td>
			<td>used to maintain animal body temp</td>
		</tr>
		<tr>
			<td>warming pad</td>
			<td>Kent Scientific</td>
			<td>TPZ-0510EA</td>
			<td>maintains animal body temperature</td>
		</tr>
		<tr>
			<td>30 gauge insulin needles</td>
			<td>BD</td>
			<td>328418</td>
			<td>IP anesthesia injection</td>
		</tr>
		<tr>
			<td>scale</td>
			<td>American Weigh Scale</td>
			<td>AWS-1KG-BLK</td>
			<td>mouse weighing</td>
		</tr>
		<tr>
			<td>cover slip (25 mm x 25 mm)</td>
			<td>VWR</td>
			<td>48366089</td>
			<td>flatten cornea to visualize mouse retina</td>
		</tr>
		<tr>
			<td>xylazine</td>
			<td>obtained from institution</td>
			<td>obtained from institution</td>
			<td>anesthesia</td>
		</tr>
		<tr>
			<td>ketamine</td>
			<td>obtained from institution</td>
			<td>obtained from institution</td>
			<td>anesthesia</td>
		</tr>
		<tr>
			<td>Volocity</td>
			<td>PerkinElmer</td>
			<td></td>
			<td>used for volumetric re-construction</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>National Institutes of Health</td>
			<td></td>
			<td>used for image analysis</td>
		</tr>
	</tbody>
</table>

a mouse model for laser-induced choroidal neovascularization

The mouse laser-induced choroidal neovascularization (CNV) model has been a crucial mainstay model for neovascular age-related macular degeneration (AMD) research. By administering targeted laser injury to the RPE and Bruch&#8217;s membrane, the procedure induces angiogenesis, modeling the hallmark pathology observed in neovascular AMD. 
First developed in non-human primates, the laser-induced CNV model has come to be implemented into many other species, the most recent of which being the mouse. Mouse experiments are advantageously more cost-effective, experiments can be executed on a much faster timeline, and they allow the use of various transgenic models. The miniature size of the mouse eye, however, poses a particular challenge when performing the procedure. Manipulation of the eye to visualize the retina requires practice of fine dexterity skills as well as simultaneous hand-eye-foot coordination to operate the laser. However, once mastered, the model can be applied to study many aspects of neovascular AMD such as molecular mechanisms, the effect of genetic manipulations, and drug treatment effects. 
The laser-induced CNV model, though useful, is not a perfect model of the disease. The wild-type mouse eye is otherwise healthy, and the chorio-retinal environment does not mimic the pathologic changes in human AMD. Furthermore, injury-induced angiogenesis does not reflect the same pathways as angiogenesis occurring in an age-related and chronic disease state as in AMD.
Despite its shortcomings, the laser-induced CNV model is one of the best methods currently available to study the debilitating pathology of neovascular AMD. Its implementation has led to a deeper understanding of the pathogenesis of AMD, as well as contributing to the development of many of the AMD therapies currently available.

La cuantificación de las lesiones CNV se puede realizar mediante el análisis de coroides-plana montada utilizando la tinción de inmunofluorescencia para etiquetar los vasos de la CNV. Los dos métodos utilizados con mayor frecuencia de la preparación del tejido son etiquetado FITC-dextrano, hecho a través de la perfusión inmediatamente antes del sacrificio de los animales, o post-mortem inmuno-tinción con un marcador de células endoteliales. Ambos métodos se han descrito anteriormente en detalle 13,14,21;<...

Watch this Scientific Journal Video about A Mouse Model for Laser-induced Choroidal Neovascularization at JoVE.com

A Mouse Model for Laser-induced Choroidal Neovascularization

Here, we present the mouse laser-induced choroidal neovascularization (CNV) protocol, an experimental model that re-creates the vascular hallmarks of neovascular age-related macular degeneration (AMD). Once mastered, it can reliably and effectively induce CNV as a model system to test various experimental measures.

Un modelo de ratón para inducida por láser neovascularización coroidea

The mouse laser-induced choroidal neovascularization (CNV) model has been a crucial mainstay model for neovascular age-related macular degeneration (AMD) ...

a-mouse-model-for-laser-induced-choroidal-neovascularization

Research

JoVE Journal

Medicine

17.5K vistas.  Northwestern University Feinberg School of Medicine. Here, we present the mouse laser-induced choroidal neovascularization (CNV) protocol, an experimental model that re-creates the vascular hallmarks of neovascular age-related macular degeneration (AMD). Once mastered, it can reliably and effectively induce CNV as a model system to test various experimental measures.

Video: A Mouse Model for Laser-induced Choroidal Neovascularization

A Laser-induced Mouse Model of Chronic Ocular Hypertension to Characterize Visual Defects

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model of ocular hypertension to mimic human high-tension glaucoma. Here laser illumination is applied to the corneal limbus to photocoagulate the aqueous outflow, inducing angle closure. The changes of IOP are monitored using a rebound tonometer before and after the laser treatment. An optomotor behavioral test is used to measure corresponding changes in visual capacity. The representative result from one mouse which developed sustained IOP elevation after laser illumination is shown. A decreased visual acuity and contrast sensitivity is observed in this ocular hypertensive mouse. Together, our study introduces a valuable model system to investigate neuronal degeneration and the underlying molecular mechanisms in glaucomatous mice.

Chronic ocular hypertension is induced using laser photocoagulation of the trabecular meshwork in mouse eyes. The intraocular pressure (IOP) is elevated for several months after laser treatment. The decrease of visual acuity and contrast sensitivity of experimental animals are monitored using the optomotor test.

Un modelo de ratón inducida por láser de la hipertensión ocular crónica a Caracterizar los defectos visibles

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model ...

Investigación

Medicina

Detecting Abnormalities in Choroidal Vasculature in a Mouse Model of Age-related Macular Degeneration by Time-course Indocyanine Green Angiography

Indocyanine Green&#160;Angiography (or ICGA) is a technique performed by ophthalmologists to diagnose abnormalities of the choroidal and retinal vasculature of various eye diseases such as age-related macular degeneration (AMD). ICGA is especially useful to image the posterior choroidal vasculature of the eye due to its capability of penetrating through the pigmented layer with its infrared spectrum. ICGA time course can be divided into early, middle, and late phases. The three phases provide valuable information on the pathology of eye problems. Although time-course ICGA by intravenous (IV) injection is widely used in the clinic for the diagnosis and management of choroid problems, ICGA by intraperitoneal injection (IP) is commonly used in animal research. Here we demonstrated the technique to obtain high-resolution ICGA time-course images in mice by tail-vein injection and confocal scanning laser ophthalmoscopy. We used this technique to image the choroidal lesions in a mouse model of age-related macular degeneration. Although it is much easier to introduce ICG to the mouse vasculature by IP, our data indicate that it is difficult to obtain reproducible ICGA time course images by IP-ICGA. In contrast, ICGA via tail vein injection provides high quality ICGA time-course images comparable to human studies. In addition, we showed that ICGA performed on albino mice gives clearer pictures of choroidal vessels than that performed on pigmented mice. We suggest that time-course IV-ICGA should become a standard practice in AMD research based on animal models.

Indocyanine Green Angiography (or ICGA) performed by tail vein injection provides high quality ICGA time course images to characterize abnormalities in mouse choroid.

La detección de anormalidades en la vasculatura coroidea en un modelo de ratón de la relacionada con la edad degeneración macular por Time-curso de indocianina verde Angiografía

Indocyanine Green&#160;Angiography (or ICGA) is a technique performed by ophthalmologists to diagnose abnormalities of the choroidal and retinal ...

An Alkali-burn Injury Model of Corneal Neovascularization in the Mouse

Under normal conditions, the cornea is avascular, and this transparency is essential for maintaining good visual acuity. Neovascularization (NV) of the cornea, which can be caused by trauma, keratoplasty or infectious disease, breaks down the so called &lsquo;angiogenic privilege' of the cornea and forms the basis of multiple visual pathologies that may even lead to blindness. Although there are several treatment options available, the fundamental medical need presented by corneal neovascular pathologies remains unmet. In order to develop safe, effective, and targeted therapies, a reliable model of corneal NV and pharmacological intervention is required. Here, we describe an alkali-burn injury corneal neovascularization model in the mouse. This protocol provides a method for the application of a controlled alkali-burn injury to the cornea, administration of a pharmacological compound of interest, and visualization of the result. This method could prove instrumental for studying the mechanisms and opportunities for intervention in corneal NV and other neovascular disorders.

Neovascularization (NV) of the cornea can complicate multiple visual pathologies. Utilizing a controlled, alkali-burn injury model, a quantifiable level of corneal NV can be produced for mechanistic study of corneal NV and evaluation of potential therapies for neovascular disorders.

Una Lesión álcalis quemar modelo de neovascularización corneal en el ratón

Under normal conditions, the cornea is avascular, and this transparency is essential for maintaining good visual acuity. Neovascularization (NV) of the ...

Renal Ischaemia Reperfusion Injury: A Mouse Model of Injury and Regeneration

Renal ischaemia reperfusion injury (IRI) is a common cause of acute kidney injury (AKI) in patients and occlusion of renal blood flow is unavoidable during renal transplantation. Experimental models that accurately and reproducibly recapitulate renal IRI are crucial in dissecting the pathophysiology of AKI and the development of novel therapeutic agents. Presented here is a mouse model of renal IRI that results in reproducible AKI. This is achieved by a midline laparotomy approach for the surgery with one incision allowing both a right nephrectomy that provides control tissue and clamping of the left renal pedicle to induce ischaemia of the left kidney. By careful monitoring of the clamp position and body temperature during the period of ischaemia this model achieves reproducible functional and structural injury. Mice sacrificed 24 hr&#160;following surgery demonstrate loss of renal function with elevation of the serum or plasma creatinine level as well as structural kidney damage with acute tubular necrosis evident. Renal function improves and the acute tissue injury resolves during the course of 7 days following renal IRI such that this model may be used to study renal regeneration. This model of renal IRI has been utilized to study the molecular and cellular pathophysiology of AKI as well as analysis of the subsequent renal regeneration.

The mouse model of renal ischaemia reperfusion injury described here comprises of a right nephrectomy that provides control tissue and clamping of the left renal pedicle to induce ischaemia that results in acute kidney injury. This model uses a midline laparotomy approach with all steps performed via one incision.

La isquemia reperfusión renal: un modelo de ratón de la lesión y regeneración

Renal ischaemia reperfusion injury (IRI) is a common cause of acute kidney injury (AKI) in patients and occlusion of renal blood flow is unavoidable ...

A Mouse Fetal Skin Model of Scarless Wound Repair

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs across species but is age dependent. The transition from a scarless to scarring phenotype occurs in the third trimester of pregnancy in humans and around embryonic day 18 (E18) in mice. However, this varies with the size of the wound with larger defects generating a scar at an earlier gestational age. The emergence of lineage tracing and other genetic tools in the mouse has opened promising new avenues for investigation of fetal scarless wound healing. However, given the inherently high rates of morbidity and premature uterine contraction associated with fetal surgery, investigations of fetal scarless wound healing in vivo require a precise and reproducible surgical model. Here we detail a reliable model of fetal scarless wound healing in the dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

During mammalian development, early gestational skin wounds heal without a scar. Here we detail a reliable and reproducible model of fetal scarless wound healing in the cutaneous dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

Un ratón fetal Modelo de la piel de la reparación de heridas sin cicatrices

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs ...

A Neonatal Mouse Spinal Cord Compression Injury Model

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal cord. A major current area of research is focused on the mechanisms of adaptive plasticity that underlie spontaneous or induced functional recovery following SCI. Spontaneous functional recovery is reported to be greater early in life, raising interesting questions about how adaptive plasticity changes as the spinal cord develops. To facilitate investigation of this dynamic, we have developed a SCI model in the neonatal mouse. The model has relevance for pediatric SCI, which is too little studied. Because neural plasticity in the adult involves some of the same mechanisms as neural plasticity in early life1, this model may potentially have some relevance also for adult SCI. Here we describe the entire procedure for generating a reproducible spinal cord compression (SCC) injury in the neonatal mouse as early as postnatal (P) day 1. SCC is achieved by performing a laminectomy at a given spinal level (here described at thoracic levels 9-11) and then using a modified Yasargil aneurysm mini-clip to rapidly compress and decompress the spinal cord. As previously described, the injured neonatal mice can be tested for behavioral deficits or sacrificed for ex vivo physiological analysis of synaptic connectivity using electrophysiological and high-throughput optical recording techniques1. Earlier and ongoing studies using behavioral and physiological assessment have demonstrated a dramatic, acute impairment of hindlimb motility followed by a complete functional recovery within 2 weeks, and the first evidence of changes in functional circuitry at the level of identified descending synaptic connections1.

This article describes a method for generating a reproducible spinal cord compression injury (SCI) in the neonatal mouse. The model provides an advantageous platform for studying mechanisms of adaptive plasticity that underlie spontaneous functional recovery.

Un ratón neonatal lesión de médula espinal de compresión Modelo

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal ...

Mouse Model for Pancreas Transplantation Using a Modified Cuff Technique

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the availability of the widest range of molecular probes and reagents to perform in vivo as well as in vitro studies. Based on our experience with various murine transplantation models, we developed a heterotopic pancreas transplantation model in mice with the intent to analyze mechanisms underlying severe ischemia reperfusion injury-associated early graft damage. In contrast to previously described techniques using suture techniques, herein we describe a new procedure using a non-suture cuff technique. 
In recent years, we have performed more than 300 pancreas transplantations in mice with an overall success rate of &gt;90%, a success rate never described before in mouse pancreas transplantation. The backbone of this non-suture cuff technique for graft revascularization consists of two major steps: (I) pulling the recipient vessel over a polyethylene/polyamide cuff and fixing it with a circumferential ligature, and (II) placing the donor vessel over the everted recipient vessel and fixing it with a second circumferential ligature. The resultant continuity of the endothelial layer results in less thrombogenic lesions with high patency rates and, finally, high success rates.
In this model, arterial anastomosis is achieved by pulling the abdominal aorta of the donor graft over the everted common carotid artery of the recipient animal. Venous drainage of the graft is achieved by pulling the portal vein of the graft over the everted external jugular vein of the recipient. This manuscript provides details and crucial steps of the organ recovery and organ implantation procedures, which will allow researchers with microsurgical skills to perform the transplantation successfully in their laboratories.

Among abdominal solid organ transplantation, pancreatic grafts are prone to develop severe ischemia reperfusion injury-associated graft damage, leading eventually to early graft loss. This protocol describes a model of murine pancreas transplantation using a non-suture cuff technique, ideally suited for analyzing these early, deleterious damages.

Modelo murino de trasplante de páncreas mediante una técnica modificada del pun ¢ o

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the ...

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among laboratories. The goal of this study was to describe the microsurgical procedure used to implant an intravesical tube and the experimental technique for recording urinary bladder pressure in an awake, freely moving mouse. In addition, experimental data is presented to show how surgery, as well as tubing type and size, affect lower urinary tract function and recording sensitivity. The effect of tube diameter on pressure recording was assessed in both polyethylene and polyurethane tubing with different internal diameters. Subsequently, the best performing tube from both materials was surgically implanted into the dome of the urinary bladder of male C57BL/6 mice. Twelve-hour, overnight micturition frequency was recorded in healthy, intact animals and animals 2, 3, 5, and 7 days post-surgery. At harvest, bladders were assessed for signs of swelling using gross observation and were subsequently processed for pathological analysis. The greatest extent of bladder swelling was observed on day 2 and 3, which correlated with behavioral voiding data showing significantly impaired bladder function. By day 5, bladder histology and voiding frequency had normalized. Based on the literature and evidence provided by our studies, we propose the following steps for in vivo recording of intravesical pressure and voided volume in an awake mouse: 1) Perform the surgery using an operating microscope and microsurgical tools, 2) Use polyethylene-10 tubing to minimize movement artifacts, and 3) Perform cystometry on post-operative day 5, when bladder swelling resolves.

This study describes the surgical procedures and experimental techniques for performing awake cystometry in a freely moving mouse. In addition, it provides experimental evidence to support its optimization and standardization.

Evaluación del procedimiento para realizar cistometría despierta en un modelo de ratón

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among ...

Puncture-Induced Iris Neovascularization as a Mouse Model of Rubeosis Iridis

We describe a model of puncture-induced iris neovascularization as a general model for noninvasive evaluation of angiogenesis. The model is also relevant for targeting neovascular glaucoma, a sight-threatening complication of diabetic retinopathy. This method is based on the induction of iris vascular response by a series of self-sealing uveal punctures on BALB/c mice and takes advantage of the postpartum maturation of mouse ocular vasculature. Mouse pups undergo uveal punctures from postnatal day 12.5, when the pups naturally open their eyes, until postnatal day 24.5. Due to the transparency of the cornea, iris vasculature can be analyzed easily through time by noninvasive in vivo methods. Furthermore, the semitransparent iris of BALB/c mice can be flatmounted for detailed immunohistologic analysis with minimal non-specific background staining. In this model, angiogenesis is mainly driven by the inflammatory and plasminogen activating systems. The puncture-induced model is the first to induce iris neovascularization in small rodents, and has the advantage of allowing direct noninvasive in vivo analysis of the angiogenic process. Moreover, the model can be combined with angiogenic modulating substances, which highlights its potential in the study of angiogenesis with an in vivo perspective.

Iris neovascularization, a common complication of ischemic retinal disease, may lead to sight-threatening neovascular glaucoma. Here, we describe a murine protocol for inducing experimental iris neovascularization that may be used for noninvasive evaluation of angiogenesis-modulating substances.

Neovascularization del diafragma como un modelo de ratón de iridis del rubeosis inducida por la punción

We describe a model of puncture-induced iris neovascularization as a general model for noninvasive evaluation of angiogenesis. The model is also relevant ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

Medición de la contractilidad isométrica de la arteria mesentérica de ratón con cable miografía

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...