J.O.B. and A.M.R. were supported by predoctoral fellowships from the National Science Foundation.

NOTA: Para este protocolo, se han centrado en el ensayo de la frecuencia stromule en la epidermis de N. deja benthamiana. Varias líneas transgénicas estables que se han generado que se puede utilizar para este propósito, incluyendo 35S PRO: FNRtp: EGFP 13 y NRIP1: Cerulean 6. Ambas líneas muestran la expresión robusta de fluoróforos en el estroma del cloroplasto de hojas cultivadas en una amplia gama de condiciones. Alternativamente, fluoróforos cloroplasto orientada se pueden expresar de forma transitoria en N. benthamiana utilizando Agrobacterium Transformaciones 13. Esto es menos ideal que las líneas transgénicas, ya que las infiltraciones de Agrobacterium inducen unas respuestas de defensa basal en N. benthamiana y las interacciones con Agrobacterium pueden alterar la frecuencia stromule en la hoja 14, lo que podría complicar la interpretación de los resultados. Por último, para visualizar la formación de stromule in vitro,cloroplastos pueden ser extraídos de cualquier especie vegetal, ya sea utilizando fluoróforos genéticamente codificados o un colorante fluorescente, tal como se describe en la sección 5 a continuación. 9,15 NOTA:. Métodos detallados de cultivo de plantas se han descrito previamente 16 Brevemente, crecer N. benthamiana plantas en macetas de 4 &quot;con cualquier mezcla de tierra profesional que ofrece un buen drenaje. Cubra plántulas con una cúpula de plástico transparente para los primeros 10-14 días para proporcionar un ambiente húmedo para la germinación. añadir cualquier mezcla de fertilizante estándar siguiendo las instrucciones del fabricante para 14- plantas días de edad. cultivar plantas bajo luz blanca, utilizando ~ 100 mmol fotones m-2 s-1 de intensidad de luz. Las plantas de agua con regularidad. 1. Preparación de muestras de hojas para la visualización NOTA: la dinámica Stromule se ven afectados por hiriendo a 8, por lo que la preparación del tejido debe llevarse a cabo inmediatamente antes de visualizar stromules por microscopía de fluorescencia confocal. Idealmente, una muestra debe ser visualizado con 15 min después de la eliminación de la planta. <ol><li> Cortar una pequeña sección de la hoja usando una hoja de afeitar muy agudo. Utilice siempre la misma región de la hoja de coherencia, y estar seguro de visualizar la misma superficie (adaxial o abaxial) en todos los experimentos. </li><li> Inmediatamente después de cortar la sección de hoja, sumergir la sección de hoja en un 5 ml jeringa sin aguja llena de agua, eliminar el aire de la jeringa, y aplicar un vacío al cubrir el orificio de la jeringa con un dedo y tirando del émbolo. <ol><li> Liberar el émbolo suavemente para evitar daños en la sección de la hoja. Repita esto dos o tres veces, o hasta que la mayor parte del aire se ha eliminado y la hoja parece ser de color verde oscuro. NOTA: Este paso elimina el aire de la hoja que puede interferir con la visualización precisa de stromules. </li></ol></li><li> Añadir una gota de agua a un portaobjetos y colocar la hoja en la sección de esta caída. A continuación, añadir otro drop de agua a la parte superior de la hoja, y colocar un cubreobjetos sobre la hoja. Si hay burbujas de aire, golpee suavemente la hoja de la cubierta hasta que se eliminan. El portaobjetos ahora está listo para la microscopía, y debe ser visualizado inmediatamente. </li></ol> 2. Visualizar Stromules con confocal de microscopía de fluorescencia <ol><li> Con luz transmitida, se centran en un campo de visión cerca del centro de la sección de la hoja (lejos de las células dañadas por la cuchilla de afeitar). Utilice un objetivo 20X de modo que muchos cloroplastos pueden ser visualizadas simultáneamente. Guardar una imagen con luz transmitida de modo que los tipos de células se pueden distinguir después, si es necesario. </li><li> Cambiar la fuente de iluminación a un láser con filtros de excitación y emisión apropiados para ser usado fluoróforo. Por ejemplo, excitar GFP con un láser de 488 nm y recoger las emisiones entre 500 nm y 525 nm. <ol><li> El uso de software del microscopio, seleccionar el canal GFP y haga clic para ajustar la apertura del agujero de alfiler de 1 Airy unidad. Seleccionar escaneo láser, para iniciar la visualización de la muestra, y luego haga clic en el canal de GFP para ajustar la intensidad del láser y la ganancia del detector para minimizar la potencia del láser al mismo tiempo visualizar claramente stromules. Una vez que se han determinado estos valores, los mantenga lo más constante posible a lo largo de todos los experimentos. </li></ol></li><li> Preparar un experimento -stack z que recogerá una serie de imágenes a través de la epidermis de la hoja en el campo de visión. <ol><li> Para el -stack z, incluir todos los cloroplastos epidérmicas en el campo de visión para evitar el sesgo (por ejemplo, si los cloroplastos cerca de la superficie de las hojas tienen más stromules en condiciones ensayadas). </li><li> Tomar imágenes en el intervalo máximo posible, que incluirá todos los stromules sino reducir al mínimo el tiempo necesario para que una -stack z. Por ejemplo, si 1 unidad de Airy es equivalente a una sección confocal de enfoque que es 2 m de profundidad, teniendo una imagen cada 2 micras es probable ideal. NOTA: La hoja epidermis es una superficie irregular, por lo que la profundidad total de la z -stack variará dependiendo de la muestra; la mayoría de -stacks z requerirán 10-20 imágenes, pero pueden incluir más. </li><li> Ajustar la velocidad de exploración y la resolución de la imagen si es necesario para asegurarse de que el -stack z es recogida rápidamente (normalmente dentro de 5-10 minutos, si es posible). Guarde el -stack z para su posterior análisis. </li></ol></li></ol> Procesamiento 3. Imagen <ol><li> Determinar la frecuencia stromule usando cualquier software de análisis de imágenes. Para este protocolo, se recomienda utilizar ImageJ, que está disponible públicamente de los Institutos Nacionales de Salud (http://imagej.nih.gov/ij/), o la actualización popular de ImageJ, Fiji (http://fiji.sc / Fiji). </li><li> Combinar la -stack z en una sola imagen con la proyección de intensidad máxima en el software. </li><li> Identificar y contar manualmente todos los cloroplastos en la imagen. </li><li> Para cada cloroplasto, determinar visualmente si una o más stromulesson vistas que se extiende desde el cloroplasto en la imagen de la z fusionada -stack. Para ejemplos, véase la figura 1. NOTA: Dado que no se conocen las funciones biológicas de stromules, ninguna, estroma llenas, extensión tubular delgada del cloroplasto puede ser considerado un &quot;stromule&quot;, independientemente de la longitud. Como regla general, una extensión de estroma llenas desde el cloroplasto es un stromule si es menos de aproximadamente 1 m de diámetro a lo largo de la mayor parte de su longitud 7. Asegúrese de que una persona se analizan todas las imágenes para controlar las diferencias subjetivas en la determinación de si o no un cloroplasto tiene una stromule. Un segundo investigador debe verificar de forma independiente las frecuencias stromule para reducir aún más los sesgos subjetivos. </li></ol> 4. Diseño Experimental y Toma de Muestras NOTA: Stromule frecuencia es muy variable entre las hojas, pero varios informes sugieren que hay poca variación en la frecuencia stromule dentro de un indivi9,17 hoja dual. <ol><li> Calcular la frecuencia hoja stromule de un solo campo de visión de una hoja. Desechar la hoja, y considerar esta una muestra independiente. No considere campos separados de vista de una hoja como muestras independientes. NOTA: No hay un fuerte consenso sobre la forma de mejores miden los cambios en la actividad stromule, y hasta que la función de stromules se define más claramente, los investigadores deben seguir siendo creativa en la cuantificación de la dinámica stromule. Para la comparación a través de la literatura, sin embargo, las normas comunes de presentación de informes se deben utilizar, además de los enfoques más creativos. Como mínimo, siempre informar de la frecuencia de cloroplastos que tienen uno o más stromules. </li><li> Determinar la frecuencia de cloroplastos que tienen stromules en un campo de visión. Utilice ImageJ para identificar todos los cloroplastos en un campo de visión. Entonces, para cada cloroplasto, determinar si el cloroplasto se ha formado al menos un stromule. Informe de frecuencia stromule como el Percentage de los cloroplastos con una stromule. NOTA: Algunos investigadores transforman el porcentaje, por ejemplo, con la transformación arcoseno, antes de informar de frecuencias stromule; mientras que esta es una opción para el análisis estadístico, el arco seno de la frecuencia stromule no es un valor intuitivo, y no necesita ser informado en el texto principal o cifras de un estudio. <ol><li> Como un enfoque alternativo, contar el número total de stromules, y se divide por el número total de los cloroplastos. NOTA: En circunstancias normales, esto va a producir resultados similares de acercarse a 4,2; puede sobreestimar la frecuencia stromule, sin embargo, si un solo cloroplasto ha formado muchos stromules. En el ejemplo mostrado en virtud de los resultados representativos, por ejemplo, varios cloroplastos tienen dos o más stromules, elevando el número de stromules por cloroplasto a 40/87 (en comparación con una frecuencia stromule de 33/87). </li><li> Como alternativa, determinar la longitud stromule lugar de la frecuencia stromule. La longitud se puede mediren ImageJ trazando el stromule con la herramienta de línea a mano alzada. NOTA: Como el papel biológico de stromules sigue siendo desconocida, no está claro que la longitud stromule afecta a su función. Reportar diferencias significativas en la duración stromule si se observan, sino también informar de las frecuencias stromule (como se describe en el apartado 4.2). NOTA: frecuencia Stromule puede ser muy variable entre diferentes hojas en las mismas condiciones de crecimiento. Por lo tanto, aunque la determinación de la frecuencia stromule puede llevar mucho tiempo, los experimentos deben ser cuidadosamente diseñadas para incluir muestras de gran tamaño. El uso de conjuntos de datos de nuestro laboratorio, se determinó que un tamaño de muestra de al menos 16 plantas para cada tratamiento es por lo general suficientemente potente para determinar si hay una diferencia estadísticamente significativa de al menos un cambio de 1,5 veces en la frecuencia stromule entre dos condiciones (con la norma α = 0,05 y β = 0,80). </li></ol></li><li> El análisis estadístico de los resultados. <ol><li> Para comparar el youn stromule frecuencias de dos tratamientos, use la prueba t de Welch (también conocida como la prueba de la prueba t o t heterocedástico suponiendo una varianza desigual). NOTA: Esta prueba no es tan poderoso como prueba de la t de Student convencional, pero la prueba t de Welch es ventajoso porque no asume que la variación en la distribución de frecuencias stromule es similar entre los tratamientos. NOTA: Dado que la frecuencia stromule es una &quot;proporción&quot;, su distribución muestral se prevé que sea binomial en lugar de lo normal, que podrían análisis estadístico teóricamente sesgo si los tamaños de las muestras son muy bajos o si la frecuencia stromule es muy bajo (cerca del 0%) o muy alta (cerca de 100%). Algunos informes han utilizado transformaciones arcoseno antes del análisis estadístico, que está destinado a transformar una distribución binomial a una distribución normal, y así satisfacer los requisitos de la norma de prueba de la t de Student o ANOVA. En la práctica, sin embargo, unarcsine transformaciones tienen poco o ningún efecto sobre el análisis estadístico, y por lo tanto no se recomiendan. En su lugar, repetir los experimentos para aumentar el tamaño de la muestra, lo que aumentará la potencia estadística y reducir los errores en la interpretación de los datos. </li><li> Cualquiera sea el enfoque estadístico se utiliza, asegúrese de informar cuidadosamente la estrategia de muestreo, tamaño de la muestra, prueba estadística, y el valor de p para cada experimento. NOTA: Al igual que con otros campos de la biología, un valor de p inferior a 0,05 se puede considerar &quot;estadísticamente significativa&quot;, aunque los investigadores deben informar sin duda el valor p obtenido precisa. Si p es ligeramente por encima de 0,05, el aumento de tamaño de la muestra con dos o tres experimentos puede ayudar a clarificar si o no la diferencia observada es reproducible y estadísticamente significativa. </li></ol></li></ol> 5. La extracción de cloroplastos intactos para visualizar Stromule Dinámica NOTA: Existen varios métodosse han utilizado para aislar los cloroplastos de las hojas, incluyendo un protocolo ligeramente diferente en un estudio reciente sobre la formación de stromule in vitro 15. El protocolo se detalla a continuación utiliza un método relativamente simple que no dió muestras cloroplasto bioquímicamente puros, pero en lugar aislar una gran cantidad de cloroplastos, sanos intactos 9,18. <ol><li> Preparar tampón de extracción frío: NaOH Hepes 50 mM, sorbitol 330 mM, EDTA 2 mM, 1,0 mM MgCl 2, y MnCl 1,0 mM 2. Ajustar el pH a 6,9 con NaOH y HCl, y refrigerar antes de su uso. NOTA: Aunque no es necesario, utilizando tampones fríos y conservación en hielo cuando sea posible dará lugar a una mayor proporción de cloroplastos intactos. </li><li> Preparar tampón de aislamiento: NaOH Hepes 50 mM, sorbitol 330 mM, EDTA 2 mM, 1,0 mM MnCl 2, mM MgCl 2 mM, KCl 10, y 1,0 NaCl 1,0 mM. Ajustar el pH a 7,6 con NaOH y HCl y refrigerar antes de su uso. </li><li> Si las hojas no sonexpresar un fluoróforo estromal codificado genéticamente, preparar diacetato de carboxifluoresceína (CFDA) solución: CFDA 50 mM solución madre en dimetil sulfóxido (DMSO) (2,3 mg CFDA por 1,0 ml DMSO). NOTA: La concentración exacta no es crítica. Mantenga CFDA de valores en la oscuridad en todo momento. Almacenar pequeñas alícuotas de CFDA 50 mM a -20 ° C. </li><li> Para el aislamiento de cloroplastos, retire ~ 5-10 g de hojas de varias plantas y enjuagar brevemente en agua fría. transferir inmediatamente a 50 ml de tampón de extracción en frío. Moler las hojas utilizando una licuadora con varios pulsos cortos. Filtrar a través de dos o tres capas de gasa para eliminar los desechos de la hoja, dividir los cloroplastos extraídos en dos tubos de 50 ml de centrífuga y centrifugar durante 1 min a 750 x g. </li><li> Eliminar el sobrenadante y resuspender los cloroplastos verdes en tampón de aislamiento 10 ml. Centrifugar de nuevo durante 1 min a 750 xg, desechar el sobrenadante, y los cloroplastos resuspender en tampón de aislamiento para llevar el volumen final a 5 ml. </li><li> Transferencia 201; l de los cloroplastos a una diapositiva, cubrir con un cubreobjetos, y comenzar la microscopía. NOTA: Los cloroplastos de las plantas transgénicas que expresan proteínas fluorescentes plastid orientada ahora están listos para la visualización por microscopía de fluorescencia confocal. </li><li> Manchar los cloroplastos de las plantas que no están expresando proteínas fluorescentes plastid orientada para visualizar stromules. <ol><li> Añadir 5 l de 50 mM CFDA a los 5 ml cloroplastos a suspender en tampón de aislamiento. Llevar la concentración final a 50 mM. </li><li> Permitir a los cloroplastos incubar durante 5 minutos, y luego transferir una pequeña alícuota (~ 50 l) a una diapositiva, cubren con un cubreobjetos, y comienzan microscopía. </li><li> Use un filtro para FITC o GFP. Utilice un objetivo de 20X para visualizar muchos cloroplastos de forma simultánea, o un objetivo superior para visualizar un solo cloroplasto aislado. NOTA: CFDA será fluorescente en el interior del estroma de los cloroplastos intactos. </li><li> Si hay una fuerte fluorescencia de fondo, lavar el Chloroplasts de nuevo en 10 ml de tampón de aislamiento después de la 5 min de incubación con CFDA, centrifugar durante 1 min a 750 xg, desechar el sobrenadante, y resuspender en 5 ml de tampón de aislamiento. </li></ol></li></ol>

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The authors have nothing to disclose.

Al investigar stromules, tres factores importantes deben ser considerados a lo largo de: (i) la manipulación del tejido de la planta deben mantenerse a un mínimo absoluto, (ii) el sistema experimental debe mantenerse constante, y (iii) las estrategias de muestreo debe ser planeada cuidadosamente para asegurar robusta, se analizan los datos reproducibles. Stromules son muy dinámicas: se pueden extender y retraer rápidamente ante los ojos de un observador bajo el microscopio. Por otra part...

Chloroplasts are dynamic organelles in plant cells responsible for photosynthesis and a host of other metabolic processes. Signaling pathways from the chloroplast also exert significant influence on plant physiology and development, coordinating plant responses to environmental stress, pathogens, and even leaf shape1-6. Recently, biologists have gained interest in a poorly understood aspect of chloroplast structure: stromules, very thin stroma-filled tubules that extend from the surface of the chloroplast7.
The biological functions of stromules remain unknown, although stromule frequency is known to vary in response to environmental stimuli7-9, and stromules may be capable of transmitting signaling molecules between organelles6. All types of plastids (not only the green, photosynthetic chloroplasts, but also clear leucoplasts, starch-filled amyloplasts, and pigmented chromoplasts, to name a few types of plastids) make stromules, and stromules are found in all land plant species that have been examined to date. Stromules can extend and retract dynamically, appearing or disappearing within seconds, or they can remain relatively stationary for long times. One of the major hurdles facing stromule biologists is that stromules are often studied using dramatically different methods, tissues, and species, making comparisons across the stromule biology literature difficult. Going forward, standard practices and thorough descriptions of the experimental systems used to study stromules will be critical to discovering the function of these ubiquitous features of chloroplast morphology.
Here we describe methods for visualizing stromule formation in the epidermal chloroplasts of Nicotiana benthamiana leaves. In the mesophyll, chloroplasts are densely packed into large, three-dimensional cells, which makes it difficult to accurately and rapidly visualize stromules by confocal microscopy. By contrast, epidermal cells are relatively flat, contain fewer chloroplasts, and are at the surface of the leaf, allowing for easy and rapid visualization of stromules. N. benthamiana is an ideal model system for these experiments because, unlike many plant species, all cells in the epidermis of N. benthamiana make chloroplasts10. In the epidermis of most plants, including Arabidopsis thaliana, only the stomatal guard cells have chloroplasts, while other epidermal cells have "leucoplasts", plastids that are clear, relatively amorphous, and nonphotosynthetic9,11,12. Thus, whereas a single field of view of an A. thaliana epidermis might show only a handful of chloroplasts in a pair of guard cells, a field of view of an N. benthamiana epidermis will include dozens or even hundreds of chloroplasts. All of the methods described here, however, can be modified to investigate other questions in stromule biology; for example, we have used the same approach to study leucoplast stromules of A. thaliana9.

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			<td height="42" style="height:42px;width:216px;">Laser Scanning Confocal Microscope&#160;</td>
			<td style="width:71px;">Carl Zeiss Inc</td>
			<td style="width:119px;">Model: LSM710</td>
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			<td height="42" style="height:42px;width:216px;">Carboxyfluorescein diacetate (CFDA)</td>
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			<td style="width:119px;">21879&#160;</td>
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			<td height="21" style="height:21px;width:216px;">Dimethyl sulfoxide (DMSO)</td>
			<td style="width:71px;">EMD</td>
			<td style="width:119px;">MX1458-6</td>
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			<td height="21" style="height:21px;width:216px;">Waring blender</td>
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			<td height="21" style="height:21px;width:216px;">Fiji</td>
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			<td>Open-source software for analyzing biological images</td>
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visualizing stromule frequency with fluorescence microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Este protocolo se utiliza para visualizar la frecuencia stromule en el día y por la noche en los cotiledones de joven N. plántulas benthamiana. Rebanadas de una pila z se fusionaron en una sola imagen (Figura 1A). Para los propósitos visuales, que la imagen fue entonces desaturado y se invierte de manera que el estroma aparece en negro (Figura 1B). Los cloroplastos se marcaron o bien como no tener stromules (asterisco verde)...

Watch this Scientific Journal Video about Visualizing Stromule Frequency with Fluorescence Microscopy at JoVE.com

Visualizing Stromule Frequency with Fluorescence Microscopy

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

La visualización de frecuencia Stromule con microscopía de fluorescencia

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

visualizing-stromule-frequency-with-fluorescence-microscopy

Research

JoVE Journal

Biology

8.0K vistas.  USDA. Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Video: Visualizing Stromule Frequency with Fluorescence Microscopy

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Visualizar el vivo Drosophila Gliales-unión neuromuscular con tintes fluorescentes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Investigación

Biología

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Visualizar solo los complejos moleculares En Vivo Uso de microscopía de fluorescencia avanzada

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

Visualización de una sola molécula de ADN de replicación con microscopía de fluorescencia

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Time-lapse de imágenes de mitosis después de la transfección de siRNA

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Electrofusión de células visualiza con microscopía de fluorescencia

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Cuantitativa de células vivas por fluorescencia microscopia análisis de la levadura de fisión

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

Visualización de las bacterias en los nematodos mediante microscopía de fluorescencia

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Análisis rápido y exploración de microscopía de fluorescencia de Imágenes

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells possess several lines of defense against damage to biologically relevant molecules like nucleic acids, lipids and proteins. In addition to organelle dynamics (fusion/fission/motility/inheritance) and tightly controlled protease activity, the degradation of surplus, damaged or compromised organelles by autophagy (cellular &#8216;self-eating&#8217;) has received much attention from the scientific community. The regulation of autophagy is quite complex and depends on genetic and environmental factors, many of which have so far not been elucidated. Here a novel method is presented that allows the convenient study of autophagy in the filamentous fungus Penicillium chrysogenum. It is based on growth of the fungus on so-called &#8216;starvation pads&#8217; for stimulation of autophagy in a reproducible manner. Samples are directly assayed by microscopy and evaluated for autophagy induction / progress. The protocol presented here is not limited for use with P. chrysogenum and can be easily adapted for use in other filamentous fungi.

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

A convenient and powerful method for studying autophagy in Penicillium chrysogenum by using starvation pads (mixtures of agarose and tap water in a microscope slide containing a central cavity) is presented here.

Análisis de la autofagia en<em&gt; Penicillium chrysogenum</em&gt; Por la inanición Pads en combinación con microscopía de fluorescencia

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells ...