Name: quantitative analysis of cell edge dynamics during cell spreading
Uploaded: 2021-05-22T15:00:00
Description: Watch this Scientific Journal Video about Quantitative Analysis of Cell Edge Dynamics during Cell Spreading at JoVE.com

Este trabajo fue apoyado por el Premio Connaught Fund New Investigator a S.P., Canada Foundation for Innovation, NSERC Discovery Grant Program (subvenciones RGPIN-2015-05114 y RGPIN-2020-05881), University of Manchester y University of Toronto Joint Research Fund, y University of Toronto XSeed Program.

1. Siembra celular
NOTA: El protocolo de propagación celular descrito se realizó utilizando fibroblastos embrionarios de ratón (MEF) que expresan PH-Akt-GFP (un marcador fluorescente para PIP3/PI(3,4)P2). Esta línea celular fue generada mediante la integración genómica de una construcción de expresión para PH-Akt-GFP (Addgene #21218) mediante la edición de genes mediados por CRISPR. Sin embargo, otros marcadores fluorescentes que se expresan de forma transitoria o ...

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J., D&#246;bereiner, H. -. G., Kieffer, N., Bresnick, A. R., Sheetz, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Periodic+Lamellipodial+Contractions+Correlate+with+Rearward+Actin+Waves.">Periodic Lamellipodial Contractions Correlate with Rearward Actin Waves.</a> Cell. 116 (3), 431-443 (2004).</li><li>Dubin-Thaler, B. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+Cell+Edge+Velocities+and+Traction+Forces+Reveals+Distinct+Motility+Modules+during+Cell+Spreading.">Quantification of Cell Edge Velocities and Traction Forces Reveals Distinct Motility Modules during Cell Spreading.</a> PLoS ONE. 3 (11), 3735 (2008).</li><li>Suraneni, P., Rubinstein, B., Unruh, J. 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El ensayo de propagación celular descrito permite el seguimiento continuo de los cambios morfológicos(por ejemplo, el tamaño y la forma de la célula) y los movimientos del borde celular(es decir, la velocidad de protuberancia y la frecuencia de retracción), que son características que faltan en la mayoría de los protocolos de propagación celular19,24. Si bien los ensayos de propagación de células de punto final utilizados comúnmente p...

Las protuberancias lamellipodiales son estructuras citoesqueléticos prominentes formadas en la parte delantera de una célula migratoria. En lamellipodia, la polimerización de actina con la ayuda del complejo Arp2/3 y las fímínas crea una malla actin ramificada de rápido crecimiento que empuja contra la membrana plasmática1,2. La fuerza de empuje generada por el trabajo de malla actin impulsa físicamente la celda hacia adelante1,3,4,5. El agotamiento del complejo Arp2/3 o la interrupción de las vías de señalización esenciales para las protuberancias lamellipodiales a menudo afectan la migración celular6, 7. Aunque la migración de células deficientes en lamellipodia también se ha reportado8,9, la importancia de la lamellipodia en la migración celular es evidente ya que el agotamiento de esta estructura sobretrusiva perturba la capacidad de la célula para moverse a través de microambientes biológicos complejos6,10.
Un obstáculo importante para entender la regulación de la lamellipodia en las células migratorias es la variabilidad natural en lamellipodial protrusión cinética, tamaño, y forma11,12,13,14. Además, estudios recientes han demostrado que la lamellipodia exhibe comportamientos protrusivos complejos, incluyendo protuberancias fluctuantes, periódicas y aceleradas14,15. En comparación con la lamellipodia altamente variable de las células migratorias6,16,lamellipodia formada durante la propagación celular son más uniformes12. Dado que la actividad protrusiva de las células de propagación y migración está impulsada por conjuntos macromoleculares idénticos, que incluyen una red actin ramificada, paquetes de actomiosina contractile y adherencias de matriz celular basadas en integrina17,18,las células de propagación han sido ampliamente utilizadas como modelo para investigar la regulación de la dinámica de lamellipodia.
La propagación celular es un proceso mecanoquímico dinámico en el que una célula en suspensión se adhiere primero a un sustrato a través de adherencias basadas en integrina17,19,20 y luego se propaga extendiendo protuberancias basadas en actin21,22,23. Durante la fase de propagación, lamellipodia que emana del cuerpo celular sobresale isotrópica y persistentemente con poca o ninguna retracción o semental12. Los protocolos de propagación celular más utilizados son los ensayos de puntos finales, donde las células de dispersión se fijan en varias ocasiones después de chapar19,24. Estos ensayos, aunque rápidos y sencillos, son limitados en su poder de diagnóstico para detectar cambios en las características dinámicas de la lamellipodia. Para determinar los mecanismos moleculares que controlan la dinámica de la lamellipodia, el grupo Sheetz fue pionero en el uso del análisis cuantitativo de células de propagación en vivo y descubrió muchas propiedades fundamentales de las protuberancias de borde celular11,12,22. Estos estudios han demostrado que el ensayo de propagación de células vivas es una técnica robusta y poderosa en la caja de herramientas de un laboratorio de biología celular. A pesar de eso, un protocolo detallado y una herramienta computacional de código abierto para un ensayo de propagación de células vivas actualmente no están disponibles para la comunidad de biología celular. Con este fin, nuestro protocolo describe los procedimientos de imágenes que propagan en vivo las células y proporciona una herramienta automatizada de análisis de imágenes. Para validar este método, utilizamos la inhibición de Arp2/3 como tratamiento experimental y demostramos que la inhibición de la función del complejo Arp2/3 no detuvo la propagación celular, sino que causó una reducción significativa en la velocidad de protuberancia celular, así como la estabilidad de las protuberancias del borde celular, dando lugar a bordes celulares irregulares. Estos datos demuestran que la combinación de imágenes de células vivas y análisis automatizado de imágenes es una herramienta útil para analizar la dinámica del borde celular e identificar componentes moleculares que regulan la lamellipodia.

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			<td>0.05% Trypsin (0.05%), 0.53 mM EDTA</td>
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			<td>Arp2/3 Complex Inhibitor I, CK-666</td>
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			<td>DMEM, 1x, 4.5 g/L Glucose, with L-Glutamine, Sodium Pyruvate and Phenol Red</td>
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			<td>Tweezers, Style 2</td>
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quantitative analysis of cell edge dynamics during cell spreading

La propagación celular es un proceso dinámico en el que una celda suspendida en medios se adhiere a un sustrato y se aplana de una forma redondeada a una forma delgada y dispersa. Después del apego de sustrato celular, la célula forma una fina lámina de lamellipodia que emana del cuerpo celular. En la lamellipodia, los monómeros de actina globular (G-actin) polimerizan en una densa malla de actina filamentosa (F-actin) que empuja contra la membrana plasmática, proporcionando así las fuerzas mecánicas necesarias para que la célula se propague. En particular, los actores moleculares que controlan la polimerización de actina en lamellipodia son esenciales para muchos otros procesos celulares, como la migración celular y la endocitosis. 
Dado que las células de dispersión forman lamellipodia continua que abarca toda la periferia celular y se expanden persistentemente hacia afuera, los ensayos de propagación celular se han convertido en una herramienta eficiente para evaluar la cinética de las protuberancias lamellipodiales. Aunque se han desarrollado varias implementaciones técnicas del ensayo de propagación celular, actualmente falta una descripción detallada del flujo de trabajo, que incluiría tanto un protocolo paso a paso como herramientas computacionales para el análisis de datos. Aquí, describimos los procedimientos experimentales del ensayo de propagación celular y presentamos una herramienta de código abierto para el análisis cuantitativo e imparcial de la dinámica del borde celular durante la propagación. Cuando se combina con manipulaciones farmacológicas y/o técnicas de silenciamiento de genes, este protocolo es apto para una pantalla a gran escala de jugadores moleculares que regulan las protuberancias lamellipodiales.

El protocolo anterior describe los procedimientos experimentales para la toma de imágenes de células vivas de células de dispersión y una herramienta computacional para el análisis cuantitativo de la dinámica de propagación celular. La herramienta computacional se puede utilizar en un formato de bajo o alto rendimiento para identificar los reproductores moleculares que regulan la maquinaria de polimerización actin en el borde de ataque celular.
La representación esquemática de los pr...

Watch this Scientific Journal Video about Quantitative Analysis of Cell Edge Dynamics during Cell Spreading at JoVE.com

Quantitative Analysis of Cell Edge Dynamics during Cell Spreading

En este protocolo, presentamos los procedimientos experimentales de un ensayo de propagación celular que se basa en microscopía de células vivas. Proporcionamos una herramienta computacional de código abierto para la segmentación imparcial de células etiquetadas fluorescentemente y análisis cuantitativos de dinámicas de lamellipodia durante la propagación celular.

Análisis cuantitativo de la dinámica del borde celular durante la propagación celular

Cell spreading is a dynamic process in which a cell suspended in media attaches to a substrate and flattens itself from a rounded to a thin and spread-out ...

El protocolo descrito en este vídeo definirá los procedimientos experimentales necesarios para la toma de imágenes a tamaño natural de las células de dispersión y el uso de una herramienta computacional que cuantifica las células que difunden la dinámica de forma imparcial y automatizada. El activo de propagación celular permite el seguimiento continuo del movimiento del borde celular y los cambios morfológicos durante la propagación de la célula, que es una característica que falta en la mayoría de los ensayos de propagación de células existentes. Además, este protocolo implementa el uso de procesamiento y análisis informáticos automatizados, proporcionando información sobre la circularidad celular, el área y los ciclos de retracción de protuberancia de las células de dispersión.Este procesamiento automatizado reduce el sesgo en el análisis de datos y proporciona un método robusto para analizar la dinámica de propagación de células para un gran número de celdas. Cuando se combina con tratamientos farmacológicos o ciencia y técnicas genéticas, este protocolo es apto para velocidades a gran escala de los jugadores moleculares que regulan las protuberancias celulares. Para el protocolo dado, las células utilizadas son fibroblastos embrionarios del ratón que codifican genéticamente PH-Akt-GFP, lo que permite el etiquetado fluorescente de la membrana plasmática.Antes del comienzo del ensayo de propagación celular, cultiva un plato de células al 90% de confluencia. Una vez que las células hayan alcanzado la confluencia adecuada, llama a un resbalón de cubierta de 22 por 22 milímetros y colóquelo en un plato de cultivo celular de 35 milímetros. Cubra el resbalón de la cubierta con fibronectina que se ha diluido en PBS a una concentración de 2,5 microgramos por mililitro.Coloque el plato con la cubierta en una incubadora de 37 grados durante una hora. Después de una hora aspirar la fibronectina asegurándose de no tocar el resbalón de la cubierta con la punta de la pipeta. Lave el plato con PBS pipendo suavemente alrededor de la tapa deslice de dos a tres veces.Para la siembra celular, comience aspirando los medios de cultivo celular desde el plato de las células. Después, lave el plato con PBS caliente. Añadir 650 microlitros de trippsina EDTA al plato e inclinar el plato para distribuir uniformemente la enzima.Coloque el plato con la trypsin en la incubadora durante un minuto. Después de la incubación primero tendrá que añadir 10 mililitros de medios de cultivo celular en un tubo centrífugo. A continuación, agregue rápidamente otros 10 mililitros de medios en el plato con el fin de apagar la trypsin.Para diluir las células que se sembrarán en el deslizamiento de la cubierta, pipetear un mililitro del contenido del plato en el tubo centrífuga. Desde el tubo, pipeta aproximadamente 500 a 1.000 microlitros de células diluidas en el plato con el resbalón de la cubierta. Asegúrese de que el deslizamiento de la cubierta esté al 10% de confluencia o a 50.000 células por mililitro y ajuste el volumen de las células diluidas según sea necesario.El propósito de estas celdas es establecer un plano de enfoque durante la adquisición de imágenes. Con las células restantes en el plato, pasa una quinta parte de las células a un plato pequeño por tratamiento. Estas serán las células que serán analizadas para la dinámica de propagación.Coloque los platos de paso y la cubierta deslice el plato en una incubadora durante ocho a 24 horas. Para probar la importancia de Arp2/3 para la propagación celular, primero agregue cinco mililitros de medios de cultivo celular en un control y un tubo centrífuga de tratamiento. A continuación añadir 20 mililitros de dmem que carece de fenol rojo en cada uno de dos tubos centrífugos más grandes.Pipeta ya sea el fármaco CK-666 que es un inhibidor del complejo Arp2/3 o el tratamiento controlado como DMSO en los tubos pequeños y grandes. Para comenzar el tratamiento farmacológico de las células, retire los platos de paso que estaban incubando durante la noche. Aspirar los medios de cultivo celular de todos los platos y lavar los platos con PBS caliente.Tome los pequeños tubos que contienen CK-666 o controle los medios complementados y agregue el contenido del tubo pertinente en cada uno de los platos del pasaje. Etiqueta cada uno de los platos con el tratamiento farmacológico correcto y luego coloca los platos de nuevo en la incubadora durante una hora adicional Después de una hora de incubación, aspirar el medicamento complementado medios de cada uno de los platos. A continuación, añadir PBS caliente a todos los platos con el fin de eliminar a fondo todos los medios rojos fenol restantes.Añadir 230 microlitros de trypsin EDTA y colocar los platos en una incubadora durante un minuto. Retire los platos trypsinizados de la incubadora y proceda a añadir cinco mililitros de la droga dmem suplementada que carece de fenol rojo en un tubo designado como tubo B.Then añadir otros cinco mililitros de los mismos medios en el plato pertinente para saciar la trippsina. Pipetear los medios arriba y abajo varias veces con el fin de separar todas las células del plato.Transfiera todo el contenido del plato a otro tubo que ya ha sido designado como tubo A.In orden para preparar una dilución celular adecuada para la toma de imágenes, transfiera un mililitro de células del tubo A al tubo B.Repita todos los pasos de dilución para cada tratamiento. Coloque todos los tubos en la incubadora durante 45 minutos para permitir que las células se recuperen de la trippsinización. Asegúrese de que todas las partes de una cámara magnética celular que puede acomodar un resbalón de cubierta de 22 por 22 milímetros se hayan limpiado a fondo antes de su uso.Retire el plato con el resbalón de la cubierta de la incubadora. Aspirar los medios de cultivo celular y lavar el resbalón de la cubierta con PBS caliente. Retire el resbalón de la cubierta con un par de fórceps y coloque suavemente el resbalón de la cubierta en la placa inferior de la cámara magnética.A continuación, coja la junta de silicona y colóquela encima del resbalón de la cubierta. Coloque el cuerpo principal de la cámara magnética en la placa inferior. Añadir un mililitro del dmem carente de fenol rojo correspondiente al tratamiento pertinente a la cámara magnética.Tome un tejido libre de pelusas y coloque cuidadosamente la carcasa entre el cuerpo principal y la placa inferior para comprobar si hay fugas. Para completar la cámara magnética, baje la cubierta transparente sobre el cuerpo principal. Por último, limpie la parte inferior de la tapa con un tejido rociado con agua y otro rociado con 70% de etanol.Precaliente la incubadora superior del escenario de un microscopio confocal a 37 grados. Coloque la cámara magnética encima del objetivo. Antes de adquirir dinámicas de propagación, establezca el foco en las células ya polarizadas en el canal fluorescente verde.Esto garantiza que las células que se extienden estarán enfocadas una vez que se adhieran al resbalón de la cubierta. Retire la cubierta transparente de la cámara magnética y la pipeta 500 microlitros del tubo B que se retiró de la incubadora. Coloque la cubierta transparente en la parte superior.Para identificar las células ideales para el análisis de propagación celular, busque halos verdes que representen células que aún no se han adjunte al resbalón de la cubierta o se encuentran en las primeras etapas de unión. Adquiera imágenes y guarde los archivos. Después de haber completado la adquisición de imágenes de propagación de celdas, comience por ejecutar un IDE de Python compatible como Spyder.Para el análisis computacional de la dinámica de propagación de celdas, busque y abra el archivo GUI de propagación de celdas proporcionado en los paquetes de script relevantes. Pulse el botón ejecutar que abrirá el panel GUI de análisis en segundo plano. La GUI alberga todos los ajustes necesarios para el análisis.Para analizar el área de celda y la circularidad, introduzca el archivo de adquisición y la configuración necesaria en la pestaña área de dispersión de celda. Los ajustes deberán realizarse en función del tipo de celda y los parámetros de adquisición de imágenes. Después de presionar ejecutar, el script creará una circularidad de celda y un área frente a un gráfico de tiempo para todas las celdas de dispersión.Para los datos del kymograph, presione el generador de kymograph y la pestaña de análisis. Este análisis requiere que los archivos de entrada se recorten pilas de imágenes de una sola celda. Después de insertar todos los ajustes y presionar ejecutar, el script creará varios kymographs para la celda dada.A la izquierda hay células representativas de los tratamientos DMSO y CK-666, segmentadas automáticamente mediante el script proporcionado. A diferencia de la morfología isotrópica y altamente circular demostrada por las células de control, Arp2/3 células inhibidas exhiben disminución de la circularidad. Además, los kymographs generados automáticamente proporcionan una resolución temporal que es fundamental para comprender la dinámica de las protuberancias.Las células de control sobresalen persistentemente con pequeñas o ninguna retracción. Por el contrario, la inhibición de Arp2/3 altera la estabilidad de las protuberancias, como lo indica el aumento de los eventos de retracción a lo largo de la propagación. Estos videos deben proporcionarle la comprensión de cómo sembrar correctamente las células, realizar tratamientos farmacológicos y preparar las herramientas de imagen y cálculo necesarias para la cuantificación de la dinámica de propagación celular.Este protocolo se puede combinar aún más con imágenes fluorescentes de citoesqueleto y ensayos de migración para identificar a los jugadores moleculares que rigen las protuberancias celulares. Gracias por ver y buena suerte con sus experimentos.

quantitative-analysis-of-cell-edge-dynamics-during-cell-spreading

Research

JoVE Journal

Biology

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

Imágenes de células vivas de la F-actina Dynamics a través de Microscopía Fluorescente Speckle (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Investigación

Biología

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Células vivas, imágenes y análisis cuantitativo de las células epiteliales embrionarias en Xenopus laevis</em

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Determining Cell Number During Cell Culture using the Scepter Cell Counter

Counting cells is often a necessary but tedious step for in vitro cell culture. Consistent cell concentrations ensure experimental reproducibility and accuracy. Cell counts are important for monitoring cell health and proliferation rate, assessing immortalization or transformation, seeding cells for subsequent experiments, transfection or infection, and preparing for cell-based assays. It is important that cell counts be accurate, consistent, and fast, particularly for quantitative measurements of cellular responses.
Despite this need for speed and accuracy in cell counting, 71% of 400 researchers surveyed1 who count cells using a hemocytometer. While hemocytometry is inexpensive, it is laborious and subject to user bias and misuse, which results in inaccurate counts. Hemocytometers are made of special optical glass on which cell suspensions are loaded in specified volumes and counted under a microscope. Sources of errors in hemocytometry include: uneven cell distribution in the sample, too many or too few cells in the sample, subjective decisions as to whether a given cell falls within the defined counting area, contamination of the hemocytometer, user-to-user variation, and variation of hemocytometer filling rate2.
To alleviate the tedium associated with manual counting, 29% of researchers count cells using automated cell counting devices; these include vision-based counters, systems that detect cells using the Coulter principle, or flow cytometry1. For most researchers, the main barrier to using an automated system is the price associated with these large benchtop instruments1.
The Scepter cell counter is an automated handheld device that offers the automation and accuracy of Coulter counting at a relatively low cost. The system employs the Coulter principle of impedance-based particle detection3 in a miniaturized format using a combination of analog and digital hardware for sensing, signal processing, data storage, and graphical display. The disposable tip is engineered with a microfabricated, cell- sensing zone that enables discrimination by cell size and cell volume at sub-micron and sub-picoliter resolution. Enhanced with precision liquid-handling channels and electronics, the Scepter cell counter reports cell population statistics graphically displayed as a histogram.

The Scepter Cell Counter is a handheld automated device that can be used to count cells, monitor cell diameter and volume, and be used to check the health and quality of cellular populations from one culture to the next.

Determinar el número de células durante el cultivo celular utilizando el contador de células Cetro

Counting cells is often a necessary but tedious step for in vitro cell culture. Consistent cell concentrations ensure experimental reproducibility and ...

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 ...

A Quantitative Fitness Analysis Workflow

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. QFA can be applied to focused observations of single cultures but is most useful for genome-wide genetic interaction or drug screens investigating up to thousands of independent cultures. The central experimental method is the inoculation of independent, dilute liquid microbial cultures onto solid agar plates which are incubated and regularly photographed. Photographs from each time-point are analyzed, producing quantitative cell density estimates, which are used to construct growth curves, allowing quantitative fitness measures to be derived. Culture fitnesses can be compared to quantify and rank genetic interaction strengths or drug sensitivities. The effect on culture fitness of any treatments added into substrate agar (e.g. small molecules, antibiotics or nutrients) or applied to plates externally (e.g. UV irradiation, temperature) can be quantified by QFA.
The QFA workflow produces growth rate estimates analogous to those obtained by spectrophotometric measurement of parallel liquid cultures in 96-well or 200-well plate readers. Importantly, QFA has significantly higher throughput compared with such methods. QFA cultures grow on a solid agar surface and are therefore well aerated during growth without the need for stirring or shaking.
QFA throughput is not as high as that of some Synthetic Genetic Array (SGA) screening methods5,6. However, since QFA cultures are heavily diluted before being inoculated onto agar, QFA can capture more complete growth curves, including exponential and saturation phases3. For example, growth curve observations allow culture doubling times to be estimated directly with high precision, as discussed previously1.
Here we present a specific QFA protocol applied to thousands of S. cerevisiae cultures which are automatically handled by robots during inoculation, incubation and imaging. Any of these automated steps can be replaced by an equivalent, manual procedure, with an associated reduction in throughput, and we also present a lower throughput manual protocol. The same QFA software tools can be applied to images captured in either workflow.
We have extensive experience applying QFA to cultures of the budding yeast S. cerevisiae but we expect that QFA will prove equally useful for examining cultures of the fission yeast S. pombe and bacterial cultures.

Quantitative Fitness Analysis (QFA) is a complementary series of experimental and computational methods for estimating microbial culture fitnesses. QFA estimates the effect of genetic mutations, drugs or other applied treatments on microbe growth. Experiments scaling from focussed analysis of single cultures to thousands of parallel cultures can be designed.

Un flujo de trabajo de Análisis Cuantitativo Gimnasio

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. ...

Surface Spreading and Immunostaining of Yeast Chromosomes

The small size of nuclei of the budding yeast Saccharomyces cerevisiae limits the utility of light microscopy for analysis of the subnuclear distribution of chromatin-bound proteins. Surface spreading of yeast nuclei results in expansion of chromatin without loss of bound proteins. A method for surface spreading balances fixation of DNA bound proteins with detergent treatment. The method demonstrated is slightly modified from that described by Josef Loidl and Franz Klein1,2. The method has been used to characterize the localization of many chromatin-bound proteins at various stages of the mitotic cell cycle, but is especially useful for the study of meiotic chromosome structures such as meiotic recombinosomes and the synaptonemal complex. We also describe a modification that does not require use of Lipsol, a proprietary detergent, which was called for in the original procedure, but no longer commercially available. An immunostaining protocol that is compatible with the chromosome spreading method is also described.

A method for surface-spreading chromosomes from budding yeast is presented. This method is derived from a method previously described by Loidl and Klein. In addition, we demonstrate a procedure for immunostaining of spread chromosomes.

Difusión de superficies y inmunotinción de cromosomas de la levadura

The small size of nuclei of the budding yeast Saccharomyces cerevisiae limits the utility of light microscopy for analysis of the subnuclear distribution ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Imágenes de alta resolución y análisis de la dinámica de microtúbulos astrales individuales de levadura en ciernes

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Vivo de la célula de microscopía de fluorescencia para observar procesos esenciales durante el crecimiento de la célula microbiana

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

Lens-free Video Microscopy for the Dynamic and Quantitative Analysis of Adherent Cell Culture

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator and that it is possible to monitor and quantify single cells along several cell cycles. We describe the full protocol used to monitor and quantify a HeLa cell culture for 2.7 days. First, cell culture acquisition is performed with a lens-free video microscope, and then the data is analyzed following a four-step process: multi-wavelength holographic reconstruction, cell-tracking, cell segmentation and cell division detection algorithms. As a result, we show that it is possible to gather a dataset featuring more than 10,000 cell cycle tracks and more than 2 x 106 cell morphological measurements.

Lens-free video microscopy enables us to monitor cell cultures directly inside the incubator. Here we describe the full protocol used to acquire and analyze a 2.7 day long acquisition of cultured HeLa cells, leading to a dataset of 2.2 x 106 measurements of individual cell morphology and 10584 cell cycle tracks.

Lente libre Video microscopia para la dinámica y el análisis cuantitativo del cultivo de células adherentes

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.