Damos las gracias a Mike W. Davidson (Universidad Estatal de Florida) para proporcionar el plásmido de ADN que codifican las proteínas fluorescentes. Este trabajo fue financiado por los Institutos Nacionales de Salud de subvención GM72048 (a DWP).

1. Preparación de muestras de proteínas fluorescentes gota Una muestra de la proteína fluorescente consiste en gotas de una emulsión de 1-octanol/water con la proteína fluorescente que residían en la fase acuosa. Esta emulsión se encuentra entre un portaobjetos de microscopio y un vaso de 22 mm cuadrados para cubrir las aplicaciones de la microscopía. <ol><li> Antes de tomar muestras de gotas fluorescentes proteína de los portaobjetos y cubreobjetos deben ser limpiados y recubiertos con un agente hidrófobo. </li><li> Limpiar la cristalería por lavado de 5 minutos con acetona y dejar secar por aire. (Si lo desea, después de limpiar el vidrio puede ser tratada durante 30 segundos en un limpiador de plasma para obtener resultados óptimos de recubrimiento). </li><li> Prepare una solución al 2% metiltrimetoxi en acetona y cubrir el material de vidrio durante una incubación de 2 minutos en esta solución. Después de quitar el revestimiento de vidrio de la solución y dejar secar por aire. Luego enjuague con etanol al 70% de una botella de spray y dejar secar de nuevo. (Si lo desea, en este punto el material de vidrio se pueden hornear durante 1 hora a 80 ° C para unir covalentemente el revestimiento de los recipientes de vidrio). Vidrio recubierto se puede almacenar por lo menos durante un mes. </li><li> Las proteínas fluorescentes se purifican como su proteína de 6 etiquetado de E. Coli 1. Medir el espectro de absorción de la proteína purificada y preparar una dilución de valores con una densidad óptica de 0,1 ~ STE en tampón (150 mM NaCl, 10 mM Tris-HCl pH 8, 1 mM EDTA), que contiene 0,1% de albúmina sérica bovina (BSA) . Además de preparar 10 ml de una mezcla de 1:1 de tampón 1-octanol y STE en un tubo cónico de 15 ml y mezclar enérgicamente. Después de dejar la mezcla hasta que la separación de fases se ha completado. La fase superior es el 1-octanol. (Atención: Debido a que 1-octanol tiene un olor fuerte que es importante utilizar un contenedor de residuos cerradas por todo lo que entra en contacto con 1-octanol). </li><li> Para hacer la emulsión pipeta 45 l 1-octanol y 5 l de proteínas fluorescentes en un tubo de microcentrífuga. Toque en el tubo de un par de veces con el dedo para comenzar la formación de la emulsión y después sonicar el tubo durante 30 segundos en un baño de ultrasonidos. En el mientras tanto conseguir un portaobjetos de microscopio y cubierta de vidrio recubierto listo. Después de la sonicación de la emulsión debe estar completamente despejado. Inmediatamente después de la sonicación pipeta de 4 l de emulsión de la mitad del tubo en un portaobjetos de microscopio recubiertas y cubra con una tapa de vidrio recubierto. </li><li> Si el procedimiento se realiza correctamente la emulsión se distribuye uniformemente entre el portaobjetos de vidrio y el objeto. En cuestión de minutos la muestra debe ser estable, que consta de aproximadamente 10 micras de espesor gotas fluorescentes con diámetro variable. Las gotas más grandes están cerca del centro de la muestra y los más pequeños se encuentran más hacia los bordes. </li></ol> 2. La creación de un experimento fotoconversión El siguiente procedimiento es una estrategia general para la creación de un experimento de la proteína fluorescente fotoconversión. Este procedimiento se puede aplicar a las proteínas purificadas, así como para las células vivas. <ol><li> Los parámetros siguientes se ofrece un punto de partida general para configurar la prueba fotoconversión: 40x aceite 1.3NA objetivo de inmersión Tamaño de la imagen = 512 x 512 píxeles Exploración zoom = 4 Pixel tiempo de espera = 6 mseg. Z-resolución (tamaño del agujero de alfiler) = 3 m </li><li> Configurar dos canales de detección de la fluorescencia inicial y photoconverted, así como un &quot;canal fotoconversión&quot;. En este ejemplo vamos a utilizar las proteínas purificadas mOrange2, que es una naranja y rojo que la proteína fluorescente fotoconvertible. Las especies de naranja se detecta utilizando 561 nm de excitación y la fluorescencia se recoge entre 570 nm y 630 nm. La especie roja photoconverted se detecta utilizando 633 nm de excitación y la fluorescencia se recoge entre 640 nm y 700 nm. Para el &quot;canal fotoconversión&quot; seleccionar 488 nm de excitación y recoger fluorescencia entre 490 nm y 540 nm. (Nota:. Imágenes del canal fotoconversión no es estrictamente necesario) </li><li> Utilice el canal para obtener imágenes de la fluorescencia inicial con la exploración continua para ajustar la potencia del láser y la ganancia del detector de la calidad de imagen óptima. </li><li> Activar el canal fotoconversión y seleccione un láser de baja potencia. Inicio de una serie de imágenes lapso de tiempo y aumentar gradualmente hasta que el láser fotoconversión importantes de blanqueo de la fluorescencia inicial se observa. Continuar la exploración hasta la fluorescencia inicial es de aproximadamente 75% blanqueado. </li><li> Desactivar el canal fotoconversión y activar el canal de detección de la fluorescencia photoconverted. Inicio de imagen con una ganancia del detector de alta y láser de baja potencia y poco a poco aumentar la potencia del láser hasta que la fluorescencia se detecta photoconverted. Una vez que detectan la fluorescencia photoconverted puede ajustar la potencia del láser y la ganancia del detector de la calidad de imagen óptima. </li><li> Por último, la potencia del láser utilizadopara fotoconversión, así como la duración de fotoconversión necesitan ser optimizados. El aumento de la potencia del láser fotoconversión se acelerará la tasa de fotoconversión, la potencia del láser sin embargo demasiada photobleach la proteína. </li><li> Una vez que el poder fotoconversión láser óptima y la duración se han determinado, estos parámetros se pueden utilizar para configurar un módulo estándar o photobleaching FRAP y el &quot;canal fotoconversión&quot; ya no es necesario. </li></ol> 3. Doble óptica con la sonda destacando mOrange2 y Dronpa Debido a la desplazada hacia el rojo propiedades espectrales, mOrange2 se puede utilizar en combinación con el verde Dronpa photoswitchable proteína fluorescente de doble sonda destacando óptico para permitir destacar selectiva de células individuales 4 (orgánulos) las poblaciones. <ol><li> Las células se cultivan en el fondo de cristal platos MatTek y transfectadas 24 horas antes de usar imágenes estándar Lipofectamine2000 transfección 1. </li><li> Configurar el microscopio en busca de mOrange2 fotoconversión como se describe en la sección 2. </li><li> Configurar el microscopio en busca de photoswitching Dronpa. Dronpa fluorescencia se pueden visualizar mediante el mOrange2 &quot;canal fotoconversión&quot; (véase el paso 2.2). (Nota: Reduzca al mínimo la potencia del láser utilizado para obtener imágenes de Dronpa, porque la potencia del láser demasiado hará que la inactivación de Dronpa.) Añadir un canal para la fotoactivación Dronpa. Usamos 800 nm de excitación de dos fotones para la fotoactivación, pero esta alternativa se puede lograr utilizando 405 nm de excitación. Determinar la potencia del láser necesaria para la imagen, foto-, y photoinactivation Dronpa de fluorescencia. </li><li> Atención: fotoconversión de mOrange2 y la inactivación de Dronpa ambos ocurren a la excitación de 488 nm. Debido a la potencia del láser de alta necesaria para mOrange2 fotoconversión esto también inactivar Dronpa fluorescencia. Por otro lado, la inactivación Dronpa ocurre ya en la potencia del láser mucho más bajo y se puede realizar sin fotoconversión mOrange2 significativo. </li><li> Una vez que los parámetros de fotoconversión mOrange2 y photoswitching Dronpa se establecen, destacando doble sonda óptica se logra a través de los siguientes pasos. En primer lugar, desactivar Dronpa fluorescencia en todo el campo de visión con nm de excitación de baja potencia 488. En segundo lugar, seleccione una región de interés y mOrange2 photoconvert con nm de excitación de alta potencia 488. Por último, seleccione una región de interés para activar Dronpa fluorescencia. </li></ol> 4. Resultados representante <img src="/files/ftp_upload/1995/1995fig1.jpg" alt="Figura 1" /> Figura 1. Gota de preparación de la muestra. A) muestra correctamente preparados de gotas. B) muestra preparada sin recubrimiento de los portaobjetos de microscopio y cubreobjetos. C) muestra preparada sin la adición de 0,1% de BSA. <img src="/files/ftp_upload/1995/1995fig2.jpg" alt="Figura 2" /> Figura 2. Efecto del poder fotoconversión láser y la duración de mOrange2 fotoconversión. Gotas único que contiene la proteína se mOrange2 photoconverted continuamente con diferentes cantidades de potencia del láser 488 nm. Potencia del láser utilizado para fotoconversión relativa fue del 10% (sólido), el 25% (línea discontinua), y el 100% (de puntos). A) naranja fluorescente especies. B) Photoconverted especie fluorescente de color rojo. <img src="/files/ftp_upload/1995/1995fig3.jpg" alt="Figura 3" /> Figura 3. Doble óptica con la sonda destacando mOrange2 y Dronpa. Un celular) que expresa mOrange2-histona H2B y Dronpa Mito-antes fotoconversión, que presentan una fluorescencia de color naranja en el núcleo y la fluorescencia verde en la mitocondria. B) Dronpa fluorescencia se apaga con nm de excitación de baja potencia 488, lo que fotoconversión mínimo de mOrange2. C) mOrange2 se photoconverted a rojo con la excitación nm de alta potencia de 488. D) Dronpa fluorescencia se conecta de nuevo con 800 nm de excitación de 2 fotones. Los paneles son superposiciones de las imágenes de fluorescencia, junto con la imagen de contraste de interferencia diferencial.

<ol>
	<li>Kremers, G. J., Hazelwood, K. L., Murphy, C. S., Davidson, M. W., Piston, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoconversion+in+orange+and+red+fluorescent+proteins.">Photoconversion in orange and red fluorescent proteins.</a> Nature Methods. 6, 355-358 (2009).</li></ol>

La muestra purificada gota proteína fluorescente es un formato de muestra muy conveniente para la caracterización fotofísicas de proteínas fluorescentes, por ejemplo, para estudiar la cinética y la cinética de photobleaching fotoconversión. El volumen de gota muy pequeña (~ 20 picolitros) facilita photobleaching y experimentos fotoconversión, que puede ser difícil de realizar en los sistemas basados ​​en la cubeta. Además, como se muestra aquí la muestra de gota es ideal para la creación de un microscop...

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<td>Microsope slides</td>
<td>&nbsp;</td>
<td>VWR Scientific</td>
<td>48312-003</td>
<td>&nbsp;</td>
<tr>
<td>22 mm cover glass</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>2940-245</td>
<td>&nbsp;</td>
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<td>1-octanol</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>O4500</td>
<td>&nbsp;</td>
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<td>methyltrimethoxysilane</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>M6420</td>
<td>&nbsp;</td>
<tr>
<td>MatTek dishes</td>
<td>&nbsp;</td>
<td>MatTek Corp.</td>
<td>P35G-1.5-14-C</td>
<td>&nbsp;</td>
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<td>Lipofectamine2000</td>
<td>&nbsp;</td>
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photoconversion of purified fluorescent proteins and dual-probe optical highlighting in live cells

Fotoconvertible proteínas fluorescentes (pc-PM) son una clase de proteínas fluorescentes con &quot;resaltador óptico&quot; capacidad, lo que significa que el color de la fluorescencia se puede cambiar por la exposición a la luz de una longitud de onda específica. Destacando óptico no invasivo permite el marcado de una subpoblación de moléculas fluorescentes, lo que es ideal para el seguimiento de las células individuales u orgánulos. Parámetros críticos para fotoconversión eficientes son la intensidad y el tiempo de exposición de la luz fotoconversión. Si la intensidad es demasiado baja, fotoconversión será lento o no ocurrir en absoluto. Por otro lado, demasiada intensidad o la exposición por mucho tiempo puede photobleach la proteína y por lo tanto reducir la eficiencia de fotoconversión. Este protocolo describe un enfoque general de cómo configurar un microscopio láser confocal de barrido para aplicaciones fotoconversión pc-FP. En primer lugar, se describe un procedimiento para la preparación de muestras purificadas gota de proteínas. Este formato de la muestra es muy conveniente para estudiar el comportamiento fotofísicas de proteínas fluorescentes bajo el microscopio. En segundo lugar, vamos a utilizar la muestra de gota de proteínas para mostrar cómo configurar el microscopio en busca de fotoconversión. Y, por último, vamos a mostrar cómo realizar destacando óptica en células vivas, incluidos los de doble óptica con la sonda destacando mOrange2 y Dronpa.

Watch this Scientific Journal Video about Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells at JoVE.com

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Este protocolo describe un método general para llevar a cabo fotoconversión de proteínas fluorescentes en un microscopio láser confocal de barrido. Se describen los procedimientos para la fotoconversión de muestras de proteínas puried, así como de doble sonda óptica destacando en células vivas con mOrange2 y Dronpa.

Fotoconversión purificada de proteínas fluorescentes y de doble sonda óptica destacar en células vivas

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

photoconversion-purified-fluorescent-proteins-dual-probe-optical

Research

JoVE Journal

Biology

12.1K vistas.  Vanderbilt University. Este protocolo describe un método general para llevar a cabo fotoconversión de proteínas fluorescentes en un microscopio láser confocal de barrido. Se describen los procedimientos para la fotoconversión de muestras de proteínas puried, así como de doble sonda óptica destacando en células vivas con mOrange2 y Dronpa.

Video: Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Etiquetado de las células madre con tintes fluorescentes para la detección no invasiva con imágenes ópticas

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Investigación

Biología

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Limpieza y cuidado apropiado del microscopio

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

Marcaje fluorescente de COS-7 Expresando etiqueta SNAP-proteínas de fusión de imágenes de células vivas

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

Translocación polarizado de proteínas fluorescentes en Xenopus Ectodermo en respuesta a la señalización de Wnt

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

Imágenes en directo de las proteínas GFP-etiquetados en<em&gt; Drosophila</em&gt; Los ovocitos

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

4D de imágenes de la agregación de proteínas en células vivas

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Microscopia intravital para formar imágenes de estructuras subcelulares en ratones vivos que expresan proteínas fluorescentes

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

En Tracking Clonal vivo de madre hematopoyéticas y células progenitoras Marcado por cinco proteínas fluorescentes utilizando confocal y Microscopía Multifotónica

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

Reconstitución funcional y la actividad del canal Las mediciones de Purificada Wildtype y mutante CFTR Proteína

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Characterization of Calcification Events Using Live Optical and Electron Microscopy Techniques in a Marine Tubeworm

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH distribution and calcium ions can be observed using live microscopy over time. This allows identification of the life stage and the tissue with the feature of interest for further electron microscopy studies. Life stage and tissues of interest are typically higher in pH and Ca signals. 
Here, using H. elegans, we present a protocol to characterize the presence of calcium carbonate structures in a biological specimen on the scanning electron microscope (SEM), using energy-dispersive X-ray spectroscopy (EDS) to visualize elemental composition, using electron backscatter diffraction (EBSD) to determine the presence of crystalline structures, and using transmission electron microscopy (TEM) to analyze the composition and structure of the material. In this protocol, a focused ion beam (FIB) is used to isolate samples with dimension suitable for TEM analysis. As FIB is a site specific technique, we demonstrate how information from the previous techniques can be used to identify the region of interest, where Ca signals are highest.

We demonstrate the use of various microscopy methods that are useful in observing the calcification of a tubeworm, Hydroides elegans, as well as locating and characterizing the first calcified material. Live microscopy and electron microscopy are used together to provide functional and material information that are important in studying biomineralization.

Caracterización de la calcificación Eventos en vivo mediante técnicas ópticas y microscopía electrónica en un Gusano Tubícola Marina

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH ...