Il nostro lavoro sulla formazione cuore tubo è stato supportato da una National Science Foundation Grant (Premio ID 0744165) per SGK

<ol><li> Appena preparare 2 ml di una soluzione contenente fissativo 12,5% glutaraldeide in tampone cacodilato 50 mM (pH7.4) in un flaconcino da 20 ml di vetro scintillazione. Aggiungere 8 ml di n-eptano e agitare vigorosamente. Lasciare le due fasi separate. Rimuovere fase superiore contenente n-eptano satura di glutaraldeide, posto in una fiala di scintillazione pulito e messo da parte. Questa sarà la soluzione di fissativo. </li><li> Raccogliere embrioni ore 0-20 utilizzando una camera di raccolta di embrioni, che è un piccolo cesto con inserto in rete asportabile. Rimuovere la membrana esterna corion immergendo embrioni in una soluzione di candeggina al 50% per 2-3 minuti, o fino a quando gli embrioni a galla della candeggina. Sciacquare abbondantemente con embrioni ddH20 e asciugare bene su un tovagliolo di carta. </li><li> Utilizzando pinze, pick up inserto in rete che è coperto con embrioni dechorionated e posto nella soluzione di fissativo, permettendo embrioni a cadere dalla mesh. Lasciare embrioni per fissare per 1,5 ore a 4 ° C sotto agitazione in un Nutator. </li><li> Preparare una capsula di Petri foderato con nastro biadesivo. Rimuovere gli embrioni dal fissativo utilizzando una pipetta Pasteur e metterli sulla superficie del nastro Petri dish.Transfer gli embrioni in una minima quantità di soluzione fissativo per evitare che il eptano nel fissativo da sciogliere la colla del nastro. Agitare la piastra Petri di fare un unico strato di embrioni. Posto in piastra Petri cappuccio fino evapora eptano. Aggiungi sufficiente Tween20 PBS + 0,1% per coprire gli embrioni. Mano devitellinize fine tappa 16 embrioni sotto un microscopio da dissezione con un ago di tungsteno smussato. Recuperare gli embrioni con una pipetta Pasteur in una provetta da microcentrifuga 1,5 ml. Passi 5-7 sono svolte in questo provetta per microcentrifuga. </li><li> Sciacquare embrioni in cacodilato 0,1 M, pH 7.4 e poi post-fix in una soluzione contenente tetrossido di osmio 1% a 0,1 M cacodilato tampone, pH 7.4 per 1 ora a temperatura ambiente. Sciacquare embrioni con 0,1 M cacodilato tampone, pH 7,4. </li><li> Disidratare embrioni in una serie graduata di etanolo e acetone. Disidratare embrioni con il 50% di etanolo per 10 minuti, seguito da etanolo al 70% per 10 minuti. Poi disidratano nel 90% acetone per 10 minuti, e due 100 passi% acetone di 10 minuti ciascuno. </li><li> Rimuovere l&#39;acetone e poi iniziare il processo di infiltrazione aggiungendo un Epon-Spurr 01:01 resina: miscela acetone per gli embrioni. Microonde per tre minuti con un forno a microonde Pelco Biowave Pro mantenendo il campione sotto pressione di vuoto. Applicare pressione di vuoto al campione per altri cinque minuti dopo che il forno a microonde è terminato. Scambio della resina 1:1, miscela acetone con il 100% Epon-Spurr resina e forno a microonde nuovo sotto pressione di vuoto. Il 100% di scambio Epon-Spurr resina l&#39;ultima volta, e forno a microonde sotto pressione vuoto. </li><li> Utilizzando una pipetta Pasteur, gli embrioni vengono trasferiti dalla provetta da microcentrifuga in uno stampo in silicone embedding. Allineare gli embrioni in una fila tale che l&#39;estremità posteriore dell&#39;embrione si allinea con il bordo dello stampo conico. Cuocere in forno a 70 ° C durante la notte. </li><li> Rimuovere il campione e prepararsi per il sezionamento. </li><li> Iniziare a tagliare sezioni 1μm utilizzando un diamante Histo coltello e Richert Ultracut E microtomo. Prendete nota di quando la sezione primo embrione viene raggiunto. Da questo punto tagliare 100μm nel campione. Rimuovere una sezione utilizzando un ciclo, e posto su un vetrino. Brevemente calore la sezione su un piatto caldo. Macchia la sezione con una goccia di blu di metilene. Sotto un microscopio ottico identificare il lume cuore. </li><li> Tagliare sezioni 90nm usando un Ultra Diamante 45 ° coltello e Richert Ultracut E microtomo. Pick up più sezioni su una griglia di rame. </li><li> Le griglie sono colorate con acetato di uranile 3% saturi in etanolo al 50% per dieci minuti e poi sciacquate tre volte in acqua bidistillata. Poi, le griglie sono colorate con citrato di piombo (0.04gms sciolto in 10ml di acqua bidistillata e 100 ml NaOH 10M) per 2,5 minuti in una camera contenente granuli di NaOH per creare un CO 2-ambiente libero. Le griglie sono lavata tre volte in acqua bidistillata. </li><li> Le sezioni sono esaminati con un microscopio elettronico JEOL 1200EX a 80Kv, e fotografato con una fotocamera digitale AMT. </li></ol>

<ol>
	<li>Tepass, U., Hartenstein, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Development+of+Cellular+Junctions+in+the+Drosophila+embryo.">The Development of Cellular Junctions in the Drosophila embryo.</a> Dev. Biol. 161, 563-596 (1994).</li><li>Sullivan, W., Ashburner, M., Hawley, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophila Protocols. , (2000).</li><li>Santiago-Martinez, E., Soplop, N. H., Patel, R., Kramer, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repulsion+by+Slit+and+Roundabout+prevents+Shotgun/E-cadherin-mediated+cell+adhesion+during+Drosophila+heart+tube+lumen+formation.">Repulsion by Slit and Roundabout prevents Shotgun/E-cadherin-mediated cell adhesion during Drosophila heart tube lumen formation.</a> J. Cell Biol. 182, 241-248 (2008).</li></ol>

La morfogenesi del tubo Drosophila cuore embrionale è emerso come sistema modello utile per studiare l&#39;adesione cellulare e cambia la forma delle cellule durante lo sviluppo embrionale. Una delle sfide affrontate nello studio di questa struttura è che il lume del tubo cuore è difficile da visualizzare in embrioni di montare tutto. La tecnica ha dimostrato in questo video, che è stato adattato da procedure descritte in precedenza 1-2, ha dimostrato di essere efficace in modo efficiente e affidabile l&#39;analisi di un notevole numero di genotipi per tubo cuore e la formazione lume 3.

preparation of embryos for electron microscopy of the drosophila embryonic heart tube

La morfogenesi della<em&gt; Drosophila</emTubo di cuore&gt; embrionale è emerso come sistema modello utile per studiare la migrazione cellulare, l&#39;adesione cellula-cellula e cambia la forma delle cellule durante lo sviluppo embrionale. Una delle sfide affrontate nello studio di questa struttura è che il lume del tubo cuore, così come le caratteristiche della membrana, che sono cruciali per la formazione del cuore del tubo, sono difficili da visualizzare in embrioni di montare tutto, a causa delle ridotte dimensioni del tubo di cuore e intra-luminale spazio rispetto per l&#39;embrione. L&#39;uso della microscopia elettronica a trasmissione consente un ingrandimento maggiore di queste strutture e dà il vantaggio di esaminare gli embrioni in sezione trasversale, che rivela facilmente la dimensione e la forma del lume. In questo video, abbiamo dettaglio il processo di fissaggio sicuro, inclusione, sezionamento e di stadio avanzato<em&gt; Drosophila</em&gt; Embrioni allo scopo di visualizzare il lume del tubo cuore così come importanti strutture cellulari compresi giunzioni cellula-cellula e la membrana basale.

Watch this Scientific Journal Video about Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube at JoVE.com

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

Descriviamo un processo di fissazione, inclusione, sezionamento, e l&#39;imaging di stadio avanzato<em&gt; Drosophila</em&gt; Embrioni per la microscopia elettronica Trasmissione del tubo cardiaco embrionale. Questa tecnica permette per la visualizzazione del lume del tubo cuore così come la membrana basale, che riveste il lume del cuore.

Preparazione di embrioni per la microscopia elettronica del Drosophila Tubo cuore embrionale

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

preparation-embryos-for-electron-microscopy-drosophila-embryonic

Research

JoVE Journal

Biology

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

15.2K Views.  UMDNJ-Graduate School of Biomedical Sciences and Rutgers: The State University of New Jersey. Descriviamo un processo di fissazione, inclusione, sezionamento, e l'imaging di stadio avanzato Drosophila Embrioni per la microscopia elettronica Trasmissione del tubo cardiaco embrionale. Questa tecnica permette per la visualizzazione del lume del tubo cuore così come la membrana basale, che riveste il lume del cuore.

Video: Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

Preparazione delle Culture neuronale da Midgastrula fase embrioni di Drosophila

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Ricerca

Biologia

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

La preparazione di Drosophila Gli embrioni per Live-Imaging Utilizzando il protocollo goccia

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

Etichette fluorescenti di Drosophila Cuore Strutture

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

Visualizzare il cuore pulsante di Drosophila</em

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

Preparazione del Drosophila Cellule S2 per la microscopia ottica

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Imaging verticale di Drosophila Embrioni

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. Thus, protocols to isolate and culture individual organs of interest are essential to provide a method of both visualizing changes in development and allowing novel treatment strategies. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel. This substrate provides enough support to maintain the three-dimensional structure but is flexible enough to allow continued contraction. In brief, hearts are excised from the embryo and placed in a mixture of cold Matrigel diluted 1:1 with growth medium. After the diluted Matrigel solidifies, growth medium is added to the culture dish. Hearts excised as late as embryonic day 16.5 were viable for four days post-dissection. Analysis of the coronary plexus shows that this method does not disrupt coronary vascular development. Thus, we present a novel method for long-term culture of embryonic hearts.

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel.

A Novel<em&gt; Ex vivo</em&gt; Metodo di Cultura per il mouse embrionale del cuore

Developmental studies in the mouse are hampered  by the inaccessibility of the embryo during gestation. Thus, protocols to  isolate and culture individual ...

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy (OCM)

Longitudinal study of the heartbeat in small animals contributes to understanding structural and functional changes during heart development. Optical coherence microscopy (OCM) has been demonstrated to be capable of imaging small animal hearts with high spatial resolution and ultrahigh imaging speed. The high image contrast and noninvasive properties make OCM ideal for performing longitudinal studies without requiring tissue dissections or staining. Drosophila has been widely used as a model organism in cardiac developmental studies due to its high number of orthologous human disease genes, its similarity of molecular mechanisms and genetic pathways with vertebrates, its short life cycle, and its low culture cost. Here, the experimental protocols are described for the preparation of Drosophila and optical imaging of the heartbeat with a custom OCM system throughout the life cycle of the specimen. By following the steps provided in this report, transverse M-mode and 3D OCM images can be acquired to conduct longitudinal studies of the Drosophila cardiac morphology and function. The en face and axial sectional OCM images and the heart rate (HR) and cardiac activity period (CAP) histograms, were also shown to analyze the heart structural changes and to quantify the heart dynamics during Drosophila metamorphosis, combined with the videos constructed with M-mode images to trace cardiac activity intuitively. Due to the genetic similarity between Drosophila and vertebrates, longitudinal study of heart morphology and dynamics in fruit flies could help reveal the origins of human heart diseases. The protocol here would provide an effective method to perform a wide range of studies to understand the mechanisms of cardiac diseases in humans.

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy OCM

Here, the experimental protocols are described for preparing Drosophila at different developmental stages and performing longitudinal optical imaging of Drosophila heartbeats using a custom optical coherence microscopy (OCM) system. The cardiac morphological and dynamical changes can be quantitatively characterized by analyzing the heart structural and functional parameters from OCM images.

Drosophila preparazione e imaging longitudinale della funzione cardiaca in vivo utilizzando Optical Coherence Microscopy (OCM)

Longitudinal study of the heartbeat in small animals contributes to understanding structural and functional changes during heart development. Optical ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Preparazione del campione e la formazione immagine di esosomi da microscopia elettronica di trasmissione

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Preparazione del campione miniaturizzati per microscopia elettronica di trasmissione

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Preparazione di embrioni per la microscopia elettronica del Drosophila Tubo cuore embrionale

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