Unsere Arbeit auf Herz tube formation wurde von der National Science Foundation Grant (Auszeichnung ID 0744165) auf SGK unterstützt

<ol><li> Frisch vorzubereiten 2ml einer Fixierlösung mit 12,5% Glutaraldehyd in 50 mM Cacodylatpuffer (pH 7,4) in einem 20 ml Glasszintillationsröhrchen. Add 8ml von n-Heptan und kräftig schütteln. Lassen Sie die zwei Phasen zu trennen. Entfernen Sie den oberen Phase mit n-Heptan mit Glutaraldehyd gesättigt, statt in ein sauberes Scintillationsfläschchen und beiseite stellen. Dies wird die Fixierlösung werden. </li><li> Sammeln 0-20 Stunden Embryonen mit einer Embryo-Entnahme-Kammer, die einen kleinen Korb mit einem herausnehmbaren Siebeinsatz ist. Entfernen Sie die äußere Membran Chorion durch Einweichen Embryonen in eine 50% ige Chlorbleichlauge für 2-3 Minuten, oder bis Embryonen schwimmen an der Oberfläche der Bleiche. Gründlich mit ddH20 und blot Embryonen auf einem Papiertuch. </li><li> Mit einer Pinzette, abholen Netzeinsatz mit dechorionated Embryonen und Stelle in die Fixierlösung bedeckt ist, so dass Embryonen aus dem Netz fallen. Lassen Embryonen für 1,5 Stunden bei 4 ° C fix beim Mischen auf einem Kolbenpendel. </li><li> Bereiten Sie eine Petrischale mit doppelseitigem Klebeband ausgekleidet. Entfernen Embryonen aus der Fixierlösung mit einer Pasteur-Pipette und legen Sie sie auf verschweißte Oberfläche Petri dish.Transfer die Embryonen in einer minimalen Menge Fixativ Lösung für das Heptan in das Fixiermittel aus Auflösung der Kleber des Bandes zu verhindern. Schütteln Sie die Petrischale zu einer einzigen Schicht von Embryonen zu machen. Legen Sie Petrischale in der Kapuze bis Heptan verdunstet. Fügen Sie genug PBS + 0,1% Tween20, die Embryonen zu decken. Hand-devitellinize späten Stufe 16 Embryonen unter einem Binokular mit einem stumpfen Wolframnadel. Recover Embryonen mit einer Pasteur-Pipette in ein 1,5 ml Mikrozentrifugenröhrchen. Schritte 5-7 sind in diesem Mikrozentrifugenröhrchen durchgeführt. </li><li> Spülen Embryonen in 0,1 M Cacodylatpuffer, pH 7,4 und dann in einer Lösung mit 1% Osmiumtetroxid in 0,1 M Cacodylatpuffer, pH 7,4 für 1 Stunde bei Raumtemperatur post-fix. Spülen Embryonen mit 0,1 M Cacodylatpuffer, pH 7,4. </li><li> Entwässern Embryonen in einer abgestuften Reihe von Ethanol und Aceton. Entwässern Embryonen mit 50% Ethanol für 10 Minuten, gefolgt von 70% Ethanol für 10 Minuten. Dann in 90% Aceton entwässern für 10 Minuten und zwei 100% Aceton Schritten von jeweils 10 Minuten. </li><li> Entfernen Aceton und dann beginnen die Infiltration durch Zugabe einer 1:1-Epon-Harz Spurr: Aceton-Gemisch, um die Embryonen. Mikrowelle für 3 Minuten mit einem Pelco Biowave Pro Mikrowelle, während die Probe unter Vakuum. Bewerben Unterdruck, um die Probe für weitere fünf Minuten nach der Mikrowelle ist beendet. Austausch der 1:1-Harz, Aceton-Gemisch mit 100% Epon-Spurr Harz und Mikrowelle wieder unter Vakuum. Tauschen Sie die 100% Epon-Spurr Harz ein letztes Mal, und Mikrowelle unter Vakuum. </li><li> Mit einer Pasteur-Pipette, sind Embryonen, die von der Mikrozentrifugenröhrchen ein Silikon Einbettform übertragen. Richten Embryonen in eine Reihe, so dass das hintere Ende des Embryos mit dem sich verjüngenden Rand der Form ausgerichtet ist. Backen Sie in einem 70 ° C im Ofen über Nacht. </li><li> Entfernen Probe und Vorbereitung für das Schneiden. </li><li> Begin Schneiden 1 &amp;mgr; Abschnitte mit einer Diamant-Histo Messer und Richert Ultracut E Mikrotom. Beachten Sie, wenn der erste Embryo Abschnitt erreicht ist. Von diesem Punkt schnitt 100 um in die Probe. Nehmen Sie einen Abschnitt mit einer Schleife, und platzieren Sie auf einen Objektträger. Kurz erhitzen Abschnitt auf einer heißen Platte. Stain Abschnitt mit einem Tropfen Methylenblau. Unter dem Lichtmikroskop zu identifizieren Herzen Lumen. </li><li> Cut 90nm Abschnitte mit einer Diamant-ultra 45 ° Messer und Richert Ultracut E Mikrotom. Pick up mehrere Abschnitte auf einer Kupfer-Netz. </li><li> Grids werden mit 3% Uranylacetat in 50% Ethanol für 10 Minuten gesättigt gefärbt und dann gespült dreimal in doppelt destilliertem Wasser. Dann werden die Gitter mit Bleicitrat (0.04gms in 10ml doppelt destilliertem Wasser und 100 ml 10 M NaOH gelöst) für 2,5 Minuten in eine Kammer mit NaOH-Pellets zu einer CO 2-freie Umgebung zu schaffen gefärbt. Die Gitter sind dreimal in doppelt destilliertem Wasser gespült. </li><li> Die Schnitte werden mit einem JEOL 1200EX Elektronenmikroskop bei 80 kV untersucht und fotografiert mit einer AMT Digitalkamera. </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>

Die Morphogenese der Drosophila embryonale Herz Rohr hat als wertvolles Modellsystem für die Untersuchung von Zell-Adhäsions-und Zell Formveränderungen während der embryonalen Entwicklung entstanden. Eine der Herausforderungen bei der Untersuchung dieser Struktur konfrontiert ist, dass das Lumen des Herzens Rohr nur schwer in whole mount Embryonen sichtbar ist. Die Technik in diesem Video gezeigt, die aus zuvor beschriebenen Verfahren 2.1 angepasst wurde, hat sich als effizient und zuverlässig in der Analyse eine erhebliche Anzahl von Genotypen für Herz-Rohr und Lumenbildung 3 erfolgreiche.

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

Die Morphogenese der<em&gt; Drosophila</em&gt; Embryonale Herz Rohr hat als wertvolles Modellsystem für die Untersuchung der Zellmigration, Zell-Zell-Adhäsions-und Zell Formveränderungen während der embryonalen Entwicklung entstanden. Eine der Herausforderungen bei der Untersuchung dieser Struktur konfrontiert ist, dass das Lumen des Herzens Rohr, sowie die Membran Funktionen, die entscheidend für Herz tube formation, schwer in whole mount Embryonen sichtbar sind, aufgrund der geringen Größe des Herzens Rohr und intra-luminale Raum relativ zu den Embryo. Die Verwendung von Transmissionselektronenmikroskopie ermöglicht höhere Vergrößerung dieser Strukturen und bietet den Vorteil der Untersuchung der Embryonen im Querschnitt, die leicht zeigt die Größe und Form des Lumens. In diesem Video, das Einbetten wir detailliert den Prozess für eine zuverlässige Fixierung und Schneiden der späten Phase<em&gt; Drosophila</em&gt; Embryonen, um die Herzen Rohr Lumen sowie wichtige zelluläre Strukturen einschließlich Zell-Zell-Verbindungen und die Basalmembran zu visualisieren.

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

Wir beschreiben ein Verfahren zur Fixierung, Einbettung, Schneiden und Bildgebung von spät<em&gt; Drosophila</em&gt; Embryonen für Trasmission Elektronenmikroskopie des embryonalen Herzens Röhre. Diese Technik ermöglicht die Visualisierung des Herzens Rohr Lumen sowie die Basalmembran, die Linien das Lumen des Herzens.

Vorbereitung der Embryonen für Elektronenmikroskopie der Drosophila Embryonale Herz Rohr

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 Aufrufe.  UMDNJ-Graduate School of Biomedical Sciences and Rutgers: The State University of New Jersey. Wir beschreiben ein Verfahren zur Fixierung, Einbettung, Schneiden und Bildgebung von spät Drosophila Embryonen für Trasmission Elektronenmikroskopie des embryonalen Herzens Röhre. Diese Technik ermöglicht die Visualisierung des Herzens Rohr Lumen sowie die Basalmembran, die Linien das Lumen des Herzens.

Serie: 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.

Vorbereitung der neuronalen Kulturen aus Midgastrula Bühne Drosophila Embryonen

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

Forschung

Biologie

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.

Die Herstellung Drosophila Embryonen für Live-Imaging mit dem Hanging-Drop-Protokoll

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.

Fluoreszenzmarkierung Drosophila Herz Structures

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.

Visualisierung des schlagenden Herzens in 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.

Vorbereitung der Drosophila S2-Zellen für Lichtmikroskopie

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 ...

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; Kultur Verfahren zur embryonalen Maus Herz

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 Vorbereitung und Longitudinal Imaging der Herzfunktion in vivo unter Verwendung optischer Kohärenzmikroskopie (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.

Probenvorbereitung und Bildgebung von Exosomen von Transmissions-Elektronenmikroskopie

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.

Miniaturisierte Probenvorbereitung für Transmissions-Elektronenmikroskopie

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

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

Biologische Probenvorbereitung durch Hochdruck gefrieren, Mikrowellen-gestützte Kontrastverstärkung und minimale Harz einbetten für Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...