Notre travail sur la formation du tube cardiaque a été soutenu par une subvention National Science Foundation (0744165 ID Award) pour SGK

<ol><li> Fraîchement préparer 2ml d&#39;une solution de fixateur contenant du glutaraldéhyde 12,5% dans un tampon cacodylate 50mM (pH 7,4) dans un flacon en verre de 20 ml à scintillation. Ajouter 8 ml de n-heptane et agiter vigoureusement. Laisser les deux phases se séparer. Retirer phase supérieure contenant le n-heptane saturés avec du glutaraldéhyde, le placer dans un flacon à scintillation propre et mis de côté. Ce sera la solution de fixation. </li><li> Recueillir des embryons de 00 à 20 heures en utilisant une chambre de collecte d&#39;embryons, qui est un petit panier avec un tamis amovible. Retirer la membrane externe du chorion par trempage des embryons dans une solution javellisée à 50% pendant 2-3 minutes, ou jusqu&#39;à ce que les embryons flottent à la surface de l&#39;eau de Javel. Rincez abondamment avec des embryons ddH20 et éponger sur un papier absorbant. </li><li> En utilisant des pinces, ramasser tamis qui est recouvert d&#39;embryons dechorionated et lieu dans la solution de fixation, permettant embryons à tomber à partir du maillage. Autoriser les embryons de fixer pendant 1,5 heures à 4 ° C tout en mélangeant sur un Nutator. </li><li> Préparez une boîte de Pétri bordée de ruban adhésif double face. Retirer le fixateur d&#39;embryons en utilisant une pipette Pasteur et les placer sur la surface enregistrée de Petri dish.Transfer les embryons dans une quantité minime de solution de fixation pour empêcher l&#39;heptane dans le fixateur de se dissoudre la colle de la bande. Secouez la boîte de Pétri pour faire une seule couche d&#39;embryons. Placez une boîte de Pétri dans le capot jusqu&#39;à évaporation d&#39;heptane. Ajouter suffisamment Tween 20 PBS + 0,1% pour couvrir les embryons. Hand-devitellinize stade tardif 16 embryons sous un microscope à dissection avec une aiguille de tungstène émoussé. Récupérer des embryons avec une pipette Pasteur dans un tube de 1,5 ml. Étapes 5-7 sont menées dans ce tube de centrifugeuse. </li><li> Rincez embryons dans 0,1 M de cacodylate pH 7,4 puis post-fixer dans une solution contenant du tétroxyde d&#39;osmium à 1% dans 0,1 M tampon cacodylate à pH 7,4 pendant 1 heure à température ambiante. Rincez embryons avec 0,1 M de cacodylate pH 7,4. </li><li> Déshydrater embryons dans une série graduée de l&#39;éthanol et l&#39;acétone. Déshydrater embryons avec de l&#39;éthanol 50% pendant 10 minutes, suivie par 70% d&#39;éthanol pendant 10 minutes. Puis se déshydrater dans l&#39;acétone à 90% pendant 10 minutes, et deux étapes 100% d&#39;acétone de 10 minutes chacune. </li><li> Enlever l&#39;acétone et ensuite commencer le processus d&#39;infiltration en y ajoutant une 01:01 Epon-Spurr résine: le mélange d&#39;acétone pour les embryons. Micro-ondes pendant trois minutes en utilisant un micro-ondes Pelco Biowave Pro tout en gardant l&#39;échantillon sous pression à vide. Appliquer une pression à vide pour l&#39;échantillon pour une période supplémentaire de cinq minutes après que le micro-ondes est terminée. Bourse de la résine 1:1, mélange acétone à 100% Epon-Spurr résine et micro-ondes à nouveau sous pression à vide. Bourse de 100% de résine Epon-Spurr une dernière fois, et micro-ondes sous pression à vide. </li><li> En utilisant une pipette Pasteur, les embryons sont transférés du tube de centrifugeuse pour un moule en silicone intégrant. Aligner les embryons dans une rangée de telle sorte que l&#39;extrémité postérieure de l&#39;embryon s&#39;aligne avec le bord du moule conique. Cuire au four à 70 ° C pendant la nuit. </li><li> Retirer l&#39;échantillon et se préparer pour la coupe. </li><li> Commencez à couper des sections 1 um en utilisant un couteau de diamant Histo et Richert Ultracut E microtome. Prenez note de quand la section premier embryon est atteint. De ce point, coupé 100 microns dans l&#39;échantillon. Supprimer une section en utilisant une boucle, et placer sur une lame de verre. Brièvement la chaleur de la section sur une plaque chauffante. Tache de la section en utilisant une goutte de bleu de méthylène. Sous un microscope optique d&#39;identifier la lumière du cœur. </li><li> Coupez les sections 90nm utilisant un ultra Diamant 45 ° Couteau et Richert Ultracut E microtome. Ramassez plusieurs sections sur une grille de cuivre. </li><li> Les grilles sont colorées à l&#39;acétate d&#39;uranyle 3% saturés dans l&#39;éthanol à 50% pendant dix minutes, puis rincés trois fois à l&#39;eau bidistillée. Ensuite, les grilles sont colorées avec du citrate de plomb (dissous dans 10ml 0.04gms eau bidistillée et 100ml de NaOH 10M) pendant 2,5 minutes dans une chambre contenant des pastilles de NaOH pour créer un sans CO 2 environnement. Les grilles sont rincés trois fois à l&#39;eau bidistillée. </li><li> Les articles sont examinés avec un microscope électronique JEOL 1200EX à 80Kv, et photographiés avec un appareil photo numérique de l&#39;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 morphogenèse du tube drosophile coeur embryonnaire a émergé comme un système modèle précieux pour étudier l&#39;adhésion cellulaire et les changements de forme cellulaire durant le développement embryonnaire. Une des difficultés rencontrées dans l&#39;étude de cette structure est que la lumière du tube cardiaque est difficile à visualiser dans des embryons de monter ensemble. La technique illustrée dans cette vidéo, qui a été adapté à partir des procédures décrites précédemment 1-2, s&#39;est révélée être réussi à façon efficace et fiable l&#39;analyse d&#39;un nombre considérable de génotypes pour tube cardiaque et la formation de lumière 3.

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

La morphogenèse de la<em&gt; Drosophile</emTube&gt; coeur embryonnaire a émergé comme un système modèle précieux pour étudier la migration cellulaire, la cellule d&#39;adhésion cellulaire et les changements de forme cellulaire durant le développement embryonnaire. Une des difficultés rencontrées dans l&#39;étude de cette structure est que la lumière du tube cardiaque, ainsi que les caractéristiques de membrane qui sont cruciaux pour la formation du tube cardiaque, sont difficiles à visualiser dans des embryons de monter ensemble, en raison de la petite taille du tube cardiaque et intra-luminal espace par rapport à l&#39;embryon. L&#39;utilisation de la microscopie électronique à transmission permet de grossissement plus élevé de ces structures et donne l&#39;avantage d&#39;examiner les embryons en coupe transversale, ce qui révèle aisément la taille et la forme de la lumière. Dans cette vidéo, nous détaillons le processus de fixation fiable, intégrant et sectionnement de stade tardif<em&gt; Drosophile</em&gt; Embryons afin de visualiser le lumen du tube cardiaque ainsi que d&#39;importantes structures cellulaires, y compris jonctions cellule-cellule et la membrane 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

Nous décrivons un processus pour la fixation, l&#39;incorporation, de sectionnement et de l&#39;imagerie du stade tardif<em&gt; Drosophile</em&gt; Embryons for Electron Microscopy Transmission du tube cardiaque embryonnaire. Cette technique permet la visualisation de la lumière du tube cardiaque ainsi que la membrane basale, qui tapisse la lumière du cœur.

Préparation d&#39;embryons pour la microscopie électronique des Drosophile Tube cardiaque embryonnaire

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 vues.  UMDNJ-Graduate School of Biomedical Sciences and Rutgers: The State University of New Jersey. Nous décrivons un processus pour la fixation, l'incorporation, de sectionnement et de l'imagerie du stade tardif Drosophile Embryons for Electron Microscopy Transmission du tube cardiaque embryonnaire. Cette technique permet la visualisation de la lumière du tube cardiaque ainsi que la membrane basale, qui tapisse la lumière du cœur.

Vidéo: 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.

Préparation des cultures de neurones d&#39;embryons de drosophile Midgastrula scène

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

Recherche

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.

La préparation des Drosophile Les embryons destinés à Live-imagerie utilisant le protocole goutte suspendue

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.

Marquage fluorescent Drosophile</emOuvrages d&#39;art> Coeur

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.

Visualiser le cœur battant Drosophile</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.

Préparation des Drosophile Les cellules S2 pour la microscopie optique

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; Méthode de la culture pour le embryonnaire Souris Coeur

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 Préparation et imagerie longitudinale de la fonction cardiaque in vivo en utilisant cohérence optique Microscopie (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.

Préparation des échantillons et l’imagerie des Exosomes par microscopie électronique à Transmission

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.

Préparation de l’échantillon miniaturisé pour la microscopie électronique à Transmission

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.

Préparation de l’échantillon biologique par congélation à haute pression, amélioration du contraste assistée par micro-ondes et minime résine enrobage pour l’imagerie de Volume

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