심장 튜브 형성에 우리 작품 SGK에 대한 국립 과학 재단 (National Science Foundation) 그랜트 (수상 ID 0744165)에 의해 지원되었다

<ol><li> 갓 20 ML 유리 섬광 병에 50mM Cacodylate 버퍼에서 12.5 %의 글루 타 알데히드 (pH7.4)를 포함하는 정착액 솔루션 2ml 준비합니다. N - 헵탄의 8ml를 추가하고 적극적으로 발라줍니다. 두 단계로 분리 수 있습니다. 깨끗한 섬광의 약병에 N - 헵탄 글루 타 알데히드와 채도를 포함하는 상위 단계, 장소를 제거하고 따로 설정합니다. 이것은 고정력있는 솔루션이 될 것입니다. </li><li> 이동식 메쉬 삽입과 작은 바구니는 배아 수집 챔버를 사용 00~20시간의 배아를 수집합니다. 2-3분의 50 % 표백 용액에 배아를 담글하여 외부 chorion 막 제거 또는 태아는 표백제의 표면에 떠까지. 종이 타월에 ddH20와 얼룩 배아로 철저히 씻어. </li><li> 포셉를 사용하여 태아의 메쉬에서 가을 있도록 정착액 솔루션으로 dechorionated 배아과 장소로 덮여 있습니다 메쉬 삽입을 선택합니다. 배아 4 1.5 시간 고칠 ° C를 Nutator에 혼합하는 동안 허용합니다. </li><li> 양면 테이프로 늘어선 페트리 접시를 준비합니다. 파스퇴르 pipet을 사용하여 정착액에서 배아를 제거하고 테이프의 접착제를 해소에서 정착액의 헵탄을 방지하기 위해 페트리 dish.Transfer의 녹화 표면 정착액 솔루션의 최소한의 배아에 그들을 놓으십시오. 배아의 단일 레이어를 만들기 위해 페트리 접시를 흔들어. 헵탄 증발 때까지 후드의 페트리 접시를 놓습니다. 배아를 커버하기에 충분한 PBS + 0.1 % Tween20를 추가합니다. 뭉툭한 텅스텐 바늘로 해부 현미경으로 직접 devitellinize 후반 16 단계 배아. 1.5 ML의 microfuge 관에 파스퇴르 pipet와 배아를 복구할 수 있습니다. 5-7 단계는이 microfuge 관에서 실시하고 있습니다. </li><li> 0.1M Cacodylate 버퍼, pH를 7.4에서 배아를 씻어 후 0.1M Cacodylate 버퍼, 실온에서 1 시간 동안 산도를 7.4에서 1 % 오스뮴의 tetroxide을 포함하는 용액에 포스트 수정. 0.1M Cacodylate 버퍼, pH를 7.4로 배아를 씻어. </li><li> 에탄올과 아세톤의 등급 일련의 배아를 탈수. 10 분 70 % 에탄올 다음 10 분 동안 50 %의 에탄올과 배아를 탈수. 그런 다음 10 분 10 분 각각의 두 100 % 아세톤 단계 90 %의 아세톤으로 탈수. </li><li> 아세톤을 제거한 다음 1:1 Epon - 스퍼 수지 추가하여 침투 과정을 시작 : 배아에 아세톤 혼합합니다. 진공 압력 아래 샘플을 유지하면서 Pelco Biowave 프로 전자 레인지를 사용하여 3 분 전자렌지. 전자 레인지가 끝난 후 추가로 5 분 동안 샘플로 진공 압력을 적용합니다. 교환 1:1 수지 100 % Epon - 스퍼 수지와 전자렌지로 다시 진공 압력 하에서 아세톤 혼합물. 진공 압력 하에서 교환 100 % Epon - 스퍼 수지 최종 시간, 전자렌지. </li><li> 파스퇴르 pipet 사용 배아는 실리콘 포함 금형에 microfuge 튜브에서 전송됩니다. 배아의 사후 끝이 금형의 테이퍼 가장자리와 일치 그러한 행을으로 배아를 맞춥니다. 70 ° C 오븐에 구워 하룻밤. </li><li> 샘플을 제거하고 sectioning 준비. </li><li> 다이아몬드 조직을 나이프와 리처트 Ultracut E 마이크로톰를 사용하여 1μm 섹션을 절단 시작합니다. 최초의 배아 부분에 도달하면 기록해 둡니다. 이 시점에서 샘플로 100μm 했네요. 루프를 사용하여 섹션을 제거하고 유리 슬라이드에 놓습니다. 간단히 뜨거운 접시에 섹션을 가열. 메틸렌 블루의 드롭을 사용하여 섹션을 얼룩. 가벼운 현미경 심장 루멘를 확인합니다. </li><li> 다이아몬드 울트라 45 ° 나이프와 리처트 Ultracut E 마이크로톰를 사용하여 90nm 부분을 잘라. 구리 격자에 여러 섹션을 선택하십시오. </li><li> 격자 10 분 50 % 에탄올에 포화 3 % uranyl 아세테이트 물들일 후 두 번 증류수로 세 번 씻어서 있습니다. 그런 다음 그리드 2 - 무료 환경 CO를 만들 수 NaOH 알약이 들어있는 챔버에 2.5 분 리드 구연 산염 (10ml 더블 증류수와 100ml 10M NaOH에 용해 0.04gms) 물들일 수 있습니다. 격자는 두 번 증류수로 세 번 씻어서 있습니다. </li><li> 섹션은 80Kv에서 JEOL 1200EX 전자 현미경으로 검사하고, AMT 디지털 카메라로 촬영하고 있습니다. </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>

Drosophila 태아의 심장 관의 morphogenesis는 배아 발달 동안 세포 접착과 세포 모양의 변화를 공부에 유용한 모델 시스템으로 등장하고 있습니다. 이 구조를 공부에 직면한 도전 과제 중 하나는 심장 튜브의 루멘 전체가 탑재 배아의 시각화하기 어려운 것입니다. 이전에 설명한 절차 1-2에서 적응되었다이 비디오에서 보여준 기술은, 효율적이고 안정적으로 심장 튜브 및 루멘 형성 3 genotypes의 상당한 수를 분석에서 성공 입증했다.

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

의 morphogenesis<em&gt; Drosophila</em&gt; 태아의 심장 관은 배아 발달 동안 셀 마이 그 레이션, 세포 - 세포 부착 및 세포 모양의 변화를 공부에 유용한 모델 시스템으로 등장하고 있습니다. 이 구조를 공부에 직면한 도전 과제 중 하나는 심장 튜브의 루멘뿐만 아니라, 심장 튜브 형성에 중요한 막 기능은 전체 탑재 배아의 시각화하기 어려운되는 심장 튜브의 작은 크기로 인해 배아에 상대적으로하고 내부 lumenal 공간. 전송 전자 현미경의 사용은 이러한 구조의 높은 배율을 허용 쉽게 루멘의 크기와 모양을 밝히지 단면에서 배아를 검사의 장점을 제공합니다. 이 비디오에서는, 우리는 세부 안정적인 고정을위한 과정은 포함하고, 늦은 단계의 sectioning<em&gt; Drosophila</em셀 - 셀 분기점 및 지하 막 포함한 심장 튜브 루멘뿐만 아니라 중요한 세포 구조를 시각화하기 위해&gt; 태아.

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

우리는 sectioning, 고정, 삽입을위한 과정을 설명하고, 늦은 단계의 이미지<em&gt; Drosophila</em&gt; 태아의 심장 튜브 Trasmission의 전자 현미경을위한 배아. 이 기술은 심장 튜브 루멘의 시각화뿐만 아니라 라인 가슴의 루멘을 지하 막에 대한 수 있습니다.

의 전자 현미경을위한 배아의 준비 Drosophila 태아의 심장 관

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

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Biology

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

15.2K 조회수.  UMDNJ-Graduate School of Biomedical Sciences and Rutgers: The State University of New Jersey. 우리는 sectioning, 고정, 삽입을위한 과정을 설명하고, 늦은 단계의 이미지 Drosophila 태아의 심장 튜브 Trasmission의 전자 현미경을위한 배아. 이 기술은 심장 튜브 루멘의 시각화뿐만 아니라 라인 가슴의 루멘을 지하 막에 대한 수 있습니다.

비디오: 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.

Midgastrula 스테이지 Drosophila의 태아에서 문화의 연결 준비

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

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생물학

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.

의 준비 Drosophila</em공중 드롭 프로토콜을 사용하여 라이브 영상을위한> 태아

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.

의 형광 라벨링 Drosophila 심장 구조

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.

에 잔 심장을 떠올리 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.

의 준비 Drosophila</em라이트 현미경을 위해> S2 세포

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.

소설<em&gt; 생체</em배아 마우스 사랑을&gt; 문화 방법

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.

초파리 준비 및 빛 간섭 현미경을 사용하여 생체 내에서 심장 기능의 세로 이미징 (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.

샘플 준비 및 전송 전자 현미경 검사 법에 의해 Exosomes의 영상

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.

전송 전자 현미경 검사 법에 대 한 소형된 샘플 준비

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.

고압 냉동, 전자 레인지를 이용한 콘트라스트 향상, 그리고 볼륨 이미지에 대 한 포함 하는 최소한의 수 지에 의해 생물학 견본 준비

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