我们感谢五Lephart女士维护飞生补助金（GM62916）和国家科学基金会的资助（MCB0817107），这项工作是由国家机构支持的股票。

聚乙烯染色体壁球准备以下协议是改编自约翰森等人所描述的程序。 （2009年）。 1。文化的第三龄果蝇幼虫为了获得高品质壁球制剂的最佳聚乙烯染色体，拥挤的培养条件是必不可少的（即大约20个鸡蛋在一个标准的4个“飞瓶和变化到一个新的瓶子铺设雌性果蝇每天），选择最胖的个人从第一次攀登，而他们仍然徘徊，但只是到化蛹前幼虫3龄作物。我们经常文化在21 ° C，但18 ° C将产生丰厚的染色体可能如一些更适合例如当乐队/带间地区需要在高分辨率显示。 2。聚乙烯壁球材料<ul><li>德拉蒙德清扫钳（2） </li><li>培养皿（60 × 15毫米） </li><li>磨砂显微镜载玻片（费舍尔号12-544-3）（多聚赖氨酸涂层） </li><li> 22 × 22毫米15号盖玻片（费舍尔第12号- 520B）（Sigmacote涂层;西格玛＃SL2） </li><li> 22 × 40毫米15号盖玻片（费舍尔12号- 530B） </li><li>金，湿巾</li><li> 20X目标相显微镜</li><li>小杜瓦（例如，真空瓶或保温瓶瓶） </li><li>长钳</li><li>剃刀刀片</li><li> Coplin坛</li><li>橡胶女佣或特百惠托盘（或同等学历密封的纸盒） </li><li>封口膜（切成22毫米平方） </li><li> 175克重量</li></ul> 3。定影液和解决方案<ol><li> 5X甲醛股票。准备一个新鲜的溶液为0.74克多聚甲醛在4.0毫升的DH 2 O 28μL1N氢氧化钾。加热至65 ° C至解散的多聚甲醛，然后储存在​​冰。 </li><li>固定液1。准备一个0.5毫升10X PBS，50μLTRITON X - 100，3.45毫升的dh 2 O和1.0毫升的新鲜的解决方案，从5倍的甲醛存量。温暖驱散TRITON X - 100，使用1小时内。 </li><li>固色剂2。准备一个1.5毫升的dh 2 O，2.5 mL冰醋酸，5倍的股票和1小时内使用甲醛和1.0毫升的新鲜解决方案。 </li><li> Lactoacetic酸溶液。准备1毫升乳酸，DH 2 O 2毫升，3毫升醋酸溶液。 </li></ol> 4。聚乙烯染色体壁球准备： <ol><li>用清水冲洗幼虫和转让，以PBS在解剖组织培养皿。 </li><li>口与一对镊子挂钩把握的提示，按住约2 / 3的方式与其他对身体，拉口钩，使唾液腺被暴露。单独从大脑的唾液腺和眼睛触角光盘，并剖析远离腺体脂肪体和任何其他相关的组织。 </li><li>新增的两个以及抑郁症的幻灯片的第一口井和200-300微升至固定液2以及两个200固定液1微升。一次传输一对唾液腺固定液1井一和孵化为目标抗原表位所需的金额，时间，一般约1-2分钟。 （注：如大多数的组蛋白修饰某些抗原表位，可能需要5分钟或更长的固定。） </li><li>使用镊子转移唾液腺固定液2以及2孵育两分钟。 </li><li> 10-30μl在一个干净的Sigmacoted盖玻片Lactoacetic溶液的传输腺体。到盖玻片，轻轻地一个多聚赖氨酸涂层显微镜幻灯片，而不采用垂直压力，拿起盖玻片，使腺体之间的滑动和盖玻片。立即方便认真抓钳盖玻片上一个边缘细胞裂解和染色体蔓延并轻轻稍微来回移动，试图以尽量减少对组织的垂直压力。另外，使用橡皮擦端甚至指尖轻轻地来回移动盖玻片。请注意，在来回移动盖玻片的任何延迟可能减少染色体蔓延，因为他们往往​​会成为一经曝光Lactoacetic溶液，更严格的。混浊的解决方案往往是一个很好的迹象细胞分离。轻轻敲击盖玻片斜刺（避免垂直压力）与橡皮擦端了几次也可能协助染色体蔓延。 </li><li>立即组织检查，以确定是否染色体是良好的传播与20或40倍的目标相差显微镜下。 </li><li>当染色体的蔓延是足够的，盖玻片一侧设置滑下一叠干净的金，湿巾。将第二金抹在上面，扁平化的染色体，由你的拇指，放在盖玻片定位和紧迫坚决避免任何盖玻片，剪切横向运动染色体。 </li><li>检查再次在显微镜下的幻灯片，以确定是否准备适合预期的目的。如果染色体移动挤压后显着，使用在后续的准备工作Lactoacetic溶液量较少。重复步骤4.1-4.8，直到已经获得了足够数量的合适的幻灯片。将盖玻片的幻灯片后的175克的重量。 </li><li>用液态氮填充一个小杜瓦（如保温瓶）。用长镊子，直到停止沸腾，浸入液态氮中的幻灯片，删除幻灯片，并立即用一个干净的刀片的边缘，在一个角落里翻转盖玻片。如果幻灯片将被立即使用，放置在一个用PBS Coplin JAR在4 ° C否则收集到的幻灯片，用95％乙醇填补了Coplin JAR。检查盖玻片，一旦霜冻已经升华了，以确认该组织坚持到幻灯片中，并没有保持与盖玻片。与其余幻灯片重复，直到所有的幻灯片都存储在任PBS（在几个小时内使用）或乙醇（期较长的存储）。 </li></ol>代表性的成果获得高质量的聚乙烯壁球编制的第一个关键的一步，是成长与大唾液腺核脂肪幼虫。二是良好的技术与传播过程中，这可能需要一些练习。一个改进传播成功的秘诀是要找到在挤压步骤lactoacetic溶液的最小体积。这将促进足够，不会产生过多的流媒体的力量，可以洗染色体臂染色体蔓延。这也是值得注意的来回移动盖玻片的任何延迟将降低染色体蔓延，因为他们变得更加刚性Lactoacetic溶液时，接触到。 如果一切顺利的话，应该有许多很好的普及聚乙烯染色体。图1展示了这样一个准备的例子，双绿化带间地区的标志和标记用的染料，污渍在蓝色的带状地区。如果获得足够的蔓延，染色体看起来像小球，如图2所示。另一方面，如果过多很多传播已经发生，chomosomes太薄，延长或分散成小块，某些情况下，如在图3所示。 <img src="/files/ftp_upload/1748/1748fig1.jpg" alt="图1" /> 图1。聚乙烯壁球准备双标记抗体JIL - 1组蛋白H3S10激酶（红色）和赫斯特（蓝色）。 <img src="/files/ftp_upload/1748/1748fig2.jpg" alt="图2" /> 图2。聚乙烯壁球不足蔓延。 <img src="/files/ftp_upload/1748/1748fig3.jpg" alt="图3" /> 图3。聚乙烯壁球太多扩频。

<ol>
	<li>Cai, W., Bao, X., Deng, H., Jin, Y., Girton, J., Johansen, J., Johansen, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+polymerase+II-mediated+transcription+at+active+loci+does+not+require+histone+H3S10+phosphorylation+in+Drosophila.">RNA polymerase II-mediated transcription at active loci does not require histone H3S10 phosphorylation in Drosophila.</a> Development. 135, 2917-2925 (2008).</li><li>DiMario, P., Rosby, R., Cui, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+visualization+of+GFP-fusion+proteins+on+polytene+chromosomes.">Direct visualization of GFP-fusion proteins on polytene chromosomes.</a> Dros. Inf. Serv. 89, 115-118 (2006).</li><li>Johansen, K. M., Cai, W., Deng, H., Bao, X., Zhang, W., Girton, J., Johansen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+studying+transcription+and+epigenetic+chromatin+modification+in+Drosophila+polytene+chromosome+squash+preparations+using+antibodies.">Methods for studying transcription and epigenetic chromatin modification in Drosophila polytene chromosome squash preparations using antibodies.</a> Methods. 48, 387-397 (2009).</li></ol>

包含在传统壁球固定协议的醋酸和乳酸，有利于带间的决议和染色体臂蔓延，但不幸的是，一些抗原决定簇不生存之本治疗。这样一个抗原表位的一个例子是H3S10ph（ 蔡等 ，2008） 。由于酸的治疗也有它的缺点是淬火GFP标记的蛋白质，DiMario等固有荧光。 （2006年）最近开发的基于甲醛的“无酸壁球技术”，允许无酸敏感的抗原决定簇的抗体检测GFP抗体标记以及GFP融合蛋白聚乙烯染色?...

preparation of drosophila polytene chromosome squashes for antibody labeling

果蝇一直喜爱的模型系统研究由于聚乙烯染色体三龄幼虫的巨大唾液腺提供细胞学优势的染色质结构和基因调控之间的关系。在这个组织中的染色体在细胞分工上升到约1000份的情况下进行多轮的复制。的DNA仍然大大扩大提供一个独特的机会，到相关的特定蛋白质的本地化染色质形态的染色体复制后每个周期相一致。因此，出现了一个高水平的界定目前在不同的基因的表观遗传修饰和转录过程的不同阶段的兴趣。为此类研究的重要工具，是与抗体的酶标记的聚乙烯染色体，转录因子，或感兴趣的组蛋白修饰。这个视频协议说明，在约翰森实验室用于制备抗体标记果蝇多线染色体的壁球技术。

Watch this Scientific Journal Video about Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling at JoVE.com

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

该视频协议说明在约翰森实验室中使用的壁球技术准备<em&gt;果蝇</em&gt;聚乙烯抗体标记染色体。

果蝇聚乙烯南瓜抗体标记染色体的制备

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological ...

preparation-drosophila-polytene-chromosome-squashes-for-antibody

Research

JoVE Journal

Biology

31.0K 次观看.  Iowa State University. 该视频协议说明在约翰森实验室中使用的壁球技术准备果蝇聚乙烯抗体标记染色体。

视频: Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

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.

从Midgastrula舞台果蝇胚胎神经文化的制备

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

科研

生物学

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.

编制果蝇实时成像的胚胎使用悬滴协议

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.

荧光标记的果蝇心结构

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

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

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

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

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

电子显微镜对胚胎的制备果蝇胚胎心管

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

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

现场剖析果蝇胚胎抗体染色筛选突变体集合的简化方法

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

发展和成人的制备果蝇大脑和视网膜实时成像

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

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.

制备果蝇 S2细胞，光学显微镜

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 ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

成人的制备果蝇</em薄切片和显微镜分析>的眼睛

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

龙虎斗动粒蛋白的动态行为与染色体运动，在生活学习有丝分裂关卡<em&gt;果蝇</em&gt;合胞胚胎

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Chromosome Preparation From Cultured Cells

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as fluorescence in situ hybridization (FISH), this assay has the powerful ability to analyze individual cells for aberrations that involve gains or losses of portions of the genome&#160;and rearrangements involving one or more chromosomes. In humans, chromosome abnormalities occur in approximately 1 per 160 live births1,2, 60-80% of all miscarriages3,4, 10% of stillbirths2,5, 13% of individuals with congenital heart disease6, 3-6% of infertility cases2, and in many patients with developmental delay&#160;and birth defects7. Cytogenetic analysis of malignancy is routinely used by researchers and clinicians, as observations of clonal chromosomal abnormalities have been shown to have both diagnostic and prognostic significance8,9.&#160; Chromosome isolation is invaluable for gene therapy and stem cell research of organisms including nonhuman primates and rodents10-13.
Chromosomes can be isolated from cells of live tissues, including blood lymphocytes, skin fibroblasts, amniocytes, placenta, bone marrow, and tumor specimens. Chromosomes are analyzed at the metaphase stage of mitosis, when they are most condensed and therefore more clearly visible. The first step of the chromosome isolation technique involves the disruption of the spindle fibers by incubation with Colcemid, to prevent the cells from proceeding to the subsequent anaphase stage. The cells are then treated with a hypotonic solution and preserved in their swollen state with Carnoy&#39;s fixative. The cells are then dropped on to slides and can then be utilized for a variety of procedures. G-banding involves trypsin treatment followed by staining with Giemsa to create characteristic light and dark bands. The same procedure to isolate chromosomes can be used for the preparation of cells for procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY)14,15.

Chromosomes can be isolated from live cells such as lymphocytes or skin fibroblasts, and from organisms including humans or mice. These chromosome preparations can be further utilized for routine G-banding and molecular cytogenetic procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY).

染色体制备培养细胞

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as ...