私たちは、ハエの株式の維持のために氏V. Lephartに感謝この作品は、健康補助金（GM62916）と国立科学財団の助成金（MCB0817107）のための国立研究所によってサポートされていました。

多糸染色体のスカッシュの準備のための以下のプロトコルは、ヨハンセン氏らに記載された手順から構成されている。 （2009）。 1。三齢ショウジョウバエの幼虫の文化高品質のスカッシュの準備に最適な多糸染色体を得るために、混雑していない培養条件は、（すなわち、20産卵標準4のメスのハエが&quot;毎日新しいボトルにボトルや変更を飛ぶ程度の場所）が不可欠です。太った人を選択します。彼らはまださまよっている間に3齢幼虫を登るの最初の収穫からではなく、蛹化直前に我々日常的に文化で21 ° C、18 ° Cはバンドは/例えばのようないくつかの目的にはより適している可能性がありますファット染色体が得られますバンド間の領域は、高分解能で可視化する必要があります。 2。多糸スカッシュ材料<ul><li>ドラモンド郭清の鉗子（2） </li><li>ペトリ皿（60 × 15 mm）の</li><li>曇らされた顕微鏡スライド（フィッシャー番号12-544-3）（ポリ-リジンコート） </li><li> 22 × 22 mmの15号のカバースリップ（フィッシャー番号12 - 520B）（Sigmacoteでコーティングした、シグマ＃SL2） </li><li> 22 × 40mmの15号のカバースリップ（フィッシャー番号12 - 530B） </li><li>金、ワイプ</li><li> 20倍対物レンズと位相差顕微鏡</li><li>小さなデュワー（例えば、真空フラスコまたは魔法瓶） </li><li>長い鉗子</li><li>剃刀の刃</li><li>コプリンジャー</li><li>ラバーメイドやタッパーウェアトレイ（または同等の密閉式トレイ） </li><li>パラフィルム（22 mmの正方形にカット） </li><li> 175グラムの重み</li></ul> 3。固定液とソリューション<ol><li> 5Xホルムアルデヒドの株式。 0.74 gの新鮮な溶液を調製しますのdH 2 O 4.0 mlのと1N KOHの28μLとパラホルムアルデヒド。 65℃くらいまで温めパラホルムアルデヒドを溶解し、氷上で保存する。 </li><li>固定液1。 5Xホルムアルデヒドの在庫から0.5ミリリットル10 × PBS、50μlのトリトンX - 100、3.45ミリリットルのdH 2 Oおよび1.0 mlの新鮮な溶液を調製します。トリトンX - 100を分散し、1時間以内に使用して暖かい。 </li><li>固定剤2。 1.5ミリリットルのdH 2 O、2.5ミリリットル氷酢酸、および1時間以内に5倍のホルムアルデヒドの株式や使用から1.0mlの新鮮な溶液を調製します。 </li><li> Lactoacetic酸溶液。 1ミリリットル乳酸、2ミリリットルのdH 2 O、および3mLの酢酸溶液を調製します。 </li></ol> 4。多糸染色体のスカッシュの準備： <ol><li>水で幼虫をすすぎ、解剖のための組織培養皿にPBSに移す。 </li><li>口の先端が鉗子一対のフック、他のペアと方法ダウンの約2 / 3体を保持つかみ、そして口を引っ張らないので唾液腺が露出しているフック。唾液、脳から腺と目触角ディスクを分離し、腺から脂肪体および他の関連する組織を離れて分析する。 </li><li>つのよくうつ病のスライドと同様に2〜固定液2の200から300マイクロリットルの最初のウェルに固定液1の200マイクロリットルを追加。よくいずれかの固定液を1に一度唾液腺の一つのペアを転送し、通常1〜2分程度の標的エピトープに必要な時間、インキュベートする。 （注：そのようなヒストン修飾のほとんどのようないくつかのエピトープは、、5分以上固定が必要な場合があります。） </li><li>使用して鉗子は、ウェル2の固定剤2に唾液腺を転送し、2分間インキュベートする。 </li><li>クリーンSigmacotedカバースリップ上でLactoacetic酸溶液の10〜30μlに腺を転送する。静かにカバースリップ上にポリリジンでコーティングされた顕微鏡スライドを下げると、腺がスライドとカバースリップの間になるように垂直方向の圧力を適用せずに、カバースリップを拾う。 直ちに慎重に一辺上にピンセットでカバーグラスを把握し、細胞溶解と拡散染色体を容易にするゆっくりと組織上の任意の縦の圧力を最小化しようとし、少し前後に移動する。あるいは静かに前後にカバースリップを移動するにも消しゴム鉛筆の側面や指先を使う。前後にカバースリップを移動するの遅れは、彼らがLactoacetic酸溶液に曝されると、より剛性になる傾向があるので染色体腕の広がりを減少させる可能性があることに注意してください。ソリューションの混濁は、多くの場合、細胞分離の良い兆候です。ゆっくりと数回にも広がって染色体を助けることができる鉛筆の消しゴム側で（垂直方向の圧力を避けるために）斜めにカバースリップをタップします。 </li><li>すぐに染色体が十分に広がっているかどうかを判断するために20または40倍の目標との位相差顕微鏡下で組織を調べます。 </li><li>拡散染色が十分であるときに、きれいな金 - ワイプのスタック上にカバースリップの側でスライドを下に設定します。せん断の場合とカバースリップのいずれかの水平移動を避け、第二の上に金がワイプとカバースリップが配置されている場所の上に親指を置き、しっかりと押すことで染色体を平坦に置きます染色体。 </li><li>準備が意図した目的に適しているかどうかを判断するために顕微鏡下で再びスライドを調べます。染色体が退治時に大幅に移動した場合は、あなたのその後の準備にLactoacetic酸溶液の少ない量を使用してください。適当なスライドの十分な数が得られるまで繰り返し、4.1から4.8を繰り返します。スライドのカバースリップ上175グラムのウェイトを置きます。 </li><li>液体窒素で小さなデュワー（例えば、魔法瓶を）埋める。長い鉗子を使用して、液体N 2へのディップスライドは、沸騰が停止するまで、スライドを削除し、そして即座にカバースリップをオフに反転する一角で、きれいなカミソリの刃のエッジを使用してください。 4℃のPBSでコプリンジャーにスライドをすぐに使用する場合、場所℃にそうでなければ95％エタノールで満たされたコプリンジャーにスライドを収集する。霜が組織をス​​ライドに付着し、カバースリップで残っていないことを確認するために離れて昇華した後にカバースリップを調べます。すべてのスライドをPBS（数時間内で使用する）または、エタノール（長期保管用）のいずれかに格納されるまで、スライドの残りの部分で繰り返します。 </li></ol>代表的な結果高品質の多糸押しつぶし標本を得るための最初の重要なステップは、大きな唾液腺の核と脂肪幼虫を成長させることです。二つ目はある程度の練習がかかる場合が拡散手続き、との良好なテクニックです。改良された拡散成功のための一つの先端は潰しステップ中lactoacetic酸溶液の最小限の量を見つけることです。これまでは染色体の腕を洗うこと、過度のストリーミング力を発生させることなく染色体の広がりを十分推進していきます。また、前後にカバースリップを動かす上でのいかなる遅延もLactoacetic酸溶液にさらされたとき、彼らはより剛性になるように染色体の腕の広がりを減少させることは注目に値する。 すべてがうまくいけば、多数のよく広がった多糸染色体があるはずです。図1は、二重緑のバンド間の領域のためのマーカーとし、色素で標識された、そのような準備の例を示している汚れ青の縞模様の地域。不十分な拡散が得られた場合、染色体は、図2に示すように、小さなボールのようになります。一方、あまり拡散が発生した場合、chomosomesは薄すぎると、拡張または図3に示すように、細かく断片化いくつかのケースになります。 <img src="/files/ftp_upload/1748/1748fig1.jpg" alt="図1" /> ダブルJIL - 1ヒストンH3S10キナーゼ（赤）とヘキスト（青色）に対する抗体で標識図1。多糸スカッシュの準備。 <img src="/files/ftp_upload/1748/1748fig2.jpg" alt="図2" /> 不十分な拡散と図2。多糸スカッシュ。 <img src="/files/ftp_upload/1748/1748fig3.jpg" alt="図3" /> 図3。すぎる広がりを持つ多糸スカッシュ。

<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>

従来のスカッシュ固定プロトコルの酢酸と乳酸を含めることは、拡散バンド間分解能と染色体腕の両方を促進するが、残念ながら、いくつかのエピトープは、この治療を生きていけない。そのようなエピトープの例は、H3S10ph（カイら 、2008）である。酸処理はまた、GFPタグ融合タンパク質、DiMario らの固有の蛍光を消光するという欠点を持っているので。 （2006）最近、GFP -抗体?...

preparation of drosophila polytene chromosome squashes for antibody labeling

ショウジョウバエは、長いクロマチン構造と三齢幼虫の巨大な唾液腺多糸染色体で提供される細胞診の利点のための遺伝子調節の関係を研究するためのお気に入りのモデルのシステムとなっています。この組織では染色体は、約1000枚を生じさせる細胞分裂がない場合の複製の多くのラウンドを受ける。 DNAは、特定のタンパク質の局在化とクロマチン形態を相関させるユニークな機会を提供大幅に拡大された染色体で、その結果、各複製サイクルの後に整列されたまま。その結果、別の遺伝子で、転写プロセスの異なる段階で存在するエピジェネティックな修飾を定義することに関心の高いレベルがあった。このような研究のための重要なツールは、酵素に対する抗体を持つ多糸染色体のラベリング、転写因子、または関心のヒストン修飾です。このビデオプロトコルは、抗体の標識にショウジョウバエ多糸染色体を準備するヨハンセンの実験室で使用されているスカッシュのテクニックを示しています。

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

このビデオプロトコルは準備するヨハンセン実験室で使用されているスカッシュのテクニックを示しています。<em&gt;ショウジョウバエ</em抗体標識用&gt;多糸染色体。

抗体標識用ショウジョウバエ多糸染色体のカボチャの準備

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 ビュー.  Iowa State University. このビデオプロトコルは準備するヨハンセン実験室で使用されているスカッシュのテクニックを示しています。ショウジョウバエ多糸染色体。

ビデオ: 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.

Midgastrulaステージショウジョウバエの胚からの神経培養の準備

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

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.

の調製ショウジョウバエ</emハンギングドロッププロトコルを使用してライブイメージングのための>胚

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.

の蛍光標識ショウジョウバエハートの構造

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.

の電子顕微鏡のための胚の調製ショウジョウバエ胎児心臓の管

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.

のライブ解剖ショウジョウバエ胚：抗体染色によってミュータントコレクションをスクリーニングするための合理化手法

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.

開発と大人の準備ショウジョウバエ</emライブイメージングのための>脳と網膜

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.

の準備ショウジョウバエ</em光学顕微鏡用> S2細胞

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.

大人の準備ショウジョウバエ</emシンセクショニングおよび顕微鏡分析のための>アイズ

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.

リビングでの動的動原体タンパク質の挙動と染色体の動きを図解して有糸分裂チェックポイントを勉強<em&gt;ショウジョウバエ</em&gt;合胞体胚

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).

培養細胞からの染色体の準備

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