この作品は、NIHの領域のプログラムからの助成金GM096076によってサポートされていました。

1。解離のハエの準備<ol><li>古い1週間未満アール所望の遺伝子（例えば、GFP標識タンパク質を発現する）の健康なハエを麻酔。丸みを帯びたクリーム色の腹部に10月15日大きなメスを選択します。 </li><li>新しい軽くyeasted食品を含むバイアルに選択されたハエや少数（3-5）男性を転送し、ハエは25℃で2日間太らせることができショウジョウバエの準備の詳細については、ワイルら 6を参照してください。肥育ハエは、ステージの様々な、特にステージ8月12日は、イメージングに利用できることが保証されます。不十分肥育またはunfattenedすると、通常はごく初期段階（1-6）と成熟（ステージ14）卵母細胞が含まれて飛ぶ。 </li></ol> 2。直立複式顕微鏡を用いたイメージングのための卵母細胞の調製<ol><li>シリコーンgreasを使用して、約1cm離れて、スライドにカバーガラス2を貼付することによって、標準ガラススライドを準備電子。唯一のカバーガラスとスライドとの間の良好なシール性を確保するのに十分なグリスを使用しています。各カバースリップをしっかりと押すと、薄く均一にグリスを広めるでしょう。カバーガラスとスライド間の接合部にいくつかのハロカーボン油27（シグマ）を追加します。彼らは、以降の手順で隔離された卵母細胞を傷つけることを防ぐためにスペーサーとして機能します。第1.5カバーガラスは、後期の卵母細胞のものと一致する平均厚さ（0.16〜0.18ミリメートル）で、うまく機能します。 </li><li>場所2-3は、22ミリメートル2カバーガラスの表面にハロカーボン油27滴。できるだけ健康な卵母細胞を得るためには、個々の女性は、口ピペッティングを用いた食品のバイアルから削除され、静かに第一麻酔なしでハロカーボン油に堆積される。 </li><li>鋭い解剖鉗子（例えばデュモン＃5など）を使用し、カバースリップに対するハエを固定し、腹部の先端をやってのける。油の下、カバースリップの表面に沿ってドラッグすることにより、卵巣を取り外します。これは、索状帯を推進していきますカバースリップに卵母細胞のn。 </li><li>そっと卵形成の少なくともステージ10Bアール卵母細胞を探し、卵巣を離れていじめる。この段階での卵母細胞は、卵母細胞、卵室の総容積の少なくとも50から60パーセントを占めている卵室を検索することで識別することができます。卵形成のこの時点では、卵母細胞を覆う包細胞は円柱状となっており、ナース細胞への接続が残っている小さな領域を除いて、すべての側面に卵母細胞を囲む。卵巣管シースから個々の卵母細胞を除去するためにピンセットを使用しています。 </li><li> 5月8日無傷の卵母細胞をカバーガラスで隔離されるまで、1〜3人の女性を使用して、進みます。任意の利用不能組織を除去する。 </li><li>慎重に卵母細胞が2つのスペーサーとの間に配置されるように、あらかじめ用意したスライド上に卵母細胞を含むカバースリップを反転させる。それは卵母細胞を傷めますので、反転カバースリップを押し下げないでください。必要であれば、tにハロカーボンオイルを少量加える&quot;サンドイッチ&quot;の彼は縁のスペースを確保するためには満たされている。 （オプション：カバースリップの沈降を可能にするために数分間スライド休息しましょう​​）のスライドは今イメージングのための準備ができています。 </li></ol> 3。倒立複合顕微鏡を用いたイメージングのための卵母細胞の調製<ol><li>セクション2で説明したようにガラス底のインサート（マテック、35ミリメートル）でその小さなペトリ皿をカバーガラスの代わりに使用された卵母細胞を除いては、基本的に解剖されています。これは、検体をカバーする必要がなく、解剖後、組織を皿で直接画像化することができますが、注意が卵母細胞は油で覆われており、完全に乾くことは許されません残っているように注意する必要があります。 </li></ol> 4。ライブイメージング<ol><li> DICや位相コントラスト光学系を用いた焦点に卵を持参し、そのような細胞質の漏出、または包細胞の膨らみなどの損傷の形跡がないかを卵母細胞を調べます。健康で損傷していない画像のみが表示された卵母細胞。 </li><li> Strea細胞質内に卵黄胞が7自家あるので銘は、GFP標識を必要とせずに、直接監視することができます。この信号は、オシロスコープのFITCの範囲に取り込むことができますが、可能性が高いGFP標識タンパク質で使用されているものよりも高いゲイン設定が必要になります。 </li><li>ストリーミングの最高のイメージがちょうどガラスカバースリップ/ペトリガラスに載っている卵母細胞の表面の下にある光のセクションから取得されます。ここでは卵母細胞は、わずかにピントの良い面のために作り、平坦化される。画像は200Xで捕捉することができます - 千X.カバースリップが卵母細胞と客観の間に障壁を形成するので、ここで説明した構成を使用して、空気の目的は、使用されるべきである。それが大幅にサンプルのドリフトや動きを最小限に抑えられるため、これが最適です。 </li><li>関心領域が合うと、brighfield光学系をオフにして、共焦点イメージングに切り替えます。ソフトウェアの高速スキャン/プレビュー機能を使用して、フォーカス調整とゲイン必要に応じて。 </li><li>キャプチャの設定は顕微鏡から顕微鏡に異なりますが、一般的なパラメータは次のとおりです。〜488nmの励起波長および〜515nmの発光波長を使用します。 Z軸はZ軸がフレームの間に10秒の遅れで、一定になるようにキャプチャを設定します。卵母細胞は静止しており、焦点面が適切であることを確認するために5から10フレームのテストシーケンスをキャプチャします。 </li><li>典型的な時間経過のシーケンスは、20の間にキャプチャされます - 50フレーム、ビデオ品質を評価するために、完成した直後に確認することができます。プロセスは、スライド内の他のセルに対して繰り返すことができる、または、必要に応じて追加の女性を解剖することができます。 </li></ol> 5。トラブルシューティングとよくある問題<ol><li>ドリフトは - 正立顕微鏡を使用する場合、細胞は時々油の拡散のためにドリフトすることができます。これは、カバーガラスの下の面積をカバーするために必要なだけのオイルを使用することによって最小限に抑えることができます。問題が解決しない場合は、少ないシリコーンgを使用するスライドに接辞スペーサーにrease。 </li></ol> 注：一部の共焦点ソフトウェアパッケージは細胞追跡を可能にするが、一般的にそれが漂って卵母細胞のイメージングを断念し、代わりに別の試験片を使用することをお勧めします。 <ol start="2"><li>ストリーミングの兆候は - ストリーミングが明らかでない場合、それはおそらく2つの原因のいずれかが原因ではない、どちらかの画像キャプチャのためのフォーカルプレーンが正しくありません（卵母細胞の内部に通常は深すぎる）または卵母細胞が損傷しています。時間と練習が両方のエラーを最小限に抑えることができます。 </li></ol>

<ol>
	<li>Becalska, A. N., Gavis, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lighting+up+mRNA+localization+in+Drosophila+oogenesis.">Lighting up mRNA localization in Drosophila oogenesis.</a> Dev. 136, 2493-2503 (2009).</li><li>He, L., Wang, X., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shining+light+on+Drosophila+oogenesis:+Live+imaging+of+egg+development.">Shining light on Drosophila oogenesis: Live imaging of egg development.</a> Curr. Opin. Gen. Dev. 21, 612-619 (2011).</li><li>King, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Ovarian development Drosophila melanogaster. , (1970).</li><li>Spradling, A. C., Bate, M., Martinez-Arias, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+genetics+of+oogenesis.">Developmental genetics of oogenesis.</a> The Development of Drosophila melanogaster. 1, 1-70 (1993).</li><li>Serbus, L. R., Cha, B. -. J., Theurkauf, W. E., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynein+and+the+actin+cytoskeleton+control+kinesin-driven+cytoplasmic+streaming+in+Drosophila+oocytes.">Dynein and the actin cytoskeleton control kinesin-driven cytoplasmic streaming in Drosophila oocytes.</a> Dev. 132, 3743-3752 (2005).</li><li>Weil, T. T., Parton, R. M., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+Individual+Drosophila+Egg+Chambers+for+Live+Imaging.">Preparing Individual Drosophila Egg Chambers for Live Imaging.</a> J. Vis. Exp. (60), e3679 (2012).</li><li>Theurkauf, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Premature+microtubule-dependent+cytoplasmic+streaming+in+cappuccino+and+spire+mutant+oocytes.">Premature microtubule-dependent cytoplasmic streaming in cappuccino and spire mutant oocytes.</a> Science. 265, 2093-2096 (1998).</li><li>Morin, X., Daneman, R., Zavortink, M., Chia, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protein+trap+strategy+to+detect+GFP-tagged+proteins+expressed+from+their+endogenous+loci+in+Drosophila.">A protein trap strategy to detect GFP-tagged proteins expressed from their endogenous loci in Drosophila.</a> PNAS. 98, 15050-15055 (2001).</li><li>R&#246;per, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rtnl1+is+enriched+in+a+specialized+germline+ER+that+associates+with+ribonucleoprotein+granule+components.">Rtnl1 is enriched in a specialized germline ER that associates with ribonucleoprotein granule components.</a> J. Cell Sci. 120, 1081-1092 (2007).</li><li>Pokrywka, N. J., Payne-Tobin, A., Raley-Susman, K. M., Swartzman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubules,+the+ER+and+Exu:+New+associations+revealed+by+analysis+of+mini+spindles+mutations.">Microtubules, the ER and Exu: New associations revealed by analysis of mini spindles mutations.</a> Mech. Dev. 126, 289-300 (2009).</li><li>Petri, W. H., Mindrinos, M. N., Lombard, M. F., Margaritis, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+development+of+the+Drosophila+chorion.ih+a+chemically+defined+organ+culture+medium.">In vitro development of the Drosophila chorion.ih a chemically defined organ culture medium.</a> Wilhelm Roux's Arch., Devl. Biol. 186, 351-362 (1979).</li></ol>

私は、開示することは何もありません。

ショウジョウバエの卵母細胞は、細胞と細胞内の成分の機能と組織に関連するさまざまな質問に対処するための汎用性の高いモデル系です。タンパク質の機能を完全に理解するには、タンパク質の挙動の空間的および時間的な側面の両方の監視が必要です。 Drosophilists利用できるイメージングシステムと遺伝の両方のツールの進歩は、それが可能であっても、小さなラボが生きた卵母細胞では、高品質のイメージング実験を実行するために作る。ここで私は、タンパク質の機能を調べるために遺伝的背景の様々な組み合わせで使用することができます後期卵形成半ばから中にGFPタグや自家蛍光タンパク質や構造物の挙動を解析するためのプロトコルの概要を示します。 このプロトコルでは、私は卵母細胞イメージングと蛍光タンパク質との構造の時間経過のビデオの取得を可能にする基本的な共焦点の設定のために準備されるプロセスを説明します。追加のdetai卵母細胞の調製上のlsは、ワイルらによって記載されています。6は 、多種多様な内因エクソントラッピング経由でタグ付けされているタンパク質を発現するハエ株の8入手可能であり、無限に多くのタンパク質変異体を設計し、ハエに再導入することができる。 生体組織での作業では、標本の健康が最も重要である。ステージ10Bから卵室は短期イメージング研究に最適です、基本的には自律ファッション11に卵形成を完了することができます。ケアは、高品質のデータを得るために、いくつかの重要な手順で注意しなければなりません。郭の間にそれはできるだけ少なく卵巣と卵母細胞を操作すると、切開及び撮像が完了するまでの時間を最小限に抑えることが重要です。理想的には、小さな切開ステーションが焦点範囲を収容する同じ部屋に配置されるべきであり、一度に少数の卵室が画像化されるべきである。ここで推奨されている数字（5-8）良い品質の画像を得ることの可能性を最大化しながら、解剖時間を最小にすることを保証します。私は、画像の質を評価し、キャプチャの設定が長い時間経過のシーケンスをキャプチャする前に、最適化されていることを確認するために、画像の短いシリーズを取り込むことをお勧めします。ほとんどのソフトウェアは個々のフレームが保存されることを可能にし、これらの画像は、ImageJの12または他のソフトウェアを使用して様々な方法で分析することができます。

<table border="1"><tbody><tr><td>名前</td><td>会社</td><td>カタログ番号</td><td>注釈</td></tr><tr><td>ハロカーボン油27 </td><td>シグマアルドリッチ</td><td> H8773-100ML </td><td></td></tr><tr><td> ＃5SF鉗子</td><td>ファイン科学ツール</td><td> 11252から00 </td><td></td></tr><tr><td>シリコングリース</td><td>シグマアルドリッチ</td><td> Z273554-1EA </td><td></td></tr><tr><td>ガラス底ペトリ皿</td><td>マテック</td><td> P35G-0-20-C </td><td></td></tr></tbody></table>

live imaging of gfp-labeled proteins in drosophila oocytes

ショウジョウバエ卵母細胞は、細胞骨格の機能、細胞組織、細胞小器官の構造と機能などの基本的な疑問を調査するための汎用性の高いシステムとして確立されている。各種のGFPタグ融合タンパク質の有用性は、多くの細胞プロセスは数分または数時間かけて生きた細胞で監視されており、この技術を用いてすることができ、そのようなRNP輸送、上皮の形態形成、組織の再構築などのプロセスが非常に詳細に記載されていることを意味しますショウジョウバエ卵母細胞の1,2インチ変異体の豊かなレパートリーと組み合わせるビデオイメージングを実行する能力は、信じられないほど詳細に検討すべき遺伝子とプロセスの巨大な様々なことができます。その一例が半ば卵形成3,4で開始する卵細胞質のストリーミングのプロセスです。質小胞のこの活発な動きは微小管とキネシン依存5であると便利sysを提供これらの段階で細胞骨格の機能を調べるためのTEM。 ここで私は、ほぼすべての共焦点顕微鏡のセットアップを使用して生きている卵母細胞のタイムラプスイメージングのためのプロトコルを提示。

GFP標識タンパク質のイメージングは​​、タグ付けされたタンパク質が発現さどれだけ強くかによって異なります。 reticulon様1（Rtnl-1）を発現する中期段階の卵母細胞は、GFP 8,9でタグ付けエクソンは、強い信号を生成する。 Rtnl-1は卵細胞質のストリーミング10（ 図1A）中に存在する長い微小管と共局在することが表示されます。 Rtnl-1は、ストリーミングを観察するための優れたマーカーであり、また、後期の卵母細胞における微小管の組織の指標である。 1フレームのキャプチャレート10秒毎（ 図1B）を使用する際に野生の卵室における卵細胞質のストリーミングは自家卵黄胞の顕著な動きとしては明らかである。 5フレームの最大投影は小胞の移動パスをマークするために生成することができます。 <img alt="図1" src="/files/ftp_upload/50044/50044fig1.jpg" /> 図1。 autofl両方のイメージングを生きるuorescentとGFPタグ構造。400XでRtnl1-GFPを発現する卵母細胞のステージ11）は、単一のタイムラプス画像。 Rtnl1-GFPの繊維はストリーミング卵母細胞における波のような動きで移動します。自家卵黄小胞が400Xの倍率で、50秒の合計10秒ごとに結像されたステージ10B卵室のB）は 5フレームの最大投影。スケールバー=30μmで

Watch this Scientific Journal Video about Live Imaging of GFP-labeled Proteins in Drosophila Oocytes at JoVE.com

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

個々のGFPタグ付きタンパク質または構造体の自家蛍光ライブイメージングのためのプロトコル<em&gt;ショウジョウバエ</em&gt;卵母細胞は、記載されている。

でGFP標識タンパク質のイメージングを生きる<em&gt;ショウジョウバエ</em&gt;卵母細胞

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

live-imaging-of-gfp-labeled-proteins-in-drosophila-oocytes

Nanyang Technological University

Research

JoVE Journal

Biology

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

11.3K ビュー.  Vassar College. 個々のGFPタグ付きタンパク質または構造体の自家蛍光ライブイメージングのためのプロトコルショウジョウバエ卵母細胞は、記載されている。

ビデオ: Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

ショウジョウバエの眼成虫ディスクに付属してグリア細胞の移行のイメージングを生きる

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

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 ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

のライブイメージングキイロショウジョウバエ胚性血球移行

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

COS - 7の蛍光標識は、生細胞イメージングのためにSNAP - Tag融合タンパク質を発現する

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

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 ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

の直立イメージングショウジョウバエ胚

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

リビングでのイメージング、細胞の形状の変更ショウジョウバエ胚

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

個々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>エッグチェンバース

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

DNA損傷応答タンパク質の2次元および3次元生細胞イメージング

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.