この作品はHYにEhrmanの交わりによってサポートされていました。 H.、DV、JRにニューヨーク大学ディーンの論文をCDにDJのフェローシップは、NIH GrantR01 EY13010とDFG交わりRJJ、NIHグラントF32EY016309に医学研究ポスドクのためのジェーン棺チャイルズメモリアル基金（RI 2208/1- 1）。我々は、原稿上のコメントのためのニーナ·フォークトとパメラBoodramに感謝します。

1。はじめに三齢幼虫、midpupalと成虫：このビデオでは、3つの定義済みの発達期間における網膜の解剖および免疫組織化学のための方法を説明します。我々のプロトコルは、他の蛹の段階のために動作しますが（それ以前の段階の詳細については、16を参照）は、1つの焦点面とその核にイメージングのためのすべてのPRに最適であるように、我々は、midpupal舞台を選んだの可視化を容易にする、簡単に識別できます転写因子の発現パターン。後半蛹網膜の郭清は成人網膜の解剖と非常によく似ています。 まず、希望の段階での幼虫や蛹を収集する方法を示しています。そこで、幼虫17-18、midpupalと成人網膜の最適解剖を入手する方法について説明します。第二に、我々は、免疫組織化学および共焦点顕微鏡を使用して、これらの段階でPRの開発を可視化する方法を示します。 昆虫の発育期間は、温度に依存しています。幼虫と蛹の再現ステージングを容易にするために、私たちは12 hr/12時間の明/暗サイクル下で25℃ですべてのフライ株式℃に上げることをお勧めします。 3齢幼虫：産卵後約96時間（25℃）、後期3齢幼虫停止送り、彼らは最終的には蛹になる食品バイアルの壁にさまよう。優しく食品用容器の壁から徘徊3齢幼虫を除去するために柔らかいブラシや鉗子を使用しています。 50％の蛹（ &#39;midpupae&#39;）は蛹化後48時間程度観測されている。正確なタイミングでは、食品バイアル上のマーカーにだけ形成された白い蛹を一周することができます。あるいは、均一なステージングのバイアル（形態学的基準を使用して蛹のステージング用に、19を参照）を得るために新しいバイアルにすべての白い蛹を移動します。 48時間後、慎重にレムピンセットを用いてバイアルからmidpupaeのオベ。 アダルトハエ：CO 2のパッドの上に麻酔の若い大人のハエ（2-5日羽化後）。鉗子を使用してヘッドを取り外し、寒い1X PBSを含むガラスウェルディッシュに保管してください。 3。解剖手順（実験1〜2時間程度） 標本を収集した後、シルガード郭清を皿に冷PBSの一滴の中に置いてください。遺伝子型10-15網膜について解剖する。 試薬や機器の適切な取り扱いのために化学物質等安全データシートだけでなく、金融機関の環境衛生および安全管理室（このプロトコールの最後にある表を参照してください）​​ご相談ください。 1。幼虫アイ成虫<ol><li>口のフックが含まれている幼虫の前端の位置を確認します。シルガード皿の底そっと幼虫の中央部を押すように鉗子を使用しています。 anothを使用erが口のフックをつかむと、体内から優しくゆっくりと引き離し鉗子。本体の主要部分を破棄し、脳に接続されたまま成虫から余分な組織を除去するために前部を保持するために口器を使用しています。 </li><li> 3ウェルガラス皿の1X PBSに鉗子で口フックをつかんで成虫と脳を転送します。固定ステップを（ ​​ステップ4.1）を実行します。 </li><li>固定後、唾液腺、脂肪組織を除去し、離れて目を触角の成虫原基から脳を引き出します。 &quot;難しい&quot;抗体のアクセシビリティを向上させるために、上皮の表面に配置されているperipodial膜は、ディスクの触角の部分を押しながら解剖ピンと離れて膜を剥離することにより除去することができる。次に、（ステップ5.1）免疫組織化学を行ってください。 </li></ol> 2。 50％の蛹化/ Midpupal網膜<ol><li>ピンセットで蛹例後端を保持します。そっと別の鉗子でキューティクルをつかんで蛹例前面を取り外し、ゆっくりと蛹殻から引き離します。頭を露出させるために離れて残留キューティクルをはがします。 </li><li>背前歯部では、基礎となる、柔らかい袋を貫通する鉗子 &quot;のヒントを参考にしてください。これは、頭部へのアクセスを提供します。優しくゆっくりP20ピペットで頭を吸引することによって胸部から頭部を分離します。新鮮な冷1X PBSにピペットの先端から頭を追放し、余分な組織を除去し、頭のキューティクルをつかむと口吻を削除します。固定ステップ（ステップ4.1）に進みます。 </li><li>ピアース着実な組織への1解剖ピンと視葉。ラミナとラミナ/視葉から網膜を分離するために網膜の間に他の解剖ピンを挿入します。免疫組織化学を行う（ステップ5.1）を実行します。 </li></ol> 3。大人の目<ol><li>鉗子で後部頭部カプセルの中央をつかむと、口の部分（吻）、脂肪TISSを削除UEと気管。 </li><li> 1鉗子で背頭のキューティクルをつかむと、そっと横に引いて、脳組織から目を分離するために、他を使用しています。ほとんどの場合、ラミナ、脳の神経網の薄くて透明なシースは、網膜に取り付けられたままになります。これは、固定後粘膜を除去する方が簡単です。 </li><li>アイマージンでキューティクルを除去しますが、さまざまなソリューションに網膜の転送を容易にするために後ろに小片にしておきます。 （ステップ4.1）。固定を行う。 </li><li>ピンセットで片側でそれをつかんで、ゆっくりと網膜を損なうことなく、他の鉗子で横に引いて、ラミナを削除します。それは実装時の網膜上に折ることができるように、残留キューティクルを取り除きます。これらの2つの手順がそうでなければ、イメージングプロセス中に、キューティクルのラミナや遺跡で隠されてしまう光受容体を、可視化するために重要である。免疫組織化学（ステップ5.1）に進みます。 </li></ol> 4。固定と和紙NG（約1時間） <ol><li>新たに固定液を（1×PBS、氷の上で渦や店舗を360μlの37％ホルムアルデヒド溶液の希釈40マイクロリットル）の準備中、氷上に置い3ウェルガラス皿に1X PBSで切開した網膜を格納します。我々は、固定プロトコールを続行すると氷（&gt; 15分）で長期保存時間を避けることをお勧めします。 </li><li> 3.7％ホルムアルデヒド溶液200μlの1×PBSを交換してください。組織を固定液に浸漬されていることを確認します。それはそうでなければ適切に修正されない場合がありますので、これは重要です。必要に応じて、ピンセットで気泡や気管を削除します。 </li><li>室温で15分間シェーカーでインキュベートする。 </li><li> 1×PBSで固定液を交換し、1×PBSで二回すすいでください。 30分（シェーカー上、室温）をPBS中で洗浄を続けます。 </li><li> PBSTでPBSを交換してください。 PBSTで二回すすぎ。 </li><li>幼虫の眼ディスク（3.1.3。）とmidpupal /成人の網膜からラミナからperipodial膜の削除を続行&lt;強い&gt;（3.2.3、3.3.4）。 </li></ol> 5。免疫組織化学<ol><li>ブロッキング：室温でシェーカーで20分間インキュベートし、PBST網膜で5％正常血清200μlのPBSを交換してください。 </li><li>一次抗体溶液を用いてブロッキング液を交換してください。パラフィルムを持つ3つのウェルディッシュを密封し、室温でシェーカー上で一晩インキュベートする。 </li><li> PBSTでソリューションを交換して、3回すすいでください。室温でシェーカーで1時間の最小値をPBSTで洗浄を続けます。 </li><li>室温で一晩振とう機にパラフィルム密封されたガラスだけでなく皿の中で二次抗体溶液とインキュベートで転送。 </li><li> PBSTで抗体溶液を交換して、3回すすいでください。室温でシェーカーで1時間の最小値をPBSTで洗浄を続けます。サンプルは数日間PBSTで続けたが、それは乾燥からそれらを防ぐためには、少なくとも1日に1回、新鮮なPBSTを追加することが重要であることができます。 </li></ol> 6。 Mounting <ol><li> 幼虫とmidpupal網膜を取り付けるために 、きれいなスライドガラスの中央に網膜を転送するためにP20を使用しています。過剰PBSTを削除します。イメージングを容易にするために、網膜を揃えるためにピンセットを使用しています。 </li><li>網膜の上にメディアをマウントする10μlを添加する。明確なマニキュア付き24x40-1カバースリップとシールで軽くカバーしています。 </li><li> 成体網膜を装着するための準備、 &#39;ブリッジ&#39; 20：ガラス面の両端に50％グリセロールの25μlの液滴を追加して、22x22のA-1カバースリップ（厚0.13〜0.17ミリメートル）でそれらをカバーしています。 </li><li> P20ピペットでPBSTを取り外し、網膜が乾燥しないように注意してください。ウェルに培地を取り付けるの20μlを添加する。 &quot;ブリッジ&quot;ガラススライドにP20と網膜を転送するために封入剤を使用しています。 </li><li>東洋に鉗子を使用します（レンズはガラススライドを向ける必要があります）とイメージングを容易にするために、網膜の位置を合わせます。必要な場合は、P20と気泡を除去。 </li><li>ゆっくり明確なマニキュアやスコッチテープでトップとシールエッジの片側から場所24x40-1カバースリップ（厚さ0.13〜0.17ミリメートル）。ストアサンプルは4℃顕微鏡スライドセーバーボックスのC（暗所におく）。 </li></ol> 7。代表的な結果このプロトコルのアプリケーションの例として、我々は、ビデオ内の3発生段階において、感光体分化を可視化するための代表的な結果を示しています。幼虫の目触角ディスクとmidpupal網膜は、それらの開発（ 図1A、B）の中に異なる光受容体の種類にラベルを付けたり、成体網膜は、2つの光受容体のサブタイプ（ 図1C）を可視化するために2ロドプシン抗体で染色した抗体で染色した。

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特別な利害関係は宣言されません。

1。トラブルシューティング我々の経験では、解剖の練習（最大〜数週間）が必要で、テーブルの上に、解剖皿に接触して指で肘や前腕を置くことによって快適な手の位置21を達成することによって促進される。そのように、唯一の親指、人差し指と中指が微妙な動きを実行します。 感光体を傷つけることなく薄層を除去することは、?...

<table border="1"><tr><td></td><td></td><td></td><td> 試薬 </td></tr><tr><td> （PBS1x、pH7.4）をリン酸緩衝生理食塩水</td><td>シグマ</td><td></td><td> 10倍原液20を準備します。室温で1×PBSでストアを取得するために蒸留水で希釈します。解剖の前に氷の上で冷やす。 </td></tr><tr><td>トリトンX-100 </td><td>シグマ</td><td> 9002-93-1 </td><td> 注意：刺激性の手袋を着用してください。 0.3パーセントのTriton X-100（PBST）で50ml 1xPBSを準備します。室温で保管してください。 </td></tr><tr><td> 37％ホルムアルデヒド溶液</td><td>フィッシャー·サイエンティフィック</td><td> F75P1GAL </td><td> 注意：有毒、推定ヒト発がん性物質は、手袋を着用してください。固定ステップの前に、新たにPBSで希釈することにより化学ヒュームフードの3.7％溶液を調製し、氷上店。 </td></tr><tr><td> 5％正常ウマ血清</td><td>ジャクソンイムノリサーチ</td><td> 008-000-001 </td><td> PBSTで5％（v / v）の希釈液を調製します。 4℃で保管してください。 </td></tr><tr><td>一次抗体および二次抗体（ 例えばロバ抗ヒツジのAlexa Fluor 488、ロバ抗ウサギのAlexa Fluor 555、ロバ抗マウスのAlexa Fluor 647） </td><td> Invitrogenのモレキュラープローブ</td><td> A11015 A31572 A31571 </td><td> 4度でPBST及び​​店舗で1:800二次抗体を希釈します。 </td></tr><tr><td>のAlexa Fluor 488ファロイジン</td><td></td><td> A12379 </td><td>二次抗体溶液で1:100に希釈します。 </td></tr><tr><td> Slowfadeゴールド褪色防止試薬</td><td> Invitrogenのモレキュラープローブ</td><td> S36936 </td><td>メディアのマウント。 -20℃で保存してください。 </td></tr><tr><td> GlyceROL </td><td>フィッシャー·サイエンティフィック</td><td> G31-1 </td><td>取付用。室温で蒸留水、店舗で50％希釈液10mlを準備します。 </td></tr><tr><td> CO 2 </td><td></td><td></td><td>ハエの成虫を麻酔するため。 </td></tr><tr><td></td><td></td><td></td><td> 機器 </td></tr><tr><td> 2つの鋭い鉗子（デュモン＃55） </td><td>ファイン科学ツール</td><td> 11255から20 </td><td></td></tr><tr><td>シルガード184シリコーンエラストマーキット</td><td>ダウコーニング</td><td></td><td>シルガード混合物でプラスチックシャーレを充填することにより、シルガード郭清皿を準備します。 </td></tr><tr><td>スリーウェルガラス解剖皿</td><td>フィッシャー·サイエンティフィック</td><td> 21から379 </td><td></td></tr><tr><td> TWO minutien解剖ピン（0.1ミリメートル径）と2 pinholders（12 cm）の</td><td>ファイン科学ツール</td><td> 26002から10 </td><td> 2 pinholdersのそれぞれに1 minutienピンを挿入し、フックを形成するために、ピンの1つを曲げる。 </td></tr><tr><td>顕微鏡カバースリップ（22x22の24x40-1とA-1） </td><td>フィッシャー·サイエンティフィック</td><td> 12-542A </td><td></td></tr><tr><td>顕微鏡スライド（25x75x1.0ミリメートル;予備洗浄） </td><td>フィッシャー·サイエンティフィック</td><td> 12-550-143 </td><td></td></tr><tr><td> 1.5 mlのマイクロチューブ</td><td></td><td></td><td></td></tr><tr><td>マニキュアやセロテープをクリア</td><td></td><td></td><td></td></tr><tr><td>オービタルシェーカー</td><td>ベルコ</td><td></td><td></td></tr><tr><td>スライドホルダーボックス</tD&gt; <td>フィッシャー·サイエンティフィック</td><td></td><td></td></tr><tr><td>パラフィルム</td><td>ビーミス</td><td> PM996 </td><td></td></tr><tr><td>細い絵筆</td><td></td><td></td><td></td></tr><tr><td> P20とP200のピペットとヒント</td><td></td><td></td><td></td></tr><tr><td>解剖顕微鏡</td><td></td><td></td><td></td></tr><tr><td>氷と一緒に小さなバケツ</td><td></td><td></td><td>ガラスウェルプレート、解剖網膜やソリューションを冷却する。 </td></tr></table>

dissection and immunohistochemistry of larval, pupal and adult drosophila retinas

キイロショウジョウバエの複眼は約750 ommatidiumの複数形（ユニット目）から構成されています。それぞれの個眼は、レンズを分泌する錐体細胞、色素細胞、毛細胞と8つの光受容体（PRS）R1〜R8 2を含む約20の細胞で構成されています。 PRSは、特殊な微絨毛構造​​、光に敏感な顔料を含有rhabdomeres、ロドプシンを（RHS）があります。 6 PRのrhabdomeres（R1〜R6）は、台形を形成し、体Rh1 3 4が含まれています。 R7とR8のrhabdomeresは台形の中心にタンデムに配置され、光の同じパスを共有しています。 R7とR8 PRSは確率的に2メインサブタイプ5のRHSのさまざまな組み合わせを表現： &#39;p&#39;のサブタイプでは、p R7sでRH3 は p R8sでRH5と結合されて、 &#39;y&#39;のサブタイプには、y R7sでRH4が関連付けられているのに対し、 yの R8sでRH6 6 7 8。 PRSとommatidiumの複数形の開発の初期仕様では幼虫の目触角成虫原基は、上皮細胞の単層で始まる。差別化の波は、ディスク9に掃引とommatidiumの複数形10月11日に未分化細胞のアセンブリを開始します。 &#39;創始細胞&#39; R8はR7を12月14日 、第1及び新入社員、R1-6と指定されています。その後、蛹の開発中に、PRの分化は感桿分体形成、シナプス形成し、最終的にrhの発現を含む広範な形態学的変化15につながる。 このプロトコルでは、網膜の形成や発達経路に関するさまざまな問題に取り組むために適用することができる網膜の開発の3つの定義された期間、網膜における解剖および免疫組織化学のための方法を説明します。ここで、我々は全体のマウント幼虫、midpupalと成人網膜の単一細胞レベルで段階的なPRの分化を視覚化するために、これらのメソッドを使用します（ <sチョン·&gt;図1）。

Watch this Scientific Journal Video about Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas at JoVE.com

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

ザ<em&gt;ショウジョウバエ</em&gt;網膜は、紋切り型の方法で生成された細胞の種類の数が少ないからなる結晶状格子です<sup&gt; 1</sup&gt;。洗練された遺伝解析への従順は、複雑な発達プログラムの研究を可能にします。このプロトコルは、感光体の分化を中心に、3つの個別の発達段階で解剖し、網膜の免疫組織化学を説明しています。

幼虫、蛹および成人の解剖と免疫組織化学<em&gt;ショウジョウバエ</em&gt;網膜

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

dissection-immunohistochemistry-larval-pupal-adult-drosophila

Research

JoVE Journal

Neuroscience

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

25.1K ビュー.  New York University. ザショウジョウバエ網膜は、紋切り型の方法で生成された細胞の種類の数が少ないからなる結晶状格子です 1。洗練された遺伝解析への従順は、複雑な発達プログラムの研究を可能にします。このプロトコルは、感光体の分化を中心に、3つの個別の発達段階で解剖し、網膜の免疫組織化学を説明しています。

ビデオ: Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

エンハンサートラップフリッパーゼの幼虫における発現と成人のマッピングとその応用ショウジョウバエ CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

神経科学

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

シングルショウジョウバエ個眼の解剖とイメージング

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear. Since there is no cell line that satisfactorily represents mammalian hair cells, research on hair cells relies on primary organ cultures. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice (Figure 1) 1-6. The utricle is a vestibular organ, and the hair cells of the utricle are similar in both structure and function to the hair cells in the auditory organ, the organ of Corti. The adult mouse utricle preparation represents a mature sensory epithelium for studies of the molecular signals that regulate the survival, homeostasis, and death of these cells.
Mammalian cochlear hair cells are terminally differentiated and are not regenerated when they are lost. In non-mammalian vertebrates, auditory or vestibular hair cell death is followed by robust regeneration which restores hearing and balance functions 7, 8. Hair cell regeneration is mediated by glia-like supporting cells, which contact the basolateral surfaces of hair cells in the sensory epithelium 9, 10. Supporting cells are also important mediators of hair cell survival and death 11. We have recently developed a technique for infection of supporting cells in cultured utricles using adenovirus. Using adenovirus type 5 (dE1/E3) to deliver a transgene containing GFP under the control of the CMV promoter, we find that adenovirus specifically and efficiently infects supporting cells. Supporting cell infection efficiency is approximately 25-50%, and hair cells are not infected (Figure 2). Importantly, we find that adenoviral infection of supporting cells does not result in toxicity to hair cells or supporting cells, as cell counts in Ad-GFP infected utricles are equivalent to those in non-infected utricles (Figure 3). Thus adenovirus-mediated gene expression in supporting cells of cultured utricles provides a powerful tool to study the roles of supporting cells as mediators of hair cell survival, death, and regeneration.

Mechanosensory hair cells are the receptor cells of the inner ear. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice. We present the dissection of the adult mouse utricle, and we demonstrate adenovirus-mediated infection of supporting cells in cultured utricles.

成体マウスの卵形嚢とアデノウイルス媒介助演細胞感染の解剖

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear.  Since there ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

幼虫給電回路の機能解析ショウジョウバエ</i

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

のライブイメージング<em&gt;ショウジョウバエ</em&gt;幼虫の神経芽細胞

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of the nervous system often produce viable larvae and adults that have locomotion defective phenotypes that are difficult to adequately describe with text or completely represent with a single photographic image. Current modes of scientific publishing, however, support the submission of digital video media as supplemental material to accompany a manuscript. Here we describe a simple and widely accessible microscopy technique for acquiring high-quality digital video of both Drosophila larval and adult phenotypes from a lateral perspective. Video of larval and adult locomotion from a side-view is advantageous because it allows the observation and analysis of subtle distinctions and variations in aberrant locomotive behaviors. We have successfully used the technique to visualize and quantify aberrant crawling behaviors in third instar larvae, in addition to adult mutant phenotypes and behaviors including grooming.

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Here we describe a simple and widely accessible microscopy technique to acquire high-quality digital video of Drosophila adult and larval mutant phenotypes from a lateral perspective.

の高品質のデジタルビデオを買収<em&gt;ショウジョウバエ</em横から見た&gt;幼虫と大人の行動

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

成体マウス蝸牛のホールマウント解剖および免疫蛍光

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

勉強するリニアアガロースチャネルを使用します<em&gt;ショウジョウバエ</em&gt;幼虫クロールの動作

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

大人のホールマウントの準備のためのシンプルなワンステップ解剖プロトコル<em&gt;ショウジョウバエ</em&gt;ブレインズ

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

キイロショウジョウバエ幼虫と大人の脳の代謝解析

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...