KBはNIH助成金GM54544によってサポートされています。

パート1：設備＆備品<ol><li>良い解剖顕微鏡は、胚の解剖が必要です。40X倍率は25倍アイピース最大限に倍率を増加させると、示唆されている。 </li><li>ファイン鉗子（番号5）は胚と胚のdevitellinizationの手動選択のために必要とされています。 </li><li>微細なガラス針を作成したり変更したりする機器が必要です。プルガラス針は解剖のために必要とされています。我々は解剖（長い最後）のために固体ガラスを好むが、標準的な肉厚ガラス管（外径1〜1.5ミリメートル）は同様に動作します。一部の人々は、10 +アンプの単巻変圧器の電池を使用して、1MのNaOHのバスで目的のシャープネスに電解細かいタングステン針を、使用してください。シャープタングステン郭清の針は、相対的に弾力性と長期的であることの利点を持っている。中空のガラス針は、（パッチクランプ電極に似て形成された）プラスチックチューブに取り付けられており、生理食塩水の吸引と追放のための解剖だけでなく、接着剤の制御された送達時に使用されます。ガラス引き手のさまざまなガラス針を製造するための用意されています。コンピュータにプログラムさ車夫は、形や大きさのさまざまなを引っ張ってプリセットすることができます。ブラウン - 炎のモデル（サッターインストゥルメント社）が広く支持されています。複合顕微鏡（20 - 40Xの目的は）使用前に電極を検査および変更できるようにする必要があります。 </li><li>一般的に他の無脊椎動物のシステムの記録のために使用されるソリューションに基づいて、1）&quot;標準&quot;（月と月、1976a）や&quot;変更された標準&quot;（Broadie、2000）サラン：ソリューションの2つのタイプは、一般的にお風呂に解剖時の胚組織を使用している、および2）&quot;体液のような&quot;（HL）サラン（Stewartら1994）、標準の生理食塩水とショウジョウバエの体液で測定イオン濃度の間の妥協。それは、これらのSalinesのはどれも正確に生体内の体液の化学組成（Broadie、2000）模倣ないことに注意しなければならない。標準の生理食塩水が含まれる（mMで）：135塩化ナトリウム、5 KClを、4のMgCl 2、1.8 CaCl 2で 、72ショ糖、5 TES、pHは7.2。実験者はまた、電気生理学的記録には適さないが、露出の組織の健康を保護する可能性が最も高い商業的に調製された昆虫の生理食塩水を（例えばシュナイダーの昆虫メディア）、検討する必要があります。 </li></ol>パート2：胚のステージングと解剖<ol><li>最適な卵のコレクションの場合は、2〜3日のための新鮮な卵ポットで（&lt;7日間）、若い、健康なオスとメスのハエを（20-40）維持する。産卵ポットは、一般的に60 mmの寒天プレートをカバーする十分な空気循環を、提供するためにミシン目を100mlのプラスチックビーカー（トライ太）で構成されています。寒天プレートは、寒天で固めリンゴやブドウジュースが含まれています。いくつかのレシピが存在しますが、次のように1つの例です：水を700ml、25〜30グラムの寒天、濃縮ジュース（ぶどうやりんご）、0.5グラムメチルパラベン（P - hydroxymethylbenzoate）を300ml、および30gの砂糖を。水と寒天をオートクレーブし、別々に砂糖やジュースでのp - hydroxymethylbenzoateを煮る。寒天とジュースのソリューションを混在させるとすぐに泡を取り除く、60 mmのプレートに注ぐ。 </li><li>採卵の前に、ハエは、少なくとも2日間に少なくとも1日2回に変更、新鮮なプレート上で酵母ペースト（7グラム4℃で保存されている水9mlののパン酵母° C）で供給されています。 3日目では、良好なポットは、時間あたり100から200の卵を生成する必要があります。より良い産卵は傷プレートの寒天とその側に維持ポットで観察されます。多くの胚は、年に置かや傷の隣になります。 </li><li>胚を大量に収集するには、静かに絵筆を使ってバスケットに寒天プ​​レートから胚を移す。バスケットは、15または50 ml遠心チューブで行うことができます。蓋とチューブの上部との間の画面（70μmのメッシュ）をトラップするのに十分な表面積を残して底部を切り取ります。のdH 2 0で胚をすすぎ、外側の絨毛膜を除去するために新鮮な50％〜100％漂白剤のペトリ皿に入れ。また、卵は細かい鉗子を使用して個別に採取し、漂白剤のドロップに、それらを手動で配置することによりdechorionatedすることができます。デchorionationは30秒から2分（新鮮な漂白剤がはるかに高速です）に連れて行くことができ、解剖顕微鏡下で監視する必要があります。絨毛膜の除去は、光沢、透明な卵黄膜を公開します。漂白剤は、急速にdevitellinized組織を破壊するとしてDechorionated卵は簡単に、すすぎ、または生理食塩水に配置する必要があります。 </li><li>それは慎重に、目的の発達年齢に胚を上演することが重要です。定義されているショウジョウバエ胚の段階では（1-17、カンポス、オルテガとハルテンシュタイン、1985）すべての機能神経筋の開発は、後期の16またはステージ17で発生するように、有用ではありません。このように、胚を25℃発育時間によって上演されている° Cの加湿インキュベーター中で、受精または後の時間で、より一般的に、産卵（AEL）の後。これらの条件下では、胚発生は21日続きます+ / - 25で1時間℃の</li><li>時限卵 - 素人（1〜2時間の卵適切に熟成されているのは）次の形態学的基準のために表示されています。のdH 2 Oで洗浄した後、dechorionated卵を表示するためのプラスチック製の培養皿（のdH 2 Oの下）に配置されます。正しいステージングは​​、解剖顕微鏡（カンポス-オルテガとハルテンシュタイン、1985）と反射光を用いて形態学的基準で確認することができます。孵化寸前の成熟後期の胚（22〜24時間AEL）は、一般的にキューティクルと膨張した気管のセグメンテーションによって認識されています。胚はまた、GFPバランサや適切なフィルターを使用した蛍光解剖顕微鏡を用いて遺伝子型を決定することができます。 </li><li>解剖のために、dechorionatedと発達段階の胚は、生理食塩水のドロップ（ステップ1.5参照）の下にカバースリップ上に（最大背側）接続されています。胚はガラスのマイクロピペットまたはタングステン針を前方または後方の端の近くに膜を穿刺し、軽く膜から胚を解放することによって卵黄膜から削除されます。解剖の準備のために、胚を背側表面上にカバースリップ上に配置されるべきである（解剖まで背側は、実験のための腹側神経系と神経筋肉組織を公開することになります。しかし、胚が明らかに他へのアクセスを容易にするために、他の方法で配置できる組織。） </li><li>初期段階の胚は、（&lt;16時間AEL）ポリリジン（; Broadieとベイト、1993a、Bポリ- L -リジン臭化水素酸塩）を塗布したガラスやガラスをきれいに直接接続。古い胚（&gt; 16時間AEL）は、キューティクル形成に続いて、Sylgard被覆（ダウコーニング）をカバーガラスに理想的に、接着されている必要があります。水重合シアノアクリレート外科または獣医接着剤は、（Broadieとベイト、1993c、d）に使用されます。接着剤は、口腔内圧で制御される接着剤の流れでゴムチューブに取り付けられた小さなガラス電極のピペット（10〜20マイクロメートル、内径の先端）を介して配信されます。接着剤は、生理食塩水の接触により急速に重合するようにケアは、接着剤の配信中に注意する必要があります。接着剤は、二価イオンが重合するために必要と接着が、無または低価のイオン濃度と生理食塩水で実行できないことにも注意してください。接着剤の少量が最初にしっかりと頭と尾を添付するために使用されます。 </li><li>胚の端はカバースリップに接着されると、切開は、ガラス電極または尖ったタングステン針で背側正中線に沿って行われます。これは、静かに最後のカット​​を行う前に切開線を穿孔によって容易になります。切開の側面は、その後ゆっくりと胚のフラットを広めるために、より多くの接着剤でカバーに取り付けられている。腸を含む内臓、脂肪体と、は、必要に応じて、気管は、ゴムチューブ（Broadieとベイト1993a - C）に接続されているガラスピペットを用いて吸引して除去する。胚は今腹中枢神経系、末梢神経、および実験のために露出身体の筋肉組織と表皮下に、カバースリップにフラット添付する必要があります。 </li></ol>

胚の正確なステージングは​​、原因だけで数時間の時間の経過とともに機能的特性の急激な成熟に重要です。いくつかの問題は、このステージングを複雑にする。最初に、ほとんどの研究者は、タイミング卵の段階の胚に産むを使用していますが、産卵時には、別の条件（Broadieら1992）で、動物から動物に非常に異なることがあります。特に、限られた食事療法で女性は前の敷設に長時間?...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
</tr>		</tbody>
		</table>
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harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

ショウジョウバエ胚発生と機能的神経科学の両方の研究のための最高の遺伝的モデルです。伝統的に、これらのフィールドは非常に大部分が独立の歴史と科学的なコミュニティと、互いに分離されています。しかし、これらは通常異なる学問分野間のインターフェイスは、神経回路形成の最終段階、機能化学シナプスの機能的電気信号特性と分化の買収の基礎となる発生プログラムです。このインタフェースは、調査のために非常に重要な領域です。 ショウジョウバエでは、機能開発のこれらのフェーズは、胚発生の最後の3分の1の間（25℃）&lt;8時間の期間中に発生します。この後期発達期間が長くタフな、不浸透性の表皮クチクラの堆積により調査に難治性と考えられた。画期的な進歩は、局所的に後期胚の制御解剖を有効にするためにキューティクルに適用できる水重合外科接着剤のアプリケーションでした。背側縦切開で、胚は実験的な調査に腹神経索と体壁の筋肉組織を露出させる、フラットなレイアウトすることができます。このシステムは、大きく胚性シナプス形成を損なうため、シナプスの接続と機能的なシナプス伝達特性の仕様と差別化を支配する分子メカニズムを明らかに遺伝的変異体を分離し、特徴付けるために使用されています。

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

この手法は、免疫組織化学または電気生理学的記録のためのショウジョウバエ胚の神経筋肉組織を公開します。それは神経筋発達の初期段階でイベントを勉強したり、孵化することができない変異体では電気生理学を実行する際に役立つ。

収穫と準備ショウジョウバエ</em電気生理学的記録とその他の手続のための>胚

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

harvesting-preparing-drosophila-embryos-for-electrophysiological

Research

JoVE Journal

Biology

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K ビュー.  University of Illinois. この手法は、免疫組織化学または電気生理学的記録のためのショウジョウバエ胚の神経筋肉組織を公開します。それは神経筋発達の初期段階でイベントを勉強したり、孵化することができない変異体では電気生理学を実行する際に役立つ。

ビデオ: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

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

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

ショウジョウバエの幼虫のNMJからシナプス電位を記録するための電気生理学的方法

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

の電気生理学的記録ショウジョウバエ胚

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 ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

ショウジョウバエの胚の生体内の遠心

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

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 ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

発現とHEK - 293T細胞における組換え電位依存性イオンチャネルの電気生理学的記録用に最適化されたトランスフェクション戦略

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

キネシンの単離及び精製からショウジョウバエ胚

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

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, ...

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.

収穫と準備ショウジョウバエ胚