我々は、これらのプロトコルの開発に貢献したゲルファントラボのメンバーに感謝。この刊行物で報告された研究は賞の数BFI-2011から295 UDCの下にVIGへの賞の数R01GM052111下の国立衛生研究所の総合医科学研究所がバスク政府教育省、大学や研究によってサポートされていました。

1。 ショウジョウバエ S2細胞<ol><li> ConAをコートしたカバーガラスの作製<ol><li>セラミックラックにカバースリップを置き、1時間クロム酸浸漬することによって、それらを洗ってください。 [注意：クロム酸は腐食性があり、目の刺激、鼻、喉、皮膚を引き起こす可能性があります]。 </li><li>酸が完全に洗い流されるまで、常に30分間のdH 2 Oを実行して徹底的にカバーグラスを洗浄します。 </li><li> 30分間（DH 2 O中0.5 mg / mlの溶液）カバーガラスを空気乾燥したConAでコーティングしてみましょう。 </li><li> 15分間のdH 2 Oでカバーグラスをすすぎ、それらを空気乾燥してみましょう。 ConAをコートしたカバーガラスは、1ヶ月まで保存することができます。 </li></ol></li><li>細胞をプレーティング<ol><li> S2細胞が指数関数的に実験に必要な細胞の数に応じて、T25やT75のCM 2フラスコで増殖させる。 </li><li>血球計数器を用いて細胞密度を計数する。追加の1ml増殖mをConAを被覆カバーガラスで35mmの組織培養皿にedium（例えば昆虫·エクスプレス）、ゆっくり皿に〜1×10 5個の細胞を移す。異なるセルからの処理が重ならないため、この細胞密度は、イメージングのために最適である。 </li><li>すぐに（2.5μMCytoDの最終濃度まで）皿に5μMCytoDと1ミリリットルの媒体を追加めっき後、プロセスの形成を誘導するために。イメージングの前に25℃で少なくとも2時間、細胞をインキュベートすることによって、プロセスの完全な開発を可能にする。 </li></ol></li></ol> 2。 ショウジョウバエ初代ニューロン培養<ol><li>神経細胞培養のための準備<ol><li> ショウジョウバエ神経細胞増殖培地（：; 5μg/ mlのインスリン; 100μg/ mlのペニシリン、100μg/ mlのストレプトマイシン、10μg/ mlのテトラサイクリン30分55ºCで不活性化し20％ウシ胎児血清、熱を補足したシュナイダー培地）を準備。このSUPplementedシュナイダーの培地は、6ヶ月間4℃で保存することができる。 </li><li> ショウジョウバエの胚（22.5グラム寒天、2リットルのフラスコ中の12.0グラムのスクロースと750ミリリットルのH 2 Oを収集するためのバイアル寒天リンゴジュースを準備し、20〜25分間のオートクレーブ、250ミリリットルのリンゴジュースを追加し、各ショウジョウバエに〜10ミリリットルを注ぐバイアル;）凝固後、コットンボールとのバイアルを接続し、食品の乾燥をオーバーを避けるためにプラスチック製の食品ラップでバイアルを包む。寒天バイアル中のりんごジ​​ュースは、4週間のためによく密封されたビニール袋に4℃で保存することができます。 </li><li> ConAを持つコート酸洗浄25ミリメートルの円形カバースリップ（ ショウジョウバエ S2細胞のプロトコルで詳細を参照）、10〜15分間、UV光の下でそれらを殺菌する。これらのカバーガラスをヶ月間保存することができる。 </li></ol></li><li> ショウジョウバエの胚性ニューロンの文化<ol><li>キープヤングアダルト胚Fを収集する前に1〜2日間乾燥酵母を補足したリンゴジュース寒天バイアルに飛ぶ又はニューロン製剤。 </li><li>収集日に、収集された胚を同期させるために、転送大人光の中で1時間のprecollection新鮮なリンゴジュース寒天バイアルへ飛行し、これらの胚を破棄。 </li><li>転送は、1時間の胚を収集し、胚の収率を高めるために暗闇の中で両方、さらに1時間、別のバイアルにハエを転送し、新しいリンゴジュース寒天バイアルに飛ぶ。 </li><li>リンゴジュース寒天バイアルから大人のハエを削除し、胚は、室温（4-6 HR-齢の胚がステージ9月11日の間にある）で段階的に別の4時間9月11日を開発してみましょう。 </li><li>バイアルに水を噴出し、小さな絵筆（少なくとも10胚が一つの良いニューロン準備のために必要とされる）とリンゴジュース寒天から胚を緩めて胚を削除します。 </li><li>水を排水するための移送ピペットを用いて100μmのナイロンメッシュセルストレーナーに胚を移し（ ショウジョウバエ胚洗浄緩衝液、0.4％のNaCl、0.03％トリトンにおけるトランスファーピペットを前すすぎX-100）。 </li><li>小さ ​​な絵筆を使ってストレーナから胚を収集し、1ミリリットルショウジョウバエ胚ウォッシュ（注：ペイントブラシまたはピペットチップを使用して、任意の寒天又は綿繊維をきれい）で満たされた1.5ミリリットルマイクロチューブにそれらを転送します。このプロトコルでは、1.5ミリリットルマイクロチューブは、一致使い捨てペレット乳棒が付属しています。 </li><li> ショウジョウバエ胚ウォッシュ3Xで胚を洗浄します。 </li><li> 10分間、新鮮な1時01市販の漂白剤の溶液および95％エタノールで胚をDechorionate [注意：漂白剤は刺激、目、鼻、喉、皮膚を引き起こす可能性があります]。 </li><li>組織培養フードに移動し、2X補足シュナイダーの培地中で胚を洗浄します。 </li><li>胚をチューブ内に200〜300μlの培地のままにして、ゆっくりと機械的に（10-15Xを挽く）、70％エタノールで滅菌されたキンブルチェース使い捨てペレット乳棒を使用して絨毛膜を除去した胚を解離する。 </li><li> 2分、TRAのために20×gで均質化された混合物を遠心分離新しいチューブに上清をnsfer。 [この手順は省略可能であり、それは胚の破片を除去するのに役立ちます] </li><li>細胞をペレットに2分間550×gで上清を遠心分離する。 </li><li> 2分間550×gで再びシュナイ​​ダー培地とスピンダウン細胞補足500μlにペレットを再懸濁し、一回の再懸濁紡糸サイクルを繰り返す。 </li><li>最終的に約100μlにペレットを再懸濁シュナイダー培地を補充し、滅菌した35ミリメートルペトリ皿中の25ミリメートルのConA-コーティングされたカバーガラス上に細胞をプレート。細胞接着を可能にするために10分間インキュベートします。 </li><li>付着した細胞をカバーガラスを水没から5μmCytoD補充したシュナイダー培地の2ミリリットルを追加します。 </li><li>一晩イメージング前に加湿した25℃のインキュベーター内の皿をインキュベートする。 </li></ol></li></ol>

<ol>
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著者は、彼らが競合する経済的利益を持っていないことを宣言します。

ここでは、 ショウジョウバエ S2細胞および一次神経細胞培養で微小管依存貨物輸送を研究するための詳細なプロトコルを提示する。両方S2プロセスや神経突起は、細胞小器官の輸送のためのトラックとして機能バンドルされた微小管アレイで満たされた構造である。 2つの戦略は、一般的に細胞小器官を標識するために使用される：ベクターのトランスフェクショ...

ショウジョウバエ S2細胞は多くの細胞プロセスを研究するための人気の高まりを得ています。これらの細胞は、もともとOregonR キイロショウジョウバエのトリプシン処理し、後期胚から得られ、1。 ショウジョウバエ S2細胞は他の細胞株に比べて多くの利点を提供するマクロファージ様な機能を維持した。それらは、CO 2なしで25℃で培養し、加熱またはガス交換を必要とせず、室温で長時間撮像することができる。より重要なことには、 ショウジョウバエ S2細胞は、目的のタンパク質の高効率ノックダウンを可能にするRNAi処理に非常に敏感である。 S2細胞は、高度にトランスフェクトされ、安定な細胞株は、目的のタンパク質の安定な異所性発現を可能にするために4週間未満に選択することができる。 S2細胞は、通常はどちらかに成長緩く付着したが、フラスコにまたは懸濁液中に広がっていない。それらは、植物レクチンコンカでプレコートしたカバースリップ上に広げるように誘導することができるnavalin A（ConAを）。スプレッド細胞の周囲は特に薄く、したがって、生細胞顕微鏡検査に最適です。 S2細胞培養のための詳細なプロトコルは、RNAi処理、生光イメージングおよび免疫染色は文献2,3に記載されています。私たちの研究室では、微小管ベースの貨物輸送を研究するためのモデル系としてS2細胞を採用しています。このようなサイトカラシンD（CytoD）として、F-アクチンの脱重合または切断する任意の薬物で処理されたショウジョウバエ S2細胞は、一様に分極微小管4で満たされた長い突起を成長さその微小管の配列に沿って貨物の動きを研究する理想的なシステムにする-10、。これらのプロセスにおける輸送の異なるパラメータ（。 すなわち軌道の長さ、速度、方向など ） を容易に自動化された粒子追跡ソフトウェアを、DiaTrack（Semasopht用いて測定することができます。http://www.microscopy.info/Organization/Details/4423;博士。パスカルロットン：http://www.csiro.au/en/Organisation-Structure/Divisions/Mathematics-Informatics-and-Statistics/PascalVallotton.aspx＃A4）。これらのプロパティは、S2細胞を組織培養細胞内の微小管に沿って貨物輸送を研究するための魅力的なシステムを作る。 S2細胞におけるパイロット実験は、 ショウジョウバエ一次ニューロンにおける貨物輸送研究の生理学的に重要なモデルに拡張することができる。 S2細胞と同様に、解離した胚から培養されたニューロンは、染料及び阻害剤のような小分子に対して透過性であり、細胞小器官輸送の高解像度のライブイメージングは​​、室温でのニューロンで行うことができる。異なる遺伝的背景を持つショウジョウバエを使って、我々はノックアウト（遺伝子の機能喪失変異が）によって一次ニューロンにおける遺伝子機能を調べることができ、ノックダウン（dsRNAを注入または式）または異所性発現（トランスジェニックハエ）。代わりに、彼らはより速く成長し、したがって、私たちは、微小管dependeを学ぶことができ、具体的には、CytoD処理を用いてアクチンフィラメントの断片化は、神経突起伸長を防ぐことはできませんアクチンフィラメント11の影響を受けずにCytoD処理したニューロンでの輸送をオルガネラNT。そのため、一次ニューロン培養は、生理学的に関連するシステム10,12に貨物輸送を研究するのに最適なシステムになり、これを組織培養細胞の利点を組み合わせると遺伝学を飛ぶ。いくつかの家族性の神経変性疾患は、に起因する、または貨物輸送の欠陥（ 例えば 、パーキンソン病13と関連付けることができたので、ハンチントン病14、アルツハイマー病15,16、筋萎縮性側索硬化症/ ALS 17、HMN7Bとペリー症候群18,19、および脊髄小脳失調タイプ5/SCA5 20）、初代神経培養は、微小管依存性輸送にこれらの疾患の原因となる特定の変異の効果の分析に役立ちます。 最後に、微小管依存性輸送の重要な特性はよく、哺乳類やハエの間で保存されているが、 <em&gt;ショウジョウバエ遺伝的および細胞培養物は、正常および病的状態におけるオルガネラ輸送の重要な側面を研究するための培養されたショウジョウバエ細胞の使用を正当化する、より安価であり、時間がかかる。

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			<td height="61" style="height:61px;width:124px;">Powder Insect cell medium (Schneider&#8217;s)</td>
			<td style="width:133px;">Sigma&#160;</td>
			<td style="width:179px;">S9895</td>
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			<td height="41" style="height:41px;width:124px;">Insect &#8211;Xpress medium</td>
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			<td height="21" style="height:21px;width:124px;">Insulin</td>
			<td style="width:133px;">Sigma</td>
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			<td height="61" style="height:61px;width:124px;">Penicillin</td>
			<td style="width:133px;">Research Products International</td>
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			<td height="61" style="height:61px;width:124px;">Streptomycin</td>
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			<td height="41" style="height:41px;width:124px;">Tetracycline hydrochloride</td>
			<td style="width:133px;">Sigma</td>
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			<td height="21" style="height:21px;width:124px;">Concanavalin A</td>
			<td style="width:133px;">Sigma</td>
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			<td height="21" style="height:21px;width:124px;">Cytochalasin D</td>
			<td style="width:133px;">VWR</td>
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			<td style="width:133px;">Molecular Probes</td>
			<td style="width:179px;">L7528</td>
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			<td height="41" style="height:41px;width:124px;">100 &#181;m Cell strainer</td>
			<td style="width:133px;">Fisher Scientific</td>
			<td style="width:179px;">22363549</td>
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			<td height="41" style="height:41px;width:124px;">25 mm Circle cover glass</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">48380 080</td>
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			<td height="41" style="height:41px;width:124px;">Drosophila Vials</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">89092-730</td>
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		<tr height="81">
			<td height="81" style="height:81px;width:124px;">Disposable pellet pestles with matching microtubes</td>
			<td style="width:133px;">Kimble Chase</td>
			<td style="width:179px;">749520-0000</td>
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organelle transport in cultured drosophila cells: s2 cell line and primary neurons.

ショウジョウバエ S2細胞は一様に分極微小管で満たされた任意のアクチン脱重合薬物形の長い分岐していないプロセスの存在下でカバーガラス上にプレー。細胞小器官は、微小管モーターによって、これらのプロセスに沿って移動する。メンテナンスが容易、安定な細胞株を作成するためのRNAiを介したタンパク質ノックダウンと効率的な手順に対する高い感受性は、ショウジョウバエ S2細胞のライブイメージングによる貨物輸送を研究するための理想的なモデルシステムを作る。解離したショウジョウバエ胚由来の初代培養ニューロンにおける軸索輸送：S2細胞で得られた結果は、さらにより生理学的に関連するシステムに適用することができる。培養されたニューロンは、S2のプロセスに非常によく似てバンドルされた微小管を充填し、長い神経突起を成長させる。 S2細胞中と同様に、培養神経細胞における細胞小器官は、初期胚に注入またはトランスジーンのいずれかで発現オルガネラ特異的蛍光色素によって、またはDNAによってコードされる細胞小器官の蛍光マーカーを用いて可視化することができるICハエ。従って、細胞小器官輸送が容易に生体イメージングを用いてガラスカバースリップ上で培養ニューロンにおいて記録することができる。ここでは、 ショウジョウバエ S2細胞および一次ニューロンで貨物輸送を培養し、可視化するための手順について説明します。私たちは、これらのプロトコルは、貨物輸送を研究するラボ用両方のシステムにアクセスできるようにと考えています。

ショウジョウバエ S2細胞は、平らな丸い「パンケーキ」形状（ 図1Aおよび1C）へのConAでコーティングされたカバースリップの広がりに取り付けられている。 S2細胞はCytoDの存在下でカバースリップ上にプレーティングし、2〜3時間拡散させしかし、それらは平行微小管（ 図1Bおよび1D）によって充填細長いプロセスを形成する。これら?...

Watch this Scientific Journal Video about Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons. at JoVE.com

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

ショウジョウバエ S2細胞および培養されたニューロンは、インビボでの電動オルガネラ輸送のイメージングのための優れたシステムである。ここでは、両方の細胞型、それらの画像化および輸送の分析を培養するための詳細なプロトコルを記載している。

培養中の細胞小器官輸送<em&gt;ショウジョウバエ</em&gt;細胞：S2細胞ラインと一次ニューロン。

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

organelle-transport-cultured-drosophila-cells-s2-cell-line-primary

Research

JoVE Journal

Biology

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

13.6K ビュー.  Feinberg School of Medicine, Northwestern University.  ショウジョウバエ S2細胞および培養されたニューロンは、インビボでの電動オルガネラ輸送のイメージングのための優れたシステムである。ここでは、両方の細胞型、それらの画像化および輸送の分析を培養するための詳細なプロトコルを記載している。 

ビデオ: Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

後期ステージショウジョウバエ蛹の脳からの初代神経培養

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

生物学

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

初代培養海馬ニューロンにおける高密度コア小胞のライブイメージング

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

の準備ショウジョウバエ</em光学顕微鏡用> S2細胞

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

培養一次マウス胚線維芽細胞におけるシグナル伝達分子のアデノウイルス媒介遺伝子除去

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

から初代細胞培養ショウジョウバエ原腸胚

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

Isolation and Primary Culture of Rat Hepatic Cells

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology and other related subjects. The method based on two-step collagenase perfusion for isolation of intact hepatocytes was first introduced by Berry and Friend in 1969 1 and, since then, has undergone many modifications. The most commonly used technique was described by Seglenin 1976 2. Essentially, hepatocytes are dissociated from anesthetized adult rats by a non-recirculating collagenase perfusion through the portal vein. The isolated cells are then filtered through a 100 &mu;m pore size mesh nylon filter, and cultured onto plates. After 4-hour culture, the medium is replaced with serum-containing or serum-free medium, e.g. HepatoZYME-SFM, for additional time to culture. These procedures require surgical and sterile culture steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol, which allow consistently in the generation of viable hepatocytes in large numbers.

Primary hepatocytes provide a valuable tool to evaluate biochemical, molecular, and metabolic functions in a physiologically relevant experimental system. We describe a reliable protocol for rat in situ liver perfusion, which consistently generates viable hepatocytes up to 1.0 &times; 108 cells per preparation with cell viability between 88 ~ 96%.

ラット肝細胞の単離及び初代培養

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

イメージングローカルCA<sup&gt; 2+</sup&gt;培養哺乳動物細胞内シグナル

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Generating a &#34;Humanized&#34; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper distribution of the genetic material requires that sister kinetochores on every chromosome become bioriented by attaching to microtubules from opposite spindle poles before progressing into anaphase. However, erroneous, non-bioriented attachment states are common and cellular pathways exist to both detect and correct such attachments during cell division. The process by which improper kinetochore-microtubule interactions are destabilized is referred to as error correction. To study error correction in living cells, incorrect attachments are purposely generated via chemical inhibition of kinesin-5 motor, which leads to monopolar spindle assembly, and the transition from mal-orientation to biorientation is observed following drug washout. The large number of chromosomes in many model tissue culture cell types poses a challenge in observing individual error correction events. Drosophila S2 cells are better subjects for such studies as they possess as few as 4 pairs of chromosomes. However, small molecule kinesin-5 inhibitors are ineffective against Drosophila kinesin-5 (Klp61F). Here we describe how to build a Drosophila cell line that effectively replaces Klp61F with human kinesin-5, which renders the cells sensitive to pharmacological inhibition of the motor and suitable for use in the cell-based error correction assay.

Generating a &quot;Humanized&quot; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

This protocol describes how to generate a Drosophila S2 cell line that is sensitive to small molecule inhibitors of kinesin-5. The use of these cells in a cell-based error correction assay is also outlined.

「ヒト化」を生成<em&gt;ショウジョウバエ</emキネシン5の薬理学的阻害に感受性&gt; S2細胞株

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper ...

Glucose Uptake Measurement and Response to Insulin Stimulation in In Vitro Cultured Human Primary Myotubes

Skeletal muscle is the largest glucose deposit in mammals and largely contributes to glucose homeostasis. Assessment of insulin sensitivity of muscle cells is of major relevance for all studies dedicated to exploring muscle glucose metabolism and characterizing metabolic alterations. In muscle cells, glucose transporter type 4 (GLUT4) proteins translocate to the plasma membrane in response to insulin, thus allowing massive entry of glucose into the cell. The ability of muscle cells to respond to insulin by increasing the rate of glucose uptake is one of the standard readouts to quantify muscle cell sensitivity to insulin. Human primary myotubes are a suitable in vitro model, as the cells maintain many features of the donor phenotype, including insulin sensitivity. This in vitro model is also suitable for the test of any compounds that could impact insulin responsiveness. Measurements of the glucose uptake rate in differentiated myotubes reflect insulin sensitivity.
In this method, human primary muscle cells are cultured in vitro to obtain differentiated myotubes, and glucose uptake rates with and without insulin stimulation are measured. We provide a detailed protocol to quantify passive and active glucose transport rates using radiolabeled [3H] 2-deoxy-D-Glucose ([3H]2dG). Calculation methods are provided to quantify active basal and insulin-stimulated rates, as well as stimulation fold.

Glucose Uptake Measurement and Response to Insulin Stimulation in In Vitro Cultured Human Primary Myotubes

In this method, human primary muscle cells are cultured in vitro to obtain differentiated myotubes and glucose uptake rates are measured. We provide a detailed protocol to quantify rates in basal and insulin-stimulated states using radiolabeled [3H] 2-deoxy-D-Glucose.

インスリン刺激に対するグルコース取り込み測定と応答<em&gt;インビトロ</em&gt;培養ヒト初代筋管

Skeletal muscle is the largest glucose deposit in mammals and largely contributes to glucose homeostasis. Assessment of insulin sensitivity of muscle ...

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

一次培養マウス腸クロム親和性細胞のセル全体を電気生理学

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...