We thank Stephan Uphoff for discussions. 
R.C. was supported by Linacre College, Oxford University. A.P. was supported by the German Academic Exchange Service (DAAD), the German National Academic Foundation and EPSRC. M.S. was supported by the Wellcome Trust. A.N.K. was supported by a UK BBSRC grant (BB/H01795X/1), and a European Research Council Starter grant (261227).

<ol>
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The authors have nothing to disclose.

多くのパラメータは、細胞のエレクトロポレーションおよび対象の生物学的システムと実験（細胞レベルまたは単一分子分析）の正確な性質に応じて、データ取得中に変化させることができる。細菌にDNAをエレクトロポレーションする際に、例えば、標識されたdsDNAフラグメントの0.25〜5ピコモル（事前に光退色を必要とすることなく、 すなわち 、）直接の単一分子の検出を可能にする、低い内在化効率をもたらす。 5 pmolのdsDNAの上方には、細胞は、単一細胞分析のための体制が適し、高負荷になる傾向がある。すべての標識されたDNAはまた、DNAのストック溶液から遊離染料（未反応のフルオロフォア）の痕跡を除去するために、先にゲル精製しなければならない。また、特にsmFRET実験のためのDNAの分解、潜在的な問題は、非天然の核酸、またはそのようなヘアピンループとしてエキソヌクレアーゼアクセス可能な末端を保護するモチーフがDNAを使用することによって対処することができる。 もうadjustablエレクトロポレーションの電子パラメータは、エレクトロポレーションの際に印加される電界強度である。低い電界強度（〜1kVの/ cm）は、単分子研究のための適切な低充填効率につながる。 （1.8 kVの/センチまで）より高い電界強度が積載効率を増加させる。しかしながら、エレクトロポレーション後の電界強度と細胞生存率との間に逆相関がある（ 図4参照）。参考のために、細菌および酵母エレクトロポレーションのために使用される通常の電界強度は〜1.5 kVの/ cmである。時定数は、すぐに任意のアーキング現象がキュベットで起こるように低下​​するので、この減衰の長さを表す時定数は、追従するための便利なパラメータである。通常の設定の下では、時定数は4ミリ秒以上でなければなりません。低い値は、低い積載効率あるいは非ロードされた損傷を受けた細胞につながる。ほとんどのエレ​​クトロポレーターは、両方の曲に変更することができる（例えば、「パルス切り捨て」または「パルス状」のような）他の自​​由度を提供セル·ローディングおよび生存。私たちは、その膜は（単一脂質二重層）は実際それほど複雑ではないので同じような手順では、また、適切なエレクトロの設定を用いて哺乳動物細胞への標識生体分子の内在化を可能にするはずである、細菌や酵母の両方にこの方法を適用し、エレクトロポレーションは、すでにこのような細胞で使用されているため21。 標識されたタンパク質を内在化すると、すべての遊離の色素は、標識されたタンパク質のストック溶液エレクトロポレーションの前に除去する必要がある。遊離色素分子は、それらの小さいサイズに、目的のタンパク質よりも優先的に内在化し、（それらの予想より速い拡散にもかかわらず）は、データ分析の間を区別するのが困難であることができる。有機的に標識されたタンパク質のサンプルのためのガイドは、エレクトロポレーションのために適切であるとして、遊離色素の残存量が2％未満であるべきである22（SDS-PAGEの蛍光スキャニングを用いて検出）。このプロセスは、特に重要であるいくつかの分子は、エレクトロポレーション、細菌や酵母の外膜に付着することがありますように。この点において、陰性対照試料は、（任意の蛍光標識された生体分子と共にインキュベートもエレクトロポレーションされていない細胞、 図2）は、空のセルの自己蛍光のレベルとして、理想的な低エレクトロポレーションした細胞よりも明らかに低い細胞当たりの蛍光強度を表示すべきである。 二本鎖DNAの場合と同様に、標識されたタンパク質の内在化効率は、エレクトロポレーションの前に細胞に加え、生体分子の量にリンクされる。しかしながら、そのような大きさおよび電荷などの他のパラメータは、内在化において役割を果たす。小さ ​​なタンパク質は、（kDaの98まで）より大きなタンパク質に対して、高い内部移行効率を示す成功し、より低い効率（ 図5）26と内部移行することができる。タンパク質の等電点、細胞膜との相互作用の可能性、また、他の物理化学的パラメータエレクトロポレーション時の影響セル·ローディング。その結果、ユーザーは標識タンパク質（&gt;50μM）の高い初期濃度が成功ロードするための最善の機会を与えることを知って、自分のシステムのための実験を​​最適化する必要があります。エレクトロポレーションも細胞（ラベルまたは非標識のいずれか）に、タンパク質および他の生体分子を導入することにより、細胞機能を撹乱し、分析するための新しいツールを提供しています。存在するT7 RNAポリメラーゼの実験（ 図5C）、我々は電気穿孔法を用いてインビボで遺伝子発現を変更することができる生体分子を導入することができ、実験のような例。 単一分子蛍光実験を行う際には、カバースリップの表面（〜100nm）を上記の薄切片内の励磁のみフルオロフォアで最高の信号対雑音比を提供するように、TIR照明は、通常、他の照明モードよりも好まれる。しかし生きた微生物の中に拡散イメージングラベル生体分子が再かもしれない（ 大腸菌用0.8ミクロンまで）帖より深い照明。高い信号対雑音比を維持しながら、より深い照明は、HILOモードで達成される。一方、広視野画像は、ユーザが高いレーザパワー全体ロードされたセルを光退色し、生産ユニタリ強度によって初期細胞の蛍光強度を割ることによって内在化される分子の数を推定された段階的な光退色の分析のために特に重要である単一分子（単一光漂白工程と、 図3）による。それらの軌道がセル全体の体積をカバーする場合でも、広視野イメージングは​​また、関心のある拡散分子を局在化するために、長期的な分子の追跡に必要とされる。 このプロトコルでは、エレクトロポレーション、細胞内で核酸を送達するための生物学者や生化学者のための標準的な技術は、種々の細胞型蛍光生体分子を送達するための簡単​​な方法を構成する方法を提示する。目小説、ハイスループット技術は、それらのネイティブ環境で標識された分子を観察するユニークなツールを提供しています。広い波長範囲をカバーするフルオロフォアで標識された生体分子の付加では、エレクトロポレーションは、非天然のヌクレオチドおよびアミノ酸、金属キレート剤、架橋剤、及びケージング基のような多くの化学基で修飾された分子を送達することができる。対象の生物学的システムは、細胞の発達に必須ではない場合には内在化後に観察されたタンパク質は、細胞内タンパク質プールのすべて（またはほとんど）を表すことを保証する、標的タンパク質をコードする遺伝子はまた、削除することができる（またはノックダウン） 。本質的には、エレクトロポレーションは、「移植」生きている細胞へのin vitroでのバイオコンジュゲートの柔軟性、したがって、合成生物学、システム生物学の取り組み、および生体内の単一分子検出に利益をもたらすことができます。

生きた細胞内部の大部分の蛍光の研究では、GFP 1などの蛍光タンパク質（FPS）、とのタンパク質融合に依存します。これらの蛍光タグは、遺伝子発現または膜輸送2-7のようなプロセスに関与するタンパク質のコピー数、拡散パターンまたは局在化の研究を可能にする。 FPSが高いラベリング特異性、容易な実装を提供し、様々な光物理的および化学的性質1を有する変異体の大規模な在庫でご利用いただけます。しかし、有機蛍光色素分子は、in vitroで彼らの大きな光安定性のための実験（最大のFPよりも安定100倍）8,9、小さいサイズ（最大のFPよりも小さなボリュームを100倍）と分子内ラベリングを容易にするために主要な選択肢のまま（主にシステイン残基の使用による）。これらの要因はすべて、単一分子蛍光のために特に重要であり、FRETが10を検討する 。 いくつかの内部移行方法のCOMBIN有機標識およびin vivo検出に有利で ​​るが過去10年間に導入されている。しかしながら、このような方法は、非天然アミノ酸15の使用を必要とするか、 例えば、（大きな単一膜の真核細胞に限定され、比較的大きなポリペプチドタグ（ 例えば 、TMP、HALO、または20kDaのSNAPタグ）11-14を使用するいずれか。 、ロード、注射器のロード、マイクロインジェクション）16-19をこすり。 このプロトコルは、新規な、 インビボでの観察と結合する有機蛍光体の長所を簡単かつ高スループットの内在化方法を記載している。この技術を開発するために、我々は、Eのように、一般に微生物をロードするために、プラスミドDNA 20,21を用いて細胞を形質転換するために使用されるエレクトロポレーションの手順を適合さcoliまたはS.有機的にラベル生体分子と出芽酵母 。プロトコルは4簡単な手順で構成されています。ラベルの生体分子と細胞のインキュベーション、エレクトロポレーション、細胞の回収、および細胞洗浄は、非内在化生体分子を除去する。ここで、我々は、細胞ベースの単一分子蛍光を研究するために、このエレクトロポレーションプロトコル、ならびに細胞イメージングおよびデータ分析プロセスを提示し、信号をFRET。 エレクトロポレーションは、生体分子（ 図1）20,21細胞に進入できるような一過性の膜孔を形成するために低イオン強度細胞懸濁液を横切って高電圧電場を放電に依存している。ただ、プラスミドDNAと細菌または酵母の形質転換と同様に、細胞は、それらのelectrocompetencyを確実にするために電気穿孔する前に用意しなければならない。この手順は、水で数回の洗浄工程からなる、膜透過性を増加させ、エレクトロポレーションキュベット中でのアーク放電を回避するために、細胞溶液のイオン強度を低下させる。このプロトコルでは、細胞は、（PROTOCOL参照：1.1）を下記のように調製することができるか、商業プロバイダから購入S。 <img alt="図1" src="/files/ftp_upload/52208/52208fig1.jpg" /> 図1：内在化プロトコルの概略図左から右へ：エレクトロコンピテント細胞（二重に標識されたDNA断片と、この例では、細菌）のアリコートに標識された生体分子の数マイクロリットルを加える。氷上で1から10分間インキュベートし、予め冷却したエレクトロポレーションキュベットに移す。エレクトロポレーションし、すぐ後に、細胞に0.5〜1ミリリットル豊富な培地を追加します。細胞が回復させるために37℃（または生物によって必要な温度、 例えば、酵母のための29℃）でインキュベート。余分な非内部化標識分子を除去するための5回の洗浄工程を行う。 PBS緩衝液およびピペットアガロースパッド上に10μlの100〜200μlの最終ペレットを再懸濁。 （広視野モードまたはHILOモード）蛍光顕微鏡での清掃カバーガラスと画像とのパッドをカバーしています。 図1）を洗浄する前に、リカバリを可能にする富栄養培地中でインキュベートする。非内在化標識生体分子の過剰な第1の塩のかなり高い濃度といくつかの界面活性剤を含有する緩衝液中で洗浄することにより除去される（3.3 PROTOCOL参照 ）。塩の存在は、そうでなければ、外膜に付着も非内部化ラベル生体分子によって形成され、非特異的な静電相互作用を破壊。同様に、洗浄緩衝液中の界面活性剤の存在は、非特異的疎水性相互作用を破壊する。 DNAの内在化は、（ 図2）簡単ですがエレクトロポレーションを用いて標識されたタンパク質を内部移行する際、予防措置が取られる必要がある。まず、有機的に標識されたタンパク質のストック·サンプルは、まだ遊離色素の小さな割合が含まれる場合があります。遊離色素分子は、タンパク質よりもはるかに小さいので、優先的に内在化されるかもしれない。観察に内在蛍光分子の大部分は、目的のタンパク質に対応するように、初期のタンパク質試料は、〜2％遊離色素（ 図5）22未満含まれている必要があります。非内在化さ標識タンパク質の過剰もエレクトロポレーション後、外側細胞膜に固執することができます。この現象は、タンパク質特異的であり、それぞれの新しいタンパク質をチェックする必要がある。我々は（：3.3.3プロトコルを参照してください）ロードされた細胞試料からの非内部化タンパク質の除去を可能にするいくつかのオプションを提案する。 最後に、細胞をリン酸緩衝液を少量に再懸濁し、蛍光顕微鏡でその画像化を可能にする、アガロースのパッド上にピペット。アガロースパッド上固定化は、SIMPLですEとそれらの完全性を損​​なうことなく、カバースリップ上の細胞を画像化する効率的な方法。パッドは、低蛍光培地が含まれている必要があります。 細胞イメージングを行うことができるいずれかの広視野、全内部反射蛍光（TIRF）またはHILO（高傾斜及び積層光学シート）を使用して顕微鏡検査。 HILO構成では、レーザビームは、TIRFよりも試料に深く浸透し、まだ大きな信号対雑音比23を可能にする、広視野用として試料全体を点灯しない。使用されるレーザパワーと時間分解能に応じて、内在化生体分子は、（段階的に、光退色分析を用いて、 図3）をカウント局在化、または24-28を追跡することができる。フルオロフォアのFRET対で二重に標識された構築物の内在化は、単一細胞または単一分子レベル（ 図6）でのFRETの両方の定量化を可能にする。 異なるパラメータを変化させることができる所望の出力に応じて、及び生物学的システムを研究。まず、細胞当たりの内在化材料の量は、標識された生体分子の濃度を変えることによって調整することができるエレクトロポレーションの前（ 図2）を細胞に添加した。エレクトロポレーションの電界強度も荷重効率および細胞生存率の両方に影響する。予想されるように、増加する電界強度と積載効率が増加しながら、エレクトロポレーションした細胞の生存率は、（ 図4A）を減少させる。両方のパラメータは、エレクトロポレーションの後にロードされ、分裂細胞​​の割合を記録することによって定量することができる。蛍光イメージングと相まって、この生存率アッセイはまた、 生きた細胞内に内在化生体分子の観察を検証し、数世代（ 図4B）にわたって連続観測を可能にします。 要約すると、このプロトコルは、に蛍光標識DNAおよびタンパク質分子の内在化を可能にする大腸菌またはS. cerevisiaeの 26。有機フルオロフォアで標識された個々の分子は、一桁長い時間スケールのためのFPよりも高い時空間解像度で追跡することができる。最後に、この方法は、広視野、TIRFおよび共焦点検出、ならびにALEX（レーザー励起28,29を交互に）としてパルス励起スキームと互換性がある。

<table border="0" cellpadding="0" cellspacing="0" width="834">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Name of Material/ Equipment</td>
			<td style="width:227px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:321px;">ElectroMax DH5-alpha Comptent cells</td>
			<td style="width:227px;">Invitrogen</td>
			<td style="width:119px;">11319-019</td>
			<td>or any other commercial or lab-mage electrocompetant bacteria or yeast.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">EZ Rich Defined Madia</td>
			<td style="width:227px;">Teknova</td>
			<td style="width:119px;">M2105</td>
			<td>low fluorescence rich media</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MicroPulser Electroporation Apparatus&#160;</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">165-2100</td>
			<td>or any classical electroporator for microorganism transformation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Certified Molecular Biology agarose</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">161-3100</td>
			<td>low fluorescence agarose for agarose pad</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:321px;">Microscope coverslips No 1.5 thickness</td>
			<td style="width:227px;">Menzel</td>
			<td style="width:119px;">BB024060SC</td>
			<td>remove background particles by heating slides in furnace at 500 &#176;C for 1h</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Single-molecule fluorescence microscope</td>
			<td style="width:227px;">Home-built</td>
			<td style="width:119px;"></td>
			<td>described in REFs</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Localization software</td>
			<td style="width:227px;">Custom-written, available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB and C++ software package that can be adapted for localization analysis.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Tracking software</td>
			<td style="width:227px;">Available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB implementation by Blair and Dufresne.&#160;</td>
		</tr>
	</tbody>
</table>

internalization and observation of fluorescent biomolecules in living microorganisms via electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Watch this Scientific Journal Video about Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation at JoVE.com

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

エレクトロポ生きた微生物で内在化および蛍光生体分子の観測

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

internalization-observation-fluorescent-biomolecules-living

Research

JoVE Journal

Biology

16.7K ビュー.  University of Oxford. Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

ビデオ: Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

抗酸菌のエレクトロポレーション

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

生物学

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

蛍光を経由してF -アクチンダイナミクスの生細胞イメージングでは、顕微鏡のスペックル（FSM）

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

初代培養およびコルチのマウス臓器のプラスミドエレクトロ。

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

Imaging Cell Shape Change in Living Drosophila Embryos

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

Imaging Cell Shape Change in Living Drosophila Embryos

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

リビングでのイメージング、細胞の形状の変更ショウジョウバエ胚

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

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

デグロンを使用した生細胞におけるユビキチン·プロテアソーム活性のモニタリング（DGN）·不安定化緑色蛍光タンパク質（GFP）ベースのレポータータンパク質

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific properties, such as binding affinity, agonist or antagonist behavior, or stability, relative to the native sequence. Protein design lies at the center of current advances drug design and discovery. Not only does protein design provide predictions for potentially useful drug targets, but it also enhances our understanding of the protein folding process and protein-protein interactions. Experimental methods such as directed evolution have shown success in protein design. However, such methods are restricted by the limited sequence space that can be searched tractably. In contrast, computational design strategies allow for the screening of a much larger set of sequences covering a wide variety of properties and functionality. We have developed a range of computational de novo protein design methods capable of tackling several important areas of protein design. These include the design of monomeric proteins for increased stability and complexes for increased binding affinity.
To disseminate these methods for broader use we present Protein WISDOM (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>), a tool that provides automated methods for a variety of protein design problems. Structural templates are submitted to initialize the design process. The first stage of design is an optimization sequence selection stage that aims at improving stability through minimization of potential energy in the sequence space. Selected sequences are then run through a fold specificity stage and a binding affinity stage. A rank-ordered list of the sequences for each step of the process, along with relevant designed structures, provides the user with a comprehensive quantitative assessment of the design. Here we provide the details of each design method, as well as several notable experimental successes attained through the use of the methods.

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

We developed computational de novo protein design methods capable of tackling several important areas of protein design. To disseminate these methods we present Protein WISDOM, an online tool for protein design (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>). Starting from a structural template, design of monomeric proteins for increased stability and complexes for increased binding affinity can be performed.

プロテインWISDOM：ベンチ用<em&gt;インシリコで</em&gt;<em&gt;デノボ</em生体分子の&gt;デザイン

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

マウス精巣の生体内マイクロインジェクションやエレクトロポレーションで

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

Tracking Drug-induced Changes in Receptor Post-internalization Trafficking by Colocalizational Analysis

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and overall cell responsiveness to ligands and is, itself, influenced by intra- and extracellular conditions, including ligand-induced signaling. Optimized for use with monolayer-plated cultured cells, but extendable to free-floating tissue slices, this protocol uses immunolabelling and colocalizational analysis to track changes in intracellular receptor trafficking following both chronic/prolonged and acute interventions, including exogenous drug treatment. After drug treatment, cells are double-immunolabelled for the receptor and for markers for the intracellular compartments of interest. Sequential confocal microscopy is then used to capture two-channel photomicrographs of individual cells, which are subjected to computerized colocalizational analysis to yield quantitative colocalization scores. These scores are normalized to permit pooling of independent replicates prior to statistical analysis. Representative photomicrographs may also be processed to generate illustrative figures. Here, we describe a powerful and flexible technique for quantitatively assessing induced receptor trafficking.

Receptor trafficking modulates signaling and cell responsiveness to ligands and is, itself, responsive to cell conditions, including ligand-induced signaling. Here, we describe a powerful and flexible technique for quantitatively assessing drug-induced receptor trafficking using immunolabeling and colocalizational analysis.

Colocalizational解析による受容体後の内在トラフィッキングにおける薬物誘発性の変更の追跡

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

リビングヒト臍帯静脈内皮細胞における高解像度タイムラプスイメージングと微小管ダイナミクスの自動解析

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. To improve the mating efficiency of B. Xylophilus and to increase the chances of mating observation, virgin adults were cultured and used for the investigation. Eggs were obtained by keeping the nematodes in water and allowing the females to lay eggs for 10 min. The second-stage juveniles (J2) were synchronized by incubating the eggs for 24 h at 25 &#176;C in the dark, and the early J4 were obtained by culturing the J2 with grey mold, Botrytis cinerea, for another 52 h. At this time point, most J4 nematodes could be clearly distinguished as being male or female using their genital morphology. The male and female J4 were collected and cultured separately in two different Petri dishes for 24 h to get virgin adult nematodes. A virgin male and a virgin female were paired in a drop of water in the well of a concave slide. The mating behavior was filmed with a video recorder under a stereomicroscope. The whole period of the mating process was 82.8 &#177;3.91 min (mean &#177;SE) and could be divided into 4 different phases: searching, contacting, copulating, and lingering. The mean minutes of duration were 21.8 &#177; 2.0, 28.0 &#177; 1.9, 25.8 &#177; 0.7 and 7.2 &#177; 0.5, respectively. Eleven sub-behaviors were described: cruising, approaching, encountering, touching, hooping, locating, attaching, ejaculating, separating, quiescence, and roaming. Interestingly, obvious intra-sexual competition was observed when one female was grouped with 3 males or one male with 3 females. This protocol is useful and valuable, not only in investigating the mating behavior of B. xylophilus, but also in acting as a reference for ethological studies of other nematodes.

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A protocol for investigating the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, is presented. Behavioral features of B. xylophilus are described in the mating process.

パインウッドセンチュウの交配挙動の観察と定量化、<em&gt;マツノザイセンチュウ</em

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. ...