著者は、SCE micromicropipetteチップの電子顕微鏡画像の未熟なニューロンとデレクダンフィールドの早送り動画キャプチャ成長のためSharminホセに感謝。

機器のセットアップ： <ol><li>この技法のために必要な電気機器は、刺激、オシロスコープ、およびマイクロピペットに挿入される銀のワイヤーを装着したマイクロピペットホルダーです。我々は一般的に軸索の楽器Axoporator 800A刺激を使用して、しかし、そのようなグラスの楽器のような他の一般的な刺激は、SD9刺激は、単一セルのエレクトロポレーションにも適しています。刺激は、内部の導電性溶液の入ったガラスのマイクロピペットの内部に配置される銀線電極とのインタフェースヘッドステージに接続されている。刺激装置から他のリードにもオタマジャクシ外部地面からの入力を受け取るオシロスコープの入力に接続されています。オタマジャクシ外部接地は、単純にエレクトロポレーション中にオタマジャクシと電気緊張接触したままで銀線です。マイクロピペットは、オタマジャクシと接触して置かれると回路が完了です。 </li><li>これが操作する余地を与え、マイクロピペットを約30-45度の角度で持ち込むことを可能にするため、単一細胞のエレクトロポレーションは、良いワーキングディスタンス（少なくとも5センチメートル）と顕微鏡が必要です。 </li></ol>マイクロピペットの作製： 適切なマイクロピペットの準備は、このプロトコルの重要なステップです。難しさは、組織を穿刺するときに容易に中断されませんいずれかの狭い先端（直径1μm以下）のバランスにあります。我々はまた、10度よりのテーパー角と先端が好ましいことを見出した。 <ol><li> 1μm以下の先端径を持つ引っ張らガラスのマイクロピペット、および10度よりも大きいの先端のテーパ角度を準備します。 <ol><li> 0.75ミリメートルの1.5ミリメートル、内径、外径と内部フィラメントとホウケイ酸ガラスは、この目的に適しています。 <ol><li>比較的厚いガラスは、先端部に剛性の程度を提供します。 </li><li>内部フィラメントは、マイクロピペットの先端に向かってバックで満たされた流体を描く上で重要です。 </li></ol></li></ol></li><li>適切なヒントの作製は、しばしば観察された結果に基づいていくつかの変更が必要になります。異なる組織がわずかに異なる形状や先端径を持つ先端が必要な場合があります。 </li></ol>単一細胞のエレクトロポレーションプロトコル： 新しいユーザーの場合、それは遺伝的に符号化された蛍光タンパク質とは異なり、彼らはすぐに落射蛍光で見ることができる、ので、蛍光デキストランの色素で開始することをお勧めします。蛍光色素の使用は、直接組織内にマイクロピペットの先端の位置を確認するために、ユーザーを可能にし、エレクトロポレーションした時点で機器が正しく設定されているかどうかに関して即座にフィードバックを与え、マイクロピペットの先端が適切であるかどうか。 <ol><li>どちらローディングシリンジを使用するか、バックを充填してエレクトロポレーションされる化合物の溶液をマイクロピペットを埋める。 <ol><li>プラスミドDNAを使用している場合は、水で調製した試料はエンドトキシンフリーであることを確認し、1-2μg/mlの間の濃度で。 </li><li>蛍光デキストランの色素の濃度は、経験的に決定されるべきである。私たちはしばしば約満たされたニューロンに水で希釈した蛍光色素を使用してください。 </li><li>通常は、0.5〜1μlののボリュームは十分です。 </li><li>ロードする前に、簡単に目詰まりにつながる可能性のあるマイクロピペットに引き込まれる粒子状破片の量を、減らすために高速でソリューションを遠心する。 </li><li>先端内の気泡は、多くの場合、現在の流れに影響を及ぼし、精力的にステージに取り付ける前にマイクロピペットをフリックすることで外れている可能があります。 </li></ol></li><li>ホルダーにマイクロピペットを挿入するか、ステージ3軸マニピュレータに取り付け。銀線電極をマイクロピペットに挿入されているか確認し、内部ソリューションとの電気緊張接触している。 </li><li>オタマジャクシ飼育培地（Steinbergの溶液、pH 7.4）で作成したMS - 222（3 - アミノ安息香酸エチルエステル）0.02％に浸漬してオタマジャクシを麻酔。 <ol><li>我々は通常、我々の実験ではステージ44から48アルビノアフリカツメガエルのオタマジャクシを使用してください。 </li><li>オタマジャクシは約5分後に通常完全に麻酔する。 </li><li>いくつかのオタマジャクシは、待機時間を削減すると同時に麻酔かもしれない、しかし、1時間を越える期間のMS - 222への暴露を避ける。 </li></ol></li><li>プラスチック製のトランスファーピペットを用いてエレクトロポレーションチャンバーに単一の麻酔オタマジャクシを転送します。 <ol><li>エレクトロポレーションチャンバーは、容易に®小さなSylgardでおたまじゃくしの形をした空洞を彫刻シリコンブロック、そしてそれはチャンバー内のオタマジャクシで電気緊張接触になるように空洞に銀のアース線を挿入することにより、社内で行うことができます。 </li></ol></li><li>湿らせた絵筆を使って位置オタマジャクシ背側を。 </li><li>それは、皮膚との接触に関心のある領域に直接隣接するまで、マイクロピペットを下ろします。 <ol><li>密集した細胞体との領域を標的にすることは大いに成功したエレクトロポレーションの可能性が増加します。我々は通常、オタマジャクシ視蓋をターゲットにしています。 </li><li> 30から45度の角度でのマイクロピペットをもたらす。急な角度はしばしば一つの視界を妨げないしながら浅く軌跡は、皮膚の穿刺をより困難なものにする。 </li></ol></li><li>続けマイクロピペットの低下は、最初に皮膚をディンプルし、基になる組織に通過します。 <ol><li>表層細胞は、in vivoイメージングのために好ましく、マイクロピペットの先端が皮膚のえくぼ形成によって徐々に低下していることを確認することでターゲットとすることができます。 </li></ol></li><li>一つは、常に一度に組織内に配置されたマイクロピペットの抵抗を監視する必要があります。 Axoporator 800A（アクソンインスツルメンツ）を使用している場合は、約10 -40mΩのの抵抗が最適です。抵抗は、先端の直径は、単一細胞電気穿孔法に適しているかどうかの良い指標です。 <ol><li>高マイクロピペットの抵抗（&gt;100MΩ）が狭すぎるのヒントが詰まってヒントまたはマイクロピペットを示すものである。 </li><li>低マイクロピペットの抵抗（&lt;10MΩ）は多くの場合、壊れた先端の結果が大きすぎる先端径の指標となる。 </li><li>直接抵抗測定を提供していない刺激を使用している場合は、マイクロピペットの抵抗を間接的にオシロスコープ（後述）によって測定される電圧パルスの振幅を観察することによって測定することができる。 </li></ol></li><li>電圧列車を適用します。 <ol><li>プラスミドDNAの場合、パルスは、訓練期間が1msの負の方形波パルスで構成される最も有効であることが判明し、500ミリ秒のパルス列の持続時間で300Hzので配信。 <ol><li>パルス電圧をオシロスコープで測定したパルスの振幅が0.75と1.5μAの間にあることが経験的にそのように決定されます。 </li></ol></li><li>蛍光デキストランの染料の場合は、300μsの期間の正の方形波パルスは10msのパルストレインの期間で300Hzの周波数で供給されています。 <ol><li> DNAのためのプロトコルと同様に、パルス電圧をオシロスコープで測定したパルスの振幅が0.75と1.5μAの間にあることが経験的にそのように決定されます。 </li><li>刺激のパラメータに追加の調整は、落射蛍光下でエレクトロポレーションの結果を観察に基づいて行うことができる。 <ol><li>エレクトロポレーションした細胞が暗く表示される場合、細胞が明るく表示されるまで、パルスパラメータは徐々に大きくする必要があります。 </li><li>細胞のクラスターラベルが付けられている場合、単一の細胞が標識されるまで、その一方で、、パルスパラメータを減らす必要があります。 </li></ol></li></ol></li></ol></li><li>マイクロピペットを撤回、組織内の別のサイトに再挿入、および電圧列車を適用する。 <ol><li>歩留まりを向上させるために、我々は一般的に標識された細胞が重複しないことを保証するために十分な間隔でオタマジャクシ視蓋の各半球で複数のサイトをエレクトロ。 </li></ol></li><li> Steinbergの飼育剤の容器中にエレクトロオタマジャクシを転送する。 <ol><li>オタマジャクシは、急速に数分以内に麻酔から回復します。 </li></ol></li><li>同じマイクロピペットは、多数のオタマジャクシに使用することができます。常にオシロスコープで測定したパルスの振幅を観測することが重要です。 <ol><li>大幅にパルス強度を増加した後、パルス振幅が低いままなら、これは可能性が一気に多くのオタマジャクシをelectroporatingする際によく見られる目詰まりしている先端の結果です。 <ol><li>マイナーチップ詰まりを取り除くために、方法は、組織にマイクロピペットを挿入した後、単一の正のパルスを適用することです。 </li><li>詰まりが外れ、または頻繁に再発を詰まらせることができない場合は、マイクロピペットを交換してください。 </li></ol></li><li>パルスの振幅が大幅にパルスパラメータを減少した後非常に大きいままの場合、これはチップが壊れ、交換が必要になっているサインです。 </li></ol></li></ol>成功したエレクトロポレーションした細胞のスクリーニング： <ol><li>エレクトロポレーション後、オタマジャクシは直立落射蛍光顕微鏡を用いて標識された細胞についてスクリーニングされる。 </li><li>蛍光色素の場合には、オタマジャクシは30分ほど後にエレクトロポレーション後にスクリーニングすることができる。 <ol><li>我々は、より長い間隔が低く、バックグラウンド蛍光レベルに関連付けられていること、しかし、発見した。 </li></ol></li><li>遺伝的に符号化された蛍光タンパク質については、スクリーニングは通常、少なくとも12時間後にエレクトロポレーションを実施しています。 <ol><li>蛍光タンパク質を発現する細胞が時間とともに明るく取得していく、そのため、薄暗い細胞は12-24時間のさらに他のインターバル後に再スクリーニングすることができる。 </li></ol></li><li>上記のようなオタマジャクシはSylgard ®チャンバー内に設置し、封入、麻酔する。個々のオタマジャクシは、落射蛍光で観察されています。 <ol><li>落射蛍光光への過度の曝露は標識細胞を殺すことができる光毒性になることに注意して下さい。したがって、細胞を蛍光灯にさらされている時間の量を最小限に抑える。 </li></ol></li><li> Steinbergの飼育のソリューションを使ってコンテナにオタマジャクシを返します。 </li></ol>

<ol>
	<li>Bestman, J. E., Ewald, R. C., Chiu, S. L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+single-cell+electroporation+for+transfer+of+DNA+and+macromolecules.">In vivo single-cell electroporation for transfer of DNA and macromolecules.</a> Nature Protocols. 1, 1267-1272 (2006).</li><li>Dunfield, D., Haas, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+electroporation.">Single cell electroporation.</a> Encyclopedia of Neuroscience (4th Ed.). , .</li><li>Haas, K., Jensen, K., Sin, W. C., Foa, L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+electroporation+in+Xenopus+tadpoles+in+vivo--from+single+cells+to+the+entire+brain.">Targeted electroporation in Xenopus tadpoles in vivo--from single cells to the entire brain.</a> Differentiation. 70, 148-154 (2002).</li><li>Haas, K., Sin, W. C., Javaherian, A., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+electroporation+for+gene+transfer+in+vivo.">Single-cell electroporation for gene transfer in vivo.</a> Neuron. 29, 583-591 (2001).</li></ol>

単一細胞電気穿孔（SCE）は、遺伝子機能を決定し、標的遺伝子操作を実行するための強力なツールです。アルビノおたまじゃくしの透明性と脳のアクセシビリティは、理想的に生きて生体内で神経細胞の成長と細胞内イベントの可視化のための最適なこのモデルシステムを作る。 SCEは、単一のニューロンの成長の可視化を可能にし、そうでなければ変更されずに脳内で細胞自律的な操作を実行する。このビ...

single cell electroporation in vivo within the intact developing brain

単一細胞電気穿孔（SCE）は、in vivoでの製剤を含む無傷の組織内の個々の細胞へのDNAや他の高分子の配信を可能にする特殊な技術です。この手法の明確な利点は、グローバルな治療から生じるものから、周囲の組織変更されず、それによって特徴的な細胞自律的な効果を残したまま、実験操作が個々のセルで実行される可能性があるということです。 in vivoイメージング技術の高度と組み合わせると、蛍光マーカーのSCEは、細胞形態、細胞増殖、および秒から数日に及ぶ時間スケール上の細胞内イベントの直接可視化を可能にします。この手法は、生体内と生体外での準備での様々な使用されているが、我々はアフリカツメガエルのオタマジャクシで使用するためにこの手法を最適化してきました。このビデオの記事で、我々は詳細アルビノアフリカツメガエルオタマジャクシの無傷の脳内の神経細胞への蛍光色素またはプラスミドDNAのSCEのための手順を。我々はまた、歩留まりを最適化する方法を議論し、蛍光SCEによって標識されたニューロンのライブ二光子蛍光イメージングの例を示します。

Watch this Scientific Journal Video about Single Cell Electroporation in vivo within the Intact Developing Brain at JoVE.com

Single Cell Electroporation in vivo within the Intact Developing Brain

単一細胞電気穿孔（SCE）は、in vivoでの製剤を含む無傷の組織内の個々の細胞へのDNAや他の高分子の配信を可能にする特殊な技術です。アフリカツメガエルオタマジャクシの無傷の脳内の神経細胞への蛍光色素またはプラスミドDNAのSCEのためここでは、詳細手順を。

無傷の開発脳内の生体内の単一細胞のエレクトロポレーション

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact ...

single-cell-electroporation-in-vivo-within-the-intact-developing-brain

Research

JoVE Journal

Biology

13.2K ビュー.  University of British Columbia - UBC. 単一細胞電気穿孔（SCE）は、in vivoでの製剤を含む無傷の組織内の個々の細胞へのDNAや他の高分子の配信を可能にする特殊な技術です。アフリカツメガエルオタマジャクシの無傷の脳内の神経細胞への蛍光色素またはプラスミドDNAのSCEのためここでは、詳細手順を。

ビデオ: Single Cell Electroporation in vivo within the Intact Developing Brain

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

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell differentiation, axon guidance, retinotopic organization and synapse formation[1]. One drawback is the inability to efficiently and reliably manipulate gene expression in RGCs in vivo, especially in the otherwise accessible murine visual pathways. Transgenic mice can be used to manipulate gene expression, but this approach is often expensive, time consuming, and can produce unwanted side effects. In chick, in ovo electroporation is used to manipulate gene expression in RGCs for examining retina and RGC development. Although similar electroporation techniques have been developed in neonatal mouse pups[2], adult rats[3], and embryonic murine retinae in vitro[4], none of these strategies allow full characterization of RGC development and axon projections in vivo. To this end, we have developed two applications of electroporation, one in utero and the other ex vivo, to specifically target embryonic murine RGCs[5, 6]. 
With in utero retinal electroporation, we can misexpress or downregulate specific genes in RGCs and follow their axon projections through the visual pathways in vivo, allowing examination of guidance decisions at intermediate targets, such as the optic chiasm, or at target regions, such as the lateral geniculate nucleus. Perturbing gene expression in a subset of RGCs in an otherwise wild-type background facilitates an understanding of gene function throughout the retinal pathway. Additionally, we have developed a companion technique for analyzing RGC axon growth in vitro. We electroporate embryonic heads ex vivo, collect and incubate the whole retina, then prepare explants from these retinae several days later. Retinal explants can be used in a variety of in vitro assays in order to examine the response of electroporated RGC axons to guidance cues or other factors. In sum, this set of techniques enhances our ability to misexpress or downregulate genes in RGCs and should greatly aid studies examining RGC development and axon projections.

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Here we present two techniques for manipulating gene expression in murine retinal ganglion cells (RGCs) by in utero and ex vivo electroporation. These techniques enable one to examine how alterations in gene expression affect RGC development, axon guidance, and functional properties.

マウス網膜神経節細胞における遺伝子発現のための子宮内および生体外でのエレクトロポレーションで

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell ...

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

使用して哺乳類の骨格筋のDNAトランスフェクションインビボでのエレクトロポレーション

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

マウスの錐体光受容体からの単一細胞サクション録音

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

サブセルラー解像度で手つかずのショウジョウバエの幼虫の in vivoイメージングで

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

アダルトゼブラフィッシュ網膜にモルフォの in vivoエレクトロポレーションに

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

リジェネアダルトゼブラフィッシュテールフィンにMorpholinos の in vivoエレクトロポレーションで

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

シングルミトコンドリアスーパーオキシド無傷ハートや点滅の共焦点イメージング<em&gt;インビボ</em

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

無傷骨格筋線維内のカルシウムスパークスの評価

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

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 ...