この作品は、難聴やその他のコミュニケーション障害に関する国立研究所（NIDCD）学内プログラムによってサポートされていました。

1。ニューロンのトランスフェクション<ol><li>ポリ- D -リジンコートマテックで文化胎生18日（E18）ラット海馬ニューロンは、35 mmのガラスボトムディッシュ1。 in vitroでの上で16〜18日（DIV）、クロンテックCalPhos哺乳類のトランスフェクションキットを用いてトランスフェニューロン。最初に、1.5 mlの培養液を交換して<a href="http://www.invitrogen.com/site/us/en/home/Products-and-Services/Applications/Cell-Culture/Mammalian-Cell-Culture/Classical_Media/dmem.html">ダルベッコ改変イーグル培地</a>トランスフェクション前に35 - mmディッシュ0.5時間ごとに（DMEM）。後で（ステップ1.6）を使用するための滅菌15 mlのチューブに元の培養培地を保存します。 </li><li>滅菌H 2 O（Clontech）および100μlの総体積は12.4μlの2 Mのカルシウム溶液（Clontech社）と10μgのpEGFPを- N1プラスミドDNAを混ぜる。 </li><li>ステップ1.2）から100μlの2 ×ボルテックス2時間HBS滴下× HBS中程度の速度で混合物を追加。 </li><li>混合物を室温で20分間座って、次にDMEMインキュベートしたニューロンにステップ1.3）から、最終的な混合物を追加してみましょう。 </li><li> 1〜1.5時間37℃のインキュベーターに戻しニューロンを置く。 </li><li>リン酸カルシウムを含有する培地を除去し、DMEMで3回細胞を洗浄。オリジナルの培地と交換、インキュベーターにDMEM培地を培養皿を返す前に。 </li></ol> 2。脊椎に関するFRAP実験<ol><li>ニューロンは、2〜4日トランスフェクション後FRAP実験に使用されます。 </li><li>すぐに（mMで）塩化ナトリウム145、KClの5、HEPES 10、グルコース10、グリシン0.005、CaCl 2の 2.6、およびMgClを含む予め温めておいたタイロード溶液を、追加することで、35mmのガラスボトムディッシュから培養液を交換してください2 1.3（NaOHで7.4に調整pHを）。 </li><li>ツァイスLSM 710共焦点顕微鏡は、FRAP実験に使用されます。ツァイスのTempModuleシステムは、温度（37℃）、湿度及び作業システムのCO 2（5％）を制御するために使用されます。 CO 2タンクが接続されていると湿度のバランスをとるために使用される水のボトルは、水で満たされていることを確認してください。 </li><li> 100 ×目標（αplan-アポクロマート100 × / 1.46オイル）でトランスフェクトされた成熟した樹状突起を見つける。の場合皿のトランスフェクションされた細胞は、その後、40 ×対物レンズ（プラン- NEOFLUAR 40 × / 1.3オイル）、および100に切り替え×画像をキャプチャする目的で、目的のセルを検索してスパースです。 </li><li> 5 ×画像へ光学ズームと256 × 256ピクセルの解像度のいくつかの棘によるデンドライトの短い断片を使用してください。画像をキャプチャするには、0.5秒のスキャンを終了するまでの公称速度9（ピクセルは、時間3.15マイクロ秒の滞留）を使用します。ピンホールは、強い蛍光を取得するために2μmのために設定されています。画像を撮影するとき、画像全体を光退色を避けるために、例1-5％を、低レーザ伝送を使用するようにしてください。 </li><li>興味の背骨を選択します。私たちの実験では、我々は〜1μmの彼らの背骨の頭の直径とキノコ棘を選択してください。 </li><li> FRAP実験を行うには、公称100％のレーザー伝送に関心の漂白剤脊椎その後、漂白前に10回を5コントロールの画像を取る[アルゴンレーザーのパワーは488レーザーラインの50％（15 MW）で、30mWです、と約3-4 mWのは488レーザーライン]を介して顕微鏡に達し、その後すぐに、漂白後の一連の画像をキャプチャ。この実験では、画像を漂白した後、15秒間1秒ごとにキャプチャされます。時間の間隔は、異なる標的タンパク質と異なる実験デザイン（ 図1）にしたがって調整する必要があります。 </li><li>画像を保存します。 </li></ol> 3。データの分析<ol><li> ImageJのソフトウェアで画像を開きます。 </li><li>興味の背骨がフロートしないようにImageJの中の - （→→&quot;翻訳&quot;および/または&quot;剛体&quot;&quot;スタック内のスライスを整列&#39;プラグイン&quot;→&quot;シャッフルスタック&#39;）は、位置合わせツールを使って画像のスタックの位置を合わせます、他の言葉で - それは、画像上の同じ位置に留まります。 </li><li> ImageJの&quot;強度Vタイムモニター&quot;ツール（&quot;プラグインを使用して、関心の背骨（F S）、トランスフェクトが無漂白地域（対照）、および時間経過の画像における背景領域（F B）の相対的蛍光強度を測定&quot;→&quot;スタック - シャッフリング&quot;→&quot;輝度Vタイムモニター&quot;）。制御領域は十分に焦点を当てた遠位樹状突起の一部である可能性があります。背景領域は、非蛍光の領域です。 </li><li>前（F C0）と（F C）光退色後の制御領域の蛍光を比較することにより、退色率（r）を計算する。 R = F C / F c0を 。 </li><li>次のようにターゲットの背骨の蛍光強度（F）を正規化： F =（F S - F B）/ R. </li><li>曲線は、一相指数関数グラフパッドプリズムソフトウェアの式（ 図2）またはパーソナルプラグインを使用してターゲット背骨の蛍光強度に適合R同様のソフトウェア。 </li><li>次の式でモバイル割合（F M）と不動の割合を（F i）を計算してください。 F M = F∞/ F 0、 ここで、F∞完全復旧後の蛍光強度であり、F 0は、退色前の蛍光強度である。 は f i = 1 - fをmと </li></ol> 4。代表的な結果： 本研究では、我々は成熟した海馬神経細胞でFRAP実験を行う。 18から22 DIVで、キノコの棘が既に形成されています。私たちの方法を使用して、このような背骨のような小さな領域での蛍光強度の動的な変更は、記録することができます。 EGFPの蛍光回復のプロセスを分析するために、我々は、漂白してから、すぐに15秒のために漂白後の1秒ごとに1つのイメージの前のコントロールなどの5枚を取る。画像の解像度は、定量分析のために十分です。タグ付き蛍光タンパク質の蛍光回復のプロファイルは、再現性の高いです。 我々はまた、簡単にImageJとグラフパッドプリズムソフトウェアを使用して、蛍光タンパク質の携帯電話と不動の分数を定義する方法を示しています。我々がここに表示FRAP法及び解析は、大きく分けて神経科学、細胞生物学と他の研究で使用することができます。 <img src="/files/ftp_upload/2568/2568fig1.jpg" alt="図1" /> 図1培養海馬ニューロンから脊椎におけるEGFP蛍光のFRAP測定。赤い矢印は退色の時間を示している。写真は光退色後の前（プレ）と0（ゼロ）の同じ面積、1、5、10、15秒を表します。脊椎の領域、制御および背景はそれぞれ、文字S、CとBとマークされます。ニューロンは、実験中に37℃に維持した。スケールバーは1μm。 <img src="/files/ftp_upload/2568/2568fig2.jpg" alt="図2" /> 図2 15秒の期間にわたってEGFP蛍光のFRAP曲線。緑の線は元の曲線を示し、赤い線は、正規化曲線を示しています。曲線上の点は、FRAPごとに1秒を示しています。曲線は、一相指数関数式でフィッティングした。退色前の平均蛍光を100％としてカウントした。この実験では、モバイル割合（MF）は94％と不動の割合は、（IF）6％です。

<ol>
	<li>Zheng, C. Y., Petralia, R. S., Wang, Y. X., Kachar, B., Wenthold, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SAP102+is+a+highly+mobile+MAGUK+in+spines.">SAP102 is a highly mobile MAGUK in spines.</a> J Neurosci. 30, 4757-4766 (2010).</li><li>Grati, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+turnover+of+stereocilia+membrane+proteins:+evidence+from+the+trafficking+and+mobility+of+plasma+membrane+Ca(2+)-ATPase+2.">Rapid turnover of stereocilia membrane proteins: evidence from the trafficking and mobility of plasma membrane Ca(2+)-ATPase 2.</a> J Neurosci. 26, 6386-6395 (2006).</li><li>Liu, L. N., Aartsma, T. J., Thomas, J. C., Zhou, B. C., Zhang, Y. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FRAP+Analysis+on+Red+Alga+Reveals+the+Fluorescence+Recovery+Is+Ascribed+to+Intrinsic+Photoprocesses+of+Phycobilisomes+than+Large-Scale+Diffusion.">FRAP Analysis on Red Alga Reveals the Fluorescence Recovery Is Ascribed to Intrinsic Photoprocesses of Phycobilisomes than Large-Scale Diffusion.</a> PLoS ONE. 4, e5295-e5295 (2009).</li><li>Wiedenmann, J., Nienhaus, G. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+imaging+with+EosFP+and+other+photoactivatable+marker+proteins+of+the+GFP+family.">Live-cell imaging with EosFP and other photoactivatable marker proteins of the GFP family.</a> Expert Rev Proteomics. 3, 361-374 (2006).</li><li>Shaner, N. C., Patterson, G. H., Davidson, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+fluorescent+protein+technology.">Advances in fluorescent protein technology.</a> J Cell Sci. 120, 4247-4260 (2007).</li><li>Sprague, B. L., McNally, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FRAP+analysis+of+binding:+proper+and+fitting.">FRAP analysis of binding: proper and fitting.</a> Trends Cell Biol. 15, 84-91 (2005).</li></ol>

利害の衝突は宣言されません。

FRAP分析は広くin vivoおよび in vitro 1月2日の研究で使用されています。この手法は、一般的にGFPを利用して<a href="http://en.wikipedia.org/wiki/Fusion_protein" title="融合タンパク質">融合タンパク質を</a> 、それはまた紅藻の融合タンパク質3を使用することもできますが、。この分析では、小文字を区別し、GFPタグ融合タンパク質の流動性を特徴付けるために使用することができます。 意味のあるFRAP解析を生成するには、それは前とFRAP実験時の不要な光退色を避けることが重要です。これを達成するために二つの方法があります。最初に、検索と実験的なニューロンを観察するプロセスは、高速でなければなりません。特に、長い時間の100倍の目標を持つ神経細胞の観察が著しく蛍光を漂白剤。第二に、高レーザパワーと頻繁にスキャンが頻繁に光退色の可能性を高める。従って、それは制御領域の光退色率を計算し、実験地域のFRAP曲線を正規化するために必要となります。正規化の方法は、（詳細については、ステップ3.3から3.5を参照）のプロトコルに記載されている。 退色のステップはまた、良好なFRAPの結果を保証するために重要です。この実験では、我々は、漂白剤100％のレーザー伝送における利益の10倍の背骨を。この条件では、漂白剤の固定の準備のバックグラウンドレベルまで背骨の蛍光に十分です。従って、我々は、光退色後0秒で背景と同じ数に蛍光強度を設定します。最初のスキャンの速度に応じて、蛍光回復のかなりの量は、すでに目的のタンパク質は、モバイル性の高いときに検出されることがあります。 多くの蛍光タンパク質プローブは、FRAPと蛋白質のダイナミクスを研究するために開発されている。補完的なアプローチは、光活性化GFP（PA - GFP）、またはphotoconvertibleバリアントを使用するために、例えば、です。一緒にFRAP技術と、これらのツールは、ライブセルイメージング研究4-6不可欠になりつつあります。

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<td>35 mm glass-bottom dishes </td>
<td>&nbsp;</td>
<td>MatTek</td>
<td>P35GC-1.0-14-C</td>
<td>&nbsp;</td>
<tr>
<td>CalPhos Mammalian Transfection Kit</td>
<td>&nbsp;</td>
<td>Clontech</td>
<td>631312</td>
<td>&nbsp;</td>
<tr>
<td>pEGFP-N1 plasmid</td>
<td>&nbsp;</td>
<td>Clontech</td>
<td>6085-1</td>
<td>&nbsp;</td>
<tr>
<td>Zeiss confocal microscope</td>
<td>&nbsp;</td>
<td>Zeiss</td>
<td>LSM 710</td>
<td>&nbsp;</td>
<tr>
<td>Zeiss TempModule system</td>
<td>&nbsp;</td>
<td>Zeiss</td>
<td>N.A.</td>
<td>&nbsp;</td>
<tr>
<td>ImageJ software</td>
<td>&nbsp;</td>
<td>NIH</td>
<td>N.A.</td>
<td>&nbsp;</td>
<tr>
<td>Graphpad Prism software</td>
<td>&nbsp;</td>
<td>Graphpad software</td>
<td>N.A.</td>
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fluorescence recovery after photobleaching (frap) of fluorescence tagged proteins in dendritic spines of cultured hippocampal neurons

FRAPは、GFPタグ融合タンパク質の流動性を定量化するために使用されています。強力な励起レーザーを用いて、GFP -タグ付きタンパク質の蛍光は、関心領域で漂白されている。地域の外から無漂白のGFP -タグタンパク質は、関心領域に拡散する際の領域の蛍光が回復します。タンパク質の移動度は、その後、蛍光回収率を測定することによって分析されます。この手法は、タンパク質の移動度と回転率を特徴付けるために使用することができる。 本研究では、我々は培養海馬ニューロンの（強化された緑色蛍光タンパク質）EGFPのベクトルを表現する。ツァイス710共焦点顕微鏡を用いて、我々は、単一スパインのGFPタンパク質の蛍光信号を光退色し、光退色後蛍光回復を記録するタイムラプス画像を撮影する。最後に、我々はImageJとグラフパッドソフトを使用して画像データを解析することにより、棘におけるGFPの携帯電話と不動画分の割合を推定する。 このFRAPプロトコルは、基本FRAP実験を実施するだけでなく、データを分析する方法を方法を示しています。

Watch this Scientific Journal Video about Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons at JoVE.com

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAPは、緑色蛍光タンパク質（GFP）タグ付き培養細胞中のタンパク質の移動度を定量化するために使用されています。我々は、光退色後蛍光回復の割合を分析することにより、GFPのモバイル/不動の分画を検討した。本研究では、FRAPは、海馬ニューロンのスパインで行われました。

培養海馬ニューロンの樹状突起棘の蛍光タグ融合タンパク質の光退色後蛍光回復（FRAP）

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

fluorescence-recovery-after-photobleaching-frap-fluorescence-tagged

Research

JoVE Journal

Neuroscience

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

54.1K ビュー.  National Institutes of Health, Bethesda. FRAPは、緑色蛍光タンパク質（GFP）タグ付き培養細胞中のタンパク質の移動度を定量化するために使用されています。我々は、光退色後蛍光回復の割合を分析することにより、GFPのモバイル/不動の分画を検討した。本研究では、FRAPは、海馬ニューロンのスパインで行われました。

ビデオ: Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

培養中枢神経系ニューロンの樹状突起スパイン形態の解析

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

神経科学

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

培養交感神経ニューロンの樹状成長を誘導する

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

培養神経細胞における膜タンパク質の側方拡散とエキソサイトーシスは、蛍光回復と蛍光損失光退色を用いて評価

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

軸索、樹状突起および脳スライスにおける個々のニューロンの樹状突起棘から電位感受性色素記録

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

イメージング初代培養マウス神経細胞における形質膜に受容体の挿入をpHluorinタグ付き

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

免疫沈降およびアシル-ビオチン所（阿部）による培養海馬神経細胞におけるタンパク質パルミトイル化の検出

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

ライブ培養神経細胞の抗体供給した後細胞表面および内在化タンパク質の異なる標識化

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

構造化照明顕微鏡を用いてラット初代海馬ニューロンの樹状突起棘を画像化

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

細胞内のCaの蛍光と生物発光イメージング<sup&gt; 2+</sup&gt;高齢者における海馬ニューロン

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.