我々は固定プロトコル、試薬や励ましを共有するための証明の原理実験、リチャード·フェッター用のPalm顕微鏡へのアクセスのためにハラルドヘスとBetzigエリックに感謝します。我々は、マイケルソン、ジェラルディンSeydoux、ステファンアイマー、ルドルフロイベ、キースNehrke、クリスチャンFrøkjr·ジェンセン、DNA構築のためのオードADA-ヌグエマとMarc Hammarlundに感謝します。我々はまた、ツァイスPAL-M、ツァイスELYRA P.1やし顕微鏡のベータ版へのアクセスを提供するためにカールツァイス社に感謝します。

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この記事への生産とフリーアクセスはカールツァイス社が主催している

ここでは、プラスチック製で、電子顕微鏡を用いた超微細構造をセクションで蛍光タンパク質をローカライズし、画像の蛍光タンパク質を保持する方法について説明します。タンパク質は、ナノメートル分解能にPALM顕微鏡を用いた回折限界以下局在していた。特定の標本にこのプロトコルを適応させるために、次のパラメータを考慮する必要があります。フルオロフォア、定量化、および配置。 蛍光タンパク質や有機蛍光色素の選択は、アプリケーションとモデルシステムによって異なります。我々は、EGFP、YFP、シトリン、mEosFP、mEos2、tdEos、mOrange、PA-mCherry、とdendronの複数形12を含む蛍光タンパク質の様々なテストをしている。各蛍光色素からの蛍光の保全は、すべての蛍光タンパク質が記載された方法を用いて保存することができることを示唆して、類似していた。それはCでうまく表現ので、我々はtdEosを選んだelegansは 、タンパク質はtdEosに融合したときに機能的に推移し、そのため光活性化特性は、ヤシの顕微鏡検査に最も適していた。しかし、tdEosの凝集したか失敗した式が、時折12観察されている。 アプリケーションに応じて、異なる蛍光団はより適しているかもしれません。多くの場合、それは光活性化蛍光性タンパク質を使用する必要はありません。単純な相関蛍光電子顕微鏡では、光活性化蛍光タンパク質を必要としません。 GFPまたは有機色素は、回折限界上記のセクションでタグ付けされたタンパク質からの蛍光画像に使用することができます。例えば、人はイメージ、蛍光顕微鏡を用いて神経網の軸索とイメージングによる電子顕微鏡写真で特定の軸索蛍光顕微鏡で蛍光を蛍光シグナルを関連付けることができます。このような誘導放出の枯渇顕微鏡（STED）12、個々の分子リターン（GSDIM）13続いて基底状態の枯渇顕微鏡のような他の超解像技術や、構造化照明顕微鏡（SIM）14は 、光活性化蛍光タンパク質を必要としません。また、有機色素9,15,16または蛍光プローブ17の固有の特性を使用した超解像イメージング技術は、容易に適用可能である。 各分子の蛍光が空間的にも時間的に分離されているので、手のひらに、分子数を定量化することができる。酸化、過少、過大、および過剰発現：しかし、定量化は、4つの理由のために誤解を招く可能性があります。まず、蛍光タンパク質の画分を変性またはサンプル処理5,12間に酸化することができる。蛍光シグナルの約90％が我々のプロトコルでの固定と包埋を通して維持されたが、試料が切削されており、表面が酸素にさらされた後に、蛍光タンパク質の酸化が発生することがあります。第二に、光活性タンパク質の活性化は、確率的であるため、複数の分子ができる所定の回折限界のスポット8で活性化。複数の分子からの蛍光は1スポットとして現れるでしょう、そして、その結果、タンパク質の総数はundercountedされます。第三に、同様の問題は過大につながることができます。手のひらに漂白することにより、それぞれの蛍光タンパク質は、ローカライズされた後、 &quot;消去&quot;。しかし、蛍光タンパク質は18永久に漂白されることなく、暗い状態から戻ることができます。このような分子は、複数回カウントされます。第四に、タグ付けされたタンパク質は、トランスジーンとして表現さ​​れ、しばしば過剰発現につながることができます複数のコピーに存在しています。したがって、Palmから定量化は、与えられた場所での分子の数を推定するが、正確には判断できないために使用することができます。 電子顕微鏡写真とヤシの画像の位置合わせもあるため、電子ビームによる光と電子顕微鏡と歪みで解像度の違いから挑戦することができます。金粒子は、しっかりとlocのサーブ電子顕微鏡写真で基準マーカーを未実現。しかし、金粒子からfluoresecenceは、光活性化ではなく、大規模な回折限界スポットとして表示されます。このように、電子顕微鏡写真上の蛍光画像の配置は推定値です。歪みもプラスチックセクションと電子の相互作用から生じる可能性があります。そのようなGMA等のアクリル系樹脂は、電子ビームの下にあまり安定しており、プラスチック製の寸法を変更することができます。このような状況下では、超微細構造と蛍光を整列する基準マーカーの非線形変換が必要な場合があります。

<table border="1"><tbody><tr><td>試薬や機器の名称</td><td>会社</td><td>カタログ番号</td><td>注釈</td></tr><tr><td>高圧冷凍庫</td><td>アブラ州</td><td> HPM 010 </td><td> WohlwendからライカやHPF-01からEMPactとHPM 100を使用することもできます。 </td></tr><tr><td>自動凍結置換ユニット</td><td>ライカ</td><td> AFSの2 </td><td> AFSの1を使用することもできます。 </td></tr><tr><td>ツァイスPALM </td><td>ツァイス</td><td> ELYRA P.1 </td><td>ニコンとVutaraはまた、商業やし顕微鏡を販売しています。 </td></tr><tr><td>走査型電子顕微鏡</td><td> FEI </td><td>ノバナノ</td><td>他の高分解能SEM顕微鏡を使用することができます。 </td></tr><tr><td>アセトン</td><td> EMSの</td><td> RT10016 </td><td></td></tr><tr><td>エタノール</td><td>シグマアルドリッチ</td><td> 459844-1L </td><td></td></tr><tr><td>オスミウムトンetroxide </td><td> EMSの</td><td> RT19134 </td><td></td></tr><tr><td>過マンガン酸カリウム</td><td> EMSの</td><td> RT20200 </td><td></td></tr><tr><td>ウシ血清アルブミンから</td><td>シグマアルドリッチ</td><td> A3059-50G </td><td></td></tr><tr><td>酢酸ウラニル</td><td>ポリサイエンス</td><td> 21447から25 </td><td>この会社の酢酸ウラニルのpHがわずかに高くなっています。 </td></tr><tr><td>グリコールメタクリレート（GMA） </td><td> SPI </td><td> 02630-AA </td><td>低酸、TEM用グレード。 </td></tr><tr><td> N、N-ジメチル-p-トルイジン</td><td>シグマアルドリッチ</td><td> D9912 </td><td></td></tr><tr><td>クライオバイアル</td><td>ナルゲン</td><td> 5000-0020 </td><td></td></tr><tr><td>ガラスバイアル</td><td> EMSの</td><td> 72632 </td><td></td></tr><tr><td> ACLARフィルム</td><td> EMSの</td><td> 50425から10 </td><td></td></tr><tr><td> BEEMカプセル</td><td> EBSciences </td><td> TC </td><td>ポリプロピレン</td></tr><tr><td> 3月8日 &quot;DISCパンチ</td><td>テッドペラ</td><td> 54741 </td><td></td></tr><tr><td>金ナノ粒子</td><td>ミクロス-nanospheres.com </td><td> 790122-010 </td><td> 2倍濃縮液を要求</td></tr></tbody></table>

nano-fem: protein localization using photo-activated localization microscopy and electron microscopy

タンパク質の分布をマッピングすると、細胞内のタンパク質の機能を理解するために不可欠です。蛍光顕微鏡は、広範囲のタンパク質局在するために使われますが、細胞内の文脈はしばしば蛍光画像では欠如しているされています。免疫電子顕微鏡法は、他の一方で、タンパク質をローカライズすることができますが、その方法は互換性のある抗体の不足によって制限され、形態学の貧しい保全とほとんどの抗原は、試料表面に露出されていないため。相関アプローチはどちら免疫蛍光または遺伝的にタグ付けされたタンパク質から、最初の全細胞からの蛍光画像を取得することができます。次に、試料を固定し、埋め込 ​​まれた電子顕微鏡観察のため、画像は1から3まで相関しているされています。しかし、低解像度の蛍光画像と基準マーカの欠如は、タンパク質の正確な局在を排除。 あるいは、蛍光イメージングは​​、プラスチック中の検体を保存した後に行うことができます。でこのアプローチは、ブロックが区分され、蛍光画像と同じ断面の電子顕微鏡写真を4-7相関している。しかし、相関画像の光の回折限界は個々の分子の位置を曖昧にし、蛍光はしばしばセルの境界を越えて延びている。 ナノ分解能蛍光電子顕微鏡（ナノFEM）を撮像することにより、ナノスケールでの光活性化ローカライゼーション顕微鏡（PALM）と電子顕微鏡を使用して同じ部分をタンパク質をローカライズするために設計されています。 Palmはイメージング個々の蛍光タンパク質による回折限界を克服し、その後、各蛍光スポット8-10の重心をマッピングする。 我々は5つのステップでナノFEM手法を概説しています。まず、試料を固定し、タグ付けされたタンパク質の蛍光を維持する条件を使用して、埋め込まれています。第二に、樹脂ブロックが搭載されている極薄のセグメント（70-80 nm）とに区画されているカバーガラス上に。第三に、蛍光はツァイスPALM顕微鏡を用いて、これらのセクションに結像される。第四に、電子密度の高い構造は、走査型電子顕微鏡を用いて、これらの同じセクションに結像される。第五に、蛍光および電子顕微鏡写真は、基準マーカーとしての金粒子を用いて整列されます。要約すると、蛍光標識タンパク質の細胞内局在は、約1週間でナノメートルの分解能で決定することができる。

Watch this Scientific Journal Video about Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy at JoVE.com

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

我々は、電子顕微鏡写真で蛍光標識タンパク質をローカライズする方法について説明します。蛍光は、第一超薄切片に光活性化ローカライゼーション顕微鏡を使ってローカライズされます。これらの画像は、同じ断面の電子顕微鏡写真に整列されます。

ナノFEM：写真活性化ローカライゼーション顕微鏡と電子顕微鏡を用いてタンパク質の局在

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

nano-fem-protein-localization-using-photo-activated-localization

Nanyang Technological University

Research

JoVE Journal

Biology

16.0K ビュー.  University of Utah. 我々は、電子顕微鏡写真で蛍光標識タンパク質をローカライズする方法について説明します。蛍光は、第一超薄切片に光活性化ローカライゼーション顕微鏡を使ってローカライズされます。これらの画像は、同じ断面の電子顕微鏡写真に整列されます。

ビデオ: Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM)

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and sensitivity in cells because the resolution of conventional light microscopy is diffraction-limited to ~250 nm. By combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM), PPIs can be visualized in cells with single molecule sensitivity and nanometer spatial resolution. BiFC is a commonly used technique for visualizing PPIs with fluorescence contrast, which involves splitting of a fluorescent protein into two non-fluorescent fragments. PALM is a recent superresolution microscopy technique for imaging biological samples at the nanometer and single molecule scales, which uses phototransformable fluorescent probes such as photoactivatable fluorescent proteins (PA-FPs). BiFC-PALM was demonstrated by splitting PAmCherry1, a PA-FP compatible with PALM, for its monomeric nature, good single molecule brightness, high contrast ratio, and utility for stoichiometry measurements. When split between amino acids 159 and 160, PAmCherry1 can be made into a BiFC probe that reconstitutes efficiently at 37 &#176;C with high specificity to PPIs and low non-specific reconstitution. Ras-Raf interaction is used as an example to show how BiFC-PALM helps to probe interactions at the nanometer scale and with single molecule resolution. Their diffusion can also be tracked in live cells using single molecule tracking (smt-) PALM. In this protocol, factors to consider when designing the fusion proteins for BiFC-PALM are discussed, sample preparation, image acquisition, and data analysis steps are explained, and a few exemplary results are showcased. Providing high spatial resolution, specificity, and sensitivity, BiFC-PALM is a useful tool for studying PPIs in intact biological samples.

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation BiFC-PALM

Protein-protein interactions are visualized in cells with nanometer spatial resolution by combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM). Described here is the use of BiFC-PALM for imaging Ras-Raf interactions in U2OS cells for visualizing the nanoscale clustering and diffusion of individual Ras-Raf complexes.

二分子蛍光相補性を持つ光活性化ローカライゼーション顕微鏡（BIFC-PALM）

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and ...

生物工学

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

生体内で1 分子追跡ショウジョウバエ運動神経がシナプス前ターミナルで

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

神経科学

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

生物学

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

ワンナノメートルの精度で蛍光イメージング（FIONA）

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

哺乳類細胞における地上州枯渇超解像イメージング

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

マイクロマニピュレーション技術形成ダイナミクスの解析と細胞骨格のレギュレータの売り上げ高

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

単一の膜タンパク質生きた哺乳類細胞の細胞内小器官のマルチカラー ローカリゼーション顕微鏡

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

超構造保存細胞における膜蛋白質局在化の電子顕微鏡分析のためのクライオAPEX法

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Confocal Fluorescence Microscopy: A Technique to Determine the Localization of Proteins in Mouse Fibroblasts

Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum Comprehensive Cancer Center, Baltimore, Maryland 21201 
2 Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, Maryland 21201
Confocal fluorescence microscopy is an imaging technique that enables increased optical resolution as compared to conventional &#34;wide-field&#34; epifluorescence microscopy. Confocal microscopes are able to achieve improved x-y optical resolution through &#34;laser scanning&#34;- typically a set of voltage-controlled mirrors (galvanometer or &#34;galvo&#34; mirrors) that direct laser-illumination to each pixel of the specimen at a time. More importantly, confocal microscopes achieve superior z-axial resolution by using a pinhole to remove out of focus light originating from locations that are not in the z-plane being scanned, thus enabling the detector to collect data from a specified z-plane. Because of the high z-resolution achievable in confocal microscopy, it is possible to collect images from a series of z-planes (also-called z-stack) and construct a 3D image through software.
Before discussing the mechanism of a confocal microscope, it is important to consider how a sample interacts with light. Light is composed of photons, packets of electromagnetic energy. A photon impinging on a biological sample can interact with the molecules comprising the sample in one of four ways: 1) the photon does not interact and passes through the sample; 2) the photon is reflected/scattered; 3) the photon is absorbed by a molecule and the absorbed energy is released as heat through processes collectively known as nonradiative decay; and 4) the photon is absorbed and the energy is then rapidly reemitted as a secondary photon through the process known as fluorescence. A molecule whose structure permits fluorescence emission is called a fluorophore. Most biological samples contain negligible endogenous fluorophores; therefore exogenous fluorophores must be used to highlight features of interest in the sample. During fluorescence microscopy, the sample is illuminated with light of the appropriate wavelength for absorption by the fluorophore. Upon absorbing a photon, a fluorophore is said to be &#34;excited&#34; and the process of absorption is referred to as &#34;excitation&#34;. When a fluorophore gives up energy in the form of a photon, the process is known as &#34;emission&#34;, and the emitted photon is called fluorescence.
The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a &#34;focal point&#34; where it is maximally focused. Beyond the focal point the light again diverges. The entering and exiting beams may be visualized as a pair of cones touching at the focal point (see Figure 1, left panel). The phenomenon of diffraction imposes a limit on how tightly a beam of light can be focused - the beam actually focuses to a spot of finite size. Two factors determine the size of the focal spot: 1) the wavelength of the light, and 2) the light-gathering ability of the objective lens, which is characterized by its numerical aperture (NA). The focal &#34;spot&#34; extends not only in the x-y plane, but also in the z direction, and is in reality a focal volume. The dimensions of this focal volume define the maximum resolution achievable by optical imaging. Although the number of photons is greatest within the focal volume, the conical light paths above and below the focus also contain a lower density of photons. Any fluorophore in the light path can thus be excited. In conventional (wide-field) epifluorescence microscopy, emission from fluorophores above and below the focal plane contribute out-of-focus fluorescence (a &#34;hazy background&#34;), which reduces image resolution and contrast, as demonstrated in Figure 1, with the red cube representing fluorophore emission above the focal plane (red star) that results in out-of-focus fluorescence (top right). This problem is ameliorated in confocal microscopy, due to the utilization of a pinhole. (Figure 2, bottom right). As depicted in Figure 3, the pinhole allows emissions originating from the focal place to reach the detector (left), while blocking the out-of-focus fluorescence (right) from reaching the detector, thus improving both resolution and contrast.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig1.jpg" /> 
Figure 1. Optical resolution of epifluorescence versus confocal microscopy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/10501/10501fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a focal volume and then diverges (left). The red star represents the focal plane of a sample that is being imaged while the red square represents fluorophore emission above the focal plane. When capturing an image of this sample using an epifluorescent microscope, the emission from the out-of-focus red square will visible and contribute to a &#34;hazy background&#34; (top right). Confocal microscopes have a pinhole which prevents the detection of light emitted outside of the focal plane, eliminating the &#34;hazy background&#34; (bottom right).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig2.jpg" /> 
Figure 2. Pinhole effect in confocal microscopy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/10501/10501fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Although the highest intensity of the excitation light is at the focal point of the lens (left, red oval), other parts of the sample not in the focal point (right, red star) will get light and fluoresce. To prevent light emitted from these out-of-focus regions to reach the detector, a screen with a pinhole is present in front of the detector. Only the in-focus light (left) emitting from the focal plane is able to travel through the pinhole and reach the detector. The out-of-focus light (right) is blocked with the pinhole and fails to reach the detector.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig3.jpg" /> 
Figure 3. Principal components of a confocal laser scanning microscope. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/10501/10501fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
For the sake of simplicity, the mechanistic description of a confocal microscope will be limited to that of the Nikon Eclipse Ti A1R. Although there may be minor technical differences between different confocal microscopes, the A1R serves well as a good model for describing confocal microscope function. The excitation light beam, produced by an array of diode lasers, is reflected by the primary dichroic mirror into the objective, which focuses the light onto the specimen being imaged. The primary dichroic mirror selectively reflects the excitation light while allowing light at other wavelengths to pass through. The light then encounters the scanning mirrors that sweep the light beam across the specimen in an x-y manner, illuminating a single (x,y) pixel at a time. Fluorescence emitted by fluorophores at the illuminated pixel is collected by the objective lens and passes through the primary dichroic mirror to reach an array of photomultiplier tubes (PMTs). Secondary dichroic mirrors direct the emission light to the appropriate PMT. Excitation light scattered by the sample back into the objective is reflected by the primary dichroic mirror back towards the specimen, and thus prevented from entering the detection light path and reaching the PMTs (see Figure 3). This allows the relatively feeble fluorescence to be quantified without contamination by light scattered from the excitation light beam, which is typically orders of magnitude more intense than the fluorescence. Because the pinhole blocks light from outside of the focal volume, the light arriving at the detector comes from a narrow, selected z-plane. Therefore, images can be collected from a series of adjacent z-planes; this series of images is often referred to as a &#39;z-stack&#39;. By using the appropriate software, a z-stack can be processed to generate a 3D image of the specimen. A particular advantage of confocal microscopy is the ability to distinguish the subcellular localization of staining. For example, the differentiation between membrane staining from intracellular staining, which is very challenging with conventional epifluorescence microscopy (1, 2, 3).
Sample preparation is an important facet of confocal imaging. A strength of optical microscopy techniques is the flexibility to image live or fixed cells. When attempting to produce 3D images, because of the number of images that must be acquired for a z-stack, the difficulty of maintaining cell health, and the movement of live cells and their organelles, the use of fixed cells is typical. The procedure for fixing and staining cells for confocal fluorescence is similar to that conventionally used in immunofluorescence. After culture in chamber slides or on coverslips, cells are fixed using paraformaldehyde to preserve cellular morphology. Non-specific antibody binding is blocked using bovine serum albumin, milk, or normal serum. In order to maintain the specificity of the secondary antibodies, the solution used should not originate from the same species in which the primary antibodies were generated. The cells are incubated with primary antibodies that bind the antigen of interest. When labeling several cellular targets, the primary antibodies must each be derived from a different species. Antibodies tagging an antigen are then bound by fluorophore-conjugated secondary antibodies. Fluorophore-conjugated secondary antibodies should be selected so that they are compatible with the wavelengths of laser excitation available in the confocal microscope. When visualizing multiple antigens, the excitation/emission spectra of the fluorophores should differ enough so that their signals can be discriminated by microscopic analysis. The stained specimen is then mounted on a slide for imaging. A mounting medium is used to prevent photobleaching and specimen dehydration. If desired, a mounting medium containing a nuclear counterstain (e.g. DAPI or Hoechst) can be used (4).
In the following protocol, mouse fibroblasts transfected to express CD1d (LCD1) were stained with antibodies recognizing CD1d and CD107a (LAMP-1). CD1d is a major histocompatibility complex 1 (MHC 1)-like receptor present on the surface of antigen presenting cells that presents lipid antigens. LAMP-1 (lysosomal associated membrane protein-1) is a transmembrane protein primarily present in lysosomal membranes. For proper antigen presentation, CD1d is trafficked through the low pH lysosomal compartment, so LAMP-1 is being used as a marker of the lysosomal compartment for this protocol. By probing the LCD1 cells with anti-CD1d and anti- LAMP-1 that were produced in different species, secondary antibodies with unique fluorophores can be used to determine the localization of each protein in the cell and whether CD1d is present in the LAMP-1 positive lysosomal compartments.

Confocal Fluorescence Microscopy and Localization of Proteins

共焦点蛍光顕微鏡:マウス線維芽細胞におけるタンパク質の局在を決定する技術

Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland School of Medicine ...

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JoVE Science Education

高等生物学

免疫学

Bimolecular Fluorescence Complementation-Coupled Photoactivated Localization Microscopy

Source: Nickerson, A. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/53154">Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM).</a> J. Vis. Exp. (2015)
This video describes BiFC-PALM &#8213; a combination of photoactivated localization microscopy and bimolecular fluorescence complementation &#8213; to assess protein-protein interactions. BiFC involves the fusion of two fluorescent protein fragments with two interacting proteins of interest. When the two proteins interact, the fragments are brought into proximity, which allows the two parts to come together and reconstitute a functional fluorescent protein. The fluorescence of a single fluorophore is localized at a single molecule level inside cells by PALM.

二分子蛍光相補結合光活性化局在顕微鏡法

Source: Nickerson, A. et al., Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM). J. Vis. Exp. (2015)
This ...

ナノFEM：写真活性化ローカライゼーション顕微鏡と電子顕微鏡を用いてタンパク質の局在

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この動画の章

二分子蛍光相補性を持つ光活性化ローカライゼーション顕微鏡（BIFC-PALM）

生体内で1 分子追跡ショウジョウバエ運動神経がシナプス前ターミナルで

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

ワンナノメートルの精度で蛍光イメージング（FIONA）

哺乳類細胞における地上州枯渇超解像イメージング

マイクロマニピュレーション技術形成ダイナミクスの解析と細胞骨格のレギュレータの売り上げ高

単一の膜タンパク質生きた哺乳類細胞の細胞内小器官のマルチカラーローカリゼーション顕微鏡

超構造保存細胞における膜蛋白質局在化の電子顕微鏡分析のためのクライオAPEX法

共焦点蛍光顕微鏡:マウス線維芽細胞におけるタンパク質の局在を決定する技術

二分子蛍光相補結合光活性化局在顕微鏡法

ナノFEM：写真活性化ローカライゼーション顕微鏡と電子顕微鏡を用いてタンパク質の局在

要約

さらに動画を探す

この動画の章

二分子蛍光相補性を持つ光活性化ローカライゼーション顕微鏡（BIFC-PALM）

生体内で1 分子追跡ショウジョウバエ運動神経がシナプス前ターミナルで

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

ワンナノメートルの精度で蛍光イメージング（FIONA）

哺乳類細胞における地上州枯渇超解像イメージング

マイクロマニピュレーション技術形成ダイナミクスの解析と細胞骨格のレギュレータの売り上げ高

単一の膜タンパク質生きた哺乳類細胞の細胞内小器官のマルチカラー ローカリゼーション顕微鏡

超構造保存細胞における膜蛋白質局在化の電子顕微鏡分析のためのクライオAPEX法

共焦点蛍光顕微鏡:マウス線維芽細胞におけるタンパク質の局在を決定する技術

二分子蛍光相補結合光活性化局在顕微鏡法

単一の膜タンパク質生きた哺乳類細胞の細胞内小器官のマルチカラーローカリゼーション顕微鏡