準備の手順インキュベーションバッファー、アッセイバッファー、および0.6 N TCAソリューションは、すぐに使えるソリューションとして提供されています。 FITC -カゼイン基質 - それはアリコートに凍結融解の繰り返しを避けるために到着時に小さなボリュームに分割しFITC -カゼイン基質をお勧めします。 FITC -カゼイン基質が凍結融解の繰り返し印加された場合、バックグラウンドでのわずかな増加は、感度を下げ、それによって、発生します。アリコートを-20℃で保存し、光から保護する必要があります。各サンプル、ブランク、またはコントロールの反応では、FITC -カゼイン基質20μlを必要とします。ミキシングや揺れ活発なだけでなく、融解/凍結とFITCは、プロテアーゼの切断のために利用できるように小さい基板を読み、原因と高いバックグラウンドで生じるカゼインから分離することがあります。 フルオレセインイソチオシアネート（FITC）コントロールソリューション- FITC制御は、適切な濃度にアッセイ緩​​衝液で再構成することができます。このソリューションは、新鮮なものと、光から保護する必要があります。 最終的なトリプシンコントロールソリューション - トリプシン、プロテアーゼ制御（商品コードT 6567）のバイアルに1mMのHClの100μlを追加する。トリプシンが溶解されるように簡単に混ぜる。インキュベーションバッファー900μLを加え、よく混ぜる。アッセイに必要に応じて別の方法として、他のバッファを使用することができます。トリプシンの最終的な作業濃度は20μg/μLのです。ときにトリプシンコントロールソリューションを準備する準備ができて、インキュベーションバッファー（9部のバッファに1部の酸性化トリプシン）の正確な量の酸性トリプシン溶液のアリコートを組み合わせる。酸性条件下でトリプシンを保存すると、トリプシンの安定性を高めます。トリプシン制御ソリューションは、温度に敏感であり、低濃度で安定していないです。 これらの準備のソリューションの貯蔵と安定性のためのキットの技術的なセキュリティ情報を確認してください。 手順<ol><li>一度にすべてのコントロール、空白、およびサンプルを調製するのが最も簡単です。各試験サンプルについて、マイクロ遠心チューブにインキュベーションバッファー20μlを、FITC -カゼイン基質液20μl、試験試料の10μlを加える。高いプロテアーゼ活性を有する試験片の場合は、サンプルの希釈が必要な場合があります。 </li><li>マイクロ遠心チューブにインキュベーションバッファー20μlを、FITC -カゼイン基質液20μl、およびコントロールサンプルの10μlを添加することにより、適切なコントロールのサンプル（コントロールサンプルを参照）を準備。 </li><li>マイクロ遠心チューブにインキュベーションバッファー20μlを、FITC -カゼイン基質液20μl、及び超純水10μlを添加してブランクサンプルを準備します。 </li><li>静かに37 60分間暗所で℃で各チューブとインキュベートを混ぜる。過度の乱流が高い蛍光バックグラウンドの原因とアッセイの感度を減少させる可能性があるので、あまりにも積極的に混合しないように注意してください。 注：インキュベーション時間は、感度を上げるために24時間まで延長することができる。高い蛍光バックグラウンドにつながる、FITC -カゼインが低下し始める可能性があるので24時間を超えないように注意してください。 </li><li>インキュベーション後、各微量遠心チューブに0.6 Nトリクロロ酢酸溶液を150μlを加える。 TCAは、非常に腐食性があり、適切な保護具を着用して処理する必要があります。 </li><li>ゆっくりと37℃で混合し、インキュベート℃で30分間、暗所で。 </li><li> 10,000 × gで10分間チューブを遠心上清は、酸可溶性、FITCラベルされた断片が含まれており、蛍光測定に使用されます。 </li></ol>蛍光測定これらのメソッドは、利用可能な計装の要件に応じて上下に拡大または縮小することができます。適切なコントロールのサンプルで作製した標準曲線との比較のために、各試験サンプルの値（FLUtest）からブランクサンプル（FLUblank）の蛍光測定値を引きます。 キュベット<ol><li>ピペット上清10μl（ステップ7）と適切なキュベットにアッセイ緩​​衝液1mlを、穏やかに混合する。注：最大24時間の蛍光を測定する前に、上清およびアッセイ緩​​衝液の溶液は2〜8℃で暗所に保存されていることがあります。 </li><li> 485nmで励起による蛍光強度を記録および535 nmの発光波長を監視する。 </li></ol>マルチウェルプレート<ol><li>上清（ステップ7）と適切なチューブまたはバイアルにアッセイ緩​​衝液1mlを、穏やかに混合注ピペットを10μl：上清およびアッセイ緩​​衝液の溶液を、最大2〜8℃で暗所に保存することができる蛍光を測定する前に24時間まで。 </li><li>黒色96ウェルプレートのウェルに200μlのを転送する。 485nmで励起による蛍光強度を記録および535 nmの発光波長を監視する。 や</li></ol><ol><li>ピペット上清2μlの（ステップ7）とアッセイB200μlの黒色96ウェルプレートのウェルにuffer。注：最大24時間の蛍光を測定する前に、上清およびアッセイ緩​​衝液の溶液は2〜8℃で暗所に保存されていることがあります。 </li><li> 485nmで励起による蛍光強度を記録および535 nmの発光波長を監視する。 </li></ol>コントロールのサンプルトリプシンコントロールソリューションは、アッセイは、検出限界を決定する、または一般的な標準曲線を作成するには、適切に実行していることを確認するために使用することができます。異なる、特定のプロテアーゼのアッセイのために、それは適切なインキュベーションバッファー中で特異的プロテアーゼを含むコントロール溶液を調製することをお勧めします。 アッセイの検出限界は、ブランク試料で得られた値を上回る大幅な蛍光測定値を生成するプロテアーゼの量です。検出限界は、計測機器の感度によって異なります。トリプシンコントロールソリューションの連続希釈を制御ソリューションを生成するために使用されることがあります。 ブランクサンプルで得られた値の120％に等しい読書を有意とされる。日常的に、トリプシンの5 ngの検出限界は、この手順で得られた。それは、少なくとも一つの5 ngのトリプシンのコントロールがそれぞれのアッセイで実行することをお勧めします。の0.5μg/ mlの濃度でトリプシンの制御ソリューションは、アッセイにおけるトリプシンの所望の5 ngのことになります。トリプシンコントロールソリューションの40倍希釈（20μg/ ml）を0.5μg/ mLと制御ソリューション、インキュベーションバッファーの39部にトリプシンコントロールソリューションの、すなわち一部の結果。 <img src="/files/ftp_upload/1514/figure1.jpg" alt="図1" /> 図1（トリプシン活性の標準曲線）：トリプシンコントロールソリューション（20μg/ mlのは）また、（図1を参照）連続希釈を行うことによって標準曲線を生成するために使用されることがあります。各コントロールのサンプルの蛍光測定値は、各コントロールのサンプル（FLUcontrol）の値からブランク試料の蛍光読書（FLUblank）を差し引くことによって修正されました。 このキットは0.15μg/ mlの（1.5 ng）を、2.5μg/ mlの（25 ng）をするまで、コントロールサンプルを使用して、トリプシン（製品コードT 6567）の典型的な標準曲線。この曲線は、ステップ4の30分のインキュベーション時間を、記載された手順に従って生成されました。蛍光測定は、マルチウェルプレートで行われた。 フルオレセインイソチオシアネートは、可能な測定器の校正（該当する製造者の指示を参照）またはFITC信号の直線性の範囲の決定のためのコントロールとして提供されています。

<ol>
	<li>Twining, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescein+Isothiocyanate-Labeled+Casein+Assay+for+Proteolytic+Enzymes.">Fluorescein Isothiocyanate-Labeled Casein Assay for Proteolytic Enzymes.</a> Anal. Biochem. 143, 30-34 (1984).</li></ol>

著者はこの記事で使用する試薬やツールを生成するシグマアルドリッチ社の従業員です。

私達はちょうど生物学的試料中のプロテアーゼ活性を検出するためにプロテアーゼ蛍光検出キットを使用して例の分析を示している。このキットは、生理的なアプリケーションで見られるプロテアーゼの多様な範囲を検出するために最適化されています。それはセリン、システイン、メタロ、およびアスパラギン酸プロテアーゼの検出に適していますが、修正がいくつかの特定のプロテアーゼを検出するために必要となる場合があります。

<table border="1">
<tr>
<td>Material Name</td>
<td>Type</td>
<td>Company</td>
<td>Catalogue Number</td>
<td>Comments (optional)</td>
</tr>
<tr>
<td><a href="http://www.sigmaaldrich.com/catalog/ProductDetail.do?lang=en&N4=PF0100|SIGMA&N5=SEARCH_CONCAT_PNO|BRAND_KEY&F=SPEC&cm_sp=affiliate-_-Jove-_-12346">Protease Fluorescent Detection Kit</a></td>
<td>&nbsp;</td>
<td>Sigma Aldrich</td>
<td>PF0100</td>
<td>&nbsp;</td>
<tr>
<td><a href="http://www.sigmaaldrich.com/catalog/ProductDetail.do?lang=en&N4=H1758|SIGMA&N5=SEARCH_CONCAT_PNO|BRAND_KEY&F=SPEC&cm_sp=affiliate-_-Jove-_-12347">Hydrochloric Acid</a></td>
<td>&nbsp;</td>
<td>Sigma Aldrich</td>
<td>H1758</td>
<td>&nbsp;</td>
</table>

use of the protease fluorescent detection kit to determine protease activity

プロテアーゼ蛍光検出キットは、既製使用するプロテアーゼ活性の存在を検出するための試薬を提供する。プロテアーゼ活性を検出するためにこの簡単なアッセイは、カゼインを基質としてフルオレセインイソチオシアネート（FITC）で標識使用しています。 プロテアーゼ活性の酸性条件下で沈殿しないより小さい断片にFITC標識カゼイン基質の開裂の結果。プロテアーゼのサンプルと基質のインキュベーション後、反応をトリクロロ酢酸（TCA）の添加により酸性化する。次いで、混合物をペレットと溶液中に残存する小規模な、酸可溶性フラグメントを形成する未消化の基質を用いて遠心分離する。上清を中和され、FITCラベルされた断片の蛍光を測定する。 記載のキットの手順では、約0.5μg/ mlの（トリプシンの5 ngの検定に追加された）の濃度でトリプシンのプロテアーゼ制御を検出します。この感度は、24時間まで、長いインキュベーション時間を増やすことができます。アッセイは、マイクロ遠心チューブ中で実行され、手続きは、キュベットまたはマルチウェルプレートのどちらかを使用して、蛍光検出のために用意されています。

Watch this Scientific Journal Video about Use of the Protease Fluorescent Detection Kit to Determine Protease Activity at JoVE.com

Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

プロテアーゼ蛍光検出キットは、蛍光を用いたプロテアーゼ活性の測定のために設計されています。また、プロテアーゼのコンタミの微量の検出に適しています。メソッドは、FITC標識カゼイン基質の独自の製剤のタンパク質分解hydroysisに基づいています。

プロテアーゼ活性を測定するためにプロテアーゼ蛍光検出キットの使用

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect ...

use-protease-fluorescent-detection-kit-to-determine-protease

Research

JoVE Journal

Biology

28.0K ビュー.  Sigma Aldrich. プロテアーゼ蛍光検出キットは、蛍光を用いたプロテアーゼ活性の測定のために設計されています。また、プロテアーゼのコンタミの微量の検出に適しています。メソッドは、FITC標識カゼイン基質の独自の製剤のタンパク質分解hydroysisに基づいています。

ビデオ: Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

Purifying Plasmid DNA from Bacterial Colonies Using the Qiagen Miniprep Kit

Plasmid DNA purification from E. coli is a core technique for molecular cloning. Small scale purification (miniprep) from less than 5 ml of bacterial culture is a quick way for clone verification or DNA isolation, followed by further enzymatic reactions (polymerase chain reaction and restriction enzyme digestion). Here, we video-recorded the general procedures of miniprep through the QIAGEN's QIAprep 8 Miniprep Kit, aiming to introducing this highly efficient technique to the general beginners for molecular biology techniques. The whole procedure is based on alkaline lysis of E. coli cells followed by adsorption of DNA onto silica in the presence of high salt. It consists of three steps: 1) preparation and clearing of a bacterial lysate, 2) adsorption of DNA onto the QIAprep membrane, 3) washing and elution of plasmid DNA. All steps are performed without the use of phenol, chloroform, CsCl, ethidium bromide, and without alcohol precipitation. It usually takes less than 2 hours to finish the entire procedure.

キアゲンミニプレップキットを用いて細菌のコロニーからプラスミドDNAを精製

Plasmid DNA purification from E. coli is a core technique for molecular cloning. Small scale purification (miniprep) from less than 5 ml of bacterial ...

生物学

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

光イメージングによる非侵襲的検出のための蛍光色素で標識幹細胞

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Sigma's Non-specific Protease Activity Assay - Casein as a Substrate

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases. Sigma's non-specific protease activity assay may be used as a standardized procedure to determine the activity of proteases, which is what we do during our quality control procedures. In this assay, casein acts as a substrate. When the protease we are testing digests casein, the amino acid tyrosine is liberated along with other amino acids and peptide fragments. Folin and Ciocalteus Phenol, or Folin's reagent primarily reacts with free tyrosine to produce a blue colored chromophore, which is quantifiable and measured as an absorbance value on the spectrophotometer. The more tyrosine that is released from casein, the more the chromophores are generated and the stronger the activity of the protease. Absorbance values generated by the activity of the protease are compared to a standard curve, which is generated by reacting known quantities of tyrosine with the F-C reagent to correlate changes in absorbance with the amount of tyrosine in micromoles. From the standard curve the activity of protease samples can be determined in terms of Units, which is the amount in micromoles of tyrosine equivalents released from casein per minute.
To view this article in Chinese, <a href="http://jove.com/index/details.stp?ID=910">click here</a>

Sigma&#039;s Non-specific Protease Activity Assay - Casein as a Substrate

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases. Sigma's non-specific protease activity assay may be used as a standardized procedure to determine the activity of proteases.

シグマの非特異的プロテアーゼの活性アッセイ - 基質としてカゼイン

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases.  Sigma's non-specific protease ...

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

酵素酢酸キナーゼのアセテート形成活性の直接検出

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Stable isotopes are essential tools in biological mass spectrometry. Historically, 18O-stable isotopes have been extensively used to study the catalytic mechanisms of proteolytic enzymes1-3. With the advent of mass spectrometry-based proteomics, the enzymatically-catalyzed incorporation of 18O-atoms from stable isotopically enriched water has become a popular method to quantitatively compare protein expression levels (reviewed by Fenselau and Yao4, Miyagi and Rao5 and Ye et al.6). 18O-labeling constitutes a simple and low-cost alternative to chemical (e.g. iTRAQ, ICAT) and metabolic (e.g. SILAC) labeling techniques7. Depending on the protease utilized, 18O-labeling can result in the incorporation of up to two 18O-atoms in the C-terminal carboxyl group of the cleavage product3. The labeling reaction can be subdivided into two independent processes, the peptide bond cleavage and the carboxyl oxygen exchange reaction8. In our PALeO (protease-assisted labeling employing 18O-enriched water) adaptation of enzymatic 18O-labeling, we utilized 50% 18O-enriched water to yield distinctive isotope signatures. In combination with high-resolution matrix-assisted laser desorption ionization time-of-flight tandem mass spectrometry (MALDI-TOF/TOF MS/MS), the characteristic isotope envelopes can be used to identify cleavage products with a high level of specificity. We previously have used the PALeO-methodology to detect and characterize endogenous proteases9 and monitor proteolytic reactions10-11. Since PALeO encodes the very essence of the proteolytic cleavage reaction, the experimental setup is simple and biochemical enrichment steps of cleavage products can be circumvented. The PALeO-method can easily be extended to (i) time course experiments that monitor the dynamics of proteolytic cleavage reactions and (ii) the analysis of proteolysis in complex biological samples that represent physiological conditions. PALeO-TimeCourse experiments help identifying rate-limiting processing steps and reaction intermediates in complex proteolytic pathway reactions. Furthermore, the PALeO-reaction allows us to identify proteolytic enzymes such as the serine protease trypsin that is capable to rebind its cleavage products and catalyze the incorporation of a second 18O-atom. Such "double-labeling" enzymes can be used for postdigestion 18O-labeling, in which peptides are exclusively labeled by the carboxyl oxygen exchange reaction. Our third strategy extends labeling employing 18O-enriched water beyond enzymes and uses acidic pH conditions to introduce 18O-stable isotope signatures into peptides.

Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Stable isotope labeling workflows employing 18O-enriched water (LeO-workflows) are versatile tools for quantitative and qualitative proteomics studies. In protease-assisted (PALeO) workflows, 18O-atoms are introduced by proteolytic cleavage and carboxyl oxygen exchange reactions mediated by proteases. In the acid-catalyzed (ALeO) workflow, 18O-atoms are introduced by carboxyl oxygen exchange at low pH.

採用プロテアーゼ及び酸触媒ラベリングワークフロー<sup&gt; 18</sup&gt; O-濃縮水

Stable isotopes are essential tools in biological mass spectrometry. Historically, 18O-stable isotopes have been extensively used to study the catalytic ...

Mouse Islet of Langerhans Isolation using a Combination of Purified Collagenase and Neutral Protease

The interrogation of beta cell gene expression and function in vitro has squarely shifted over the years from the study of rodent tumorigenic cell lines to the study of isolated rodent islets. Primary islets offer the distinct advantage that they more faithfully reflect the biology of intracellular signaling pathways and secretory responses. Whereas the method of islet isolation using tissue dissociating enzyme (TDE) preparations has been well established in many laboratories1-4, variations in the consistency of islet yield and quality from any given rodent strain limit the extent and feasibility of primary islet studies. These variations often occur as a result of the crude partially purified TDEs used in the islet isolation procedure; TDEs frequently exhibit lot-to-lot variations in activity and often require adjustments to the dose of enzyme used. A small number of reports have used purified TDEs for rodent cell isolations5, 6, but the practice is not widespread despite the routine use and advantages of purified TDEs for human islet isolations. In collaboration with VitaCyte, LLC (Indianapolis, IN), we developed a modified mouse islet isolation protocol based on that described by Gotoh7, 8, in which the TDEs are perfused directly into the pancreatic duct of mice, followed by crude tissue fractionation through a Histopaque gradient9, and isolation of purified islets. A significant difference in our protocol is the use of purified collagenase (CIzyme MA) and neutral protease (CIzyme BP) combination. The collagenase was characterized by the use of a6 fluorescence collagen degrading activity (CDA) assay that utilized fluorescently labeled soluble calf skin fibrils as substrate6. This substrate is more predictive of the kinetics of collagen degradation in the tissue matrix because it relies on native collagen as the substrate. The protease was characterized with a sensitive fluorescent kinetic assay10. Utilizing these improved assays along with more traditional biochemical analysis enable the TDE to be manufactured more consistently, leading to improved performance consistency between lots. The protocol described in here was optimized for maximal islet yield and optimal islet morphology using C57BL/6 mice. During the development of this protocol, several combinations of collagenase and neutral proteases were evaluated at different concentrations, and the final ratio of collagenase:neutral protease of 35:10 represents enzyme performance comparable to Sigma Type XI. Because significant variability in average islet yields from different strains of rats and mice have been reported, additional modifications of the TDE composition should be made to improve the yield and quality of islets recovered from different species and strains.

A detailed description of mouse islet isolation is described using the technique of in situ pancreatic ductal cannulation and perfusion of a combination of purified collagenase and neutral protease.

コラゲナーゼと中性プロテアーゼの精製の組合せを用いて単離ランゲルハンスマウスの膵島

The interrogation of beta cell gene expression and function in vitro has squarely shifted over the years from the study of rodent tumorigenic cell lines ...

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

蛍光プロテアーゼ保護を使用して膜タンパク質のトポロジの決定（FPP）

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro and in vivo. Fluorescent probes are particularly useful in determining the bio-distribution of administered large molecules, where the addition of a small-molecule fluorescent label is unlikely to affect the kinetics or bio-distribution of the compound. A variety of methods exist to examine bio-distribution that vary significantly in the amount of effort required and whether the resulting measurements are fully quantitative, but using multiple methods in conjunction can provide a rapid and effective system for analyzing bio-distributions.
Ex vivo whole-organ imaging is a method that can be used to quickly compare the relative concentrations of fluorescent molecules within tissues and between multiple types of tissues or treatment groups. Using an imaging platform designed for live-animal or whole-organ imaging, fluorescence within intact tissues can be determined without further processing, saving time and labor while providing an accurate picture of the overall bio-distribution. This process is ideal in experiments attempting to determine the tissue specificity of a compound or for the comparison of multiple different compounds. Quantitative tissue histology on the other hand requires extensive further processing of tissues in order to create a quantitative measure of the labeled compounds. To accurately assess bio-distribution, all tissues of interest must be sliced, scanned, and analyzed relative to standard curves in order to make comparisons between tissues or groups. Quantitative tissue histology is the gold standard for determining absolute compound concentrations within tissues.
Here, we describe how both methods can be used together effectively to assess the ability of different administration methods and compound modifications to target and deliver fluorescently labeled molecules to the central nervous system1.

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Ex vivo whole-organ imaging is a rapid method for determining the relative concentrations of fluorescently labeled compounds within and between tissues or treatment groups. Conversely, quantitative fluorescence histology, while more labor intensive, allows for the quantification of the absolute tissue levels of labeled molecules.

の用法<em&gt; ex vivoで</em&gt;全体の臓器イメージングと定量的組織組織学は、蛍光標識分子のバイオ分布を決定します

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro ...

The Lactate Dehydrogenase Sequestration Assay &#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a simple protocol to monitor this process is described. Moreover, typical results and experimental validation of the method under autophagy-inducing conditions in various types of cultured mammalian cells are provided. During bulk autophagy, autophagosomes sequester cytosol, and thereby also soluble cytosolic proteins, alongside other autophagic cargo. LDH is a stable and highly abundant, soluble cytosolic enzyme that is non-selectively sequestered into autophagosomes. The amount of LDH sequestration therefore reflects the amount of bulk autophagic sequestration. To efficiently and accurately determine LDH sequestration in cells, we employ an electrodisruption-based fractionation protocol that effectively separates sedimentable from cytosolic LDH, followed by measurement of enzymatic activity in sedimentable fractions versus whole-cell samples. Autophagic sequestration is determined by subtracting the proportion of sedimentable LDH in untreated cells from that in treated cells. The advantage of the LDH sequestration assay is that it gives a quantitative measure of the autophagic sequestration of endogenous cargo, as opposed to other methods that either involve ectopic expression of sequestration probes or semi-quantitative protease protection analyses of autophagy markers or receptors.

The Lactate Dehydrogenase Sequestration Assay &amp;#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Here a simple and well-validated protocol for measuring bulk autophagic sequestration activity in mammalian cells is described. The method is based on quantifying the proportion of lactate dehydrogenase (LDH) in sedimentable cell fractions compared to total cellular LDH levels.

隔離の乳酸脱水素酵素試金-哺乳類細胞における一括 Autophagic 隔離活動を決定するシンプルで信頼性の高い方法

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a ...

Multiplexed Fluorescent Immunohistochemical Staining of Four Endometrial Immune Cell Types in Recurrent Miscarriage

Immunohistochemistry is the most commonly used method for the identification and visualization of tissue antigens in biological research and clinical diagnostics. It can be used to characterize various biological processes or pathologies, such as wound-healing, immune response, tissue rejection, and tissue-biomaterial interactions. However, the visualization and quantification of multiple antigens (especially for immune cells) in a single tissue section using conventional immunohistochemical (IHC) staining remains unsatisfactory. Hence, multiplexed technologies were introduced in recent years to identify multiple biological markers in a single tissue sample or an ensemble of different tissue samples. 
These technologies can be especially useful in differentiating the changes in immune cell-to-cell interactions within the endometrium between fertile women and women with recurrent miscarriages during implantation. This paper describes a detailed protocol for multiplexed fluorescence IHC staining to investigate the density and clustering of four major immune cell types simultaneously in precisely timed endometrial specimens during embryo implantation. The method includes sample preparation, multiplex optimization with markers for immune cell subtypes, and the scanning of the slides, followed by data analysis, with specific reference to detecting endometrial immune cells. 
Using this method, the density and clustering of four major immune cell types in the endometrium can be simultaneously analyzed in a single tissue section. In addition, this paper will discuss the critical factors and troubleshooting to overcome possible fluorophore interference between the fluorescent probes being applied. Importantly, the results from this multiplex staining technique can help provide an in-depth understanding of the immunologic interaction and regulation during embryo implantation.

Despite the advancements in multiplex immunohistochemistry and multispectral imaging, characterizing the density and clustering of major immune cells simultaneously in the endometrium remains a challenge. This paper describes a detailed multiplex staining protocol and imaging for the simultaneous localization of four immune cell types in the endometrium.

再発流産における4種類の子宮内膜免疫細胞型の多重化蛍光免疫物質化学的染色

Immunohistochemistry is the most commonly used method for the identification and visualization of tissue antigens in biological research and clinical ...