細胞：ラット（またはマウス）の主海馬ニューロン培養（in vitroで14から28日間）。 刺激：電気、二つの白金電極を介して配信、70から90 mVの顕微鏡：倒立CCD蛍光顕微鏡で60倍の油レンズ。 ソフトウェア：Slidebook（インテリジェントイメージングイノベーション、サンタモニカ、カリフォルニア州） 詳細については、補足的な方法のページを参照してください。 <ol><li>暖かいHEPES緩衝室温に生理食塩水（HBS）。グルタミン酸受容体拮抗薬APV（50μMの最終濃度）とCNQX（10μM最終）を追加します。各FMの実験は、通常20 mLのHBSが必要です。 10μMFM 4から64（Molecular Probes社）の濃度を操作するための、（拮抗薬が追加された）HBSのストック溶液1:1000に希釈する。各実験のために2 mLのFM solnを行います。光暴露を防止するために箔を備えたソリューションをカバーする。 </li><li>電極室にニューロンを含むカバースリップをマウントします。チャンバー内のメディアは、チャンバーの上部と同じ高さと完全に電極を覆っていることを確認してください。カバースリップの下から余分な溶液を除去。レンズ（油のレンズを使用している場合）にオイルを追加。 </li><li>灌流装置を設定します。水、エタノール、水、そしてHBS：最初の指定された順序（次を追加する前に、徹底的にそれぞれの流れを聞かせ）で、それぞれ次の数mLですすいでください。次のHBSを追加する（今後のステップに手作業で追加するには、数mlを保存）し、血流を閉じます。カバースリップの端で反対側の血流のコンセントと吸引を設定します。灌流のコンセントは、吸引力が非常に上部に休息する場所として、チャンバー内に達すると、それが最善です。 HBSは（どちらか灌流によるまたは手で）、吸引を開いて追加するとき、それは完全に電極を覆う右だけのレベルでメディアを維持するようにその位置を調整します。 </li><li>あなたがイメージすると、大体の焦点に持参しようとするとカバースリップ上の領域を探します。最適なエリアは多くのシナプスが含まれていますが、個々のプロセスがつかなくなるようなので、高密度であってはならない。ないから最小限の細胞体、余分な膜（例えば、星状膠細胞の塊のような）、または他の非特異的な材料（糸くずなど）がないはずです。 FMの染料は、非特異的にこれらに付着する場合があります。 </li><li> （緑色の光で励起し、赤から遠赤放射を収集する）&quot;WG&quot;フィルターキューブを使用してください。イメージ0の発症（一般に300 900のAPにこのローディングの刺激のために使用される）により10Hzで900 APを提供するSlideview&quot;キャプチャ設定&quot;を設定します。画像ウィンドウで、&quot;蛍光 - 可視 - 赤&quot;のフィルタ設定を選択して、100ミリ秒に露光時間を変更する。任意の既存のタイムラプスの設定をオフにします。あなたが刺激を開始する準備が整うまでの任意の画像を取ってはいけない。 </li><li>吸引はオン/開いていることを確認してください。素早く吸引の反対側の端にある部屋にFM soln（2 mL）を加える。すぐに画像を取り、刺激を開始するために画像のウィンドウでは大丈夫押してください。刺激後30から45秒待ってから、すぐに（私はピペットで手でこれを行うが、これは血流によっても行うことができます）HBSの〜2 mLを加えることによりFMを洗い流す。 </li><li> 〜10分間灌流を経由してHBSとFMを洗ってください。流速は1から1.5 mL /分にする必要があります。この洗浄ステップの間、刺激の発症が始まるように刺激の設定を変更する@画像10（通常は900から1200のAP @ 10 Hzのがこの脱色刺激のために使用されています）。洗浄に7分程度、二重設定を（画像10の刺激の発症）確認してください。今すぐ小さなフォーカスウィンドウを使用すると、サブ領域を選択し、重点を置いています。刺激の設定が次のステップに進む前に変更されていることを確認してください。洗浄が完了したかどうかを確認するために単一のイメージ（個々のシナプスが明確にpunctatedする必要がある）を取る。されていない場合は、流量の追加0.5mL/minを増加させ、別の2分間待ちます。 </li><li>一度洗浄が完了すると、まだ良い、明確な信号を（通常は50から100の間にミリ秒。露光時間はFMの急速な退色による最小化する必要がある）を受信するために、必要に応じて露光時間を調整します。その後、38個のイメージを5秒ごとに取ることタイムラプスプリファレンスを設定する。 HBSは、さらに5分間十分あるように血流を確認し、血流が流れるままに。脱色を開始するボックスに追加開始。 </li><li>脱色した後、イメージ0時1200年のAP @ 10Hzのを（通常は1200〜2000のAPがこの最後のステップのために使用されている）を提供するための刺激の設定を変更してください。タイムラプスの設定をオフにして、刺激を開始するために単一のイメージを取る。血流は、まだこの段階ではベースライン/&quot;総放出可能&quot;蛍光を決定するために、残りの小胞FM色素を除去に役立つ通過している必要があります。刺激が終了した後、焦点に領域を持って、刺激の設定をオフにしてください。灌流と吸引を閉じますしてから、2つの最終的なベースラインの画像を撮影する。単一の画像やZ -スタック画像（0.5μmのステップ）のどちらかを取ることができる。実験時の焦点面のずれがあった場合には、Z -スタックが便利です。 </li><li>あなたのFM実験が行われます。すぐに多くの実験を行っている場合、レンズとチャンバーは各試行の間に掃除が、灌流装置は、最後の実験後まで掃除する必要はありませんする必要があります。その後、水、エタノールで完全に装置を洗浄します。 </li></ol>

<ol>
	<li>Betz, W. J., Bewick, G. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+analysis+of+synaptic+vesicle+recycling+at+the+frog+neuromuscular+junction.">Optical analysis of synaptic vesicle recycling at the frog neuromuscular junction.</a> Science. 255, 200-203 (1992).</li></ol>

The authors have nothing to disclose.

小胞のリサイクルの少なくとも一つのラウンドを誘導するのに十分である300活動電位（AP）のように、300台のAPまたは複数のローディング刺激は、しばしば小胞のリサイクルプールにラベルを付けるために与えられます。など900のAPのようなより強いローディング刺激が、、標識する追加の小胞の名目上の数を可能にするかもしれないが、これはまた、より非特異的な染色につながる、色素は細胞外膜に埋め?...

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measuring exocytosis in neurons using fm labeling

小胞放出の動態を測定する能力は、神経伝達の基本のいくつかの洞察を提供することができます。ここでは、シナプス小胞放出の速度を監視するためのFM色素で標識された小胞のリアルタイムイメージングを使用していました。 FM4 - 64は、エンドサイトーシスを刺激されているシナプス小胞の膜に埋め込まれて赤色蛍光両親媒性のスチリル色素です。親油性の相互作用は、色素が大きく従って小胞と公称細胞外液に関連付けられている明るいシグナルを発し、蛍光の増加を引き起こす。洗浄工程が細胞膜内に外部色素を除去するために使用された後、残りのFMは、小胞内に濃縮し、その後、エキソサイトーシスは、電気刺激の別のラウンドにより誘導されたときに排出されます。小胞の放出速度は、蛍光の結果減少から測定されます。 FM色素が外部と一時的に適用できるので、それは、トランスフェクションしたシナプスと隣接する制御boutons間の金利を比較する場合は特に、ニューロン培養でエキソサイトーシスの速度を決定するための便利なツールです。

Watch this Scientific Journal Video about Measuring Exocytosis in Neurons Using FM Labeling at JoVE.com

Measuring Exocytosis in Neurons Using FM Labeling

小胞放出の動態を測定する能力は、神経伝達の基本のいくつかの洞察を提供することができます。ここでは、海馬ニューロン培養における前シナプス小胞の放出速度を測定するために赤色蛍光色素FM 4〜64で標識された小胞のリアルタイムイメージングを使用していました。

FMラベリングを用いた神経細胞の測定エキソサイトーシス

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

measuring-exocytosis-in-neurons-using-fm-labeling

Research

JoVE Journal

Biology

19.3K ビュー.  Harvard. 小胞放出の動態を測定する能力は、神経伝達の基本のいくつかの洞察を提供することができます。ここでは、海馬ニューロン培養における前シナプス小胞の放出速度を測定するために赤色蛍光色素FM 4〜64で標識された小胞のリアルタイムイメージングを使用していました。

ビデオ: Measuring Exocytosis in Neurons Using FM Labeling

Using Laser Tweezers For Manipulating Isolated Neurons In Vitro

In this paper and video, we describe the protocols used in our laboratory to study the targeting preferences of regenerating cell processes of adult retinal neurons in vitro. Procedures for preparing retinal cell cultures start with the dissection, digestion and trituration of the retina, and end with the plating of isolated retinal cells on dishes made especially for use with laser tweezers. These dishes are divided into a cell adhesive half and a cell repellant half. The cell adhesive side is coated with a layer of Sal-1 antibodies, which provide a substrate upon which our cells grow. Other adhesive substrates could be used for other cell types. The cell repellant side is coated with a thin layer of poly-HEMA. The cells plated on the poly-HEMA side of the dish are trapped with the laser tweezers, transported and then placed adjacent to a cell on the Sal-1 side to create a pair. Formation of cell groups of any size should be possible with this technique. "Laser-tweezers-controlled micromanipulation" means that the investigator can choose which cells to move, and the desired distance between the cells can be standardized. Because the laser beam goes through transparent surfaces of the culture dish, cell selection and placement are done in an enclosed, sterile environment. Cells can be monitored by video time-lapse and used with any cell biological technique required. This technique may help investigations of cell-cell interactions.

This video describes the manipulation of cultured neurons using laser tweezers in vitro.

体外で分離された神経細胞を操作するためのレーザーピンセットを使用して

In this paper and video, we describe the protocols used in our laboratory to study the targeting preferences of regenerating cell processes of adult ...

生物学

Calcium Imaging of Cortical Neurons using Fura-2 AM

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium indicator dyes, which are BAPTA-based organic molecules that change their spectral properties in response to the binding of Ca2+ ions. Calcium indicator dyes fall into two categories, ratio-metric dyes like Fura-2 and Indo-1 and single-wavelength dyes like Fluo-4. Ratio-metric dyes change either their excitation or their emission spectra in response to calcium, allowing the concentration of intracellular calcium to be determined from the ratio of fluorescence emission or excitation at distinct wavelengths.  The main advantage of using ratio-metric dyes over single wavelength probes is that the ratio signal is independent of the dye concentration, illumination intensity, and optical path length allowing the concentration of intracellular calcium to be determined independently of these artifacts. One of the most common calcium indicators is Fura-2, which has an emission peak at 505 nM and changes its excitation peak from 340 nm to 380 nm in response to calcium binding.  Here we describe the use of Fura-2 to measure intracellular calcium elevations in neurons and other excitable cells.

Calcium signals play a key role in many cellular processes including gene expression, survival and differentiation. Here we demonstrate how to perform calcium imaging using Fura-2 AM. Calcium imaging is a valuable tool to study the regulation of intracellular calcium in real time and its regulation of signaling cascades.

AMフラ-2を用いて皮質ニューロンのカルシウムイメージング

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium ...

Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume. A specially designed differential pressure transducer and an occlusion tail-cuff measure the total blood volume in the tail without the need to obtain the individual pulse signal. Special attention is afforded to the length of the occlusion cuff in order to derive the most accurate blood pressure readings. VPR can easily obtain readings on dark-skinned rodents, such as C57BL6 mice and is MRI compatible. The CODA system provides you with measurements of six (6) different blood pressure parameters; systolic and diastolic blood pressure, heart rate, mean blood pressure, tail blood flow, and tail blood volume. Measurements can be made on either awake or anesthetized mice or rats. The CODA system includes a controller, laptop computer, software, cuffs, animal holders, infrared warming pads, and an infrared thermometer. There are seven different holder sizes for mice as small as 8 grams to rats as large as 900 grams.

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. This tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume.

ボリューム圧力記録、テールカフ法を用いたマウスにおける測定血圧

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA ...

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

全反射顕微鏡による網膜双極細胞におけるイメージングのエキソサイトーシス

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Measuring Fast Calcium Fluxes in Cardiomyocytes

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to extracellular agents is predominantly regulated by the phospholipase C&beta;- G&alpha;q pathway localized on the plasma membrane which is stimulated by agents such as acetylcholine 3,4. We have recently found that plasma membrane protein domains called caveolae5,6 can entrap activated G&alpha;q 7. This entrapment has the effect of stabilizing the activated state of G&alpha;q and resulting in prolonged Ca2+ signals in cardiomyocytes and other cell types8. We uncovered this surprising result by measuring dynamic calcium responses on a fast scale in living cardiomyocytes. Briefly, cells are loaded with a fluorescent Ca2+ indicator. In our studies, we used Ca2+ Green (Invitrogen, Inc.) which exhibits an increase in fluorescence emission intensity upon binding of calcium ions. The fluorescence intensity is then recorded for using a line-scan mode of a laser scanning confocal microscope. This method allows rapid acquisition of the time course of fluorescence intensity in pixels along a selected line, producing several hundreds of time traces on the microsecond time scale. These very fast traces are transferred into excel and then into Sigmaplot for analysis, and are compared to traces obtained for electronic noise, free dye, and other controls. To dissect Ca2+ responses of different flux rates, we performed a histogram analysis that binned pixel intensities with time. Binning allows us to group over 500 traces of scans and visualize the compiled results spatially and temporally on a single plot. Thus, the slow Ca2+ waves that are difficult to discern when the scans are overlaid due to different peak placement and noise, can be readily seen in the binned histograms. Very fast fluxes in the time scale of the measurement show a narrow distribution of intensities in the very short time bins whereas longer Ca2+ waves show binned data with a broad distribution over longer time bins. These different time distributions allow us to dissect the timing of Ca2+fluxes in the cells, and to determine their impact on various cellular events.

We present a method to isolate rapid (microsecond) calcium events from slower fluxes in living cells using laser scanning confocal microscopy. The method measures fluorescence intensity fluctuations of calcium indicators by recording line scans of several hundred pixels in a cell. Histogram analysis allows us to isolate the time scales of different calcium fluxes.

心筋細胞における高速カルシウムフラックスの測定

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

代謝標識とフローサイトメトリーで細胞周期の進行速度を測定

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

拘束されていない全身プレチスモグラフィを使用したマウスにおける呼吸機能の測定

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

ハイドラ内グルタチオンによって誘発される摂食応答を測定する

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

の剛性を測定します<em&gt; ex vivoで</em原子間力顕微鏡を使用&gt;マウス大動脈

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Imaging FITC-dextran as a Reporter for Regulated Exocytosis

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated exocytosis is fundamental for intercellular communication and is a key mechanism for the secretion of neurotransmitters, hormones, inflammatory mediators, and other compounds, by a variety of cells. At least three distinct mechanisms are known for regulated exocytosis: full exocytosis, where a single SG fully fuses with the plasma membrane, kiss-and-run exocytosis, where a single SG transiently fuses with the plasma membrane, and compound exocytosis, where several SGs fuse with each other, prior to or after SG fusion with the plasma membrane. The type of regulated exocytosis undertaken by a cell is often dictated by the type of secretory trigger. However, in many cells, a single secretory trigger can activate multiple modes of regulated exocytosis simultaneously. Despite their abundance and importance across cell types and species, the mechanisms that determine the different modes of secretion are largely unresolved. One of the main challenges in investigating the different modes of regulated exocytosis, is the difficulty in distinguishing between them as well as exploring them separately. Here we describe the use of fluorescein isothiocyanate (FITC)-dextran as an exocytosis reporter, and live cell imaging, to differentiate between the different pathways of regulated exocytosis, focusing on compound exocytosis, based on the robustness and duration of the exocytic events.

Here we detail a method for live cell imaging of regulated exocytosis. This method utilizes FITC-dextran, which accumulates in lysosome-related organelles, as a reporter. This simple method also allows distinguishing between different modes of regulated exocytosis in cells that are difficult to manipulate genetically.

規制エキソサイトーシスの記者として FITC デキストランをイメージング

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated ...