Cellules: Rat (ou la souris) primaire cultures de neurones hippocampiques (14-28 jours in vitro). Stimulation: électrique, livré par deux électrodes de platine; 70-90 mV Microscopie: lentille d&#39;huile 60x sur un microscope inversé CCD fluorescentes. Logiciel: Slidebook (Intelligent Imaging Innovations, Santa Monica, Californie) Voir la page supplémentaire méthodes pour plus de détails <ol><li> Chaud une solution saline tamponnée HEPES (HBS) à température ambiante. Ajouter les antagonistes des récepteurs de glutamate APV (50 uM de concentration finale) et CNQX (10 pM final). Chaque expérience FM nécessite généralement 20-30 ml de HBS. Pour travailler la concentration de 10 uM FM 4-64 (Molecular Probes), diluer 1:1000 solution stock dans HBS (avec des antagonistes ajoutée). Faire 2 ml FM soln pour chaque expérience. Couvrir avec une feuille solution pour éviter l&#39;exposition lumineuse. </li><li> Montez la lamelle contenant les neurones sur la chambre de l&#39;électrode. Vérifiez que les médias dans la chambre est au niveau haut de la chambre et le couvre complètement les électrodes. Retirez tout excès de solution de fond de la lamelle. Ajouter l&#39;huile de la lentille (si vous utilisez une lentille de pétrole). </li><li> Mettre en place l&#39;appareil de perfusion. Rincer d&#39;abord quelques mL de chaque année suivant l&#39;ordre donné dans le (laissez chaque flux une profondeur avant d&#39;ajouter le suivant): l&#39;eau, l&#39;éthanol, l&#39;eau, puis HBS. Ensuite, ajoutez l&#39;EBM (sauf quelques mL d&#39;ajouter à la main dans les étapes futures) et proche de la perfusion. Mettre en place la prise de perfusion et de l&#39;aspiration sur les côtés opposés sur les bords de la lamelle. Il est préférable que la prise de perfusion atteint dans la chambre, où, comme l&#39;aspiration doit reposer tout en haut. Tout en ajoutant HBS (soit par perfusion ou à la main), ouvrez l&#39;aspiration et d&#39;ajuster sa position afin qu&#39;il conserve les médias au niveau de la droite, juste recouvrant entièrement les électrodes. </li><li> Cherchez la zone sur la lamelle à l&#39;image que vous souhaitez et d&#39;essayer de le mettre à peu près au point. Zones optimales contiennent nombreuses synapses, mais ne devrait pas être si denses telles que les processus individuels deviennent indiscernables. Il ne devrait pas être à un minimum les corps cellulaires, les membranes extérieures (telles que des touffes d&#39;astrocytes), ou d&#39;autres matériaux non spécifiques (tels que les peluches). Le colorant non spécifique FM peuvent adhérer à ces derniers. </li><li> Utilisez le &quot;GT&quot; cube filtre (qui excite avec la lumière verte et recueille rouge-à-jour des émissions en rouge). Réglez &quot;préférences de capture&quot; SlideView pour livrer 900 AP à 10Hz sur l&#39;apparition de l&#39;image de 0 (généralement de 300 à 900 points d&#39;accès sont utilisés pour cette relance de chargement). Dans la fenêtre d&#39;image, sélectionnez l&#39;option &quot;fluo-vis-rouge&quot; des paramètres de filtre et changer le temps d&#39;exposition à 100 ms. Éteignez tous les paramètres timelapse pré-existante. Ne prenez pas toutes les images jusqu&#39;à ce que vous êtes prêt à commencer la stimulation. </li><li> Assurez-vous que l&#39;aspiration est sur / ouvert. Ajouter rapidement le soln FM (2 ml) à la chambre à l&#39;extrémité opposée de l&#39;aspiration. Appuyez immédiatement sur OK dans la fenêtre d&#39;image pour prendre une image et de commencer la stimulation. Attendre 30 à 45 secondes après la stimulation, puis rapidement laver FM en ajoutant ~ 2 ml de HBS (je le fais à la main avec une pipette, mais cela peut aussi être fait par perfusion). </li><li> Laver la FM avec HBS par perfusion pour ~ 10 minutes. Le débit devrait être de 1 à 1,5 ml / min. Durant cette étape de lavage, changer les préférences de stimulation de sorte que le début de la stimulation commence @ 10 images (généralement de 900 à 1200 points d&#39;accès @ 10 Hz sont utilisées pour ce stimulus décoloration). Environ 7 min dans le lavage, vérifier les préférences (apparition de stimulus à l&#39;image 10). Maintenant, en utilisant la fenêtre de petit foyer choisir et de se concentrer une sous-région. Assurez-vous que les paramètres de stimulation ont été changés avant de procéder à l&#39;étape suivante. Prenez une seule image pour voir si le lavage est terminé (synapses individuelles doivent être clairement pointillée). Si non, augmenter le débit une 0.5mL/min supplémentaires, et d&#39;attendre encore 2 minutes. </li><li> Une fois de lavage est terminé, régler le temps d&#39;exposition nécessaires pour continuer de recevoir un bon signal clair (généralement entre 50-100 ms. Temps d&#39;exposition doit être minimisée en raison de photoblanchiment rapide de la FM). Ensuite définir les préférences timelapse de prendre 38 images toutes les 5 secondes. Vérifiez la perfusion de faire il ya assez de HBS pendant 5 minutes, et de laisser couler la perfusion. Démarrer timelapse pour commencer décoloration. </li><li> Après Décolorer, modifier les paramètres de stimulation pour livrer 1200 @ 10Hz AP (généralement de 1200 à 2000 points d&#39;accès sont utilisés pour cette étape finale) sur l&#39;image 0. Désactivez les paramètres timelapse et de prendre une seule image pour commencer la stimulation. La perfusion doit être encore couler Cette étape permettra d&#39;éliminer toute vésiculaires reste de teinture FM, afin de déterminer de base / «total libérable&quot; par fluorescence. Après la relance est terminée, éteignez les paramètres de stimulation, mettre la région dans le foyer. Fermer la perfusion et aspiration, Puis prendre deux images de référence finale. Soit des images simples ou des images Z-stack (0,5 um étapes) peuvent être prises. Z-piles sont utiles dans le cas où il a été un changement dans le plan de mise au point pendant l&#39;expérience. </li><li> Votre expérience FM est fait. Si immédiatement faire plus d&#39;expériences, la lentille et la chambre doit être nettoyée entre chaque essai, mais l&#39;appareil de perfusion ne doit pas être nettoyée qu&#39;après l&#39;expérience finale. Rincer l&#39;appareil entièrement avec de l&#39;eau, puis de l&#39;éthanol. </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.

Comme 300 des potentiels d&#39;action (AP) est suffisante pour induire au moins un tour de recyclage des vésicules, une relance de chargement de 300 points d&#39;accès ou plus est souvent donnée à l&#39;étiquette de la piscine de recyclage des vésicules. Bien que les stimuli de chargement plus intenses, tels que 900 points d&#39;accès, peut permettre à un nombre nominal de vésicules supplémentaire à être étiquetés, ce qui augmente également la quantité de temps, le colorant peut intégrer dans les membranes extracellulaires,...

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measuring exocytosis in neurons using fm labeling

La capacité de mesurer la cinétique de libération des vésicules peut aider à fournir un aperçu de certaines des bases de la neurotransmission. Ici nous avons utilisé l&#39;imagerie en temps réel des vésicules étiquetés avec FM colorant pour surveiller le taux de libération des vésicules présynaptiques. FM4-64 est un colorant fluorescent rouge amphiphiles styryle qui embarque dans les membranes des vésicules synaptiques que l&#39;endocytose est stimulée. Interactions lipophiles provoquer la teinture d&#39;augmenter fortement la fluorescence, émettant ainsi un signal lumineux lorsqu&#39;ils sont associés à des vésicules et un nominal quand dans le liquide extracellulaire. Après une étape de lavage est utilisée pour aider à enlever le colorant externes au sein de la membrane plasmique, le reste FM est concentrée dans les vésicules et est ensuite expulsé lors exocytose induite par un autre tour de la stimulation électrique. Le taux de libération de vésicules est mesurée par la diminution résultant de la fluorescence. Depuis FM colorant peut être appliqué externe et transitoire, il est un outil utile pour déterminer les taux de l&#39;exocytose dans les cultures de neurones, surtout quand on compare les taux entre les synapses et les boutons de contrôle transfectées voisins.

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

Measuring Exocytosis in Neurons Using FM Labeling

La capacité de mesurer la cinétique de libération des vésicules peut aider à fournir un aperçu de certaines des bases de la neurotransmission. Ici nous avons utilisé l&#39;imagerie en temps réel des vésicules étiquetés avec le rouge FM colorant fluorescent 4-64 pour mesurer le taux de libération des vésicules présynaptiques dans les cultures de neurones hippocampiques.

Mesure exocytose dans les neurones en utilisant un étiquetage FM

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

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Biology

19.3K vues.  Harvard. La capacité de mesurer la cinétique de libération des vésicules peut aider à fournir un aperçu de certaines des bases de la neurotransmission. Ici nous avons utilisé l'imagerie en temps réel des vésicules étiquetés avec le rouge FM colorant fluorescent 4-64 pour mesurer le taux de libération des vésicules présynaptiques dans les cultures de neurones hippocampiques.

Vidéo: 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.

L&#39;aide de pincettes laser pour la manipulation neurones isolés 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 ...

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Biologie

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.

Imagerie calcique des neurones corticaux utilisant Fura-2 heures

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.

Mesurer la pression sanguine chez des souris en utilisant enregistrement de la pression du volume, une méthode Queue-coiffe

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.

L&#39;exocytose imagerie dans les cellules bipolaires rétine par microscopie TIRF

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.

Mesurer les flux de calcium dans les cardiomyocytes rapide

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.

Mesure cinétique cellulaire avec la progression du cycle marquage métabolique et cytométrie de flux

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.

Mesure de la fonction respiratoire chez la souris utilisant effrénée pléthysmographie corps entier

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.

Mesure induite glutathion-réponse d&#39;alimentation à Hydra

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.

Mesure de la rigidité de<em&gt; Ex Vivo</em&gt; Souris aortes Utiliser 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 ...

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.

L’imagerie FITC-dextran comme Reporter pour l’exocytose régulée

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