Ce travail a été financé par un ministère de la Défense de l&#39;ère Scholar Award Espoir (n ° W81XWH-05-0396) et chercheur au Pew le Prix des sciences biomédicales à Edward B. III Brown.

1. Aligner le système optique. L&#39;équipement clé pour une expérience MP-FRAP comprennent: un mode bloqué source laser, cellule de Pockels (pour le faisceau de modulation), générateur d&#39;impulsions, dichroïques, objectif, émission de fluorescence du filtre, tube photomultiplicateur fermée, et un système d&#39;enregistrement des données (photon contre et multicanaux scaler). 2. Déterminer les pouvoirs de surveiller en toute sécurité. <ol><li> Atténuer le laser à un faible, mais raisonnable, le pouvoir de générer la fluorescence dans l&#39;échantillon. </li><li> Concentrer au sein de l&#39;échantillon. </li><li> Définissez un compteur de photons à intégrer sur une échelle de temps beaucoup plus long que la récupération de fluorescence prévu. Avec le volume focal maintenue immobile dans l&#39;échantillon (sur notre logiciel de microscope cela est réalisé par un balayage point), d&#39;enregistrer un nombre raisonnable de données de comptage de photons. </li><li> Augmenter la puissance et prendre une autre série de données de comptage de photons. Répétez jusqu&#39;à ce que l&#39;augmentation du nombre de photons semble diminuer de façon significative par rapport à l&#39;augmentation de puissance. </li><li> Terrain log (chiffres de photons) en fonction de log (puissance). Un écart par rapport à une pente de deux indique blanchiment au cours de l&#39;écran. Choisissez comme votre alimentation du moniteur une puissance qui est en dessous de ce coupé, mais qui permet un signal raisonnables par rapport au bruit. </li></ol> 3. Déterminer les paramètres d&#39;entrée au générateur d&#39;impulsions et multicanaux détartreur. <ol><li> Déterminer le dos de l&#39;enveloppe--estimations du coefficient de diffusion et de temps de récupération basé sur la littérature et / ou l&#39;équation de Stokes-Einstein et l&#39;équation de diffusion. </li><li> Réglez les valeurs des paramètres nécessaires sur le générateur d&#39;impulsions et le zoom. <ol class="lalpha"><li> Sur le générateur d&#39;impulsions, régler le temps de pré-blanchiment et de l&#39;eau de Javel. Une bonne série de règles de base est que l&#39;eau de Javel préalable devrait être 1-2 fois plus grand que le temps de récupération moitié prévu, et le temps de l&#39;eau de Javel doit être un dixième de la moitié du temps prévu de récupération. </li><li> Sur le scaler, définir la largeur bin à environ une moitié de la durée de javel, et définir le nombre de bacs par enregistrement tels que le produit de la largeur de votre poubelle et les bacs sélectionnés par enregistrement est supérieure ou égale à environ un rétablissement complet attendu et pré -eau de Javel et parfois l&#39;eau de Javel. </li><li> Retour au générateur d&#39;impulsions et de définir la fréquence de la séquence pre-bleach/bleach/monitor égale à une valeur juste plus petit que l&#39;inverse du produit de l&#39;époque largeur ben le nombre de bacs par enregistrement. * </li><li> Enfin, définissez le nombre de dossiers / scan sur le scaler basée sur l&#39;intensité du signal d&#39;alimentation du moniteur à votre choix. * Il est très important que le temps d&#39;une séquence complète pre-bleach/bleach/monitor (1/frequency) mis sur le générateur d&#39;impulsions est supérieur au temps de la collecte des données (ben x largeur des bacs / enregistrement) mis sur le mesureur. Si ce critère n&#39;est pas atteint, déclenche l&#39;eau de Javel multiples peuvent apparaître dans un dossier unique sur le mesureur. </li></ol></li><li> Réglez l&#39;alimentation du moniteur à un niveau acceptable, préalablement déterminée. Réglez la puissance javel à 200 mW ou plus au-dessus du seuil de blanchissement déterminée lors de l&#39;essai de surveiller le blanchiment. Ces pouvoirs sont tous deux définis comme des tensions différentes à travers le cristal PC. </li><li> Sceau du microscope, éteindre les lumières, allumez le PMT, une numérisation point sur le logiciel de microscope, puis démarrez le générateur d&#39;impulsions et détartreur. </li><li> Analyser la courbe de récupération résultant en traçant la courbe et le marquage trois points importants: 1) la fluorescence pré-blanchiment, 2) la fluorescence immédiatement après l&#39;eau de Javel, et 3) la fluorescence à la fin de la série de données. Utilisez les valeurs de fluorescence avant et immédiatement après l&#39;eau de javel afin de mieux estimer le temps de demi-redressement. </li><li> Avec cette estimation du temps de demi-recouvrement, utiliser les règles de base décrites précédemment pour déterminer de nouvelles valeurs pour le pré-blanchiment, eau de javel, et des durées largeur de bac. En outre, l&#39;utilisation de la fluorescence pré-blanchiment et de fluorescence à la fin de la série de données afin de mieux estimer le temps nécessaire pour parvenir au recouvrement complet. En règle générale, les données doivent être collectées pour une période de temps qui permet la récupération complète pour être signalée pour la dernière moitié des données. </li><li> Réglez les paramètres sur le générateur d&#39;impulsions et détartreur, de prendre une nouvelle courbe, et continuent peaufiner les paramètres jusqu&#39;à ce que votre courbe présente une eau de Javel et la reprise vigoureuse en douceur. Gardez à l&#39;esprit que la durée de blanchiment peut, et doit, être réduit en dessous de la règle de base, si possible. La puissance javel peut aussi être ajustée si elle est trop peu profonde, ou trop profonde, une eau de Javel est atteint. </li></ol> 4. Test de la saturation d&#39;excitation. <ol><li> Utiliser moniteur approprié et les pouvoirs eau de javel, déterminée précédemment, prendre une série de MP-FRAP courbes. Analysez chaque reprise, normalisée à la fluorescence pré-blanchiment, en utilisant le formulaire approprié mathématique et une méthode des moindres carrés non-linéaires algorithme d&#39;ajustement (voir «Analyse des données» ci-dessous). Enregistrer la valeur ajustée de l&#39;eau de Javel paramètre de profondeur.</li><li> Continuez de prendre et d&#39;analyser des séries successives de MP-FRAP courbes à des valeurs croissantes de la puissance javel. </li><li> Log Plot (paramètre de profondeur de Javel) comme une fonction de log (puissance). Tout écart par rapport à une pente de deux indique la saturation d&#39;excitation. Choisissez une puissance qui donne l&#39;eau de Javel une eau de Javel forte, mais ne cause pas de saturation. </li></ol> 5. Continuez de prendre le MP-FRAP courbes. <ol><li> Continuez de prendre des courbes, comme ci-dessus, seront tous les paramètres correctement réglés. </li></ol> 6. Analyse des données. <ol><li> Supprimez les données pré-blanchiment et de l&#39;eau de Javel à partir des données de fluorescence définir et ajuster les données de temps tel que t = 0 correspond à la valeur de fluorescence immédiatement après l&#39;eau de Javel. </li><li> Normaliser la récupération de fluorescence résultant par rapport à la moyenne pré-blanchiment par fluorescence. </li><li> Monter la courbe normalisée en utilisant le modèle mathématique appropriée et d&#39;un algorithme non-linéaire des moindres, des carrés de montage, telles que la fonction lsqcurvefit dans MATLAB. Le modèle présenté ici représente recouvrements de fluorescence à la fois influencé par la diffusion et le flux de convection: <img src="/files/ftp_upload/1636/1636eq.jpg" alt="Équation" /> où F o est la fluorescence pré-blanchiment moyenne, β est le paramètre de profondeur de l&#39;eau de Javel, τ D est la caractéristique de temps de récupération diffusif, R est le rapport du carré de l&#39;axiale (w z) radiale (r w) largeurs de la deux photons volume focal, τ vx = w r / v x, τ vy = w r / v y, τ vz = w z / v z et v 2 = v x 2 + v y 2 + z 2 v est la vitesse de l&#39;écoulement convectif. Pour un écoulement limité au plan d&#39;imagerie, τ → ∞ vz et 1 / τ vx + 1 / τ vy peut être remplacé par une vr / τ. En l&#39;absence d&#39;écoulement convectif, les deux exponentielles dans le numérateur passer à 1 et l&#39;expression est fortement réduite, à seulement deux paramètres d&#39;ajustement, β et τ D. Le coefficient de diffusion est facilement calculé comme D = w D r 2 / 8τ. </li></ol> 7. Des résultats représentatifs. <img src="/files/ftp_upload/1636/1636fig1.jpg" alt="Figure 1" /> Figure 1. Javel et la courbe représentant la récupération pour FITC-BSA. Une courbe présente une bonne récupération eau de Javel et la reprise vigoureuse en douceur, avec la fluorescence à la récupération complète étant signalée pour la dernière moitié des données. <img src="/files/ftp_upload/1636/1636fig2.jpg" alt="Figure 2" /> Figure 2. Courbe de rétablissement normalisé pour FITC-BSA (rouge) et l&#39;associé des moindres carrés (en noir). Cet ajustement donne un coefficient de diffusion de 52,9 UM2 / s, compatible avec la littérature.

<ol>
	<li>Denk, W., Strickler, J. H., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+laser+scanning+fluorescence+microscopy.">Two-photon laser scanning fluorescence microscopy.</a> Biophys J. 60, 73-76 (1990).</li><li>Brown, E. B., Wu, E. S., Zipfel, W., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+molecular+diffusion+in+solution+by+multiphoton+fluorescence+photobleaching+recovery.">Measurement of molecular diffusion in solution by multiphoton fluorescence photobleaching recovery.</a> Biophys J. 77, 2837-2849 (1999).</li><li>Sullivan, K. D., Sipprell, W. H. B. r. o. w. n., Jr, E. B., Brown, E. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+model+of+fluorescence+recovery+expands+the+application+of+multiphoton+fluorescence+recover+after+photobleaching+in+vivo.">Improved model of fluorescence recovery expands the application of multiphoton fluorescence recover after photobleaching in vivo.</a> Biophys J. 96, 5082-5094 (2009).</li></ol>

La puissance du multi-photons de recouvrement de fluorescence après photoblanchiment réside dans sa capacité à sonder des échantillons épais avec une résolution 3D. Depuis son développement dans les années 1990, MP-FRAP a été utilisée pour déterminer le coefficient de diffusion (ou les paramètres de transport analogue) dans les corps cellulaires, ex vivo des tissus tranches épaisses, et dans les tissus in vivo et l&#39;interstitium. Dans cet article, nous avons présenté l&#39;...

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<td>Ti:sapphire laser</td>
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measuring diffusion coefficients via two-photon fluorescence recovery after photobleaching

Multi-fluorescence après photoblanchiment de récupération est une technique de microscopie utilisée pour mesurer le coefficient de diffusion (ou les paramètres de transport analogues) des macromolécules, et peut être appliqué à la fois<em&gt; In vitro</em&gt; Et<em&gt; In vivo</em&gt; Les systèmes biologiques. Multi-fluorescence après photoblanchiment de récupération est effectuée par photoblanchiment une région d&#39;intérêt dans un échantillon fluorescent à l&#39;aide d&#39;un flash laser intense, puis d&#39;atténuer le faisceau et le suivi de la fluorescence comme encore fluorescentes molécules de l&#39;extérieur de la région de diffusion intérêt à remplacer les molécules photobleached . Nous commencerons notre démonstration en alignant le faisceau laser à travers la cellule de Pockels (laser modulateur) et le long du chemin optique à travers la boîte de balayage laser et la lentille de l&#39;objectif de l&#39;échantillon. Pour plus de simplicité, nous utiliserons un échantillon de colorant fluorescent aqueuse. Nous allons ensuite déterminer les paramètres appropriés expérimental pour notre échantillon, y compris, de surveiller et de blanchiment des pouvoirs, la durée de javel, des largeurs de bac (pour le comptage de photons), et le temps de récupération de fluorescence. Ensuite, nous allons décrire la procédure à suivre pour prendre les courbes de récupération, un processus qui peut être largement automatisée par le biais de LabVIEW (National Instruments, Austin, TX) pour un débit amélioré. Enfin, le coefficient de diffusion est déterminé en ajustant les données de récupération pour le modèle mathématique approprié en utilisant un algorithme d&#39;ajustement des moindres carrés, facilement programmable à l&#39;aide de logiciels tels que MATLAB (The Mathworks, Natick, MA).

Watch this Scientific Journal Video about Measuring Diffusion Coefficients via Two-photon Fluorescence Recovery After Photobleaching at JoVE.com

Measuring Diffusion Coefficients via Two-photon Fluorescence Recovery After Photobleaching

Dans cet article nous allons décrire la procédure de mesure de coefficients de diffusion multi-photons en utilisant la récupération de fluorescence après photoblanchiment. Nous allons commencer par aligner le laser le long du chemin optique à l&#39;échantillon et la détermination des paramètres expérimentaux appropriés, puis continuer à générer et enfin l&#39;ajustement de courbes de recouvrement de fluorescence.

Mesure des coefficients de diffusion par deux photons de récupération de fluorescence après photoblanchiment

Multi-fluorescence recovery after photobleaching is a microscopy technique used to measure the diffusion coefficient (or analogous transport parameters) ...

measuring-diffusion-coefficients-via-two-photon-fluorescence-recovery

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JoVE Journal

Biology

11.1K vues.  University of Rochester. Dans cet article nous allons décrire la procédure de mesure de coefficients de diffusion multi-photons en utilisant la récupération de fluorescence après photoblanchiment. Nous allons commencer par aligner le laser le long du chemin optique à l'échantillon et la détermination des paramètres expérimentaux appropriés, puis continuer à générer et enfin l'ajustement de courbes de recouvrement de fluorescence.

Vidéo: Measuring Diffusion Coefficients via Two-photon Fluorescence Recovery After Photobleaching

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Deux photons axotomie et time-lapse imagerie confocale dans embryons de poissons zèbres vivent

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Recherche

Biologie

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Imagerie time-lapse de la mitose après transfection de siRNA

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Deux-Photon-Based photoactivation dans Live embryons de poissons zèbres

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

Dans la quantification in vivo des interactions des protéines G récepteurs couplés utilisant résolution spectrale microscopie à deux photons

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

Photobleaching Assays (FRAP &amp; FLIP) to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with good spatial and temporal resolution. Here we describe how to perform FRAP and FLIP assays of chromatin proteins, including H1 and HP1, in mouse embryonic stem (ES) cells. In a FRAP experiment, cells are transfected, either transiently or stably, with a protein of interest fused with the green fluorescent protein (GFP) or derivatives thereof (YFP, CFP, Cherry, etc.). In the transfected, fluorescing cells, an intense focused laser beam bleaches a relatively small region of interest (ROI). The laser wavelength is selected according to the fluorescent protein used for fusion. The laser light irreversibly bleaches the fluorescent signal of molecules in the ROI and, immediately following bleaching, the recovery of the fluorescent signal in the bleached area - mediated by the replacement of the bleached molecules with the unbleached molecules - is monitored using time lapse imaging. The generated fluorescence recovery curves provide information on the protein's mobility. If the fluorescent molecules are immobile, no fluorescence recovery will be observed. In a complementary approach, Fluorescence Loss in Photobleaching (FLIP), the laser beam bleaches the same spot repeatedly and the signal intensity is measured elsewhere in the fluorescing cell. FLIP experiments therefore measure signal decay rather than fluorescence recovery and are useful to determine protein mobility as well as protein shuttling between cellular compartments. Transient binding is a common property of chromatin-associated proteins. Although the major fraction of each chromatin protein is bound to chromatin at any given moment at steady state, the binding is transient and most chromatin proteins have a high turnover on chromatin, with a residence time in the order of seconds. These properties are crucial for generating high plasticity in genome expression1. Photobleaching experiments are therefore particularly useful to determine chromatin plasticity using GFP-fusion versions of chromatin structural proteins, especially in ES cells, where the dynamic exchange of chromatin proteins (including heterochromatin protein 1 (HP1), linker histone H1 and core histones) is higher than in differentiated cells2,3.

Photobleaching Assays FRAP &amp; FLIP to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

We describe photobleaching methods including Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) to monitor chromatin protein dynamics in embryonic stem (ES) cells. Chromatin protein dynamics, which is considered to be one of the means to study chromatin plasticity, is enhanced in pluripotent cells.

Photobleaching essais (FRAP et FLIP) pour mesurer la dynamique des protéines de la chromatine dans Vivre cellules souches embryonnaires

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with ...

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing (MTT)

Our goal is to obtain a comprehensive description of molecular processes occurring at cellular membranes in different biological functions. We aim at characterizing the complex organization and dynamics of the plasma membrane at single-molecule level, by developing analytic tools dedicated to Single-Particle Tracking (SPT) at high density: Multiple-Target Tracing (MTT)1. Single-molecule videomicroscopy, offering millisecond and nanometric resolution1-11, allows a detailed representation of membrane organization12-14 by accurately mapping descriptors such as cell receptors localization, mobility, confinement or interactions.
We revisited SPT, both experimentally and algorithmically. Experimental aspects included optimizing setup and cell labeling, with a particular emphasis on reaching the highest possible labeling density, in order to provide a dynamic snapshot of molecular dynamics as it occurs within the membrane. Algorithmic issues concerned each step used for rebuilding trajectories: peaks detection, estimation and reconnection, addressed by specific tools from image analysis15,16. Implementing deflation after detection allows rescuing peaks initially hidden by neighboring, stronger peaks. Of note, improving detection directly impacts reconnection, by reducing gaps within trajectories. Performances have been evaluated using Monte-Carlo simulations for various labeling density and noise values, which typically represent the two major limitations for parallel measurements at high spatiotemporal resolution.
The nanometric accuracy17 obtained for single molecules, using either successive on/off photoswitching or non-linear optics, can deliver exhaustive observations. This is the basis of nanoscopy methods17 such as STORM18, PALM19,20, RESOLFT21 or STED22,23, which may often require imaging fixed samples. The central task is the detection and estimation of diffraction-limited peaks emanating from single-molecules. Hence, providing adequate assumptions such as handling a constant positional accuracy instead of Brownian motion, MTT is straightforwardly suited for nanoscopic analyses. Furthermore, MTT can fundamentally be used at any scale: not only for molecules, but also for cells or animals, for instance. Hence, MTT is a powerful tracking algorithm that finds applications at molecular and cellular scales.

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing MTT

Multiple-Target Tracing is a homemade algorithm developed for tracking individually labeled molecules within the plasma membrane of living cells. Efficiently detecting, estimating and tracing molecules over time at high-density provide a user-friendly, comprehensive tool to investigate nanoscale membrane dynamics.

Cartographie Diffusion moléculaire dans la membrane plasmique par multiple-cible de Recherches (MTT)

Our goal is to obtain a comprehensive description of molecular processes occurring at cellular membranes in different biological functions. We aim at ...

Mouse Sperm Cryopreservation and Recovery using the I&middot;Cryo Kit

Thousands of new genetically modified (GM) strains of mice have been created since the advent of transgenesis and knockout technologies. Many of these valuable animals exist only as live animals, with no backup plan in case of emergency. Cryopreservation of embryos can provide this backup, but is costly, can be a lengthy procedure, and generally requires a large number of animals for success. Since the discovery that mouse sperm can be successfully cryopreserved with a basic cryoprotective agent (CPA) consisting of 18&#37; raffinose and 3&#37; skim milk, sperm cryopreservation has become an acceptable and cost-effective procedure for archiving, distributing and recovery of these valuable strains.
 Here we demonstrate a newly developed I&bull;Cryo kit for mouse sperm cryopreservation. Sperm from five commonly-used strains of inbred mice were frozen using this kit and then recovered. Higher protection ratios of sperm motility (&gt; 60&#37;) and rapid progressive motility (&gt; 45&#37;) compared to the control (basic CPA) were seen for sperm frozen with this kit in 5 inbred mouse strains. Two cell stage embryo development after IVF with the recovered sperm was improved consistently in all 5 mouse strains examined. Over a 1.5 year period, 49 GM mouse lines were archived by sperm cryopreservation with the I&bull;Cryo kit and later recovered by IVF.

Mouse Sperm Cryopreservation and Recovery using the I&amp;middot;Cryo Kit

Here we demonstrate the newly developed I&#x2022;Cryo kit for mouse sperm cryopreservation. Two-cell stage embryo development with frozen-thawed sperm was improved consistently in 5 mouse strains with the use of this kit. Over a 1.5 year period, 49 genetically modified mouse lines were archived by sperm cryopreservation with the I&#x2022;Cryo kit and later successfully recovered by IVF.

Cryoconservation du sperme de souris et de récupération utilisant les I · Cryo Kit

Thousands of new genetically modified (GM) strains of mice have been created since the advent of transgenesis and knockout technologies.  Many of these ...

Easy Measurement of Diffusion Coefficients of EGFP-tagged Plasma Membrane Proteins Using k-Space Image Correlation Spectroscopy

Lateral diffusion and compartmentalization of plasma membrane proteins are tightly regulated in cells and thus, studying these processes will reveal new insights to plasma membrane protein function and regulation. Recently, k-Space Image Correlation Spectroscopy (kICS)1&#160;was developed to enable routine measurements of diffusion coefficients directly from images of fluorescently tagged plasma membrane proteins, that avoided systematic biases introduced by probe photophysics. Although the theoretical basis for the analysis is complex, the method can be implemented by nonexperts using a freely available code to measure diffusion coefficients of proteins. kICS calculates a time correlation function from a fluorescence microscopy image stack after Fourier transformation of each image to reciprocal (k-) space. Subsequently, circular averaging, natural logarithm transform and linear fits to the correlation function yields the diffusion coefficient. This paper provides a step-by-step guide to the image analysis and measurement of diffusion coefficients via kICS.
First, a high frame rate image sequence of a fluorescently labeled plasma membrane protein is acquired using a fluorescence microscope. Then, a region of interest (ROI) avoiding intracellular organelles, moving vesicles or protruding membrane regions is selected. The ROI stack is imported into a freely available code and several defined parameters (see Method section) are set for kICS analysis. The program then generates a &#34;slope of slopes&#34; plot from the k-space time correlation functions, and the diffusion coefficient is calculated from the slope of the plot. Below is a step-by-step kICS procedure to measure the diffusion coefficient of a membrane protein using the renal water channel aquaporin-3 tagged with EGFP as a canonical example.

This paper provides a step by step guide to the fluctuation analysis technique k-Space Image Correlation Spectroscopy (kICS) for measuring diffusion coefficients of fluorescently labeled plasma membrane proteins in live mammalian cells.

Facile mesure de coefficients de diffusion de EGFP-marqués plasma protéines membranaires en utilisant K-image de l&#39;espace spectroscopie de corrélation

Lateral diffusion and compartmentalization of plasma membrane proteins are tightly regulated in cells and thus, studying these processes will reveal new ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Recouvrement des adultes coeurs de poisson zèbre à haut débit pour des applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on changes in intracellular calcium ([Ca2+]i) and nitric oxide (NO). In order to understand how [Ca2+]i, NO and downstream molecules are handled by a blood vessel in response to vasoconstrictors and vasodilators, we developed a novel technique that applies calcium-labeling (or NO-labeling) dyes with two photon microscopy to measure calcium handling (or NO production) in isolated blood vessels. Described here is a detailed step-by-step procedure that demonstrates how to isolate an aorta from a rat, label calcium or NO within the endothelial or smooth muscle cells, and image calcium transients (or NO production) using a two photon microscope following physiological or pharmacological stimuli. The benefits of using the method are multi-fold: 1) it is possible to simultaneously measure calcium transients in both endothelial cells and smooth muscle cells in response to different stimuli; 2) it allows one to image endothelial cells and smooth muscle cells in their native setting; 3) this method is very sensitive to intracellular calcium or NO changes and generates high resolution images for precise measurements; and 4) described approach can be applied to the measurement of other molecules, such as reactive oxygen species. In summary, application of two photon laser emission microscopy to monitor calcium transients and NO production in the endothelial and smooth muscle cells of an isolated blood vessel has provided high quality quantitative data and promoted our understanding of the mechanisms regulating vascular function.

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Vascular cell functiondepends on activity of intracellular messengers. Described here is an ex vivo two photon imaging method that allows the measurement of intracellular calcium and nitric oxide levels in response to physiological and pharmacological stimuli in individual endothelial and smooth muscle cells of an isolated aorta.

Deux photons imagerie intracellulaire de Ca<sup&gt; 2+</sup&gt; Manipulation et production d&#39;oxyde nitrique endothélial et en cellules de muscle lisse d&#39;un Rat aorte isolée

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on ...