Nous remercions Andreas Schönle, Max-Planck-Institut de chimie biophysique, l&#39;Allemagne et David J. Sandstrom, National Institute of Mental Health, National Institutes of Health, Bethesda, MD, Etats-Unis pour des conseils techniques. Nous tenons à remercier Frank Kötting, European Neuroscience Institute, Göttingen pour la construction de la chambre de l&#39;imagerie et l&#39;appareil anesthésie. Ce travail a été soutenu par des subventions de l&#39;Initiative d&#39;Alzheimer Forschung à TMRYZ a été soutenue par une bourse du Conseil des bourses de Chine, le SBH par une bourse de la Graduate School pour les Neurosciences cellulaires et moléculaires, Université de Tübingen.

A) de l&#39;Assemblée de la chambre de l&#39;imagerie <ol><li> Sélectionnez une larve du stade choisi (par exemple au début 3 ème stade larvaire en laissant un intervalle d&#39;observation d&#39;environ 24 heures à 25 ° C jusqu&#39;à errance scène). </li><li> Rincer la larve à l&#39;eau pour le nettoyer du milieu de culture et de la limande sécher. </li><li> Enduire le centre de l&#39;élément de fond de la chambre, dans la région pour faire face à la larve, avec un mince film d&#39;huile sur les halocarbures. </li><li> Mettez la larve avec la face ventrale face à l&#39;objectif du microscope dans la chambre (permet jonction neuromusculaire (JNM) 26 et 27 à imager) (figure 1 AE). </li><li> Placer l&#39;entretoise en plastique sur la couche d&#39;huile, avec les fentes d&#39;air de la face vers le haut de grille (Figure 1). S&#39;il vous plaît vérifier à cette étape: La hauteur de la cale est à peu près la moitié du diamètre des larves, la largeur de la fente doit être environ deux fois le diamètre de la larve. </li><li> Placez la chambre d&#39;imagerie sur le microscope </li></ol> B) Anesthésie de la larve <ol start="7"><li> Relier les deux entrées de la chambre avec une anesthésie appropriée périphérique / vaporisateur. S&#39;il vous plaît vérifier à l&#39;avance: La concentration efficace de desflurane dans la salle de l&#39;air ne devrait jamais dépasser celle spécifiée par les règlements de sécurité! Seul un très petit volume total de desflurane est nécessaire pour anesthésier les larves. L&#39;application de 15% (v / v) d&#39;une desflurane s&#39;est avéré être un bon point de départ pour déterminer la concentration idéale pour être utilisée dans une expérience. Pour des raisons de sécurité nous suggérons que le vaporisateur ne devrait jamais contenir plus que nécessaire desflurane pour 2-4 heures de l&#39;imagerie in vivo. </li><li> Immerger l&#39;autre extrémité du tube fixé à la sortie de la chambre dans un verre d&#39;eau, puis ouvrir les vannes de régulation du débit de l&#39;anesthésique dans la chambre pendant environ cinq secondes. S&#39;il vous plaît vérifier à cette étape: Ne bulles d&#39;air montent dans l&#39;eau? Si ce n&#39;est pas la chambre est percé. </li><li> Fermer les vannes pendant environ trois secondes. </li><li> Surveiller les mouvements résiduels de larves dans le microscope. Vérifiez la fois le rythme cardiaque et les mouvements musculaires. Si nécessaire soupapes s&#39;ouvrent et se ferment comme décrit dans l&#39;étape 8 et 9 jusqu&#39;à ce que l&#39;anesthésie de la larve est terminée, puis fermez tous les robinets et commencer l&#39;imagerie. S&#39;il vous plaît vérifier à cette étape: Mouvements musculaires résiduelles ou pouls indique que la larve n&#39;est pas correctement anesthésié. Anesthésie complète est vitale pour des images haute résolution. La larve ne devrait normalement pas être anesthésiés pendant plus de 15-20 minutes à une heure donnée. </li></ol> C) Imagerie <ol start="11"><li> Identifier la position correcte et l&#39;image de la structure d&#39;intérêt (figure 2-4). </li></ol> D) La récupération d&#39;une anesthésie <ol start="12"><li> Après l&#39;imagerie est terminée laisser entrer l&#39;air dans la chambre. S&#39;il vous plaît vérifier à cette étape: Vérifiez les contractions musculaires et des battements de cœur de la larve par une lumière halogène transmis. Comme les muscles commencent traitance qu&#39;il est sécuritaire d&#39;enlever la larve de la chambre de l&#39;imagerie. </li><li> Détachez la chambre de l&#39;appareil anesthésie et de le retirer du microscope. Démonter la chambre avec soin et placez la larve dans un milieu de culture de volée écrasée. </li><li> Rangez la vaisselle dans un incubateur à la température appropriée. </li></ol> Séries E) <ol start="15"><li> Répétez la procédure jusqu&#39;à 2-14 points de temps suffisantes ont été obtenues. Alternative protocole: Si la larve est à imager plus d&#39;une fois dans un intervalle de 30 minutes il peut être pratique de le laisser dans la chambre d&#39;imagerie jusqu&#39;au point de temps anesthésie prochaine. Répétez la procédure jusqu&#39;à 7-14 points de temps suffisantes ont été obtenues. S&#39;il vous plaît vérifier à cette étape: La larve est de ne pas être conservé dans la chambre de l&#39;imagerie depuis plus de deux heures à la fois, ni d&#39;être anesthésiés pendant plus de 15-20 minutes à une heure. </li></ol><img src="/files/ftp_upload/2249/2249fig1.jpg" alt="Figure 1" /> La figure 1 de l&#39;Assemblée de la chambre de l&#39;imagerie. (A) Placer la larve et l&#39;entretoise en plastique sur la couche d&#39;huile. (B) Placez un 22 x 22 mm lamelle sur l&#39;entretoise et insérez la bague de guidage en plexiglas dans la chambre, à côté (C) fixer la position de la larve avec l&#39;anneau de métal et (D) à proximité de la chambre. (E) Maintenant, la chambre est prête à être montée sur le microscope. <img src="/files/ftp_upload/2249/2249fig2.jpg" alt="Figure 2" /> Figure 2 Corps-mur muscles de larves de drosophile. Muscles et NMJs ont été visualisées par l&#39;expression d&#39;une protéine de fusion GFP-CD8-Sh 8. Les muscles sont présentés comme observé lors de la focalisation à la face ventrale à travers la cuticule dans la larve. Dans (AC) le muscle le plus superficiel, 27 ans, est montré, dans (KL) les muscles les plus intérieurs, 6 et 7, sont représentés. Barre d&#39;échelle: 100 um, Az entre deux tranches est de 2 um. <img src="/files/ftp_upload/2249/2249fig3.jpg" alt="Figure 3" /> Figure 3 Identité de tégument muscles de larves de drosophile. L&#39;identité des muscles en L3 drosophile larves, A3 segment, est montré. Muscles et NMJs ont été visualisées par l&#39;expression d&#39;une protéine de fusion GFP-CD8-Sh 8. Les muscles sont affichés comme observé lors de la focalisation à la face ventrale à travers la cuticule dans les larves. En AC le muscle le plus superficiel, 27 ans, est montré, dans KL les muscles les plus intérieurs, 6 et 7, sont représentés. Barre d&#39;échelle: 100 um, Az entre deux tranches est de 2 um. <img src="/files/ftp_upload/2249/2249fig4.jpg" alt="Figure 4" /> Figure 4 Identité des jonctions neuromusculaires des larves de drosophile. (AD) NMJs ont été visualisés par expression d&#39;une protéine de fusion DGluRIIA-MRFP 1. Le NMJs sont présentés comme observé lors de la focalisation à la face ventrale à travers la cuticule dans la larve. Dans (AB) le plus superficiel JNM, 27 ans, est montré, dans le CD du NMJs plus intérieure, 6 et 7, sont représentés. Barre d&#39;échelle: 100 um, Az entre deux tranches est de 5 um. (E) et (F) l&#39;identité du superficiel (E) et plus intérieure (F) NMJs est donnée à titre de référence.

<ol>
	<li>Rasse, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutamate+receptor+dynamics+organizing+synapse+formation+in+vivo.">Glutamate receptor dynamics organizing synapse formation in vivo.</a> Nat Neurosci. 8, 898-905 (2005).</li><li>Rasse, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> In vivo imaging of long-term changes in the Drosophila neuromuscular system [dissertation]. , (1988).</li><li>Fuger, P., Behrends, L. B., Mertel, S., Sigrist, S. J., Rasse, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+synapse+development+and+measuring+protein+dynamics+using+two-color+fluorescence+recovery+after+photo-bleaching+at+Drosophila+synapses.">Live imaging of synapse development and measuring protein dynamics using two-color fluorescence recovery after photo-bleaching at Drosophila synapses.</a> Nat Protoc. 2, 3285-3298 (2007).</li><li>Wang, X., Schwarz, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+axonal+transport+of+mitochondria.">Imaging axonal transport of mitochondria.</a> Methods in enzymology. 457, 319-333 (2009).</li><li>Vinegoni, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mesoscopic+fluorescence+tomography+for+in-vivo+imaging+of+developing+Drosophila.">Mesoscopic fluorescence tomography for in-vivo imaging of developing Drosophila.</a> J Vis Exp. , (2009).</li><li>Lin, C. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+imaging+of+Drosophila+larva+by+multiphoton+autofluorescence+and+second+harmonic+generation+microscopy.">Label-free imaging of Drosophila larva by multiphoton autofluorescence and second harmonic generation microscopy.</a> Journal of biomedical optics. 13, 050502-050502 (2008).</li><li>Levitan, E. S., Lanni, F., Shakiryanova, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+imaging+of+vesicle+motion+and+release+at+the+Drosophila+neuromuscular+junction.">In vivo imaging of vesicle motion and release at the Drosophila neuromuscular junction.</a> Nat Protoc. 2, 1117-1125 (2007).</li><li>Zito, K., Parnas, D., Fetter, R. D., Isacoff, E. Y. &. a. m. p. ;. a. m. p., Goodman, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Watching+a+synapse+grow:+noninvasive+confocal+imaging+of+synaptic+growth+in+Drosophila.">Watching a synapse grow: noninvasive confocal imaging of synaptic growth in Drosophila.</a> Neuron. 22, 719-729 (1999).</li></ol>

La méthode présentée a été initialement développé pour étudier les synapses glutamatergiques sur les muscles de la paroi du corps de Drosophila melanogaster larves. La drosophile jonction neuromusculaire (JNM) est caractérisé par une architecture de cyto-stéréotypée des muscles et les neurones et est donc idéal pour l&#39;imagerie in vivo. Cependant, le protocole décrit anesthésie n&#39;est pas limité à l&#39;imagerie de la JNM, la transparence des larves de drosophile facilite l&#39;adaptation du protocole décrit pour étudier le développement des organes, la migration des cellules, le transport de fret axonale et sub-cellulaire de réorganisation au sein des cellules.

Réactifs: <ul><li> Desflurane (Suprane, Baxter, Unterschleißheim, Allemagne) </li><li> D&#39;huile sur les halocarbures (ex Voltalef H10S huile / Atofina, Puteaux, France) </li><li> Fly milieu de culture </li></ul> Equipement: <ul><li> Inversé microscope confocal </li><li> Microscope binoculaire (par exemple Stemi 2000, Carl Zeiss, Jena, Allemagne) </li><li> Petit pinceau comme utilisé pour trier les mouches </li><li> Incubateur </li><li> Personnalisé de chambre construite imagerie (Pour les dessins mécaniques voir réf. 3) </li><li> Vaporisateur / anesthésie appareil (Pour plus de détails voir Réf. 3) </li></ul>

in vivo imaging of intact drosophila larvae at sub-cellular resolution

Des améliorations récentes dans l&#39;imagerie optique, les fluorophores génétiquement codés et des outils génétiques permettant l&#39;établissement efficace de lignées d&#39;animaux transgéniques souhaités ont permis de processus biologiques à étudier dans le contexte d&#39;une vie, et même dans certains cas se comporter, de l&#39;organisme. Dans ce protocole, nous allons décrire comment anesthésier intacte larves de drosophile, en utilisant les anesthésiques volatils desflurane, pour suivre le développement et la plasticité synaptique des populations à une résolution sub-cellulaire 1-3. Alors que d&#39;autres méthodes utiles pour anesthésier les larves de Drosophila melanogaster ont été décrits précédemment 4,5,6,7,8, le protocole présentés ici démontrent des améliorations significatives pour les raisons suivantes combinées principales caractéristiques: (1) Un très haut degré de l&#39;anesthésie, même les battement de coeur est arrêté permettant la résolution latérale de 150 nm 1, (2) un taux de survie élevé des&gt; 90% par cycle anesthésie, permettant l&#39;enregistrement de plus de cinq points de temps sur une période de deux heures à plusieurs jours et ( 3) une sensibilité élevée qui nous permet dans 2 cas à étudier la dynamique des protéines exprimées à des niveaux physiologiques. Dans le détail, nous avons été en mesure de visualiser la sous-unité du récepteur du glutamate postsynaptique GluR-IIA exprimé via le promoteur endogène dans une écurie lignées transgéniques et la ligne de trappe exon FasII-GFP 1. (4) Contrairement aux autres méthodes 4,7 larves peuvent être imagées non seulement vivant, mais aussi intacte (c&#39;est à dire non disséqué) permettant l&#39;observation de se produire sur un nombre de jours 1. Les détails vidéo d&#39;accompagnement de la fonction des différentes parties de la chambre de l&#39;imagerie in vivo 2,3, le montage correct de la larve, la procédure de l&#39;anesthésie, la manière de ré-identifier des postes spécifiques au sein d&#39;une larve et l&#39;élimination sûre des larves de l&#39;imagerie Chambre.

Watch this Scientific Journal Video about In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution at JoVE.com

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Ce protocole décrit une méthode fiable pour l&#39;anesthésie et l&#39;imagerie de intactes<em&gt; Drosophila melanogaster</emLarves&gt;. Nous avons utilisé les anesthésiques volatils desflurane pour permettre l&#39;imagerie répétitive au sub-cellulaire de résolution et de ré-identification des structures pour un maximum de quelques jours<sup&gt; 1</sup&gt;.

L&#39;imagerie in vivo chez la drosophile larves Intact au sub-cellulaire Résolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

in-vivo-imaging-of-intact-drosophila-larvae-at-sub-cellular-resolution

Research

JoVE Journal

Biology

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

14.5K vues.  University of Tübingen. Ce protocole décrit une méthode fiable pour l'anesthésie et l'imagerie de intactes Drosophila melanogaster. Nous avons utilisé les anesthésiques volatils desflurane pour permettre l'imagerie répétitive au sub-cellulaire de résolution et de ré-identification des structures pour un maximum de quelques jours 1.

Vidéo: In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Haute résolution de mesure du comportement Odor-Driven dans les larves de drosophile

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Recherche

Biologie

Labeling hESCs and hMSCs with Iron Oxide Nanoparticles for Non-Invasive in vivo Tracking with MR Imaging

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative medicine. A non-invasive method to monitor the transplanted stem cells repeatedly in vivo would greatly enhance our ability to understand the mechanisms that control stem cell death and identify trophic factors and signaling pathways that improve stem cell engraftment. MR imaging has been proven to be an effective tool for the in vivo depiction of stem cells with near microscopic anatomical resolution. In order to detect stem cells with MR, the cells have to be labeled with cell specific MR contrast agents. For this purpose, iron oxide nanoparticles, such as superparamagnetic iron oxide particles (SPIO), are applied, because of their high sensitivity for cell detection and their excellent biocompatibility. SPIO particles are composed of an iron oxide core and a dextran, carboxydextran or starch coat, and function by creating local field inhomogeneities, that cause a decreased signal on T2-weighted MR images. This presentation will demonstrate techniques for labeling of stem cells with clinically applicable MR contrast agents for subsequent non-invasive in vivo tracking of the labeled cells with MR imaging.

For the evaluation of new stem cell therapies it is important to non-invasively track the injected cells in vivo. This video will show you how to label human mesenchymal and embryonic stem cells with iron oxide based contrast agents in vivo for subsequent MR imaging in vivo.

Étiquetage CSEh et hMSCs d&#39;oxyde de fer nanoparticules pour les non-invasives dans le suivi in ​​vivo avec IRM

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Suivi vidéo haute résolution de la locomotion chez les adultes Drosophila melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Mésoscopique fluorescence tomographie pour In vivo L&#39;imagerie du développement Drosophile</em

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

In vivo d&#39;embryons de drosophile Centrifugation

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Changement de forme d&#39;imagerie cellulaire en vie Drosophile Embryons

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

Microgavage de larves de poisson zèbre

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

Vivre imagerie de dédouanement cellulaire par apoptose au cours<em&gt; Drosophila</em&gt; Embryogenèse

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An ex vivo technique for imaging mouse airway epithelia for quantitative analysis of motile cilia function important for insight into mucociliary clearance function has been established. Freshly harvested mouse trachea is cut longitudinally through the trachealis muscle and mounted in a shallow walled chamber on a glass-bottomed dish. The trachea sample is positioned along its long axis to take advantage of the trachealis muscle to curl longitudinally. This allows imaging of ciliary motion in the profile view along the entire tracheal length. Videos at 200 frames/sec are obtained using differential interference contrast microscopy and a high speed digital camera to allow quantitative analysis of cilia beat frequency and ciliary waveform. With the addition of fluorescent beads during imaging, cilia generated fluid flow also can be determined. The protocol time spans approximately 30 min, with 5 min for chamber preparation, 5-10 min for sample mounting, and 10-15 min for videomicroscopy.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described. This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

Ex méthode in vivo pour l&#39;imagerie haute résolution de la motilité Cilia dans Rongeur épithélium des voies respiratoires

An ex vivo technique for imaging mouse airway epithelia for quantitative  analysis of motile cilia function important for insight into mucociliary  ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Imagerie confocale de simples mitochondriales superoxyde Flashs à coeur Intact ou<em&gt; In Vivo</em

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...