Ce travail a été soutenu par une subvention du NIH décerné à R. Brewster (1R01GM085290-01A1).

1.Microinjection <ol><li> Diluer plasmide codant pour la membrane-cible Green Fluorescent Protein (mGFP, gracieuseté de Richard Harland) à une concentration de 40 ng / ml. L&#39;ADN est préparé par linéarisation plasmide (mGFP/PCS2 +, gracieuseté de Richard Harland) purifiés en utilisant le kit Qiagen Maxi Prep. Ajout de rouge de phénol (dilué 1 / 10 volume total) à la solution est préférable de visualiser la solution. Conserver la solution d&#39;ADN sur la glace. </li><li> En vertu d&#39;un stéréomicroscope, calibrer l&#39;aiguille d&#39;injection (90mm capillaires en verre à partir Narishinge scientifique; Cat n ​​° AN-GD-1) en alignant l&#39;aiguille sur une lame de diplôme et tapotant doucement la partie de l&#39;aiguille dont le diamètre est 1 um en utilisant un rasoir propre lame (pour obtenir un bout de la taille appropriée). </li><li> Pour calibrer l&#39;aiguille pour injecter un volume connu (2NL), connectez l&#39;aiguille de l&#39;appareil micro-injection (PCI 100 microinjecteur; Harvard Apparatus) et avant de remplir l&#39;aiguille avec 0.5μl d&#39;une solution contenant du rouge de phénol et d&#39;eau placé sur un morceau de la paraffine. La solution peut être distribuée en utilisant une pédale. Régler la pression à 7 kPa et de varier le temps (10msec 30msec) pour qu&#39;il prenne 250 pédales à dispenser 0.5μl de la solution. </li><li> Placez délicatement l&#39;aiguille calibrée sur une bande de cire dans un plat humidifié à 100 mm de Pétri. Pour humidifier le plat, placer Kimwipes humides sur les bords (ce qui empêche l&#39;évaporation de la solution dans l&#39;aiguille). </li><li> Utiliser une micropipette pour charger 0,5 1 microlitre d&#39;ADN dans la fin d&#39;aiguille (l&#39;extrémité qui n&#39;était pas étalonnée). </li><li> Une fois la solution a atteint la pointe de l&#39;aiguille, connectez l&#39;extrémité de l&#39;aiguille qui a été utilisé pour le chargement d&#39;un appareil de micro-injection. Réglez les paramètres sur la microinjecteur livrer environ 2 nl / injection à l&#39;aide d&#39;une pédale. </li><li> Ensuite, les embryons de poisson zèbre position au milieu d&#39;une boîte de Pétri remplie de milieu d&#39;embryon (Stock 1,0 ml de Hank # 1, 0,1 ml de bouillon de Hank # 2, Stock 1,0 ml de Hank # 4, 95,9 ml ddH 2 O, Stock 1,0 ml de Hank # 5 , Stock 1,0 ml frais de Hank # 6, Utiliser environ 10 gouttes de NaOH 1 M à pH 7,2). </li><li> Une fois que les embryons se sont développés à l&#39;étape désirée, doucement l&#39;emprise du chorion avec des pinces fines et de situer l&#39;aiguille dans la zone ciblée pour l&#39;injection. Pour une expression large éventail mGFP, injecter de l&#39;ADN dans le jaune ou le cytoplasme au stade 1CELL. Pour restreindre l&#39;expression de mGFP à une plus petite région, injecter de l&#39;ADN à la 4 (ou plus) stade de cellules dans le cytoplasme d&#39;une cellule unique. </li><li> Appuyez sur la pédale de l&#39;appareil de micro-injection et de livrer environ 2 nl, en s&#39;assurant que la solution (visible en raison de l&#39;ajout de rouge de phénol) est correctement injecté. </li><li> Retirer délicatement l&#39;aiguille de l&#39;embryon. </li><li> Répétez l&#39;opération pour tous les embryons dans la boîte de Pétri. </li><li> Autoriser les embryons se développer à 28,5 ° C à l&#39;étape désirée. </li><li> Embryons Dechorionate par un léger décollement du chorion en utilisant une pince fine (si les embryons sont plus jeunes que 24hpf, dechorionating dans un plat en verre de Petri augmente le taux de survie des embryons). </li><li> Retirer chorion vide de la boîte de Pétri à l&#39;aide d&#39;une pipette en verre. </li></ol> 2. Imagerie mGFP marqué sections vibratome <ol><li> Utiliser une pipette en verre pour le transport d&#39;embryons dans fixateur (paraformaldéhyde 4% dans une solution tampon phosphate 1X (PBS). Fix embryons à 4 ° C pendant la nuit. </li><li> Lavez embryons dans du PBS 1X 3 fois pendant 5 minutes chacune. </li><li> Placer les embryons dans une boîte de Pétri et, à l&#39;aide de pinces fines, retirez les jaunes sans abîmer l&#39;embryon. Enlever le jaune autant que possible. </li><li> Préparer 4% d&#39;agarose faible point de fusion de H 2 O. </li><li> Placez agarose fondu dans le moule en plastique de sectionnement. </li><li> Utilisez une pince fine pour soulever un embryon injecté à partir de la boîte de Pétri et de placer et d&#39;orienter dans l&#39;agarose. L&#39;embryon doit être positionné vers une extrémité du moule. </li><li> Une fois que l&#39;agarose est durcie, utilisez un outil de surface plane, comme une spatule, d&#39;enlever le bloc d&#39;agarose du moule. </li><li> Coupez l&#39;extrémité inférieure du bloc d&#39;agarose avec une lame de rasoir afin qu&#39;elle est encore. </li><li> Ajouter une goutte de super glue sur la scène vibratome et placer le bloc d&#39;agarose sur elle (le côté du bloc qui a été stabilisée avec le rasoir doit faire face à la colle). Pour sectionner plusieurs embryons, monter plusieurs agarose-embedded spécimens sur la scène. </li><li> Fixez le stade de la vibratome vibratome (Vibratome, Inc.) </li><li> Définissez les paramètres de l&#39;vibratome (paramètres communs: vitesse-3, épaisseur 50 microns-). Si sectionner plusieurs embryons, aligner des blocs d&#39;agarose sur la scène et de s&#39;assurer que la lame passe à travers l&#39;embryon entier, mais pas toute la largeur des blocs d&#39;agarose (tels que les articles correspondant à l&#39;embryon sera de même rester ensemble). </li><li> Verser du PBS 1X à l&#39;étape de vibratome. </li><li> Section. </li><li> Une fois que les embryons sont sectioNed, retirez le bloc d&#39;agarose (s) de la scène vibratome. </li><li> Soigneusement séparer chaque section par la coupe à l&#39;arrière du bloc d&#39;agarose (s). </li><li> Utilisez une pince fine de sélectionner les sections désiré et le placer dans un PBS 1X rempli multi-même plat (utiliser différents puits pour les sections correspondant aux différents embryons). </li><li> Répétez l&#39;opération pour chaque embryon. </li><li> Procéder à l&#39;étiquetage des anticorps, les étapes 19 mais 23, pour améliorer l&#39;imagerie mGFP (cette étape peut-être pas nécessaire si les articles sont imagés dans un ou deux jours de préparation). </li><li> Traiter sections pendant 30 minutes à température ambiante avec une solution de blocage dans du PBS 1X à bloquer la liaison non spécifique. </li><li> Sections Incuber avec l&#39;anticorps primaire (lapin une-GFP, Invitrogen Cat n ​​° A11122) dilué @ 1:200 dans la solution de blocage sur un agitateur pendant 5 heures à température ambiante ou une nuit à 4 ° C. </li><li> Laver quatre fois (10 min, 15 min, 30 min et 60 min, chaque) dans 1% BSA/1X PBS / 1% solution de DMSO. </li><li> Traiter avec un anticorps secondaire sections appropriées dans une solution de blocage pendant 5 heures à température ambiante ou une nuit à 4 ° C. </li><li> Laver quatre fois (10 min, 15 min, 30 min et 60 min, chaque) dans 1% BSA/1X PBS / 1% solution de DMSO. </li><li> Traiter les sections au DAPI (0,1% en DAPI 50ml PBS 1X; Invitrogen Cat n ​​° D1306) pendant 30 minutes pour colorer les noyaux. </li><li> Laver trois fois dans du PBS 1X pendant 5 minutes chacune. </li><li> Mont sur la diapositive. </li><li> Image en utilisant la microscopie confocale. </li></ol> 3. En direct d&#39;imagerie <ol><li> Dechorionate vivants, d&#39;embryons injectés 30 minutes avant qu&#39;elles n&#39;atteignent le stade désiré de développement. Placer les embryons dans une solution de sédatif (0,01% en moyenne tricaïne embryon) pendant 15 min. </li><li> Déposer une goutte de 1% d&#39;agarose sur une boîte de culture à fond de verre 35mm (MatTek; Partie n ° P35G-o-20-C). </li><li> Autoriser agarose à se solidifier. </li><li> Le tube capillaire sur la flamme de chaleur pour faire trois trous dans agarose (d&#39;autres objets tels que des conseils micropipette peut être utilisé pour faire les trous). Assurez-vous qu&#39;il n&#39;y a aucune résiduelle agarose dans le fond du trou. </li><li> Remplissez la boîte de Pétri avec des moyennes d&#39;embryon. </li><li> Utiliser une pipette en verre pour placer l&#39;embryon dans le trou. </li><li> Ensuite, d&#39;orienter l&#39;embryon à l&#39;aide de pinces fines pour que la zone à imager (tête dorsale) est en contact avec le verre. </li><li> Soigneusement le transport des boîtes de Pétri à la platine du microscope (placer le couvercle sur la boîte de Pétri pour empêcher l&#39;évaporation). </li><li> Alors que l&#39;imagerie, s&#39;assurer que les embryons sont à 28,5 ° C. </li></ol> 4. Résultats Nous décrivons ici une approche simple pour les cellules seule image dans le tube neural de poisson zèbre en utilisant l&#39;expression de la mosaïque. En dépit de la transparence optique de l&#39;embryon de poisson zèbre, nous avons constaté que la visualisation des cellules neurales est grandement améliorée dans les sections obtenues en utilisant un vibratome. Ces sections sont épais (50 mm) permettant à l&#39;imagerie d&#39;extension cellulaire d&#39;une cellule donnée dans les différents plans focaux à l&#39;aide d&#39;un microscope confocal. Résultats utilisant cette méthodologie ont été publiés ailleurs 2, mais des images représentatives de cellules dans le tube neural d&#39;un embryon WT (figure 1) et une N-cadhérine (N-CAD) mutant (figure 2) sont fournis. Ce dernier révèle que les cellules dans les régions latérales de la plaque neurale en N-cad mutants ne parviennent pas à s&#39;orienter correctement vers la ligne médiane (figure 2). Contrairement à cet étiquetage mosaïque, immunomarquage neuroépithélium avec un marqueur de surface cellulaire générales telles que la bêta-caténine ne permet pas la morphologie des cellules individuelles pour être visualisées (figure 3). Imagerie time-lapse d&#39;embryons vivants complété l&#39;étiquetage des échantillons fixés mosaïque, permettant une meilleure compréhension de la dynamique cellulaire qui se déroulent pendant la neurulation. Film 1 a révélé que les cellules WT migrent activement vers la ligne médiane par l&#39;extension des protubérances membranaires médiane orientée. <img src="/files/ftp_upload/1976/1976fig1.jpg" alt="Figure 1" /> Figure 1. MGFP d&#39;expression dans le tube neural d&#39;un embryon de poisson zèbre WT. <img src="/files/ftp_upload/1976/1976fig2.jpg" alt="Figure 2" /> Figure 2. MGFP d&#39;expression dans le tube neural d&#39;un embryon de poisson zèbre N-cadhérine. <img src="/files/ftp_upload/1976/1976fig3.jpg" alt="Figure 3" /> Figure 3. B-caténine coloration immuno du neuroépithélium.

<ol>
	<li>N&#252;sslein-Volhard, C., Dahm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Zebrafish Practical approach series. , (2002).</li><li>Hong, E., Brewster, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=N-cadherin+is+required+for+the+polarized+cell+behaviors+that+drive+neurulation+in+the+zebrafish.">N-cadherin is required for the polarized cell behaviors that drive neurulation in the zebrafish.</a> Development. 133, 3895-3905 (2006).</li><li>Andersen, E., Asuri, N., Clay, M., Halloran, 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+Cell+Motility+and+Actin+Cytoskeleton+of+Individual+Neurons+and+Neural+Crest+Cells+in+Zebrafish+Embryos.">Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos.</a> J Vis Exp. , (2010).</li></ol>

En conclusion, les techniques de marquage décrites ici permettent une analyse simple cellule de processus morphogénétiques dans l&#39;embryon de poisson zèbre. L&#39;accent principal de ce protocole est sur les méthodes d&#39;imagerie des cellules étiquetées dans le tube neural en utilisant mGFP sous le contrôle d&#39;un promoteur ubiquitaire. Pour des applications supplémentaires de ce test d&#39;expression transitoire de lecteurs devraient se référer à un document récent de Andersen et al. 3...

A correction was made to <a href="http://www.jove.com/index/Details.stp?ID=1976">Labeling and Imaging Cells in the Zebrafish Hindbrain</a>. There was an error in the authors affiliations. The authors have been corrected to:
Pradeepa Jayachandran1, Elim Hong2, Rachel Brewster1 1Department of Biological Sciences, University of Maryland, Baltimore County 2Center for Neuroscience, Children's National Medical Center
instead of:
Pradeepa Jayachandran1, Elim Hong2, Rachel Brewster2 1Center for Neuroscience, Children's National Medical Center 2Department of Biological Sciences, University of Maryland Baltimore County

A correction to the author affiliations has been made for the article Labeling and Imaging Cells in the Zebrafish Hindbrain.

Erratum: Labeling and Imaging Cells in the Zebrafish Hindbrain

Solutions <ul><li> Hank Stock # 1: 8,0 g de NaCl, 0,4 g de KCl dans 100 ml H 2 O dd </li><li> Hank Stock # 2: 0,358 g de Na 2 HPO 4 anhydre, 0,60 g de KH 2 PO 4 dans 100 ml d&#39;O 2 ddH </li><li> Hank Stock # 4: 0,72 g de CaCl2 dans 50 ml d&#39;O 2 ddH </li><li> Hank Stock # 5: 1,23 g MgSO 4 x7H 2 O dans 50 ml H 2 O dd </li><li> 1 X de tampon phosphate: 0,8% de NaCl, KCl 0,02%, 0,02 M PO4, pH 7,3 </li><li> Solution de blocage: 2% de sérum de chèvre normal, 1% d&#39;albumine sérique bovine (BSA), diméthylsulfoxyde à 1% (DMSO) </li></ul>

labeling and imaging cells in the zebrafish hindbrain

La clé de la compréhension des processus morphogénétiques qui façonnent l&#39;embryon vertébré précoce est la capacité de cellules d&#39;image à haute résolution. Dans les embryons de poisson zèbre, l&#39;injection des résultats d&#39;ADN plasmidique dans l&#39;expression de la mosaïque, permettant la visualisation de cellules individuelles ou de petits amas de cellules<sup&gt; 1</sup&gt;. Nous décrivons comment l&#39;injection de l&#39;encodage d&#39;ADN plasmidique membrane cible Green Fluorescent Protein (mGFP) sous le contrôle d&#39;un promoteur ubiquitaire peut être utilisé pour les cellules de l&#39;imagerie subissant la neurulation. Au centre de ce protocole est la méthodologie utilisée pour l&#39;imagerie des cellules marquées à haute résolution dans les sections et aussi en temps réel. Ce protocole implique l&#39;injection d&#39;ADN dans des embryons de poisson zèbre mGFP jeunes. Les embryons sont ensuite traitées pour sectionner vibratome, l&#39;étiquetage d&#39;anticorps et d&#39;imagerie avec un microscope confocal. Alternativement, les embryons vivants exprimer mGFP peuvent être imagées par time-lapse microscopie confocale. Nous avons déjà utilisé cette approche simple pour analyser les comportements cellulaires qui déterminent la formation du tube neural dans la région du cerveau postérieur de l&#39;embryon de poisson zèbre<sup&gt; 2</sup&gt;. Les préparatifs fixes a permis pour la visualisation sans précédent des formes de cellules et de l&#39;organisation dans le tube neural, tandis que l&#39;imagerie vivre complété cette approche permettant une meilleure compréhension de la dynamique cellulaire qui se déroulent pendant la neurulation.

Watch this Scientific Journal Video about Labeling and Imaging Cells in the Zebrafish Hindbrain at JoVE.com

Labeling and Imaging Cells in the Zebrafish Hindbrain

La clé de la compréhension des processus morphogénétiques qui façonnent l&#39;embryon précoce est la capacité de cellules d&#39;image à haute résolution. Nous décrivons ici une technique de marquage des cellules individuelles ou de petits amas de cellules dans des embryons de zebrafish entier avec membrane cible protéine fluorescente verte.

L&#39;étiquetage et l&#39;imagerie des cellules dans le cerveau postérieur Zebrafish

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish ...

labeling-and-imaging-cells-in-the-zebrafish-hindbrain

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11.2K vues.  University of Maryland, Baltimore County. La clé de la compréhension des processus morphogénétiques qui façonnent l'embryon précoce est la capacité de cellules d'image à haute résolution. Nous décrivons ici une technique de marquage des cellules individuelles ou de petits amas de cellules dans des embryons de zebrafish entier avec membrane cible protéine fluorescente verte.

Vidéo: Labeling and Imaging Cells in the Zebrafish Hindbrain

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Imagerie time-lapse en direct de relation clonale cellules progénitrices neurales dans le cerveau antérieur développement Zebrafish

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

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Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Imagerie des cellules vivantes des cellules sensorielles de précurseurs d&#39;organes en Intact Drosophile Les chrysalides

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Electroporation of the Hindbrain to Trace Axonal Trajectories and Synaptic Targets in the Chick Embryo

Electroporation of the chick embryonic neural tube has many advantages such as being quick and efficient for the expression of foreign genes into neuronal cells. In this manuscript we provide a method that demonstrates uniquely how to electroporate DNA into the avian hindbrain at E2.75 in order to specifically label a subset of neuronal progenitors, and how to follow their axonal projections and synaptic targets at much advanced stages of development, up to E14.5. We have utilized novel genetic tools including specific enhancer elements, Cre/Lox - based plasmids and the PiggyBac-mediated DNA transposition system to drive GFP expression in a subtype of hindbrain cells (the dorsal most subgroup of interneurons, dA1). Axonal trajectories and targets of dA1 axons are followed at early and late embryonic stages at various brainstem regions. This strategy contributes advanced techniques for targeting cells of interest in the embryonic hindbrain and for tracing circuit formation at multiple stages of development.

How neuronal networks are established in the embryonic brain is a fundamental question in developmental neurobiology. Here we combined an electroporation technique with novel genetic tools, such as Cre/Lox&#x2013;plasmids and PiggyBac-mediated DNA transposition system in the avian hindbrain to label dorsal interneurons and track their axonal projections and synaptic targets at various developmental stages.

Électroporation de la Rhombencéphale tracer des trajectoires axonales et cibles synaptiques dans l&#39;embryon de poulet

Electroporation of  the chick embryonic neural tube has many advantages such as being quick and efficient  for the expression of foreign genes into ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

Étiquetage des cellules isolées dans le système nerveux central des<em&gt; Drosophila melanogaster</em

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

Marquage rétrograde des cellules ganglionnaires de la rétine à l&#39;âge adulte de poisson zèbre avec des colorants fluorescents

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

The Mouse Hindbrain As a Model for Studying Embryonic Neurogenesis

The mouse embryo forebrain is the most commonly employed system for studying mammalian neurogenesis during development. However, the highly folded forebrain neuroepithelium is not amenable to wholemount analysis to examine organ-wide neurogenesis patterns. Moreover, defining the mechanisms of forebrain neurogenesis is not necessarily predictive of neurogenesis in other parts of the brain; for example, due to the presence of forebrain-specific progenitor subtypes. The mouse hindbrain provides an alternative model for studying embryonic neurogenesis that is amenable to wholemount analysis, as well as tissue sections to observe the spatiotemporal distribution and behavior of neural progenitors. Moreover, it is easily dissected for other downstream applications, such as cell isolation or molecular biology analysis. As the mouse hindbrain can be readily analyzed in the vast number of cell lineage reporter and mutant mouse strains that have become available, it offers a powerful model for studying the cellular and molecular mechanisms of developmental neurogenesis in a mammalian organism. Here, we present a simple and quick method to use the mouse embryo hindbrain for analyzing mammalian neural progenitor cell (NPC) behavior in wholemount preparations and tissue sections.

This article demonstrates how the mouse embryonic hindbrain can be used as a model for studying developmental neurogenesis in both whole organ and tissue section preparations.

Le rhombencéphale de souris comme un modèle pour étudier la neurogenèse embryonnaire

The mouse embryo forebrain is the most commonly employed system for studying mammalian neurogenesis during development. However, the highly folded ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

En imagerie calcique Vivo des cellules ciliées ligne latérale chez le poisson zèbre larvaire

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

Identifying Cell Surface Markers of Primary Neural Stem and Progenitor Cells by Metabolic Labeling of Sialoglycan

Neural stem and progenitor cells (NSPCs) are the cellular basis for the complex structures and functions of the brain. They are located in specialized niches in vivo and can be isolated and expanded in vitro, serving as an important resource for cell transplantation to repair brain damage. However, NSPCs are heterogeneous and not clearly defined at the molecular level or purified due to a lack of specific cell surface markers. The protocol presented, which has been previously reported, combines a neural-endothelial co-culture system with a metabolic glycan labeling method to identify the surface sialoglycoproteome of primary NSPCs. The NSPC-endothelial co-culture system allows self-renewal and expansion of primary NSPCs in vitro, generating a sufficient number of NSPCs. Sialoglycans in cultured NSPCs are labeled using an unnatural sialic acid metabolic reporter with bioorthogonal functional groups. By comparing the sialoglycoproteome from self-renewing NSPCs expanded in an endothelial co-culture with differentiating neural culture, we identify a list of membrane proteins that are enriched in NSPCs. In detail, the protocol involves: 1) set-up of an NSPC-endothelial co-culture and NSPC differentiating culture; 2) labeling with azidosugar per-O-acetylated N-azidoacetylmannosamine (Ac4ManNAz); and 3) biotin conjugation to modified sialoglycan for imaging after fixation of neural culture or protein extraction from neural culture for mass spectrometry analysis. Then, the NSPC-enriched surface marker candidates are selected by comparative analysis of mass spectrometry data from both the expanded NSPC and differentiated neural cultures. This protocol is highly sensitive for identifying membrane proteins of low abundance in the starting materials, and it can be applied to marker discovery in other systems with appropriate modifications

Presented here is a protocol that combines an in vitro neural-endothelial co-culture system and metabolic incorporation of sialoglycan with bioorthogonal functional groups to expand primary neural stem and progenitor cells and label their surface sialoglycoproteins for imaging or mass-spectrometry analysis of cell surface markers.

Identification des marqueurs de surface cellulaire des cellules neurales primaires de tige et de progéniture par l'étiquetage métabolique du sialoglycan

Neural stem and progenitor cells (NSPCs) are the cellular basis for the complex structures and functions of the brain. They are located in specialized ...

Labeling and Imaging of Amyloid Plaques in Brain Tissue Using the Natural Polyphenol Curcumin

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). Therefore, detection of the presence of A&#946; in AD brain tissue is a valuable tool for developing new treatments to prevent the progression of AD. Several classical amyloid binding dyes, fluorochrome, imaging probes, and A&#946;-specific antibodies have been used to detect A&#946; histochemically in AD brain tissue. Use of these compounds for A&#946; detection is costly and time consuming. However, because of its intense fluorescent activity, high-affinity, and specificity for A&#946;, as well as structural similarities with traditional amyloid binding dyes, curcumin (Cur) is a promising candidate for labeling and imaging of A&#946; plaques in postmortem brain tissue. It is a natural polyphenol from the herb Curcuma longa. In the present study, Cur was used to histochemically label A&#946; plaques from both a genetic mouse model of 5x familial Alzheimer&#39;s disease (5xFAD) and from human AD tissue within a minute. The labeling capability of Cur was compared to conventional amyloid binding dyes, such as thioflavin-S (Thio-S), Congo red (CR), and Fluoro-jade C (FJC), as well as A&#946;-specific antibodies (6E10 and A11). We observed that Cur is the most inexpensive and quickest way to label and image A&#946; plaques when compared to these conventional dyes and is comparable to A&#946;-specific antibodies. In addition, Cur binds with most A&#946; species, such as oligomers and fibrils. Therefore, Cur could be used as the most cost-effective, simple, and quick fluorochrome detection agent for A&#946; plaques.

Curcumin is an ideal fluorophore for labeling and imaging of amyloid beta protein plaques in brain tissue due to its preferential binding to amyloid beta protein as well as its structural similarities with other traditional amyloid binding dyes. It can be used to label and image amyloid beta protein plaques more efficiently and inexpensively than traditional methods.

Étiquetage et imagerie des plaques amyloïdes dans les tissus cérébraux à l'aide de la curcumine polyphénol naturelle

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Imagerie in vivo de tissu cérébral entièrement actif chez des larves et des juvéniles de poissons-zèbres éveillés par enlèvement du crâne et de la peau

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...