Diese Arbeit wurde durch ein NIH Zuschuss für R. Brewster (1R01GM085290-01A1) unterstützt.

1.Microinjection <ol><li> Verdünnen Plasmid kodiert Membran gezielte Green Fluorescent Protein (mGFP, mit freundlicher Genehmigung von Richard Harland) zu einer Konzentration von 40 ng / ml. DNA wird durch Linearisierung Plasmid (mGFP/PCS2 +, mit freundlicher Genehmigung von Richard Harland) unter Verwendung des Qiagen Maxi Prep Kit vorbereitet. Die Zugabe von Phenolrot (verwässert 1 / 10 Volumen), um die Lösung bevorzugt, die Lösung zu visualisieren. Keep DNA-Lösung auf Eis. </li><li> Unter einem Stereomikroskop, kalibrieren Sie die Injektionsnadel (90mm Glaskapillaren aus Narishinge Scientific; Kat.-Nr. NA-GD-1), indem Sie die Nadel auf einer abgestuften Rutsche und leichtes Antippen der Teil der Nadel, wo der Durchmesser beträgt 1 &amp;mgr; mit einer Rasierklinge Klinge (ein Tipp von der richtigen Größe zu erhalten). </li><li> Um die Nadel zu kalibrieren, um ein bekanntes Volumen (2nl) spritzen, verbinden Sie die Nadel auf die Mikroinjektion Geräten (PCI 100 Mikroinjektor; Harvard Apparatus) und Front füllen Sie die Nadel mit 0.5μl einer Lösung mit Phenolrot und Wasser auf ein Stück platziert von Paraffin. Die Lösung kann verzichtet mit einem Fußpedal werden. Stellen Sie den Druck bis 7 kPa und variieren Sie die Zeit (10msec 30msec), so dass es 250 Pedale nimmt zu verzichten 0.5μl der Lösung. </li><li> Legen Sie das kalibrierte Nadel auf einem Streifen von Wachs in einem befeuchteten 100 mm Petrischale. Um befeuchten die Schüssel legen nassen Kimwipes an den Rändern (dies verhindert die Verdunstung der Lösung in die Nadel). </li><li> Verwenden einer Mikropipette zu 0,5-1μl der DNA in die Nadel Ende (das Ende, das nicht kalibriert wurde) zu laden. </li><li> Sobald die Lösung der Nadelspitze erreicht hat, verbinden Sie das Ende der Nadel, die für das Laden einer Mikroinjektion Apparat verwendet wurde. Passen Sie die Einstellungen auf dem Mikroinjektor auf ca. 2 nl / Injektion mit einem Fußpedal zu liefern. </li><li> Als nächstes Position Zebrafischembryonen in der Mitte einer Petrischale mit Embryo Medium (1,0 ml Hanks Stock # 1, 0,1 ml Hanks Lager Nr. 2, 1,0 ml Hanks Stock # 4, 95,9 ml ddH 2 O, 1,0 ml Hanks Stock # 5 gefüllt , 1,0 ml frisches Hanks Stock # 6, Verwenden ca. 10 Tropfen 1 M NaOH auf pH 7,2). </li><li> Sobald die Embryonen auf die gewünschte Stufe entwickelt haben, vorsichtig Griff das Chorion mit feinen Pinzetten und situieren die Nadel in den Bereich für die Injektion gezielt. Für breite Palette mGFP Ausdruck, injizieren DNA in den Dotter oder das Zytoplasma der 1CELL Bühne. Um Ausdruck mGFP auf eine kleinere Region beschränken, injizieren DNA an die 4 (oder mehr) Zellen der Bühne in das Zytoplasma einer einzelnen Zelle. </li><li> Drücken Sie das Fußpedal der Mikroinjektion Apparate und liefern ca. 2 nl, sicherzustellen, dass die Lösung (sichtbar durch die Zugabe von Phenolrot) richtig gespritzt wird. </li><li> Entfernen Sie vorsichtig die Nadel aus dem Embryo. </li><li> Wiederholen Sie dies für alle Embryonen in der Petrischale. </li><li> Lassen Embryonen bei 28,5 ° C zu entwickeln, um gewünschte Bühne. </li><li> Dechorionate Embryonen durch leichtes Abziehen des Chorion mit feinen Pinzette (wenn Embryonen jünger als 24hpf sind, dechorionating in einem Glas-Petrischale erhöht die Überlebensrate der Embryonen). </li><li> Entfernen Sie leere Chorion aus der Petrischale mit einer Glaspipette. </li></ol> 2. Imaging mGFP-markierten Vibratom Abschnitte <ol><li> Verwenden Sie eine Glaspipette an Embryonen in Fixativ (4% Paraformaldehyd in 1X Phosphat-Pufferlösung (PBS) zu transportieren. Fix Embryonen bei 4 ° C über Nacht. </li><li> Wash Embryonen in 1X PBS 3 mal für jeweils 5 Minuten. </li><li> Legen Sie die Embryonen in Petrischalen und mit feinen Pinzetten, entfernen Sie das Eigelb ohne Beschädigung des Embryos. Entfernen Sie so viel wie möglich Eigelb. </li><li> Bereiten Sie 4% Low-Melt Agarose in H 2 O. </li><li> Legen geschmolzene Agarose in Kunststoff Schneiden Form. </li><li> Verwenden feinen Pinzette zu einem injizierten Embryo aus der Petrischale heben und zu platzieren und zu orientieren in der Agarose. Der Embryo muss zu einem Ende der Form positioniert werden. </li><li> Sobald die Agarose ausgehärtet ist, verwenden Sie eine ebene Fläche Werkzeug, wie einem Spatel, bis die Agarose Block von der Form zu entfernen. </li><li> Schneiden Sie das untere Ende des Agarose-Block mit einer Rasierklinge, so dass es gerade ist. </li><li> Fügen Sie einen Tropfen Superkleber auf die Vibratom Bühne und platzieren Sie die Agarose-Block auf sie (die Seite des Blocks, die aus mit dem Rasiermesser wurde geebnet sollte der Kleber face). Für Schnitte mehrere Embryonen, mount mehrere Agarose eingebettete Proben auf der Bühne. </li><li> Bringen Sie die Vibratom Bühne, um die Vibratom (Vibratom, Inc.). </li><li> Stellen Sie die Parameter des Vibratom (allgemeine Einstellungen: speed-3, Dicke-50 um). Wenn Schnitte mehrere Embryonen, richten Agarose Blöcke auf der Bühne und sicherzustellen, dass die Klinge durch den ganzen Embryo geht aber nicht über die gesamte Breite der Agarose-Blöcke (wie die Abschnitte, die dem gleichen Embryo zusammen bleiben). </li><li> Gießen 1X PBS in der Vibratom Bühne. </li><li> Section. </li><li> Sobald die Embryonen Sectioned, entfernen Sie die Agarose-Block (s) aus dem Vibratom Bühne. </li><li> Sorgfältig trennen Sie die einzelnen Kapitel, indem die Rückseite der Agarose-Block (s). </li><li> Verwenden feinen Pinzette gewünschten Abschnitte und in einen 1X PBS gefüllten Multi-Well-Platte (mit unterschiedlichen Bohrungen für die Abschnitte, die den verschiedenen Embryonen) zu wählen. </li><li> Wiederholen Sie für jeden Embryo. </li><li> Fahren Sie mit Antikörpermarkierung, wenn die Schritte 19 23, zu mGFP Bildgebung (dieser Schritt nicht notwendig sein, wenn Abschnitte innerhalb eines Tages oder zwei der Vorbereitung sind abgebildet) zu verbessern. </li><li> Gönnen Abschnitte für 30 Minuten bei Raumtemperatur mit Blockierungslösung in 1X PBS, um unspezifische Bindung zu blockieren. </li><li> Inkubieren Abschnitte mit primärem Antikörper (Kaninchen a-GFP, Invitrogen Kat. Nr. A11122) verdünnt @ 1:200 in Blocking-Lösung auf einem Schüttler für 5 Stunden bei Raumtemperatur oder über Nacht bei 4 ° C. </li><li> Wash viermal (10 min, 15 min, 30 min und 60 min, jeweils) in 1% BSA/1X PBS / 1% DMSO-Lösung. </li><li> Gönnen Abschnitte mit entsprechenden Sekundär-Antikörper in Blocking-Lösung für 5 Stunden bei Raumtemperatur oder über Nacht bei 4 ° C. </li><li> Wash viermal (10 min, 15 min, 30 min und 60 min, jeweils) in 1% BSA/1X PBS / 1% DMSO-Lösung. </li><li> Behandeln Sie die Abschnitte mit DAPI (0,1% DAPI in 50 ml 1X PBS, Invitrogen Kat. Nr. D1306) für 30 Minuten, um die Kerne zu färben. </li><li> Dreimal in 1X PBS für jeweils 5 Minuten. </li><li> Berg auf dem Objektträger. </li><li> Bild mit Hilfe der konfokalen Mikroskopie. </li></ol> 3. Live-Bildgebung <ol><li> Dechorionate leben, injizierten Embryonen 30 Minuten, bevor sie das gewünschte Entwicklungsstadium. Legen Sie die Embryonen in sedative Lösung (0,01% Tricaine in Embryo Medium) für 15 min. </li><li> Geben Sie einen Tropfen 1% Agarose auf einem 35mm Glasboden Kulturschale (MatTek; Art.-Nr. P35G-o-20-C). </li><li> Lassen Agarose verfestigt. </li><li> Hitze Kapillare über Flamme 3 Löcher in Agarose (andere Objekte wie Mikropipettenspitzen verwendet werden, um die Löcher zu machen) zu machen. Stellen Sie sicher, dass es keine Rest-Agarose in das Loch unten. </li><li> Füllen Sie die Petrischale mit Embryo Medium. </li><li> Verwenden Sie eine Glaspipette, den Embryo in das Loch Platz. </li><li> Als nächstes Orientierung des Embryos mit feinen Pinzetten, so dass die Fläche abgebildet werden (dorsal Kopf) ist in Kontakt mit dem Glas. </li><li> Vorsichtig transportieren die Petrischale auf dem Mikroskoptisch (legen Sie den Deckel über der Petrischale, um Verdunstung zu verhindern). </li><li> Während Bildgebung, sicherzustellen, dass die Embryonen bei 28,5 ° C liegen </li></ol> 4. Ergebnisse Wir beschreiben hier eine einfache Lösung, um Bild Einzelzellen im Zebrafisch Neuralrohr mit Mosaik Ausdruck. Trotz der optischen Transparenz des Zebrafisch-Embryos, haben wir festgestellt, dass die Visualisierung von Nervenzellen stark in Querschnitten unter Verwendung eines Vibratom verbessert. Diese Abschnitte sind dick (50 mm) ermöglicht für die Bildgebung von Zell-Erweiterung aus einer Zelle in verschiedenen Brennebenen mit einem konfokalen Mikroskop. Ergebnisse mit dieser Methode wurden an anderer Stelle 2, veröffentlicht, aber repräsentative Bilder von Zellen in das Neuralrohr eines WT Embryo (Abb. 1) und ein N-Cadherin (N-cad)-Mutante (Abbildung 2) sind vorhanden. Letzteres zeigt, dass Zellen in seitlichen Bereichen der Neuralplatte in N-cad-Mutanten nicht richtig zu orientieren in Richtung der Mittellinie (Abbildung 2). Im Gegensatz zu diesem Mosaik Etikettierung ist Immunfärbung der Neuroepithel mit einer allgemeinen Zelloberflächenmarker wie Beta-Catenin nicht zulassen, dass die Morphologie der einzelnen Zellen visualisiert werden (Abb. 3). Zeitraffer-Aufnahme von Live-Embryonen ergänzt das Mosaik Kennzeichnung von festen Proben und ermöglicht ein besseres Verständnis der zellulären Dynamik, die während Neurulation. Movie 1 ergab, dass WT-Zellen aktiv an der Mittellinie wandern durch die Ausweitung medial orientierten Membran Vorsprünge. <img src="/files/ftp_upload/1976/1976fig1.jpg" alt="Abbildung 1" /> Abbildung 1. MGFP Ausdruck in das Neuralrohr eines WT Zebrafischembryo. <img src="/files/ftp_upload/1976/1976fig2.jpg" alt="Abbildung 2" /> Abbildung 2. MGFP Ausdruck in das Neuralrohr eines N-Cadherin Zebrafischembryo. <img src="/files/ftp_upload/1976/1976fig3.jpg" alt="Abbildung 3" /> Abbildung 3. B-Catenin Immun-Färbung der Neuroepithel.

<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>

Abschließend beschrieb die Kennzeichnung Techniken hier erlauben einzelne Zelle Analyse von morphogenetischen Prozesse im Zebrafisch Embryo. Der primäre Schwerpunkt dieses Protokoll wird auf Methoden zur Darstellung von markierten Zellen in das Neuralrohr mit mGFP unter der Kontrolle eines ubiquitären Promoters. Weitere Anwendungen dieser transiente Expression Assays sollte sich der Leser zu einer neueren Arbeit von Andersen et al verweisen. 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

Lösungen <ul><li> Hank Stock # 1: 8,0 g NaCl, 0,4 g KCl in 100 ml dd H 2 O </li><li> Hank Stock # 2: 0,358 g Na 2 HPO 4 Wasserfreies, 0,60 g KH 2 PO 4 in 100 ml ddH 2 O </li><li> Hank Stock # 4: 0,72 g CaCl 2 in 50 ml ddH 2 O </li><li> Hank Stock # 5: 1,23 g MgSO 4 x7H 2 O in 50 ml dd H 2 O </li><li> 1 X-Phosphat-Puffer: 0,8% NaCl, 0,02% KCl, 0,02 M PO4, pH 7,3 </li><li> Blocking Solution: 2% normalem Ziegenserum, 1% Rinderserumalbumin (BSA), 1% Dimethylsulfoxid (DMSO) </li></ul>

labeling and imaging cells in the zebrafish hindbrain

Schlüssel zum Verständnis der morphogenetischen Prozesse, die die frühen Wirbeltierembryo Form ist die Möglichkeit, Bild-Zellen in hoher Auflösung. In Zebrafisch-Embryos, Injektion von Plasmid-DNA führt zu Mosaik-Ausdruck, so dass für die Visualisierung von einzelnen Zellen oder kleine Gruppen von Zellen<sup&gt; 1</sup&gt;. Wir beschreiben, wie Injektion von Plasmid-DNA, die Membran-bezogene Green Fluorescent Protein (mGFP) unter der Kontrolle eines ubiquitären Promotor für Imaging-Zellen, die eine Neurulation eingesetzt werden kann. Im Mittelpunkt dieses Protokolls ist die Methode für die Bildgebung markierten Zellen mit hoher Auflösung in Abschnitte und auch in Echtzeit. Das Protokoll beinhaltet die Injektion von DNA in mGFP junge Zebrabärblinge. Die Embryonen werden dann für Vibratom Schneiden, Antikörpermarkierung und Bildgebung mit einem konfokalen Mikroskop verarbeitet. Alternativ können lebende Embryonen zum Ausdruck mGFP aufgenommen mit Zeitraffer-konfokalen Mikroskopie werden. Wir haben früher dieser naive Ansatz verwendet, um die zelluläre Verhaltensweisen, Neuralrohr Bildung Laufwerk im Hinterhirn Region Zebrafischembryonen analysieren<sup&gt; 2</sup&gt;. Die feste Vorbereitungen für beispiellose Visualisierung von Zellformen und Organisation in das Neuralrohr erlaubt während der Live-Bildgebung dieser Ansatz ermöglicht ein besseres Verständnis der zellulären Dynamik, die während Neurulation ergänzt.

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

Schlüssel zum Verständnis der morphogenetischen Prozesse, die der frühe Embryo Form ist die Möglichkeit, Bild-Zellen in hoher Auflösung. Wir beschreiben hier eine Methode zur Kennzeichnung einzelner Zellen oder kleine Gruppen von Zellen in ganzen Zebrafisch-Embryos mit Membran-bezogene Green Fluorescent Protein.

Labeling und Imaging Cells in der Zebrafisch Hinterhirn

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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Neuroscience

11.2K Aufrufe.  University of Maryland, Baltimore County. Schlüssel zum Verständnis der morphogenetischen Prozesse, die der frühe Embryo Form ist die Möglichkeit, Bild-Zellen in hoher Auflösung. Wir beschreiben hier eine Methode zur Kennzeichnung einzelner Zellen oder kleine Gruppen von Zellen in ganzen Zebrafisch-Embryos mit Membran-bezogene Green Fluorescent Protein.

Serie: 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.

Zeitraffer-Live-Imaging von klonal verbundenen neuronalen Vorläuferzellen in der Entwicklung von Zebrafisch Vorderhirnneuronen

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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Neurowissenschaften

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

Live-Cell-Imaging von Sinnesorgan Vorläuferzellen im Intact Drosophila Puppen

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.

Elektroporation des Hinterhirn zu Axonal Trajectories und Synaptic Treffer in der Embryo-Küken Trace

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.

Beschriften einzelner Zellen im zentralen Nervensystem<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.

Retrograde Markierung von retinalen Ganglienzellen im erwachsenen Zebrafisch mit Fluoreszenzfarbstoffen

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.

Die Maus Hinterhirn als Modell für die Erforschung embryonaler Neurogenese

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.

In Vivo Calcium Imaging der Seitenlinie Haarzellen im Larvenstadium Zebrafisch

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.

Identifizieren von Zelloberflächenmarkern von primären neuralen Stamm- und Vorläuferzellen durch metabolische Kennzeichnung von 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.

Etikettierung und Bildgebung von Amyloid-Plaques im Gehirngewebe mit dem natürlichen 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). ...

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.

In vivo Bildgebung von vollaktivem Hirngewebe bei wachen Zebrafischlarven und Jungtieren durch Schädel- und Hautentfernung

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