This work was supported by an NIH grant awarded to R. Brewster (1R01GM085290-01A1 ).

1.Microinjection
<ol>
 <li>Dilute plasmid encoding membrane-targeted Green Fluorescent Protein (mGFP, courtesy of Richard Harland) to a concentration of 40 ng/ml. DNA is prepared by linearizing plasmid (mGFP/PCS2+, courtesy of Richard Harland) purified using the Qiagen Maxi Prep Kit. Addition of phenol red (diluted 1/10 total volume) to the solution is preferred to visualize the solution. Keep DNA solution on ice.</li>
 <li>Under a stereomicroscope, calibrate the injection needle (90mm glass capillaries from Narishinge Scientific; Cat No. NA-GD-1) by aligning the needle on a graduated slide and gently tapping the part of the needle where the diameter is 1&mu;m using a clean razor blade (to obtain a tip of the appropriate size).</li>
 <li>In order to calibrate the needle to inject a known volume (2nl), connect the needle to the microinjection apparatus (PCI 100 Microinjector; Harvard Apparatus) and front fill the needle with 0.5&mu;l of a solution containing phenol red and water placed on a piece of paraffin. The solution can be dispensed using a foot pedal. Set the pressure to 7 KPa and vary the time (10msec 30msec) so that it takes 250 pedals to dispense 0.5&mu;l of the solution.</li>
 <li>Carefully place the calibrated needle on a strip of wax in a humidified 100 mm Petri dish. To humidify the dish, place wet Kimwipes around the edges (this prevents evaporation of the solution in the needle).</li>
 <li>Use a micropipette to load 0.5-1&mu;l of DNA into the needle end (the end that was not calibrated).</li>
 <li>Once the solution has reached the needle tip, connect the end of the needle that was used for loading to a microinjection apparatus. Adjust the settings on the microinjector to deliver approximately 2 nl/injection using a foot pedal.</li>
 <li>Next, position zebrafish embryos in the middle of a Petri dish filled with Embryo Medium (1.0 ml Hank's Stock #1, 0.1 ml Hank's Stock #2, 1.0 ml Hank's Stock #4, 95.9 ml ddH2O, 1.0 ml Hank's Stock #5, 1.0 ml fresh Hank's Stock #6, Use about 10 drops 1 M NaOH to pH 7.2).</li>
 <li>Once the embryos have developed to the desired stage, gently grip the chorion with fine forceps and situate the needle into the area targeted for injection. For broad range mGFP expression, inject DNA into the yolk or the cytoplasm at the 1cell stage. To restrict expression of mGFP to a smaller region, inject DNA at the 4 (or more) cells stage into the cytoplasm of a single cell.</li>
 <li>Press the foot pedal of the microinjection apparatus and deliver approximately 2 nl, ensuring that the solution (visible due to the addition of phenol red) is properly injected.</li>
 <li>Carefully remove the needle from the embryo.</li>
 <li>Repeat for all embryos in the Petri dish.</li>
 <li>Allow embryos to develop at 28.5&deg;C to desired stage.</li>
 <li>Dechorionate embryos by gently peeling off the chorion using fine forceps (if embryos are younger than 24hpf, dechorionating in a glass Petri dish increases the survival rate of embryos).</li>
 <li>Remove empty chorion from the Petri dish using a glass pipette.</li>
</ol>
2. Imaging mGFP-labeled vibratome sections
<ol>
 <li>Use a glass pipette to transport embryos into fixative (4% paraformaldehyde in 1X phosphate buffer solution (PBS). Fix embryos at 4&deg;C overnight.</li>
 <li>Wash embryos in 1X PBS 3 times for 5 minutes each.</li>
 <li>Place the embryos in Petri dish and, using fine forceps, remove the yolk without damaging the embryo. Remove as much yolk as possible.</li>
 <li>Prepare 4% low-melt agarose in H2O.</li>
 <li>Place molten agarose into plastic sectioning mold.</li>
 <li>Use fine forceps to lift an injected embryo from the Petri dish and to place and orient in the agarose. The embryo should be positioned towards one end of the mold.</li>
 <li>Once the agarose is hardened, use a flat surface tool, such as a spatula, to remove the agarose block from the mold.</li>
 <li>Cut the bottom end of the agarose block with a razor blade so that it is even.</li>
 <li>Add a drop of super glue to the vibratome stage and place the agarose block on it (the side of the block that was leveled off with the razor should face the glue). For sectioning multiple embryos, mount several agarose-embedded specimens on the stage.</li>
 <li>Attach the vibratome stage to the vibratome (Vibratome, Inc.).</li>
 <li>Set the parameters of the vibratome (common settings: speed-3; thickness-50&mu;m). If sectioning multiple embryos, align agarose blocks on the stage and ensure that the blade goes through the whole embryo but not through the entire width of the agarose blocks (such that sections corresponding to the same embryo will remain together).</li>
 <li>Pour 1X PBS in the vibratome stage.</li>
 <li>Section.</li>
 <li>Once the embryos are sectioned, remove the agarose block(s) from the vibratome stage.</li>
 <li>Carefully separate each section by cutting the back of the agarose block(s).</li>
 <li>Use fine forceps to select desired sections and place into a 1X PBS-filled multi-well dish (use different wells for sections corresponding to different embryos).</li>
 <li>Repeat for each embryo.</li>
 <li>Proceed with antibody labeling, steps 19 though 23, to enhance mGFP imaging (this step may not be necessary if sections are imaged within a day or two of preparation).</li>
 <li>Treat sections for 30 minutes at room temperature with blocking solution in 1X PBS to block non-specific binding.</li>
 <li>Incubate sections with primary antibody (rabbit a-GFP, Invitrogen Cat No. A11122) diluted @ 1:200 in the blocking solution on a shaker for 5 hours at room temperature or overnight at 4&deg;C.</li>
 <li>Wash four times (10 min, 15 min, 30 min and 60 min, each) in 1% BSA/1X PBS/ 1% DMSO solution.</li>
 <li>Treat sections with appropriate secondary antibody in blocking solution for 5 hours at room temperature or overnight at 4&deg;C.</li>
 <li>Wash four times (10 min, 15 min, 30 min and 60 min, each) in 1% BSA/1X PBS/ 1% DMSO solution.</li>
 <li>Treat the sections with DAPI (0.1% DAPI in 50ml 1X PBS; Invitrogen Cat No. D1306) for 30 mins to stain the nuclei.</li>
 <li>Wash three times in 1X PBS for 5 minutes each.</li>
 <li>Mount on slide.</li>
 <li>Image using confocal microscopy.</li>
</ol>
3. Live imaging
<ol>
 <li>Dechorionate live, injected embryos 30 minutes before they reach the desired developmental stage. Place the embryos in sedative solution (0.01% Tricaine in Embryo Medium) for 15 min.</li>
 <li>Place a drop of 1% agarose on a 35mm glass bottom culture dish (MatTek; Part No. P35G-o-20-C).</li>
 <li>Allow agarose to solidify.</li>
 <li>Heat capillary tube over flame to make 3 holes in agarose (other objects such as micropipette tips may be used to make the holes). Ensure that there is no residual agarose in the hole bottom.</li>
 <li>Fill the Petri dish with Embryo Medium.</li>
 <li>Use a glass pipette to place the embryo in the hole.</li>
 <li>Next, orient the embryo using fine forceps so that the area to be imaged (dorsal head) is in contact with the glass.</li>
 <li>Carefully transport the Petri dish to the microscope stage (place the lid over the Petri dish to prevent evaporation).</li>
 <li>While imaging, ensure that the embryos are at 28.5&deg;C.</li>
</ol>
4. Results
We describe here a straightforward approach to image single cells in the zebrafish neural tube using mosaic expression. Despite the optical transparency of the zebrafish embryo, we have found that visualization of neural cells is greatly enhanced in cross sections obtained using a vibratome. These sections are thick (50 mm) allowing for imaging of cellular extension from a given cell in different focal planes using a confocal microscope. Results using this methodology have been published elsewhere 2, but representative images of cells in the neural tube of a WT embryo (Figure 1) and an N-cadherin (N-cad) mutant (Figure 2) are provided. The latter reveals that cells in lateral regions of the neural plate in N-cad mutants fail to orient properly towards the midline (Figure 2). In contrast to this mosaic labeling, immunostaining the neuroepithelium with a general cell surface marker such as beta-catenin does not allow the morphology of individual cells to be visualized (Figure 3).
Time lapse imaging of live embryos complemented the mosaic labeling of fixed specimens, enabling a better understanding of the cellular dynamics that take place during neurulation. Movie 1 revealed that WT cells actively migrate towards the midline by extending medially-oriented membrane protrusions.
<img src="/files/ftp_upload/1976/1976fig1.jpg" alt="Figure 1" /> 
 Figure 1. mGFP expression in the neural tube of a WT zebrafish embryo.
<img src="/files/ftp_upload/1976/1976fig2.jpg" alt="Figure 2" /> 
 Figure 2. mGFP expression in the neural tube of a N-cadherin zebrafish embryo.
<img src="/files/ftp_upload/1976/1976fig3.jpg" alt="Figure 3" /> 
 Figure 3. b-catenin immuno staining of the neuroepithelium.

<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>

No conflicts of interest declared.

In conclusion, the labeling techniques described here allow for single cell analysis of morphogenetic processes in the zebrafish embryo. The primary emphasis of this protocol is on methods for imaging labeled cells in the neural tube using mGFP under the control of a ubiquitous promoter. For additional applications of this transient expression assay readers should refer to a recent paper by 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's Stock #1: 8.0 g NaCl , 0.4 g KCl in 100 ml dd H2O</li> <li>Hank's Stock #2: 0.358 g Na2HPO4 Anhydrous, 0.60 g KH2PO4 in 100 ml ddH2O</li> <li>Hank's Stock #4: 0.72 g CaCl2 in 50 ml ddH2O</li> <li>Hank's Stock #5: 1.23 g MgSO4x7H2O in 50 ml dd H2O</li> <li>1 X Phosphate Buffer: 0.8% NaCl, 0.02% KCl, 0.02 M PO4, pH 7.3</li> <li>Blocking Solution: 2% normal goat serum, 1% bovine serum albumin (BSA), 1% dimethysulfoxide (DMSO)</li> </ul>

labeling and imaging cells in the zebrafish hindbrain

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish embryos, injection of plasmid DNA results in mosaic expression, allowing for the visualization of single cells or small clusters of cells 1 . We describe how injection of plasmid DNA encoding membrane-targeted Green Fluorescent Protein (mGFP) under the control of a ubiquitous promoter can be used for imaging cells undergoing neurulation. Central to this protocol is the methodology for imaging labeled cells at high resolution in sections and also in real time. This protocol entails the injection of mGFP DNA into young zebrafish embryos. Embryos are then processed for vibratome sectioning, antibody labeling and imaging with a confocal microscope. Alternatively, live embryos expressing mGFP can be imaged using time-lapse confocal microscopy. We have previously used this straightforward approach to analyze the cellular behaviors that drive neural tube formation in the hindbrain region of zebrafish embryos 2. The fixed preparations allowed for unprecedented visualization of cell shapes and organization in the neural tube while live imaging complemented this approach enabling a better understanding of the cellular dynamics that take place during 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

Key to understanding the morphogenetic processes that shape the early embryo is the ability to image cells at high resolution. We describe here a technique for labeling single cells or small clusters of cells in whole zebrafish embryos with membrane-targeted Green Fluorescent Protein.

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

11.3K ビュー.  University of Maryland, Baltimore County. 初期胚を形成する形態形成のプロセスを理解するための鍵は、高解像度での画像セルにできることです。ここでは、細胞膜をターゲットとする緑色蛍光タンパク質で全体のゼブラフィッシュの胚では細胞の単一細胞または小さなクラスターを標識するためのテクニックを説明します。

ビデオ: 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.

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

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

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.

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.

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.

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.

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.

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.

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.

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.

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