The project was supported by NIH R01NS120218, the UCSF Mary Anne Koda-Kimble Seed Award for Innovation, and Chan Zuckerberg Biohub.

We have used the AB wild type line and transgenic line Tg [ef1a:Myr-Tdtomato] for the study. All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of California, San Francisco, USA (Approval Number: AN179000).
1. Preparation of zebrafish embryos
<ol>
	<li>Set up fish crossing tanks in the afternoon before 5:00 p.m., with one female wild-type fish and one male Tg [ef1a:Myr-Tdtomato] fish by using dividers to separate them in each tank.</li>
	<li>Remove the dividers before 11:00 a.m. from all crossing tanks the next morning. Keep quiet and do not disturb the fish while they are mating. Spawning typically occurs within 30 min after removing the dividers. The fertilized eggs remain on the bottom of the tank.
	<ol>
		<li>Collect fertilized eggs from the tanks with a mesh filter. Transfer the eggs by washing them off into a Petri dish full of egg water and examine the embryos under a dissecting microscope at a 10x to 20x magnification.</li>
	</ol>
	</li>
	<li>Transfer the fertilized embryos to a clean Petri dish containing approximately 20 mL of embryo medium (100 mL of 1,000x stock solution contains 29.4 g of NaCl, 1.27 g of KCl, 4.85 g of CaCl2.2H2O, and 8.13 g of MgSO4.7H2O) with large-bore glass Pasteur pipettes. Keep the embryos at 28 &#176;C. Keep 50 fertilized embryos per Petri dish with 30 mL of embryo medium at 28.5 &#176;C. 
	&#8203;NOTE: One pair of fish normally produces 100-300 embryos, and the fertilization and survival rate of healthy embryos is over 95% for each mating.</li>
	<li>The next morning, take the embryos out of the incubator at 8:00 a.m., when the embryos have reached the developmental stage of ~18-20 h post-fertilization (hpf). Observe them under an epifluorescence microscope at a 20x magnification, using white light at first. At that time, the zebrafish embryos are at the 20 somite stage.
	<ol>
		<li>Discard dead embryos that are cloudy or ruptured under the microscope using a glass pipette.</li>
	</ol>
	</li>
	<li>Turn on the fluorescent lamp and choose the RFP filter setting of the microscope. Then, select the embryos with strong red fluorescence and transfer them to a new dish with egg water.
	<ol>
		<li>Dechorionate the embryos manually with two fine forceps (the tips of forceps should be sharp and undamaged) under white light (Table of Materials).</li>
		<li>Hold the chorion with one pair of forceps and make a tear in the chorion with the other forceps. Open the tear carefully using the forceps, and make it large enough for the embryo to pass through by gently pushing the embryo with the tips of the other forceps.</li>
	</ol>
	</li>
	<li>Transfer the dechorionated embryos to a new Petri dish with fresh embryonic medium for a quick rinse before microinjection.</li>
</ol>
2. Preparation of microinjection
<ol>
	<li>Use the capillaries (1.2 mm OD, 0.9 mm ID, with filament) for pulling fine injection needles on a puller. Design and optimize the pulling program according to the handbook.</li>
	<li>Open the tip of the needle with forceps under a stereo dissection microscope. Make the diameter of the tip around 10 &#181;m, and the taper angle around 30&#176;.</li>
	<li>Conjugate the mouse monoclonal anti-Dld antibody with the anti-Mouse-IgG-Atto647N before injection.
	<ol>
		<li>For each injection experiment, mix 0.5 &#181;L of the anti-Dld antibody (0.5 mg/mL) with 2 &#181;L of the anti-Mouse-IgG-Atto647N antibody (1 mg/mL) by pipetting 5-10 times. Then, incubate at room temperature for at least 30 min (or on ice for 2-3 h).</li>
		<li>After incubation, add 2.5 &#181;L of blocking buffer (10 mg/mL mouse IgG) and 0.5 &#181;L of 0.5% phenol red to the antibody mixture, and pipette 10x to mix thoroughly, to block any unconjugated antibodies remaining in the mixture. 
		NOTE: For each trial, prepare one extra mixture without the anti-Dld antibody and use it as a control.</li>
	</ol>
	</li>
	<li>Prepare 1% low-melting point agarose in the embryo medium. Heat the agarose contained mixture at 70 &#176;C till the mixture turns transparent.
	<ol>
		<li>Aliquot the agarose solution in 2 mL microcentrifuge tubes. Keep the aliquots in a heat block at ~40 &#176;C.</li>
	</ol>
	</li>
	<li>Rinse the embryo in the tube containing 1% low-melting point agarose for 3 s.</li>
	<li>Place the embryo on an inverted plastic Petri dish lid together with individual drops of agarose (~30-40 &#181;L) to mount each embryo separately on the lid as shown in Figure 1. Lay the embryos flat laterally in the agarose and keep this position until the agarose has solidified at room temperature.</li>
	<li>Mount 12 embryos in three rows one by one as above. Cover all the mounted embryos in the agarose with egg medium.</li>
</ol>
3. Microinjection
<ol>
	<li>Put the embedded embryos under the stereomicroscope and cover the agarose with egg water.</li>
	<li>Set the air pressure injector together with micromanipulators, placing them close to the microscopes as shown in Figure 1. Use the steel gas cylinder containing gaseous nitrogen (N2) under high pressure as the air resource.
	<ol>
		<li>Open the gas valve only after the embryos have been mounted in the agarose. Then, front load the prepared glass needle with 2 &#181;L of antibody mixture on the micromanipulator, as shown in Figure 1, when using the front fill module of the microinjector.</li>
		<li>Tune the input pressure to ~80-90 psi, and the injection pressure to ~20 psi.</li>
	</ol>
	</li>
	<li>Calibrate the injection volume by using a micrometer under the microscope, as described previously15. Set the tune time duration to be from 10 ms to 120 ms, according to the size of needle opening. Deliver each pulse of injection by tapping the paddle. Tune the injection volume of each delivery to ~4-5 nL.</li>
	<li>Poke the tip of the microinjection needle through the dorsal roof plate of the hindbrain posterior to the r0/r1 hinge point, and inject about 10 nL (two or three pulses) of antibody mixture without hitting the brain tissue. Observe the flowing of red fluids in the brain ventricle. 
	NOTE: The hindbrain ventricle is posterior to the midbrain hindbrain boundary. The injected phenol red-containing antibody mixture fills up the brain ventricle from the hindbrain to the forebrain immediately by diffusing with cerebrospinal fluid.</li>
	<li>After injection, remove the tip of the needle from the embryo swiftly by rotating the knob of the micromanipulator. For a successful injection, the red dye of the injected mixture remains in the brain ventricle stably without leaking into the surrounded agarose.</li>
	<li>Move the mounting plate under the microscope to locate another mounted embryo at a suitable position for repeats.
	<ol>
		<li>After injecting six to eight embryos, peel the agarose with a microsurgical knife to release the embryos from the embedded agarose. Transfer the microinjected embryos to a fresh dish with 30 mL of embryo medium, and place them at room temperature for the next steps.</li>
	</ol>
	</li>
</ol>
4. Mounting and time-lapse live imaging
<ol>
	<li>After 30 min, transfer the selected embryos to 10 mL of embryo medium. Add 420 &#181;L of tricaine stock (4 mg/mL) to 10 mL of embryo medium to anesthetize the embryos.</li>
	<li>To mount the embryos, prepare 0.8% low melting point agarose containing the same concentration of tricaine in the tube. Keep the aliquots in a heat block at ~40 &#176;C.</li>
	<li>Use a glass pipette to immerse the injected embryos in the warm agarose for 3 s. Then, immediately remove the embryos from the agarose with the same glass pipette and place the embryos on the center of 35 mm glass bottom culture dishes with a drop of agarose from the tube. Place only one embryo per drop on the glass.</li>
	<li>Orientate the embryos gently with a fiber probe or loading tip to keep the dorsal side of the embryonic brain as close to the glass bottom as possible. Tune the embryo position gently to extend the embryo without curling as the agarose solidifies gradually.</li>
	<li>Afterward, check the embryo position by flipping the glass bottom dish over. Ensure that the whole dorsal forebrain with the correctly mounted embryos can be seen under the microscope.</li>
	<li>Add 2-3 mL of 28.5 &#176;C preheated embryonic medium containing tricaine to cover the embryo. Place the dish properly on the temperature-controlled stage of the confocal microscope, as shown in Figure 1. Adjust the temperature of the imaging chamber to be at 28.5 &#176;C. The embryo is now ready for imaging.</li>
	<li>Perform time-lapse live imaging with a fixed time interval using a 40x water immersion objective.
	<ol>
		<li>Use the imaging and microscope control software (&#181;MANAGER: https://micro-manager.org/)16.</li>
		<li>Select the imaging channels of 564 nm and 647 nm for imaging the membrane fluorescence of MyR-Tdtomato transgenic zebrafish and endosomal anti-Dld-Atto647N in the cell (Figure 1).</li>
		<li>Set the laser power at 30% for both channels. Use an exposure time per z-plane of 200 ms for each channel.</li>
		<li>For each mounted zebrafish embryo, use a scanning z-step of 1 &#181;m for 20 z-planes in total, and a scanning cycle of 100 timeframes.</li>
		<li>Set the time interval between each scanning cycle at 20 s and the scanning mode as channel, slice.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We have developed an antibody uptake assay for labeling and imaging endosomal Notch/Delta trafficking in zebrafish radial glia progenitors with high efficiency. Compared to previous methods used for tracking labeled anti-DeltaD antibody in Drosophila SOP cells7,8, our method used microinjection instead of incubation of samples in the conjugated antibody. Fluorescently conjugated anti-Dld antibodies were microinjected into the hindbrain ventricle; this technique d...

Notch signaling controls cell fate decision and patterning during development in metazoans1, and recent studies have shown that Notch signaling in stem cell division mainly depends on endocytic trafficking2,3. Endocytosed Notch/Delta can activate Notch signaling in the nucleus and enhance the transcription of Notch target genes4,5,6. Directional Notch/Delta endosomal trafficking was first observed in Drosophila sensory organ precursor (SOP) cells during its asymmetric cell division (ACD), resulting in a higher Notch signaling activity in pIIa than in pIIb7,8. Antibody uptake assays with anti-Delta and anti-Notch antibodies have been applied to monitor the endocytic process in mitotic SOP cells. Notch/DeltaD endosomes move along with a kinesin motor protein to the central spindle during cytokinesis, and are asymmetrically translocated into the pIIa cell due to the antiparallel array of the asymmetric central spindle at the last moment of cell division3,8. These studies have shed light on the molecular mechanisms regulating asymmetric division in Drosophila SOP cells, but it is unclear whether similar endocytic processes occur in vertebrate radial glia progenitors (RGPs).
Moreover, the molecular mechanisms that regulate asymmetric Notch/DeltaD signaling during vertebrate RGP division are not well understood. In zebrafish, it has been reported that the interaction of Notch and Delta facilitates the endocytosis of the DeltaD ligand9. It is unknown whether DeltaD endocytosis can impact the cell fate choice of daughter cells in the developing vertebrate brain. Recent studies show injecting fluorescently conjugated anti-DeltaD antibodies into the neural tube could label Sara endosomes specifically in neuroepithelial cells, and anti-DeltaD containing Sara endosomes preferentially segregate into proliferating daughter cells10. It has been suggested that Notch signaling from the endosomes could regulate daughter cell fate. Previous results have shown that most zebrafish RGP cells in the developing forebrain undergo ACD, and the daughter cell fate determination is dependent on intralineage Notch/DeltaD signaling11. In order to elucidate the nature of the intralineage Notch/DeltaD signaling in zebrafish RGPs, we have developed the anti-DeltaD antibody uptake assay in the zebrafish developing brain. Using this protocol, we have successfully performed live labeling and imaging of DeltaD endocytic trafficking in mitotic RGPs.
The fluorescently labeled anti-DeltaD-antibody is efficiently internalized into the RGPs along the forebrain ventricle. It has greatly facilitated the discovery of directional trafficking of DeltaD endosomes in the asymmetrically dividing RGPs12,13. Compared to previous antibody uptake protocols developed for Drosophila notum cultures and the zebrafish spinal cord10, this protocol has achieved long-lasting and highly efficient anti-DeltaD labeling in the brain ventricle cell layer, specifically with less than 10 nL of microinjected antibody mixture. The hindbrain ventricle injection is very convenient for applying the antibody uptake assay in the developing brain, as the hindbrain ventricle is well expanded in zebrafish embryos and filled up with cerebrospinal fluid at early development stage14. By injecting the antibody mixture into the hindbrain ventricle without injuring any crucial developing tissues, the protocol has minimized possible damage to the imaging zone in the forebrain as much as possible. The reduced dosage of the injected primary antibody has also avoided potential side-effects of interfering with endogenous Delta-Notch signaling in vivo. This protocol can be easily combined with other pharmacological or genetic perturbations, utilized at different developmental stages, and possibly adapted to the adult brain as well as human pluripotent stem cell-derived 2D/3D brain organoids. Taken together, the protocol has made it possible to understand how and when Notch signaling asymmetry is established during ACD. The main challenge for successful implementation of this protocol is to achieve precise delivery of appropriate concentrations of the antibody based on specific experimental conditions.

<table><tbody><tr><td>35mm glass bottom culture dish&amp;nbsp;</td><td>MatTek corporation</td><td>P35GC-1.5-10-C</td><td></td></tr><tr><td>air pressure injector&amp;nbsp;</td><td>Narishige</td><td>IM300</td><td></td></tr><tr><td>Anti-Mouse-IgG-Atto647N&amp;nbsp;</td><td>Sigma-Aldrich</td><td>50185</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2.&lt;/sub&gt;2H&lt;sub&gt;2&lt;/sub&gt;O&amp;nbsp;</td><td>Sigma-Aldrich</td><td>C3306</td><td></td></tr><tr><td>Capillaries, 1.2 mm OD, 0.9 mm ID, with filament</td><td>World Precision Instruments</td><td>1B120F-6</td><td></td></tr><tr><td>CSU-W1 Spinning Disk/High Speed Widefield</td><td>Nikin</td><td>N/A</td><td>Nikon&amp;nbsp;Ti inverted fluorescence microscope with CSU-W1 large field of view confocal.&amp;nbsp;</td></tr><tr><td>Dumont Medical Tweezers Style 5</td><td>Thomas Scientific</td><td>72877-D</td><td></td></tr><tr><td>Flaming-Brown P897 puller</td><td>Sutter Instruments</td><td>N/A</td><td>https://www.sutter.com/manuals/P-97-INT_OpMan.pdf</td></tr><tr><td>KCl</td><td>Millipore</td><td>529552</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4.&lt;/sub&gt;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>M2773</td><td></td></tr><tr><td>micromanipulators</td><td>World Precision Instruments</td><td>WPI M3301R</td><td></td></tr><tr><td>Mouse anti-Dld&amp;nbsp;</td><td>Abcam</td><td>AB_1268496</td><td></td></tr><tr><td>Mouse IgG blocking buffer from Zenon</td><td>Thermofisher Scientific</td><td>Z25008</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S3014</td><td></td></tr><tr><td>Phenol red</td><td>Sigma-Aldrich</td><td>P0290</td><td></td></tr><tr><td>Stemi 2000&amp;nbsp;&amp;nbsp;</td><td>Zeiss&amp;nbsp;</td><td>N/A</td><td></td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich</td><td>E10521</td><td></td></tr><tr><td>UltraPureTM low melting point agarose&amp;nbsp;</td><td>Invitrogen</td><td>16520050</td><td></td></tr></tbody></table>

antibody uptake assay for tracking notch/delta endocytosis during the asymmetric division of zebrafish radial glia progenitors

Asymmetric cell division (ACD), which produces two daughter cells of different fates, is fundamental for generating cellular diversity. In the developing organs of both invertebrates and vertebrates, asymmetrically dividing progenitors generates a Notchhi self-renewing and a Notchlo differentiating daughter. In the embryonic zebrafish brain, radial glia progenitors (RGPs)-the principal vertebrate neural stem cells-mostly undergo ACD to give birth to one RGP and one differentiating neuron. The optical clarity and easy accessibility of zebrafish embryos make them ideal for in vivo time-lapse imaging to directly visualize how and when the asymmetry of Notch signaling is established during ACD. Recent studies have shown that dynamic endocytosis of the Notch ligand DeltaD plays a crucial role in cell fate determination during ACD, and the process is regulated by the evolutionarily conserved polarity regulator Par-3 (also known as Pard3) and the dynein motor complex. To visualize the in vivo trafficking patterns of Notch signaling endosomes in mitotic RGPs, we have developed this antibody uptake assay. Using the assay, we have uncovered the dynamicity of DeltaD-containing endosomes during RGP division.

In Figure 2A, the embryos injected with Atto647N, without binding with the primary antibody, showed background fluorescence in the brain ventricle. Very few engulfed fluorescent particles can be observed in the cells. The anti-Dld-Atto647N injected zebrafish embryos showed large amounts of internalized fluorescent particles in most cells of the developing forebrain (Figure 2A, right panel). After zooming in to focus on mitotic RGPs, as shown in <strong class="xf...

Watch this Scientific Journal Video about Antibody Uptake Assay for Tracking Notch/Delta Endocytosis During the Asymmetric Division of Zebrafish Radial Glia Progenitors at JoVE.com

Antibody Uptake Assay for Tracking Notch/Delta Endocytosis During the Asymmetric Division of Zebrafish Radial Glia Progenitors

This work develops an antibody uptake assay for imaging intra-lineage Notch/DeltaD signaling in dividing radial glia progenitors of the embryonic zebrafish brain.

Asymmetric cell division (ACD), which produces two daughter cells of different fates, is fundamental for generating cellular diversity. In the developing ...

antibody-uptake-assay-for-tracking-notchdelta-endocytosis-during

Research

JoVE Journal

Developmental Biology

1.9K Views.  Chan Zuckerberg Biohub. Notch-signalering regelt sulfaatbeslissing en patronen tijdens de ontwikkeling in metazoën. We hebben een antilichaamopnametest ontwikkeld voor het labelen van de notch ligand delta-D en het in beeld brengen van de endocytische handel in radiale glia-voorlopers in de zich ontwikkelende voorhersenen van zebravissen. De zebravis verborgen hersenventrikelinjectie maakt selectieve Anti-Dld-Atto-647N-opname mogelijk door myototische radiale glia-voorlopers die de hersenventrikels in ontwikkeling ...

Video: Antibody Uptake Assay for Tracking Notch/Delta Endocytosis During the Asymmetric Division of Zebrafish Radial Glia Progenitors

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.

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 ...

Neuroscience

Affinity Labeling Detection of Endogenous Receptors from Zebrafish Embryos

By combining the powers of Affinity Labeling and Immunoprecipitation (AFLIP), a technique for the detection of low abundance receptors in zebrafish embryos has been implemented. This technique takes advantage of the selectivity and sensitivity conferred by affinity labeling of a given receptor by its ligand with the specificity of the immunoprecipitation. We have used AFLIP to detect the type III TGF-&#946; receptor (TGFBR3), also know as betaglycan, during early zebrafish development. AFLIP was instrumental in validating the efficacy of a TGFBR3 morphant zebrafish phenotype. In the first step, embryo protein extracts are prepared and used to generate 125I-TGF-&#946;2-TGFBR3 complexes that are purified by immunoprecipitation. Later, these complexes are covalently cross-linked and revealed using SDS-PAGE separation and autoradiography detection. This technique requires the availability of a labeled ligand for, and a specific antibody against, the receptor to be detected, and shall be easily adapted to identify any growth factor or cytokine receptor that meets these requirements.

A novel technique for the detection of low abundance endogenous receptors present in zebrafish embryos is described. We have named it AFLIP because it consists of affinity labeling of the receptor by its ligand linked to immunoprecipitation.

By combining the powers of Affinity Labeling and Immunoprecipitation (AFLIP), a technique for the detection of low abundance receptors in zebrafish ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Biology

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Elucidating Microglia Behavior in Zebrafish Optic Tectum Development

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and function. Externally developing zebrafish embryos possess optical transparency, which along with well-characterized transgenic reporter lines that fluorescently label microglia, make zebrafish an ideal vertebrate model for such studies. In this paper, we take advantage of the unique features of the zebrafish model to visualize the dynamics of microglia cells in vivo and under physiological conditions. We use confocal microscopy to record a timelapse of microglia cells in the optic tectum of the zebrafish embryo and then, extract tracking data using the IMARIS 10.0 software to obtain the cells' migration path, mean speed, and distribution in the optic tectum at different developmental stages. This protocol can be a useful tool to elucidate the physiological significance of microglia behavior in various contexts, contributing to a deeper characterization of these highly motile cells.

Author Spotlight: Using Zebrafish to Explore Microglia Migration During Brain Development

We demonstrate a method that takes advantage of fast scanning confocal microscopy to perform live imaging of microglia cells in the developing zebrafish optic tectum, allowing for the analysis of the dynamics of these cells in vivo.

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and ...

Immunology and Infection

Monitoring On-Target Signaling Responses in Larval Zebrafish - Z-REX Unmasks Precise Mechanisms of Electrophilic Drugs and Metabolites

Reactive metabolites and related electrophilic drugs are among the most challenging small molecules to study. Conventional approaches to deconstruct the mode of action (MOA) of such molecules leverage bulk treatment of experimental specimens with an excess of a specific reactive species. In this approach, the high reactivity of electrophiles renders non-discriminate labeling of the proteome in a time- and context-dependent manner; redox-sensitive proteins and processes can also be indirectly and often irreversibly affected. Against such a backdrop of innumerable potential targets and indirect secondary effects, linking phenotype to specific target engagement remains a complex task. Zebrafish targeting reactive electrophiles and oxidants (Z-REX)-an on-demand reactive-electrophile delivery platform adapted for use in larval zebrafish-is designed to deliver electrophiles to a specific protein of interest (POI) in otherwise unperturbed live fish embryos. Key features of this technique include a low level of invasiveness, along with dosage-, chemotype-, and spatiotemporally-controlled precision electrophile delivery. Thus, in conjunction with a unique suite of controls, this technique sidesteps off-target effects and systemic toxicity, otherwise observed following uncontrolled bulk exposure of animals to reactive electrophiles and pleiotropic electrophilic drugs. Leveraging Z-REX, researchers can establish a foothold in the understanding of how individual stress responses and signaling outputs are altered as a result of specific reactive ligand engagement with a specific POI, under near-physiologic conditions in intact living animals.

Zebrafish targeting reactive electrophiles and oxidants (Z-REX) is a chemical biology-based method for the investigation of reactive small-molecule signaling. This technique can be applied to live fish of different developmental stages. Here, we couple standard assays in zebrafish with Z-REX for signaling pathway analysis.

Reactive metabolites and related electrophilic drugs are among the most challenging small molecules to study. Conventional approaches to deconstruct the ...

Antibody Uptake Assay for Tracking Notch/Delta Endocytosis During the Asymmetric Division of Zebrafish Radial Glia Progenitors

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