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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	<li>Baonza, A., Garcia-Bellido, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notch+signaling+directly+controls+cell+proliferation+in+the+Drosophila+wing+disc.">Notch signaling directly controls cell proliferation in the Drosophila wing disc.</a> Proceedings of the National Academy of Sciences. 97 (6), 2609-2614 (2000).</li><li>Chitnis, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Developmental Dynamics. 235 (4), 886-894 (2006).</li><li>Daeden, A., Gonzalez-Gaitan, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endosomal+trafficking+during+mitosis+and+notch-dependent+asymmetric+division.">Endosomal trafficking during mitosis and notch-dependent asymmetric division.</a> Progress in Molecular and Subcellular Biology. 57, 301-329 (2018).</li><li>Le Borgne, R., Schweisguth, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notch+signaling:+endocytosis+makes+delta+signal+better.">Notch signaling: endocytosis makes delta signal better.</a> Current Biology. 13 (7), 273-275 (2003).</li><li>Chapman, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notch1+endocytosis+is+induced+by+ligand+and+is+required+for+signal+transduction.">Notch1 endocytosis is induced by ligand and is required for signal transduction.</a> Biochimica et Biophysica Acta. 1863 (1), 166-177 (2016).</li><li>Schroeter, E. H., Kisslinger, J. A., Kopan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notch-1+signalling+requires+ligand-induced+proteolytic+release+of+intracellular+domain.">Notch-1 signalling requires ligand-induced proteolytic release of intracellular domain.</a> Nature. 393 (6683), 382-386 (1998).</li><li>Coumailleau, F., F&#252;rthauer, M., Knoblich, J. A., Gonz&#225;lez-Gait&#225;n, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Directional+Delta+and+Notch+trafficking+in+Sara+endosomes+during+asymmetric+cell+division.">Directional Delta and Notch trafficking in Sara endosomes during asymmetric cell division.</a> Nature. 458 (7241), 1051-1055 (2009).</li><li>Derivery, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polarized+endosome+dynamics+by+spindle+asymmetry+during+asymmetric+cell+division.">Polarized endosome dynamics by spindle asymmetry during asymmetric cell division.</a> Nature. 528 (7581), 280-285 (2015).</li><li>Matsuda, M., Chitnis, A. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interaction+with+Notch+determines+endocytosis+of+specific+Delta+ligands+in+zebrafish+neural+tissue.">Interaction with Notch determines endocytosis of specific Delta ligands in zebrafish neural tissue.</a> Development. 136 (2), 197-206 (2009).</li><li>Kressmann, S., Campos, C., Castanon, I., F&#252;rthauer, M., Gonz&#225;lez-Gait&#225;n, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Directional+Notch+trafficking+in+Sara+endosomes+during+asymmetric+cell+division+in+the+spinal+cord.">Directional Notch trafficking in Sara endosomes during asymmetric cell division in the spinal cord.</a> Nature Cell Biology. 17 (3), 333-339 (2015).</li><li>Dong, Z., Yang, N., Yeo, S. -. Y., Chitnis, A., Guo, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intralineage+directional+Notch+signaling+regulates+self-renewal+and+differentiation+of+asymmetrically+dividing+radial+glia.">Intralineage directional Notch signaling regulates self-renewal and differentiation of asymmetrically dividing radial glia.</a> Neuron. 74 (1), 65-78 (2012).</li><li>Zhao, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polarized+endosome+dynamics+engage+cytoplasmic+Par-3+that+recruits+dynein+during+asymmetric+cell+division.">Polarized endosome dynamics engage cytoplasmic Par-3 that recruits dynein during asymmetric cell division.</a> Science Advances. 7 (24), (2021).</li><li>Zhao, X., Garcia, J., Royer, L. A., Guo, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Colocalization+analysis+for+cryosectioned+and+immunostained+tissue+samples+with+or+without+label+retention+expansion+microscopy+(LR-ExM)+by+JACoP.">Colocalization analysis for cryosectioned and immunostained tissue samples with or without label retention expansion microscopy (LR-ExM) by JACoP.</a> Bio-Protocol. 12 (5), 4336 (2022).</li><li>Gutzman, J. H., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+brain+ventricle+injection.">Zebrafish brain ventricle injection.</a> Journal of Visualized Experiments. (26), e1218 (2009).</li><li>Sive, H. L., Grainger, R. M., Harland, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calibration+of+the+injection+volume+for+microinjection+of+Xenopus+oocytes+and+embryos.">Calibration of the injection volume for microinjection of Xenopus oocytes and embryos.</a> Cold Spring Harbor Protocols. 2010 (12), (2010).</li><li>Edelstein, A., Amodaj, N., Hoover, K., Vale, R., Stuurman, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Computer+control+of+microscopes+using+&#181;Manager.">Computer control of microscopes using &#181;Manager.</a> Current Protocols in Molecular Biology. , (2010).</li><li>Lukinavi&#269;ius, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorogenic+probes+for+multicolor+imaging+in+living+cells.">Fluorogenic probes for multicolor imaging in living cells.</a> Journal of the American Chemical Society. 138 (30), 9365-9368 (2016).</li></ol>

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>35mm glass bottom culture dish&#160;</td>
			<td>MatTek corporation</td>
			<td>P35GC-1.5-10-C</td>
			<td></td>
		</tr>
		<tr>
			<td>air pressure injector&#160;</td>
			<td>Narishige</td>
			<td>IM300</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Mouse-IgG-Atto647N&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>50185</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2.2H2O&#160;</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&#160;Ti inverted fluorescence microscope with CSU-W1 large field of view confocal.&#160;</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>MgSO4.7H2O</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&#160;</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&#160;&#160;</td>
			<td>Zeiss&#160;</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&#160;</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

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Developmental Biology

1.8K Views.  Chan Zuckerberg Biohub. This work develops an antibody uptake assay for imaging intra-lineage Notch/DeltaD signaling in dividing radial glia progenitors of the embryonic zebrafish brain.

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

Ex Vivo Culture of Pharyngeal Arches to Study Heart and Muscle Progenitors and Their Niche

The pharyngeal mesoderm of developing embryos contributes to broad regions of head and heart musculature. We have developed a novel method to study head and heart progenitor cell development with pharyngeal arches (also known as branchial arches) ex vivo. Using this method, we have recently described that the second pharyngeal arch contains self-renewing heart progenitors and serves as a microenvironment for expansion of the progenitors during mouse heart development. The progenitor cells remain undifferentiated and expansive inside the arch, but quickly become functional cardiomyocytes as they migrate out of the arch. We also reported that first pharyngeal arch contains muscle progenitors giving rise to myotubes after leaving the arch. Here, we demonstrate the procedure for the dissection and ex vivo culture of first and second pharyngeal arches from developing mouse embryos. The method enables one to study head and heart progenitor/muscle development, including cardiomyocyte and myotube formation in detail ex vivo.

Ex Vivo Culture of Pharyngeal Arches to Study Heart and Muscle Progenitors and Their Niche

Here, we present a protocol to culture pharyngeal arches to study the biology of heart and muscle progenitor cells and their microenvironment.

The pharyngeal mesoderm of developing embryos contributes to broad regions of head and heart musculature. We have developed a novel method to study head ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Directed Differentiation of Primitive and Definitive Hematopoietic Progenitors from Human Pluripotent Stem Cells

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem cells (hPSCs). Until recently, efforts to differentiate hPSCs into HSCs have predominantly generated hematopoietic progenitors that lack HSC potential, and instead resemble yolk sac hematopoiesis.&#160;These resulting hematopoietic progenitors may have limited utility for in vitro disease modeling of various adult hematopoietic disorders, particularly those of the lymphoid lineages. However, we have recently described methods to generate erythro-myelo-lymphoid multilineage definitive hematopoietic progenitors from hPSCs using a stage-specific directed differentiation protocol, which we outline here. Through enzymatic dissociation of hPSCs on basement membrane matrix-coated plasticware, embryoid bodies (EBs) are formed. EBs are differentiated to mesoderm by recombinant BMP4, which is subsequently specified to the definitive hematopoietic program by the GSK3&#946; inhibitor, CHIR99021. Alternatively, primitive hematopoiesis is specified by the PORCN inhibitor, IWP2. Hematopoiesis is further driven through the addition of recombinant VEGF and supportive hematopoietic cytokines. The resulting hematopoietic progenitors generated using this method have the potential to be used for disease and developmental modeling, in vitro.

Here, we present human pluripotent stem cell (hPSC) culture protocols, used to differentiate hPSCs into CD34+ hematopoietic progenitors. This method uses stage-specific manipulation of canonical WNT signaling to specify cells exclusively to either the definitive or primitive hematopoietic program.

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Histological Analyses of Acute Alcoholic Liver Injury in Zebrafish

Alcoholic Liver Disease (ALD) refers to damage to the liver due to acute or chronic alcohol abuse. It is among the leading causes of alcohol-related morbidity and mortality and affects more than 2 million people in the United States. A better understanding of the cellular and molecular mechanisms underlying alcohol-induced liver injury is crucial for developing effective treatment for ALD. Zebrafish larvae exhibit hepatic steatosis and fibrogenesis after just 24 h of exposure to 2% ethanol, making them useful for the study of acute alcoholic liver injury. This work describes the procedure for acute ethanol treatment in zebrafish larvae and shows that it causes steatosis and swelling of the hepatic blood vessels. A detailed protocol for Hematoxylin and Eosin (H&#38;E) staining that is optimized for the histological analysis of the zebrafish larval liver, is also described. H&#38;E staining has several unique advantages over immunofluorescence, as it marks all liver cells and extracellular components simultaneously and can readily detect hepatic injury, such as steatosis and fibrosis. Given the increasing usage of zebrafish in modeling toxin and virus-induced liver injury, as well as inherited liver diseases, this protocol serves as a reference for the histological analyses performed in all these studies.

This protocol describes histological analyses of the livers from zebrafish larvae that have been treated with 2% ethanol for 24 h. Such acute ethanol treatment results in hepatic steatosis and swelling of the hepatic vasculature.

Alcoholic Liver Disease (ALD) refers to damage to the liver due to acute or chronic alcohol abuse. It is among the leading causes of alcohol-related ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Defining the Program of Maternal mRNA Translation during In vitro Maturation using a Single Oocyte Reporter Assay

Events associated with oocyte nuclear maturation have been well described. However, much less is known about the molecular pathways and processes that take place in the cytoplasm in preparation for fertilization and acquisition of totipotency. During oocyte maturation, changes in gene expression depend exclusively on the translation and degradation of maternal messenger RNAs (mRNAs) rather than on transcription. Execution of the translational program, therefore, plays a key role in establishing oocyte developmental competence to sustain embryo development. This paper is part of a focus on defining the program of maternal mRNA translation that takes place during meiotic maturation and at the oocyte-to-zygote transition. In this method paper, a strategy is presented to study the regulation of translation of target mRNAs during in vitro oocyte maturation. Here, a Ypet reporter is fused to the 3&#39; untranslated region (UTR) of the gene of interest and then micro-injected into oocytes together with polyadenylated mRNA encoding for mCherry to control for injected volume. By using time-lapse microscopy to measure reporter accumulation, translation rates are calculated at different transitions during oocyte meiotic maturation. Here, the protocols for oocyte isolation and injection, time-lapse recording, and data analysis have been described, using the Ypet/interleukin-7 (IL-7)-3&#39; UTR reporter as an example.

This protocol describes a reporter assay to study the regulation of mRNA translation in single oocytes during in vitro maturation.

Events associated with oocyte nuclear maturation have been well described. However, much less is known about the molecular pathways and processes that ...

Primary Cell Cultures to Study the Regeneration Potential of Murine M&#252;ller Glia after MicroRNA Treatment

M&#252;ller glia (MG) are the predominant glia in the neural retina and can function as a regenerative source for retinal neurons. In lower vertebrates such as fish, MG-driven regeneration occurs naturally; in mammals, however, stimulation with certain factors or genetic/epigenetic manipulation is required. Since MG comprise only 5% of the retinal cell population, there is a need for model systems that allow the study of this cell population exclusively. One of these model systems is primary MG cultures that are reproducible and can be used for a variety of applications, including molecule/factor screening and identification, testing of compounds or factors, cell monitoring, and/or functional tests. This model is used to study the potential of murine MG to convert into retinal neurons after supplementation or inhibition of microRNAs (miRNAs) via transfection of artificial miRNAs or their inhibitors. The use of MG-specific reporter mice in combination with immunofluorescent labeling and single-cell RNA sequencing (scRNA-seq) confirmed that 80%-90% of the cells found in these cultures are MG. Using this model, it was discovered that miRNAs can reprogram MG into retinal progenitor cells (RPCs), which subsequently differentiate into neuronal-like cells. The advantages of this technique are that miRNA candidates can be tested for their efficiency and outcome before their usage in in vivo applications.

M&#252;ller glia primary cultures obtained from mouse retinas represent a very robust and reliable tool to study the glial conversion into retinal progenitor cells after microRNA treatment. Single molecules or combinations can be tested before their subsequent application of in vivo approaches.

M&#252;ller glia (MG) are the predominant glia in the neural retina and can function as a regenerative source for retinal neurons. In lower vertebrates ...