We thank members of the Rosenblatt laboratory for scientific discussions, suggestions, and comments. We would also like to thank Mary Halloran who kindly provided the plasmid encoding RFP-UtrCH and David Grunwald who provided the CK:GFP transgenic zebrafish. Thanks also to Gretchen King and the staff of the Centralized Zebrafish Resource at the University of Utah for excellent maintenance and care of the zebrafish. This work was supported by NIH-NIGMS NIH Director&#x2019;s New Innovator Award 1 DP2 OD002056-01 to JR. GTE was supported by NIH Multidisciplinary Cancer Training Program Grant 5T32 CA03247-8.

Basic Workflow for the Visualization of Actin Dynamics During Cell Extrusion in the Epidermis of Developing Zebrafish 
The epidermis of the developing zebrafish is comprised of two distinct layers, the surface layer (or periderm) and a basal layer of cells 
that contact the basement membrane2. The cells of the outer surface layer undergo apoptosis and are eliminated from the tissue by
 extrusion3 (Figure 1). To visualize this process in real time, we inject RNA encoding red fluorescent protein fused to the
 calponin homology domain of utrophin, an actin binding protein (RFP-UtrCH) 4,5, into one-cell stage transgenic zebrafish expressing green fluorescent protein (GFP) under the stratified epithelia promoter cytokeratin 86 (Figure 2A). Although RFP-UtrCH is ubiquitously expressed in the animal after RNA injection, we focus on the superficial epidermal cells that express both RFP and GFP to follow actin dynamics specifically in the epidermis (Figure 2B). We then treat the larvae with G418 to induce apoptotic cell extrusion and image the epidermis and actin filaments using a spinning disc confocal microscope and acquire 4D datasets. 
Prior to Starting the Experiment
<ol>
	<li>In vitro transcribe the RNA from a linearized DNA template using the mMessage mMachine SP6 kit (Ambion) and purify the capped RNA using the RNeasy Mini kit (Qiagen). Dilute the RNA to &tilde;60ng/ul using sterile water, make 3-5&mu;l aliquots and store at -80&deg;C until ready for injection. Note: Avoid repetitive freeze/thawing to prevent degradation of the RNA.</li>
 <li>Make a mold to hold the embryos during injection by pouring 40mL of 2% agarose in E3 embryo medium (E3) into a 100 x 15mm culture plate and 
	placing a microinjection mold (Adaptive Science Tools, # I-34) in this tray. Once the agarose has solidified, remove the mold to expose the troughs in the 
	agarose. Store the plate at 4&deg;C until ready for use. </li>
 <li>Pull capillary needles for injection using borosilicate glass with filaments (OD 1.0mm, ID 0.78, 10cm length) and a Sutter P-97 Pipette Puller at the following settings (Heat 485, Pull 50, Velocity 60, Delay/Time 90, Pressure 200). Note: Different types of glass will require optimization of the listed conditions.</li>
 <li>Make up 1% Low Melt (LM) agarose in E3 and store 1mL aliquots at 42&deg;C. Be careful as evaporation of E3 from the stocks can increase the agarose concentration.</li>
 <li>Maintain adult zebrafish under standard laboratory conditions, with a regular light/dark cycle of 14 hours light and 10 hours of darkness7. The night before the experiment set up fish to mate by placing 3 female and 2 males in a tank separated by a divider. The next morning, change the water in the tank and pull the divider when the lights come on.</li>
</ol>
1. Injection of RNA Encoding the Fluorescently Tagged Actin Binding Protein
<ol>
	<li>Thaw the RNA on ice. Add 0.5% Phenol Red to the RNA at a 1:1 ratio for a final RNA concentration of &tilde;30ng/&mu;l and load 1&mu;l solution into a pulled capillary needle by back-filling. </li>
 <li>Collect &tilde;75 1-cell stage8 CK:GFP embryos in E3. Pipet the collected embryos into the microinjection mold with a 5 &frac34; Pasteur pipet and gently orient the embryos in the trough with FST Dumont #5 forceps. Note: Take care to work quickly as to avoid injecting into a multi-cell stage embryo, which can result in mosaic expression of the injected RNA.</li>
 <li>Under a standard laboratory dissecting stereomicroscope, inject the RNA into the yolk of the embryos using a pressure-controlled microinjector (Harvard Apparatus PLI-100 Pico-Injector). A red spot (phenol red) should be visible in the embryo after injection. Inject at least 50 embryos for each experiment. (See the previous JoVE protocol for a detailed protocol on RNA injection into zebrafish embryos9). 
	Note: 
 <ol type="A">
 	<li>A volume of &tilde;2nl should be used for injection of RNA into one-cell stage embryos. To determine the amount of RNA injected, dispense a bolus of the RNA into a drop of mineral oil on a calibrated micrometer slide or under a microscope with a scale bar built into the eyepieces. The droplet of RNA should be a perfect sphere. Calculate the amount delivered by using the equation: Volume (in nl) = 4/3&pi;r3. Adjust the volume delivered by changing the injection pressure or time on the instrument. </li>
 <li>Care should also be taken to inject the lowest amount of RNA that allows visualization of the fluorophore, as high concentrations of RNA can be toxic to the developing embryo. To determine appropriate expression level, a dose-curve experiment should be performed before running the experiment.</li>
 
 </ol>
</li>
<li>Sort the embryos to remove any unfertilized or damaged eggs and place all remaining embryos at 28.5&deg;C. </li>
<li>After 24 hours, use a fluorescent dissecting microscope to select zebrafish embryos that have a high level of GFP (epidermis) and RFP (actin) fluorescence. Place the selected embryos at 28.5&deg;C until ready to proceed with the drug treatment to induce apoptosis.</li>
</ol>

2. Induction of Cell Extrusion in Zebrafish Larvae using G418
We have found that exposure to the aminoglycoside antibiotic G418, or Geneticin, causes apoptosis and extrusion of epidermal cells in developing zebrafish larvae3 by an unknown mechanism. Importantly, this treatment only works on larvae that are 4 days post fertilization and older. We find that &tilde; 5-25 cells can be found extruding at any given time from the fin epithelial of G418 treated larval zebrafish. As the amount of apoptotic extruding cells can vary from fish to fish, we recommend mounting multiple fish for imaging.
<ol>
	<li>Collect and place 4 day post fertilization (dpf) zebrafish larvae exhibiting both GFP and RFP fluorescence into a 35 x 10mm culture dish containing 3 milliliters (mLs) of E3. One can treat up to 25 larvae in this size culture dish. </li>
 <li>Carefully replace the media with 3mLs of E3 containing 1mg/mL of G418 and incubate larvae in the drug for 4 hours at 28.5&deg;C. Note that G418 is light sensitive, so take care to keep the culture dish covered or in the dark. </li>
 <li>Remove media and replace with pre-warmed 28.5&deg;C E3 media containing 0.02% Tricaine and proceed to mounting the larvae. </li>
</ol>

3. Mounting Zebrafish Larvae for Imaging

<ol>
	<li>Transfer 3 larvae to a 1.5 ml eppendorf tube and remove most of the E3 after the fish sink to bottom of tube. We have found that up to 3 animals can be properly mounted before the agarose solidifies. </li>
 <li>Using a cut pipet tip, pipet 120ul of 1% LM-agarose (42&deg;C) into the tube, mix, and then gently pipet the larvae and agarose mixture onto the coverglass in the bottom of a MatTek culture dish.</li>
 <li>Using a FST pin holder with an insert pin or tungsten needle attached, quickly orient the larvae so they are straight and flat against the coverglass of the MatTek dish. 
	Note: We typically use a straight pin to first orient the larvae and then an L-shaped pin to ensure they are flat against the coverglass while preventing damage to the specimens. 
</li>
 <li>Once the agarose has solidified, gently fill the dish with E3 containing 0.02% Tricaine.
	Note: Alternatively, G418 can be added to the E3 in the imaging dish to continue inducing extrusion, although long-term exposure to 1mg/mL G418 will kill the larvae. 
</li>
</ol>
4. Live Imaging of Actin Dynamics and Individual Epithelial Cell Behaviors During Cell Extrusion using a Spinning Disc Confocal Microscope

We have found that the entire process of apoptotic cell extrusion from the epidermis of developing zebrafish larvae takes approximately 20 minutes3. Here we demonstrate how to setup a time-lapse imaging experiment to collect a series of z planes through a single layer of the zebrafish epidermis to visualize the extrusion process. We have found that typical widefield fluorescent microscopes cause significant photobleaching of the actin filaments and do not allow for timelapse imaging. Therefore, to maximize our resolution and prevent photobleaching, we use a spinning disc confocal microscope. Below we describe how to set up the experiment using the Andor IQ software with a Nikon microscope, although the principles discussed can be applied to similar microscope set-ups. 
<ol>
	<li>First, turn on the Argon (for excitation of GFP at 488nm) and HeNe (for excitation of RFP at 561nm) lasers, microscope, camera and epifluorescent lamp. </li>
 <li>Within the Andor IQ software (version 1.10.1), create a time series protocol containing the wavelengths you want to image, the frequency of the intervals between image captures and the number times the capture should be repeated. </li>
 <li>Gently place the MatTek dish containing your specimen on the microscope stage and use a 20x objective with transmitted light to focus on the larvae.</li>
 <li>Move the 40X water immersion objective (N.A. 0.8) into the correct position. Using this objective, we can film approximately 20-25 cells per field, which allows us to follow cellular behaviors during extrusion while also resolving sub-cellular actin dynamics. Note: The same mounting and imaging strategies can be applied to upright microscopes, although a 40X water immersion objective with a long working distance must be used. </li>
 <li>Carefully place a drop of water on the top of the 40x objective and quickly place the MatTek dish on top. Note: Zeiss Immersol immersion solution can also be used if evaporation of the water is a problem. </li>
 <li>Identify the specimen using phase or DIC illumination and then use 488nm epifluorescence to focus specifically on the GFP positive epidermis. </li>
 <li>Switch to confocal scanning mode. Adjust the laser intensity and exposure time for each of the channels (488nm and 561nm) accordingly. Take care to balance the laser power and time of exposure for optimal detection of your signal and to prevent any photobleaching or toxicity. </li>
 <li>To identify extruding cells, use epifluorescence to visualize the GFP labeled epidermis and identify a group of cells in a classic rosette pattern, 
	consisting of a collection of cells surrounding a small round cell in the middle. Using confocal scanning mode, there should also be an accumulation 
	of the RFP-labeled actin visible as a ring around the small round cell in the middle, a hallmark of cell extrusion. Once you have identified an extruding 
	cell, quickly set up the microscope to acquire 4D (X-Y-Z-Time) confocal datasets. </li>
 <li>Set the upper and lower points within the z plane to visualize a single layer of the epidermis, including the extruding cell, and select the step size. For time-lapse imaging, set the intervals for image capture and number of repetitions. For example, we typically acquire a z-series spanning 5-10&mu;m, with a 1 &mu;m step size, every 1-2 minutes for 30 minutes. </li>
 <li>To view the data, we typically make 3-D projections of the z series and save the data set as a movie file that is compatible with Quick Time (Apple). In this manner, we can visualize contraction of the actin ring and epithelial behaviors that occur around the dying cell over time. </li>
</ol>
5. Representative Results
<img src="/files/ftp_upload/2689/2689fig1.jpg" alt="Figure 1"/> 
Figure 1. Schematic of apoptotic cell extrusion. Actin filaments are shown in red, GFP positive epidermal cells are green.
<img src="/files/ftp_upload/2689/2689fig2.jpg" alt="Figure 2"/> 
Figure 2. Experimental Workflow. A. Schematic of the workflow for the experiment. B. Expression of RFP-UtrCH in the epidermis of a 4dpf CK:GFP zebrafish. 
<img src="/files/ftp_upload/2689/2689fig3.jpg" alt="Figure 3"/> 
Figure 3. Still frames (z-projections) from a time-lapse movie that follows live actin dynamics during the process of epithelial cell extrusion. Arrows denote the ring of actin formed by neighboring cells which contracts and closes over the course of 2 minutes.

<ol>
	<li>Rosenblatt, J., Raff, M. C., Cramer, L. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+epithelial+cell+destined+for+apoptosis+signals+its+neighbors+to+extrude+it+by+an+actin-+and+myosin-dependent+mechanism.">An epithelial cell destined for apoptosis signals its neighbors to extrude it by an actin- and myosin-dependent mechanism.</a> Curr Biol. 11, 1847-1857 (2001).</li><li>Le Guellec, D., Morvan-Dubois, G., Sire, J. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skin+development+in+bony+fish+with+particular+emphasis+on+collagen+deposition+in+the+dermis+of+the+zebrafish+(Danio+rerio).">Skin development in bony fish with particular emphasis on collagen deposition in the dermis of the zebrafish (Danio rerio).</a> Int J Dev Biol. 48, 217-231 (2004).</li><li>Slattum, G., McGee, K. M., Rosenblatt, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=P115+RhoGEF+and+microtubules+decide+the+direction+apoptotic+cells+extrude+from+an+epithelium.">P115 RhoGEF and microtubules decide the direction apoptotic cells extrude from an epithelium.</a> J Cell Biol. 186, 693-702 (2009).</li><li>Burkel, B. M., Dassow, G. v. o. n., Bement, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Versatile+fluorescent+probes+for+actin+filaments+based+on+the+actin-binding+domain+of+utrophin.">Versatile fluorescent probes for actin filaments based on the actin-binding domain of utrophin.</a> Cell Motil Cytoskeleton. 64, 822-832 (2007).</li><li>Berndt, J. D., Clay, M. R., Langenberg, T., Halloran, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rho-kinase+and+myosin+II+affect+dynamic+neural+crest+cell+behaviors+during+epithelial+to+mesenchymal+transition+in+vivo.">Rho-kinase and myosin II affect dynamic neural crest cell behaviors during epithelial to mesenchymal transition in vivo.</a> Dev Biol. 324, 236-244 (2008).</li><li>Gong, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=fluorescent+protein+expression+in+germ-line+transmitted+transgenic+zebrafish+under+a+stratified+epithelial+promoter+from+keratin8.">fluorescent protein expression in germ-line transmitted transgenic zebrafish under a stratified epithelial promoter from keratin8.</a> Dev Dyn. 223, 204-215 (2002).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book, A Guide for the Laboratory Use of Zebrafish (Danio rerio). , (2000).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Dev Dyn. 203, 253-310 (1995).</li><li>Yuan, S., Sun, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+mRNA+and+morpholino+antisense+oligonucleotides+in+zebrafish+embryos.">Microinjection of mRNA and morpholino antisense oligonucleotides in zebrafish embryos.</a> J Vis Exp. , (2009).</li><li>Curado, S., Stainier, D. Y., Anderson, 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=Nitroreductase-mediated+cell/tissue+ablation+in+zebrafish:+a+spatially+and+temporally+controlled+ablation+method+with+applications+in+developmental+and+regeneration+studies.">Nitroreductase-mediated cell/tissue ablation in zebrafish: a spatially and temporally controlled ablation method with applications in developmental and regeneration studies.</a> Nat Protoc. 3, 948-954 (2008).</li></ol>

Animal Welfare Assurance
All procedures performed in this protocol using the zebrafish, Danio rerio, have been approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Utah (Animal Welfare Assurance #10-07017).

The protocol described here demonstrates a simple and straightforward method to visualize the process of extrusion in a live epithelial tissue. This type of experiment allows us to examine subtleties of actin dynamics not previously appreciated in fixed tissue analyses, and therefore, complements common immunofluorescent methods. We can use this protocol in combination with chemical inhibitors or genetic mutants to better analyze the extrusion process and study disease states associated with the failure to remove and c...

<table border="1">
 <tbody>
 <tr>
 <th>Name</th>
 <th>Type</th>
 <th>Company</th>
 <th>Catalog Number</th>
 <th>Comments</th>
 </tr>
 <tr>
 <td>RNeasy Mini Kit</td>
 <td>Reagent</td>
 <td>Qiagen</td>
 <td>74104</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>mMessage mMachine SP6 Kit</td>
 <td>Reagent</td>
 <td>Ambion</td>
 <td>AM1340</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Crossing Tanks</td>
 <td>Tool</td>
 <td>Thoren Aquatic Systems</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>The Pipet Pump</td>
 <td>Tool</td>
 <td>Bel Art Products</td>
 <td>F3789</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>5 &frac34; Pasteur Pipet </td>
 <td>Tool</td>
 <td>VWR</td>
 <td>14672-608</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microinjection Mold</td>
 <td>Tool</td>
 <td>Adaptive Science Tools</td>
 <td>I-34</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>100x15mm Culture Plate</td>
 <td>Tool</td>
 <td>Fisher</td>
 <td>08-757-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Flamming/Brown Micropipet Puller</td>
 <td>Tool</td>
 <td>Sutter Instrument Company</td>
 <td>Model P97</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Borosilicate Glass Capillaries with Filament </td>
 <td>Tool</td>
 <td>Sutter Instrument Company</td>
 <td>B1400-78-10</td>
 <td>OD 1.0mm, ID 0.78mm, 10cm length</td>
 </tr>
 <tr>
 <td>Electrode Storage Jar</td>
 <td>Tool</td>
 <td>World Precision Instruments</td>
 <td>E210</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Phenol Red</td>
 <td>Reagent</td>
 <td>Sigma</td>
 <td>P0290</td>
 <td>0.5% in DPBS</td>
 </tr>
 <tr>
 <td>Pressure-Controlled Microinjector and Micromanipulator</td>
 <td>Tool</td>
 <td>Harvard Apparatus</td>
 <td>PLI-100</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>35x10mm Culture Dish</td>
 <td>Tool</td>
 <td>Cellstar</td>
 <td>627-160</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>G418 (Geneticin)</td>
 <td>Reagent</td>
 <td>GIBCO</td>
 <td>10131-035</td>
 <td>50 mg/mL in dH20</td>
 </tr>
 <tr>
 <td>Dumont #5 Forceps</td>
 <td>Tool</td>
 <td>Fine Science Tools</td>
 <td>11253-20</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>12cm Pin Holder </td>
 <td>Tool</td>
 <td>Fine Science Tools</td>
 <td>26016-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Insert Pins</td>
 <td>Tool</td>
 <td>Fine Science Tools</td>
 <td>26007-02 and-03</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Glass Bottom Culture Dish with 1.5 coverglass</td>
 <td>Tool</td>
 <td>MatTek</td>
 <td>P35G-1.5-10-C</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Low Melt Agarose</td>
 <td>Reagent</td>
 <td>Fisher</td>
 <td>BP1360-100</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tricaine</td>
 <td>Reagent</td>
 <td>Sigma</td>
 <td>A-5040</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dissecting Microscope</td>
 <td>Microscope</td>
 <td>Leica</td>
 <td>MZ6</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dissecting Microscope</td>
 <td>Microscope</td>
 <td>Nikon</td>
 <td>SMZ645</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fluorescent Dissecting Microscope</td>
 <td>Microscope</td>
 <td>Olympus</td>
 <td>S2X12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Spinning Disc Confocal Microscope</td>
 <td>Microscope</td>
 <td>Nikon</td>
 <td>Eclipse Ti</td>
 <td>Outfitted with a Yokagawa spinning disc head</td>
 </tr>
 <tr>
 <td>CCD Camera</td>
 <td>Camera</td>
 <td>Andor</td>
 <td>DV-885-VP</td>
 <td>1002x1004x8 micron square pixels</td>
 </tr>
 
 </tbody>
</table>

live imaging of cell extrusion from the epidermis of developing zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Watch this Scientific Journal Video about Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish at JoVE.com

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

live-imaging-cell-extrusion-from-epidermis-developing

Research

JoVE Journal

Biology

14.6K 次观看.  University of Utah. 死亡的细胞被挤压协调一致的收缩邻近的细胞，上皮组织，而不破坏的屏障功能。发展中国家斑马鱼的光学清晰度提供了一个极好的系统，以可视化挤出生活上皮。在这里，我们描述的方法来诱导和幼虫斑马鱼的表皮细胞决议形象挤压。

视频: Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Dissection of Organs from the Adult Zebrafish

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental analysis of the embryo and larva is extensive and the morphology has been well documented, descriptions of adult zebrafish anatomy and studies of development of the adult structures and organs, together with techniques for working with adults are lacking. The organs of the larva undergo significant changes in their overall structure, morphology, and anatomical location during the larval to adult transition.  Externally, the transparent larva develops its characteristic adult striped pigment pattern and paired pelvic fins, while internally, the organs undergo massive growth and remodeling.  In addition, the bipotential gonad primordium develops into either testis or ovary.  This protocol identifies many of the organs of the adult and demonstrates methods for dissection of the brain, gonads, gastrointestinal system, heart, and kidney of the adult zebrafish. The dissected organs can be used for in situ hybridization, immunohistochemistry, histology, RNA extraction, protein analysis, and other molecular techniques.  This protocol will assist in the broadening of studies in the zebrafish to include the remodeling of larval organs, the morphogenesis of organs specific to the adult and other investigations of the adult organ systems.

This protocol describes a procedure for identifying and dissecting organs from the adult zebrafish.

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental ...

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 ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.