We thank Emilie Menant for fish care, the Polytechnique Bioimaging Facility, in particular Pierre Mahou,&#160;for assistance with live imaging on their equipment partly supported by R&#233;gion Ile-de-France (interDIM) and Agence Nationale de la Recherche (ANR-11-EQPX-0029 Morphoscope2, ANR-10-INBS-04 France BioImaging). This work was supported by the ANR grants 15-CE13-0016-1, 18-CE13-0024, 20-CE13-0016, and the European Union&#39;s Horizon 2020 research and innovation programme under the Marie Sk&#322;odowska-Curie grant agreement No 840201, the Minist&#232;re de l&#39;Enseignement Sup&#233;rieur et de la Recherche and the Centre National de la Recherche Scientifique.

All animal work was approved by the Ethical Committee N 59 and the Minist&#232;re de l&#39;Education Nationale, de l&#39;Enseignement Sup&#233;rieur et de la Recherche under the file number APAFIS#15859-2018051710341011v3. Some of the steps described below are specific to our equipment and software but could be easily adapted to different equipment.
1. Injection preparation
<ol>
	<li>Prepare 75 mL of 1% agarose solution in Embryo Medium (EM).</li>
	<li>Place the injecting mold in a 90 mm Petri dish and pour approximately 50 mL of agarose, enough for the mold to float. Let the agarose solidify and remove the injecting mold.</li>
	<li>Prepare an agarose-coated dish by pouring 1 mL of agarose in a 30 mm Petri dish.</li>
	<li>Prepare 4 &#181;L of 30 ng/&#181;L Histone2B-mCherry mRNA solution by diluting the stock solution in RNase-free water and keep on ice. 
	&#8203;NOTE: Take care to wear gloves while manipulating mRNA to avoid RNase-mediated degradation.</li>
	<li>Pull an injection needle from a capillary using the micropipette puller.</li>
</ol>
2. Embryo preparation
<ol>
	<li>Once fishes have laid eggs, collect, rinse, and harvest in a 90 mm Petri dish in EM. Place the embryos in a 28.5 &#176;C incubator.</li>
	<li>Wait 20 min for the first cell to become visible.</li>
	<li>Transfer 30 embryos to the injection plate filled with EM. Squeeze embryos in the grooves using slightly blunt forceps and orient them with the animal pole up.</li>
	<li>Using a microloader tip, fill an injection needle with 2 &#181;L of mRNA solution. Insert the needle in the capillary holder placed in a micro-manipulator connected with polytetrafluoroethylene (PTFE) tubing to an air injector.</li>
	<li>Under the stereomicroscope, carefully break the tip of the needle.</li>
	<li>Inject the mRNA solution in the 1-cell stage embryos by inserting the needle in the cell. 
	NOTE: The volume injected is approximately one-third of the cell volume.</li>
	<li>Place back injected embryos in the 28.5 &#176;C incubator.</li>
</ol>
3. Preparation of the two-photon microscope
NOTE: Two lasers are used in this protocol. One is used to image GFP (at 920 nm) and perform ablations (at 820 nm). It will be referred to as the green/ablation laser. The other is used at 1160 nm to image mCherry. It will be referred to as the red laser.
<ol>
	<li>Set the green/ablation laser to 820 nm (ablation wavelength) and the red laser to 1160 nm (mCherry excitation).</li>
	<li>Using movable mirrors on the optical path, align green/ablation and red laser beams both at the entry and exit of the scan head. 
	NOTE: This increases the laser beam focus and minimizes focal volume for excitation and ablation.</li>
	<li>Measure the maximum power of the green/ablation laser at 820 nm under the objective. To do so, place the power meter under the objective, close the black chamber, set green/ablation laser power to 100%, and open the shutters. Compute the percentage of laser power needed to reach 300 mW.</li>
	<li>Set back the green/ablation laser to 920 nm (GFP excitation) and set the laser power to 7%. Set the red laser power to 15%.</li>
	<li>Activate epi-PhotoMultiplier Tubes (PMT) detectors for green and red lines; set green and red line PMT sensitivity to 65.</li>
	<li>Set the field of view to 400 x 400 &#181;m, image resolution to 512 x 512 pixels, and scanning frequency to 800 Hz.</li>
	<li>Select 3D Timelapse Imaging mode. Then, create a folder and activate Autosave for data after each acquisition.</li>
	<li>Assemble the heating chamber and set it to 28 &#176;C. Wait at least 10 min for the chamber and the objective to warm.</li>
</ol>
4. Mounting the embryo
<ol>
	<li>Under a fluorescence stereomicroscope, identify embryos at 70% epiboly that express GFP. 
	NOTE: Select embryos with a bright signal in the axial mesoderm and no background fluorescence for better imaging quality.</li>
	<li>Transfer three to four selected embryos in the agarose coated dish (step 1.3) using a plastic Pasteur pipette and carefully dechorionate them using fine forceps. 
	NOTE: Dechorionated embryos are very delicate and will burst upon contact with air or plastic.</li>
	<li>Pour 1 mL of 0.2% agarose in 1x penicillin-streptomycin EM in a small glass vial. Place the vial in a preheated 42 &#176;C dry block heater. 
	NOTE: The following steps must be performed quickly to allow embryo orientation before agarose sets.</li>
	<li>Transfer a dechorionated embryo in the 0.2% agarose glass vial using a fire-polished glass pipette. Take care not to add too much EM in the agarose to avoid diluting it. Discard the remaining EM from the pipette and aspirate the embryo back along with enough agarose to cover the slide of the glass bottom dish before the embryo falls out of the pipette.</li>
	<li>Blow the agarose and the embryo on the glass slide of the dish. Take care not to let the embryo touch the air or the plastic side of the dish. Next, fill the chamber around the glass slide with agarose.</li>
	<li>Use an eyelash to orient the embryo so that the targeted region is at the top (Figure 1B). 
	NOTE: When orienting embryos, take care to only touch the blastoderm, not the very fragile yolk. Agarose will set in around 1 min, depending on room temperature.</li>
	<li>Wait ~5 min for the agarose to set completely, and then add a few drops of penicillin-streptomycin EM.</li>
</ol>
5. Locating the embryo and pre-ablation imaging
<ol>
	<li>Place the glass bottom dish under the objective in the heated chamber. Immerse the objective in penicillin-streptomycin EM and close the heated chamber.</li>
	<li>Move the slider to set the light path to oculars. Then, using oculars, fluorescent lamps, and stage control, find an embryo and set the focus to the surface of the embryo.</li>
	<li>Turn the fluorescence lamp off, set the light path to PMTs, and close the black chamber. 
	NOTE: Be careful to turn off all light sources in the black chamber as it might damage the PMTs.</li>
	<li>Start live imaging and locate axial mesoderm. Adjust the green/ablation and red laser powers to have a good signal (i.e., between 1,000 and 20,000 photons per pixel for GFP expressing areas). Use the red channel to move the stage to the very top of the embryo and set this position as Z = 0.</li>
	<li>Choose a time-step of 1 min and a Z-step of 2 &#181;m. A Z-course of 110 &#181;m is sufficient to encompass the whole polster and is acquired in less than 1 min with these settings. Set the first slice 15 &#181;m above the axial mesoderm (in the more superficial ectoderm). 
	&#8203;NOTE: The polster moves along a curved line so that the bottom slice of the Z-stack should be set 30 &#181;m deeper than the polster deepest position to accommodate its movement during the time-lapse imaging (Figure 1E).</li>
	<li>Record 10-15 min of pre-ablation movie.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62815/62815fig01.jpg" /> 
Figure 1: Successful outcome of laser ablations. (A) Scheme of a gastrulating embryo at 70% epiboly in dorsal view; pAM: posterior axial mesoderm; black arrow marks the direction of polster migration; black square indicates a typical field of view for ablations in the polster. (B) Scheme of embryo mounting for polster severing. Lateral view. The embryo is mounted such that the plane of the polster is perpendicular to the optical axis. (C) survival and (D)&#160;morphology&#160;of control and ablated embryos at 24 h post-fertilization. Scale bar is 300 &#181;m. (E) Time sequence from laser ablation in the polster of a Tg(gsc:GFP) embryo expressing Histone2B-mCherry. Views with the green channel only are maximum projections. The close-up displays the ablated area containing cell debris. Views with green and red (displayed as magenta) channels are XY and XZ slices before and after ablation (the green lightning bolt represents ablation). XZ slices show that the overlying tissues (magenta nuclei without GFP expression) have not been affected by the ablation of underlying structures. The yellow dashed box corresponds to the ROI selected for laser ablation treatment. The scale bar is 50 &#181;m in large views and 25 &#181;m in the close-up. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62815/62815fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Target location and laser ablation
<ol>
	<li>Locate the polster contour on live imaging and, using the Electro-Optic Modulator Region of Interest (EOM ROI) tool, draw a 20 pixel (15 &#181;m) large rectangle that spans the width of the polster. Place this rectangle in the middle of the polster (Figure 1E).</li>
	<li>Note the axial position of the highest and lowest planes containing polster cells. Ablations will be performed every 10 &#181;m in between these two planes. Take care that the ROI does not overlap the yolk cell on any of these planes.</li>
	<li>Place the stage at the lowest Z position of the interval. Ablations must be performed bottom-up as debris absorb light.</li>
	<li>Set the green/ablation laser wavelength to 820 nm and set the Power Percentage to obtain an exit power of 300 mW (step 3.3).</li>
	<li>Set the Imaging Frequency to 200 Hz.</li>
	<li>Set green/ablation laser imaging EOM to 0 and select ROI-Treat mode.</li>
	<li>Turn on the EOM and set the treatment to start immediately (after 0 frame).</li>
	<li>Set the Imaging Mode to Timelapse and de-activate Autosave.</li>
	<li>Set the Time Step to Fast mode.</li>
	<li>Set the Number of Treatment Frames and Number of Frames to the value corresponding to the targeted depth (Table 1).</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Depth (&#181;m)</td>
			<td>Treatment frames</td>
		</tr>
		<tr>
			<td>-30</td>
			<td>1</td>
		</tr>
		<tr>
			<td>-35</td>
			<td>1-2</td>
		</tr>
		<tr>
			<td>-40</td>
			<td>1-2</td>
		</tr>
		<tr>
			<td>-45</td>
			<td>2</td>
		</tr>
		<tr>
			<td>-50</td>
			<td>2-3</td>
		</tr>
		<tr>
			<td>-55</td>
			<td>3</td>
		</tr>
		<tr>
			<td>-60</td>
			<td>3-4</td>
		</tr>
		<tr>
			<td>-65</td>
			<td>4</td>
		</tr>
		<tr>
			<td>-70</td>
			<td>4</td>
		</tr>
		<tr>
			<td>-75</td>
			<td>4-5</td>
		</tr>
		<tr>
			<td>-80</td>
			<td>4-5</td>
		</tr>
		<tr>
			<td>-85</td>
			<td>5</td>
		</tr>
		<tr>
			<td>-90</td>
			<td>5</td>
		</tr>
		<tr>
			<td>-95</td>
			<td>5-6</td>
		</tr>
		<tr>
			<td>-100</td>
			<td>6</td>
		</tr>
		<tr>
			<td>-105</td>
			<td>6</td>
		</tr>
	</tbody>
</table>
Table 1: Suggested number of laser treatment frames as a function of targeted cell depth in the embryo (0 being the embryo&#39;s surface).
<ol start="11">
	<li>Start imaging. The acquisition is black as the shutter to PMT closes during EOM treatment.</li>
	<li>Move up the stage to the next Z position of the list (step 6.2).</li>
	<li>Repeat steps 6.10 to 6.12 until the top of the polster is reached.</li>
</ol>
7. Post-ablation verification and imaging
<ol>
	<li>Set the green/ablation laser to 920 nm and 5% power. Set the green/ablation laser imaging EOM to 100 and select the Fullfield mode.</li>
	<li>Set the Imaging Frequency to 800 Hz. Turn EOM off.</li>
	<li>Go through the whole stack in live mode to check whether every plane has been ablated. If this is not the case, go back to step 6.2. 
	NOTE: Ablation sometimes induces a vertical shift of neighboring tissues so that the Z-stack might have to be redefined.</li>
	<li>Set the Imaging Mode to 3D Timelapse and re-activate Autosave. Record 40-60 min of post-ablation movie.</li>
	<li>Check, in the post-ablation movie, whether the targeted cells were effectively ablated. Fluorescence recovery, or targeted cells occupying space and preventing follower cells from moving through, indicate that targeted cells were only photobleached and not ablated (Figure 1E and Figure 2A).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62815/62815fig02.jpg" /> 
Figure 2: Negative results of laser ablations. (A) Typical examples of potential failures in laser ablation. Large XY views are maximum projections, XZ view is a reconstructed section. Laser treated area is located between the two white arrowheads. Three focal planes are highlighted in the reconstructed section and displayed on the right. They correspond to three different kinds of failures. Plane 1 shows that cells above the polster have been ablated. This can be identified by the presence of autofluorescent debris on this focal plane (see close-up) above the polster (see position of plane 1 on the reconstructed section). This likely results from an incorrect definition of the region to be ablated. Plane 2 shows cells that have been bleached but not ablated. They can be identified as the low fluorescence signal still reveals intact cell contours (see close-up). Plane 3 displays intact cells, which have hardly been bleached by laser treatment. This could result from an incorrect definition of the region to be ablated or from poor treatment. In the situations depicted in planes 2 and 3, it is possible to re-apply the ablation treatment&#160;to the non-ablated&#160;targeted cells. The scale bar is 50 &#181;m in large views and 20 &#181;m in close-ups. (B) A typical example of bubbles (marked by white asterisks) formed by cavitation because of a too intense laser treatment. Such bubbles are not limited to a Z-plane, sometimes even spanning the full height of the polster, deforming neighboring tissues. The scale bar is 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62815/62815fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
8. Data analysis
<ol>
	<li>Open time-lapse series with the image analysis software and set correct pixel size.</li>
	<li>In the Spot function, set the Object Size to 10 &#181;m, as this is the average nucleus size during gastrulation. Then, run the Spot function to detect and track the nuclei. 
	NOTE: Detection may be slightly improved by considering the lower axial resolution, fitting a 12 &#181;m long ellipsoidal shape along the Z-axis.</li>
	<li>Use filters to remove false positives. In the Tg(Gsc:GFP) line, cells from the embryonic axis and some endodermal cells are labeled in green. Hence, filtering on green intensity allows a quick selection of these cells (Figure 3A).</li>
	<li>Set the maximal distance between consecutive points to a value compatible with the speed of the cells. 
	NOTE: Be careful to consider the time interval between two frames. Polster cells migrate at 2.8 &#177; 0.8 &#181;m/min. Hence, allowing 4 &#181;m of maximum displacement for a time step of 1 min removes most artefactual tracks.</li>
	<li>Allowing gaps over one or two time points provides longer continuous tracks but may introduce tracking errors. If a nucleus is not detected correctly at a one-time point, consider re-running spot detection with different parameters/filters.</li>
	<li>Visually check tracks and, if necessary, correct them.</li>
	<li>Export the results as a .xlsx file. Process the file using published spreadsheet routines24 (Figure 3B) and custom routines on data analysis software (available on request).</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62815/62815fig03.jpg" /> 
Figure 3: Isolation of the anterior half of the polster affects cell directionality. (A) 3D reconstructions a Tg(gsc:GFP) embryo expressing Histone2B-mCherry (displayed in magenta), before and after a laser ablation severing the polster in its middle. Nuclei belonging to the anterior half of the polster are marked with a magenta dot and tracked over time before and after ablation (see Movie S1). The scale bar is 50 &#181;m. (B) As a measure of migration persistence, direction auto-correlation of cells belonging to the anterior part of the polster before and after ablation. Cells display a continuous motion before ablation, which drastically decreases after ablation, indicating loss of collective-oriented migration. Direction auto-correlation was also measured on cells forming the anterior half of the polster of a non-ablated embryo, as a control. The graph envelopes indicate standard error. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62815/62815fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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A., Fielding, T., Houart, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dkk1+controls+cell-cell+interaction+through+regulation+of+non-nuclear+&#946;-Catenin+pools.">Dkk1 controls cell-cell interaction through regulation of non-nuclear &#946;-Catenin pools.</a> Developmental Cell. 51 (6), 775-786 (2019).</li><li>Gorelik, R., Gautreau, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+and+unbiased+analysis+of+directional+persistence+in+cell+migration.">Quantitative and unbiased analysis of directional persistence in cell migration.</a> Nature Protocols. 9 (8), 1931-1943 (2014).</li><li>Grill, S. W., Howard, J., Sch&#228;ffer, E., Stelzer, E. H. K., Hyman, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+distribution+of+active+force+generators+controls+mitotic+spindle+position.">The distribution of active force generators controls mitotic spindle position.</a> Science. 301 (5632), 518-521 (2003).</li><li>Desprat, N., Supatto, W., Pouille, P. -. A. A., Beaurepaire, E., Farge, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+deformation+modulates+twist+expression+to+determine+anterior+midgut+differentiation+in+Drosophila+embryos.">Tissue deformation modulates twist expression to determine anterior midgut differentiation in Drosophila embryos.</a> Developmental Cell. 15 (3), 470-477 (2008).</li><li>Farhadifar, R., R&#246;per, J. -. C., Aigouy, B., Eaton, S., J&#252;licher, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+influence+of+cell+mechanics,+cell-cell+interactions,+and+proliferation+on+epithelial+packing.">The influence of cell mechanics, cell-cell interactions, and proliferation on epithelial packing.</a> Current Biology. 17 (24), 2095-2104 (2007).</li><li>Willier, B. H., Oppenheimer, J. 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> Foundations of Experimental Embryology. , (1964).</li><li>Ashby, W. J., Zijlstra, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Established+and+novel+methods+of+interrogating+two-dimensional+cell+migration.">Established and novel methods of interrogating two-dimensional cell migration.</a> Integrative Biology: Quantitative Biosciences from Nano to Macro. 4 (11), 1338-1350 (2012).</li><li>Bosze, B., 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=Pcdh18a+regulates+endocytosis+of+E-cadherin+during+axial+mesoderm+development+in+zebrafish.">Pcdh18a regulates endocytosis of E-cadherin during axial mesoderm development in zebrafish.</a> Histochemistry and Cell Biology. 154 (5), 463-480 (2020).</li></ol>

The authors declare no competing interests.

Here, we describe a protocol that uses non-linear optics to perform deep and spatially well-defined volume ablations. The most critical step of the protocol is to find treatment conditions that provide sufficient energy to allow ablations, but not too much energy, to avoid excessive debris or cavitation. The amount of delivered energy at the target site mainly depends on: (1) the laser exit power, (2) the quality of laser alignment, (3) the nature of the tissue through which the light passes to reach the ablation plane, (4) the depth of the ablation plane. Therefore, before each experiment, it is crucial to measure laser exit power, adjust it to a reference value (300 mW at 820 nm in our protocol), and ensure proper laser alignment. Under these assumptions, treatment conditions should be reproducible from one experimental day to another. We recommend performing extensive tests to define optimal parameters (laser power and number of treatment frames) for a specific sample type. These parameters can then be used in all similar experiments. In the example described here (severing of the polster during gastrulation), we have, for instance, established treatment conditions for ablations at different depths within the embryo (Table 1) and now rely on this chart when performing experiments. Of note, the 820 nm wavelength was chosen as it is, on our system, the wavelength providing the highest peak energies (due to laser and optics properties). Shorter or longer wavelengths could be used depending on system properties6,11,12. Depth of the targeted tissue being a critical parameter, embryo mounting is also a crucial step, as incorrect mounting may increase tissue thickness that light must pass through to reach the target volume.
One of the original features of the described protocol is to ablate an entire volume by performing successive ablations on different focal planes. Since ablations will generate debris that absorbs light, we identified that it is crucial to start ablation on the most profound plane and sequentially ablate from the deepest to the most superficial plane.
This protocol describes the ablation of deep and large volumes and the recording of neighboring tissue response within minutes after ablation to over an hour. One of the potential limitations of the protocol is the time required to perform ablation and restart imaging. Two factors limit this delay. The first one is the time needed to perform ablations on multiple focal planes. On our system, severing of the polster is performed by a trained user in 2-3 min. This could be reduced by optimizing the software to automate the ablation on different planes. Still, total ablation time will equal the time required to scan the target region, times the number of repetitions on each focal plane, times the number of focal planes, which, in our conditions, is about 1 min. Considering cell migration speed, this means that some cells may enter or exit the targeted area during the ablation procedure. In our case, this did not prove to be an issue but could be if an absolute precision in cell targeting is required. The second limiting factor is that the same laser is used to perform ablation (at 820 nm) and excite green fluorophore (at 920 nm). The delay between the last ablation and the start of recording is thus defined by the time required to tune the laser from 820 nm to 920 nm, ranging from 30 s to 1 min.
In some cases, in particular, for smaller ablations (single-cell ablation, ablations of subcellular components such as cytoskeleton elements), recording the immediate response of the cell/tissue may be critical to infer its mechanical state25,26,27. In such cases, the limitation could be circumvented, either by imaging with another laser (here, for instance, recording only red signals with the 1160 nm laser or&#160;using a third laser line) or imaging green fluorophore at 820 nm. This is not the optimal wavelength for imaging (limited excitation of fluorophores, strong photo-toxic effects) but could be used over short periods to record immediate tissue response.
Few techniques are available to eliminate cells and see how this affects the rest of the embryo. The two main options are to remove cells physically or to destroy them, as in laser ablations. Compared to physical removal, cell destruction may release cytoplasmic content, which can influence neighboring cells. This was historically highlighted by the controversy and diverging results obtained by Wilhelm Roux and Hans Driesch regarding the mosaic or regulative development of the frog and sea urchin embryo28. More recently, differences have been observed in wound healing assays, depending on whether the wound is created by scratch (which destroys some cells) or removing an insert29. However, physically removing cells without damaging other tissues is only possible for cells at the very surface of the embryo and cells that are not too adherent to their neighbors, thus limiting the range of such approaches. Consequently, different strategies have been developed to destroy cells, laser ablations being the most employed. UV laser ablations have been and are increasingly used, in particular, to perform small, superficial ablations, and observe immediate tissue response.
We here described the use of infrared light to perform deeper and spatially well-defined ablations. The main limitation of this protocol is the requirement for an infrared pulsed laser and a two-photon imaging setup. However, such equipment is becoming more and more frequent on imaging platforms. In addition, the EOM used here to ablate one region in the image selectively could be replaced by a Fluorescence Recovery After Photobleaching (FRAP) module. Though less convenient, it could even be possible to perform the protocol without EOM nor FRAP modules by simply zooming on the targeted area10. Using a pulsed infrared laser brings two main advantages compared to most classical ablation protocols. First, thanks to the efficient penetration of infrared light in living tissues, deep focal planes can be reached with laser powers that do not induce out-of-focus damages. This allowed us to target cells as deep as 120 &#181;m, out of reach with one-photon excitation protocols. Second, the use of non-linear optics ensures excellent axial resolution, permitting precisely controlled 3D ablations, even at depth in the tissue. Combining these two advantages allows ablation of specifically defined, deep, and eventually large volumes.
We describe using a two-photon microscope to sever the polster, an experiment we and others recently performed30. With few adjustments, the proposed protocol could, however, be adapted to many different samples. We have, for instance, successfully used it to perform complete ablation of the polster, ablations within the lateral mesoderm during gastrulation, or ablations of individual Schwan cells during their migration on their associated axon, without affecting the axon. We, therefore, believe this protocol is a valuable and versatile tool, which should be helpful in many experimental systems to analyze the impact of some cells/tissues on the behavior and development of the neighboring structures.

Cell-cell interactions play vital roles in development. Cells provide signals that their direct neighbors, or cells further away, can perceive, thereby influencing their fate and/or behavior. Many of these signals are chemical in nature. For instance, in the well-characterized induction events, one cell group produces diffusible molecules affecting the fate of another cell population1. Other signals, however, are mechanical; cells exert forces and constraints on their neighbors, which the neighbors perceive and respond to2.
One way of studying the importance of these cell-cell interactions in vivo is to eliminate some cells and observe subsequent development. Unfortunately, available techniques to remove or destroy cells are limited. Cells can be removed surgically3,4, using needles or small wires, but such treatments are invasive, not very precise, and usually performed under a stereomicroscope, preventing immediate imaging under a microscope. Furthermore, targeting deep cells implies piercing a hole in overlying tissues, creating unwanted perturbations. Genetically encoded photosensitizers, such as KillerRed, have been used to induce cell death via light illumination5. Photosensitizers are chromophores that generate reactive oxygen species upon light irradiation. Their main limitation is that they require long light illuminations (around 15 min), which may be difficult to achieve if cells are moving, and that they induce cell death through apoptosis, which is not immediate.
Finally, laser ablations have been developed and widely used in the past 15 years6,7,8,9,10,11,12. A laser beam is focused on the targeted cell/tissue. It induces its ablation through heating, photoablation, or plasma-induced ablation; the involved process depends on the power density and exposure time13. Most ablation protocols use UV lasers for their high energy. However, UV light is both absorbed and scattered by biological tissues. Thus, targeting deep cells requires a high laser power, which then induces damages in more superficial, out-of-plane tissues. This limits the use of UV lasers to superficial structures and explains their relatively low axial resolution. Non-linear optics (so-called two-photon microscopy) uses non-linear properties of light to excite a fluorophore with two photons of approximately half-energy in the infrared domain. When applied to ablations, this has three main advantages. First, the infrared light is less scattered and less absorbed than UV light by biological tissues14, allowing to reach deeper structures without increasing the required laser power. Second, the use of a femtosecond pulsed laser provides very high power densities, creating an ablation through plasma induction, which, contrary to heating, does not diffuse spatially15. Third, the power density inducing plasma formation is reached at the focal point only. Thanks to these properties, two-photon laser ablations can be used to precisely target deep cells without affecting the surrounding tissue environment.
Collective migrations are an excellent example of developmental processes in which cell-cell interactions are fundamental. Collective migrations are defined as cell migrations in which neighboring cells&#160;influence the behavior of one cell16. The nature of these interactions (chemical or mechanical) and how they affect cell migration can vary greatly and is often not entirely understood. The ability to remove cells and observe how this affects the others is critical in further unraveling these collective processes. A few years ago, we established &#8212; using surgical approaches &#8212; that the migration of the polster during zebrafish gastrulation is a collective migration17. The polster is a group of cells that constitutes the first internalizing cells on the dorsal side of the embryo18. These cells, labeled in green in the Tg(gsc:GFP) transgenic line, are located deep in the embryo, below several layers of epiblast cells. During gastrulation, this group leads the extension of the axial mesoderm, migrating from the embryonic organizer to the animal pole19,20,21,22,23 (Figure 1A). We established that cells require contact with their neighbors to orient their migration in the direction of the animal pole. However, better understanding the cellular and molecular bases of this collective migration involves removing some cells to see how this influences the remaining ones. We, therefore, developed ablations of large and deep volumes using a two-photon microscopy setup. Here, we demonstrate the use of this protocol to sever the polster in its middle and observe the consequences on cell migration by tracking nuclei labeled with Histone2B-mCherry.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>25x water immersion objective</td>
			<td>Olympus</td>
			<td>XLPLN25XWMP2</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>PanReac AppliChem</td>
			<td>A8963,0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Data analysis software : Matlab</td>
			<td>Math Works</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electro-optic modulator (EOM)</td>
			<td>ConOptics</td>
			<td>350-80LA</td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo Medium (EM) solution</td>
			<td></td>
			<td></td>
			<td>Westerfield, M. The Zebrafish Book. A Guide for the Laboratory Use of Zebrafish (Danio rerio), 5th Edition. University of Oregon Press, Eugene (Book). (2000).</td>
		</tr>
		<tr>
			<td>Environmental chamber chamber</td>
			<td>Okolab</td>
			<td>H201-T-UNIT-BL</td>
			<td></td>
		</tr>
		<tr>
			<td>EOM driver</td>
			<td>ConOptics</td>
			<td>302RM</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence source</td>
			<td>Lumencor</td>
			<td>SOLA</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass bottom dishes</td>
			<td>MatTek</td>
			<td>P35G-0-10-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass capillaries</td>
			<td>Harvard Apparatus</td>
			<td>300085</td>
			<td>Outside diameter 1.0 mm, inside diameter 0.58&#160;mm</td>
		</tr>
		<tr>
			<td>Glass pipettes</td>
			<td>Volac</td>
			<td>D810</td>
			<td>Tip should be fire polished</td>
		</tr>
		<tr>
			<td>Green/ablation laser</td>
			<td>Spectra Physics</td>
			<td>Mai Tai HP DeepSee</td>
			<td></td>
		</tr>
		<tr>
			<td>Histone2B-mCherry mRNA</td>
			<td></td>
			<td></td>
			<td>Synthesized from pCS2-H2B-mCherry plasmid (Dumortier&#38; al. 2012)</td>
		</tr>
		<tr>
			<td>Image analysis software: IMARIS</td>
			<td>Bitplane</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImSpector software</td>
			<td>Abberior Instruments Development Team</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Injection mold</td>
			<td>Adapative Science Tools</td>
			<td>I-34</td>
			<td></td>
		</tr>
		<tr>
			<td>Microloader tips</td>
			<td>Eppendorf</td>
			<td>5242956003</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Narishige</td>
			<td>MN-151</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter</td>
			<td>P-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>mMESSAGE mMACHINE SP6 Transcription Kit</td>
			<td>Invitrogen</td>
			<td>AM1340</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Thermofisher</td>
			<td>15140-122</td>
			<td>10 000 units penicillin and 10 mgstreptomycin per ml</td>
		</tr>
		<tr>
			<td>Photomultiplier tube (PMT)</td>
			<td>Hammamatsu</td>
			<td>H7422-40</td>
			<td></td>
		</tr>
		<tr>
			<td>PicoPump (Air injector)</td>
			<td>World Precision Instrument</td>
			<td>PV820</td>
			<td></td>
		</tr>
		<tr>
			<td>Red laser</td>
			<td>Spectra Physics</td>
			<td>OPO/Insight DeepSee</td>
			<td></td>
		</tr>
		<tr>
			<td>RNAse free water for injection</td>
			<td>Sigma</td>
			<td>W3500</td>
			<td></td>
		</tr>
		<tr>
			<td>Spreadsheet software: Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Nikon</td>
			<td>SMZ18</td>
			<td></td>
		</tr>
		<tr>
			<td>Tg(gsc:GFP) zebrafish line</td>
			<td></td>
			<td></td>
			<td>Doitsidou, M. et al. Guidance of primordial germ cell migration by the chemokine SDF-1. Cell. 111 (5), 647&#8211;59, doi: doi.org/10.1016/S0092-8674(02)01135-2 (2002).</td>
		</tr>
		<tr>
			<td>TriM Scope II microscope</td>
			<td>La Vision Biotech</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

deep and spatially controlled volume ablations using a two-photon microscope in the zebrafish gastrula

Morphogenesis involves many cell movements to organize cells into tissues and organs. For proper development, all these movements need to be tightly coordinated, and accumulating evidence suggests this is achieved, at least in part, through mechanical interactions. Testing this in the embryo requires direct physical perturbations. Laser ablations are an increasingly used option that allows relieving mechanical constraints or physically isolating two cell populations from each other. However, many ablations are performed with an ultraviolet (UV) laser, which offers limited axial resolution and tissue penetration. A method is described here to ablate deep, significant, and spatially well-defined volumes using a two-photon microscope. Ablations are demonstrated in a transgenic zebrafish line expressing the green fluorescent protein in the axial mesendoderm and used to sever the axial mesendoderm without affecting the overlying ectoderm or the underlying yolk cell. Cell behavior is monitored by live imaging before and after the ablation. The ablation protocol can be used at different developmental stages, on any cell type or tissue, at scales ranging from a few microns to more than a hundred microns.

To sever the polster in its middle, a Tg(gsc:GFP) embryo, injected with Histone2B-mCherry mRNAs was mounted at the 70% epiboly stage, as described in step 4. The polster was identified by GFP expression, and the embryo was mounted so that the plane of the polster is perpendicular to the optical axis (Figure 1B). Tilting the embryo away from this position will complicate the procedure. The light will have to go through more tissues to reach the ablation planes, and ablation planes will be tilted relative to embryonic axes. Having verified that all the cell nuclei are correctly labeled, a 10 min pre-ablation time-lapse was recorded to capture cell movements before ablation (Figure 1E and Figure 3A, Movie S1).
The polster was morphologically identified, and a rectangular area of 15 &#181;m x 200 &#181;m, located in its middle, was ablated on five focal planes to ensure severing on the whole depth of the polster (Figure 1E,&#160;Movie S1). Imaging was restarted right after ablation and used to monitor the efficiency of the procedure. If successful, the ablation will have eliminated all cellular structures and GFP and mCherry fluorescence so that the ablated volume should appear as a signal-free volume. However, some debris may be created. This debris is autofluorescent, both in the green and red channels, and usually displays irregular elongated shapes parallel to the direction of ablation (Figure 1E). A too intense treatment will form a large amount of debris that may act as an obstacle&#160;and perturb cell behavior. Stronger treatments will even induce cavitation, marked by the formation of bubbles in the tissue (Figure 2B). Cavitation is associated with a mechanical shock wave propagating in tissues and may induce damages out of the targeted volume13,15. Embryos with cavitation bubbles should be discarded, and treatment should be tuned down by performing fewer treatment frames.
Conversely, too little treatment may photobleach fluorophores without inducing plasma formation, hence without ablating (Figure 2A). Incomplete photobleaching can easily be spotted by the presence of dim fluorescence with a characteristic cellular shape (Figure 2A). Such embryos should be discarded or treated again performing more treatment frames. Complete photobleaching is more challenging to differentiate from successful ablation, as both would result in a signal-free volume. Photobleaching can, however, be identified retrospectively, as fluorescence will progressively recover in the course of the post-ablation imaging. This, however, implies that non-ablated embryos are imaged for at least half an hour, which is time-consuming. We, therefore, suggest adjusting the treatment intensity (by increasing the number of treatments) to induce the formation of few visible debris, which will not affect cell behavior but immediately confirm effective ablation. Finally, the absence of damage in cells surrounding the ablation volume should be checked on the first post-ablation images (Figure 2A). When laser treatment is tuned correctly (formation of few debris), damages in neighboring tissues are unlikely to result from the spatial spreading of the ablation, which is very well spatially defined, but rather result from inaccurate selection of the region to be ablated and/or tissue movements in the time between target selection and ablation. Embryos with affected neighboring tissues should be discarded.
After successful ablation, Z-stacks were captured every minute for 40 min, recording both the GFP cytoplasmic signal and the mCherry nuclear signal. Nuclei were then tracked, and their movement was used as a proxy for cell movement. Tracks corresponding to polster cells were identified on their strong GFP signal&#160;(Figure 3A, Movie S1). The persistence of cell movement was measured by computing the cell direction auto-correlation24. Focusing on polster cells located in the anterior half of the polster revealed that severing the polster in its middle, thus separating these cells from the posterior part of the polster, decreased their direction autocorrelation (Figure 3B), demonstrating that proper migration of polster cells requires integrity of the whole polster, in line with its demonstrated collective migration17.
After acquiring the post-ablation movie, embryos can be unmounted, carefully extracting them from the agarose using fine forceps, and incubated at 28.5 &#176;C until they reach 24 h post-fertilization. Again, embryos should survive and should not present any apparent morphological defect (Figure 1C,D).
Movie S1: Successful laser ablation. Laser ablation in the middle of the polster of a Tg(gsc:GFP) embryo expressing Histone2B-mCherry. Nuclei from the anterior part of the polster are tracked over time and marked by magenta dots. Tracks are time color-coded (Figure 3). Empty frames correspond to laser ablation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62815/MovieS1.mp4" target="_blank">Please click here to download this Movie.</a>

Watch this Scientific Journal Video about Deep and Spatially Controlled Volume Ablations using a Two-Photon Microscope in the Zebrafish Gastrula at JoVE.com

Deep and Spatially Controlled Volume Ablations using a Two-Photon Microscope in the Zebrafish Gastrula

Embryonic development requires large-scale coordination of cell motion. Two-photon excitation mediated laser ablation allows the spatially controlled 3-dimensional ablation of large groups of deep cells. In addition, this technique can probe the reaction of collectively migrating cells in vivo to perturbations in their mechanical environment.

Morphogenesis involves many cell movements to organize cells into tissues and organs. For proper development, all these movements need to be tightly ...

deep-spatially-controlled-volume-ablations-using-two-photon

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

1.9K Views.  CNRS UMR7645, INSERM U1182, Institut Polytechnique de Paris. Embryonic development requires large-scale coordination of cell motion. Two-photon excitation mediated laser ablation allows the spatially controlled 3-dimensional ablation of large groups of deep cells. In addition, this technique can probe the reaction of collectively migrating cells in vivo to perturbations in their mechanical environment.

Video: Deep and Spatially Controlled Volume Ablations using a Two-Photon Microscope in the Zebrafish Gastrula

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

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 ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many myopathies affecting skeletal muscle function can be quickly and easily assessed in zebrafish over the first few days of embryogenesis. By 24 hr post-fertilization (hpf), wildtype zebrafish spontaneously contract their tail muscles and by 48 hpf, zebrafish exhibit controlled swimming behaviors. Reduction in the frequency of, or other alterations in, these movements may indicate a skeletal muscle dysfunction. To analyze swimming behavior and assess muscle performance in early zebrafish development, we utilize both touch-evoked escape response and locomotion assays.
Touch-evoked escape response assays can be used to assess muscle performance during short burst movements resulting from contraction of fast-twitch muscle fibers. In response to an external stimulus, which in this case is a tap on the head, wildtype zebrafish at 2 days post-fertilization (dpf) typically exhibit a powerful burst swim, accompanied by sharp turns. Our method quantifies skeletal muscle function by measuring the maximum acceleration during a burst swimming motion, the acceleration being directly proportional to the force produced by muscle contraction.
In contrast, locomotion assays during early zebrafish larval development are used to assess muscle performance during sustained periods of muscle activity. Using a tracking system to monitor swimming behavior, we obtain an automated calculation of the frequency of activity and distance in 6-day old zebrafish, reflective of their skeletal muscle function. Measurements of swimming performance are valuable for phenotypic assessment of disease models and high-throughput screening of mutations or chemical treatments affecting skeletal muscle function.

Zebrafish are an excellent model to study muscle function and disease. During early embryogenesis zebrafish begin regular muscle contractions producing rhythmic swimming behavior, which is altered when the muscle is disrupted. Here we describe a touch-evoked response and locomotion assay to examine swimming performance as a measure of muscle function.

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Confocal Microscope-Based Laser Ablation and Regeneration Assay in Zebrafish Interneuromast Cells

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally replenished in non-mammalian vertebrates such as zebrafish. The zebrafish lateral line system is a useful model for characterizing sensory hair cell regeneration. The lateral line is comprised of hair cell-containing organs called neuromasts, which are linked together by a string of interneuromast cells (INMCs). INMCs act as progenitor cells that give rise to new neuromasts during development. INMCs can repair gaps in the lateral line system created by&#160;cell death. A method is described here for selective INMC ablation using a conventional laser-scanning confocal microscope and transgenic fish that express green fluorescent protein in INMCs. Time-lapse microscopy is then used to monitor INMC regeneration and determine the rate of gap closure. This represents an accessible protocol for cell ablation that does not require specialized equipment, such as a high-powered pulsed ultraviolet laser. The ablation protocol may serve broader interests, as it could be useful for the ablation of additional cell types, employing a tool set that is already available to many users. This technique will further enable the characterization of INMC regeneration under different conditions and from different genetic backgrounds, which will advance the understanding of sensory progenitor cell regeneration.

Laser ablation is a widely applicable technology for studying regeneration in biological systems. The presented protocol describes use of a standard laser-scanning confocal microscope for laser ablation and subsequent time-lapse imaging of regenerating interneuromast cells in the zebrafish lateral line.

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally ...