These studies were funded by grants received from the MEXT, JSPS KAKENHI Grant Numbers JP25290009, JP25650120, JP17K07494, and JP17H05984.

1. Ablation of a Subpopulation of Neurons Using a Two-photon Laser Microscope &#12288;
Note: If users plan on performing Ca imaging following ablation, use the UAShspzGCaMP6s line21. If users plan on performing behavioral recording following ablation, use the UAS:EGFP line, as the ablation of EGFP-positive cells is easier to perform than of GCaMP6s-expressing cells.
<ol>
	<li>Begin by setting up the mating of a Gal4 line that labels specific neurons to be studied and UAS:EGFP or UAShspzGCaMP6s. Be sure to use the nacre background for both parents so that nacre homozygotes can be obtained. 
	Note: Homozygotes of nacre contain no melanophores on the body surface (Figure 1B). In the present study, a Gal4 line gSAIzGFFM119B (which labels the pretectum) and another Gal4 line hspGFFDMC76A (which labels the ILH) are used21.</li>
	<li>On the following day of the mating setup, collect the zebrafish eggs, and raise them to the larval stage at which point the neurons of interest will express fluorescent reporters.</li>
	<li>Mount the zebrafish larva in 2% low melting point-agarose (Figure 1B).
	<ol>
		<li>Begin by preparing 2% agarose stock solution: dissolve 2 g of low melting-point agarose powder in 100 mL system water (i.e., the water used to maintain adult zebrafish in a circulating water system)22 or E3 water23, by bringing the water to the boiling point in a microwave, and mixing vigorously.</li>
		<li>Microwave the 2% low melting-point agarose stock until it boils. Pour 3.5-4.0 mL of the agarose onto the lid of a 6-cm Petri dish. Wait until the agarose cools so that it is warm to the touch before proceeding.</li>
		<li>Next, place a single larva (not anesthetized at this step) in the agarose. Keep adjusting the position and orientation of the larva so that it is upright, using a dissecting needle (Figure 1C), until the agarose solidifies to ensure the proper positioning of the larva. 
		Note: The larva will continue to swim around while the agarose solidifies.</li>
		<li>Once the larva is positioned, allow 5 min for the agarose to solidify completely.</li>
		<li>Pour 5 mL of tricaine solution (0.4% at 1x working concentration) over the agarose. 
		Note: To avoid movements by the larvae during laser ablation, the larvae must be kept anesthetized throughout the ablation procedure.</li>
	</ol>
	</li>
	<li>Ablate the larva using the bleaching function in a two-photon laser microscopy system.
	<ol>
		<li>Place the agarose-embedded larva under a two-photon laser scanning microscope and start the microscopy system.</li>
		<li>Start the image acquisition software and click the &#34;On&#34; button on the &#34;Laser&#34; tab to turn on the laser (Figure 1D). Wait for the &#34;Status&#34; of the laser to change from &#34;Busy&#34; to &#34;Mode-locked.&#34;</li>
		<li>On the &#34;Channels&#34; tab, set the laser&#39;s wavelength at 880 nm (maximum power: approximately 2,300 mW) for EGFP ablation, or at 800 nm (maximum power: approximately 2,600 mW) for GCaMP ablation.</li>
		<li>Select the 20X objective lens by manually moving the lens revolver. Click the &#34;Locate&#34; tab, click &#34;GFP&#34; to change the optical pathways, and directly view the fluorescence by eye.</li>
		<li>Locate the zebrafish larval brain at the center of view; then click the &#34;Acquisition&#34; tab to go back to the two-photon microscopy.</li>
		<li>As a record of the &#39;before ablation&#39; condition, take a z-stack image with the 20X objective lens at fast scanning-speed (to avoid heat-damage). Later compare the images taken before and after ablation experiments (Figure 2A). To obtain a z-stack, click &#34;Z-Stack&#34; to select the z-stack option and expand the tab. Set the lower limit (click the &#34;Set First&#34; button&#34;) after focusing on the ventral end of the larval brain, and set the upper limit (click the &#34;Set Last&#34; button) after focusing on the dorsal most-surface of the brain. Then, click the &#34;Start Experiment&#34; button to run the z-stack image acquisition.</li>
		<li>Select the 63X objective lens by manually moving the lens revolver.</li>
		<li>Click the &#34;Locate&#34; tab to switch to the epi-fluorescent microscopy, locate the cells to be ablated at the center of view by eye, then click the &#34;Acquisition&#34; tab to go back to laser scanning microscopy.</li>
	</ol>
	</li>
	<li>On the &#34;Acquisition Mode&#34; tab, set the &#34;Frame Size&#34; at 256 x 256. Click &#34;Live&#34; and observe the neurons of interest.</li>
	<li>Starting from the dorsal-most side, find a focal plane in which the cells to be ablated are in focus.</li>
	<li>Mark a small, circular area about one third the cell&#39;s size in diameter on each neuron as a region of interest (ROI) using the &#34;Regions&#34; function.</li>
	<li>Set the scan speed to 13.93 s/256 x 256 pixels (134.42 &#181;m x 134.42 &#181;m), which corresponds to a laser dwell time of approximately 200 &#181;s/pixel, or 200 &#181;s/0.5 &#181;m. Also, set 4 repetitions (&#39;Iteration cycle: 4&#39;). Alternatively, optimize the number of iterations and scanning speed so that sufficient ablation is achieved.</li>
	<li>Perform the &#39;Bleaching&#39; function for ablation, in combination with the &#39;Time series (2 cycles)&#39; to image the sample (before and after ablation) and &#39;Regions&#39; (to set the ROIs) or equivalent functions in the software used (Figure 1D).</li>
	<li>By comparing the first image (before ablation) and the second image (after ablation) in the Time series cycle, ensure that after bleaching, the fluorescence in the targeted cells decreases to that observed at the background level (Figure 2B).</li>
	<li>If fluorescence is still present after ablation, increase the number of iterations.</li>
	<li>Next, move the focal plane slightly deeper (i.e., towards the ventral side), and choose the next focal plane where un-ablated cells appear.</li>
	<li>Perform the bleaching function as described above (step 1.9). Repeat these steps until all the cells in the neural structure of interest have been ablated.</li>
	<li>After going through all the focal planes covering the neural structures to be ablated, check the cells for abolished fluorescence. If some cells are found to still be fluorescent due to insufficient ablation, laser-irradiate them again as described above.</li>
	<li>If the larva needs to be recovered from agarose after laser irradiation, do not try to force it out of the agarose because this might damage the larva. Instead, carefully make tiny cuts in the agarose around the larva perpendicularly to the surface of its body. Then, wait for the larva to swim out of agarose on its own when the anesthetic wears off.</li>
	<li>Proceed to the next larva for ablation or perform control experiments using another population of neurons that are uninvolved in the behavior under investigation. 
	Note: Here, as a control, olfactory bulb neurons in UAS:EGFP; gSAIzGFFM119B larvae were laser-ablated using the same protocol described above (Figure 2A, right).</li>
	<li>Allow several hours or 1 day for recovery before proceeding to Ca imaging (Section 2) or behavioral recording (Section 3). During this recovery time, check the larval health (i.e., normal spontaneous swimming, no apparent heat-damage around the ablated area, etc.) and remove larvae showing any signs of poor health.</li>
</ol>
2. Calcium Imaging to Record Prey-evoked Neuronal Activity in the Pretectum-ablated Zebrafish Larvae
<ol>
	<li>To prepare a recording chamber, put a secure-seal hybridization chamber gasket on a glass slide, with the adhesive side facing on the glass, and remove the top seal. 
	Note: This creates a small recording chamber that is 9 mm in diameter and 0.8 mm in depth (Figure 3A).</li>
	<li>Next, place a nylon mesh (size of opening: 32 &#181;m) on a 50-mL tube that has the bottom cut open. Use the cap to attach the mesh on the tube (Figure 3B).</li>
	<li>Prepare the paramecia (which is the prey to be used as the visual stimulus for larva). 
	Note: Prepare the Paramecium culture (steps 2.3.1 and 2.3.2) beforehand.
	<ol>
		<li>Obtain paramecia seed culture from commercial sources or academic bio-resource centers24 in advance.</li>
		<li>Culture the paramecium for several days in water containing autoclaved rice straw and a pellet of dry beer-yeast22 (Figure 3C).</li>
		<li>Pour the paramecium culture through 2 sieves successively (one with a 150-&#181;m aperture, followed by another with a 75-&#181;m aperture) to remove debris from the culture.</li>
		<li>Collect paramecia in the flow-through from the previous step on a nylon mesh (13-&#181;m aperture).</li>
		<li>Re-suspend the paramecia on the nylon mesh into a glass beaker using a squeeze bottle of system water (Figure 3D).</li>
	</ol>
	</li>
	<li>Rinse the mesh (Figure 3B) in system water 2x (Figure 3E). 
	Note: The purpose of this step is to remove any possible odorants and tastants contained in the paramecium culture medium, as the goal of the present study is to observe visually driven neuronal activity.</li>
	<li>Place a zebrafish larva in the tricaine solution to anesthetize.</li>
	<li>Mount a single larva in 2% low melting-point agarose.
	<ol>
		<li>First, microwave 2% low melting-point agarose and add 1/10 volume of 10x concentrated tricaine to the agarose.</li>
		<li>Place one drop of the tricaine-containing agarose onto the recording chamber (Figure 3A).</li>
		<li>After confirming the agarose is not too hot, place the anesthetized larva (from step 2.5) into the agarose, immediately removing any excess agarose so that the surface of the agarose looks slightly convex.</li>
		<li>Quickly orient the larva into an upright position with a dissecting needle (Figure 1C). 
		Note: These procedures must be completed within several seconds before the agarose solidifies.</li>
		<li>Once the larva is properly positioned, allow 5 min for the agarose to solidify completely. Then, pour a drop of 1x tricaine solution on the agarose.</li>
	</ol>
	</li>
	<li>Remove the agarose around the head of the zebrafish larva and make space for the paramecium to swim using a surgical knife. To do this, first, make a few small cuts in the agarose (Figure 3F). Then, carefully remove the excess agarose by gently pouring system water on the chamber. 
	Note: At this step, the tricaine will be washed out.</li>
	<li>Next, put one paramecium in the recording chamber to serve as a visual stimulus. 
	Note: When the paramecium swims near the head of the zebrafish larva, it will evoke a neuronal response (Figure 3G).</li>
	<li>Place the recording chamber under an epi-fluorescence microscope equipped with a scientific CMOS camera (Figure 3H) and select a 2.5X objective lens (Figure 3I).</li>
	<li>Collect time-lapse recordings of the GCaMP fluorescence signals.
	<ol>
		<li>Start the image acquisition application for the equipped camera.</li>
		<li>Click &#34;Sequence Pane&#34; and select &#34;Hard Disk Record&#34; on the pane. In the Scan Settings section, set the &#34;Frame Count&#34; to 900 or any other total frame number desired. 
		Note: If available, use time-lapse recording software with a module (here, &#34;Hard Disk Record&#34;) that enables efficient saving of the data onto a hard drive.</li>
		<li>In the &#34;Capture&#34; pane, set the exposure time at 30-ms (i.e., 33.3 fps), and Binning 2 x 2.</li>
		<li>On the microscope controlling touch panel, choose a filter set for GFP, as the GCaMP has similar excitation/emission spectra to GFP.</li>
		<li>Click &#34;Live&#34; on the Capture pane to locate the larva in the camera view and focus (Figure 4A), and then click &#34;Stop Live&#34; and click the &#34;Start&#34; button. Save the movie in .cxd or any other format that holds information about the acquisition condition.</li>
	</ol>
	</li>
	<li>After the recordings, analyze the GCaMP6s fluorescent signals (Ca signals) by obtaining mean pixel values for the ROI set on a brain structure in the time-lapse movie using the free software Fiji25. 
	Note: Fiji is a version of ImageJ with many useful plug-in&#39;s preinstalled and can read .cxd format image files with the Bio-Formats plug-in.</li>
	<li>If the larva move slightly during the recording, perform image registration by running the TurboReg plug-in26 in Fiji. From the menu, select &#39;Plugins&#39;, &#39;Registration&#39;, and &#39;TurboReg&#39;.</li>
	<li>Consider how to analyze the data according to the objective of the study. The following is an example.
	<ol>
		<li>Calculate the F/F0 (fluorescence intensity at each time, divided by the fluorescence intensity at rest or before any Ca transient has occurred) as follows:</li>
		<li>Set ROIs on the pretectum or the ILH using the ROI Manager: in the menu, select &#34;Analyze,&#34; &#34;Tools,&#34; &#34;ROI Manager&#34;, and &#34;Add&#34; to the list of the ROIs (Figure 4A).</li>
		<li>Calculate the mean pixel values for the ROIs (i.e., fluorescence intensities of GCaMP6s) by selecting on the ROI Manager panel: &#34;More,&#34; &#34;Multi-Measure&#34;, and &#34;OK.&#34; Save the Results as a spreadsheet file.</li>
		<li>As the signals are noisy, average the signals for 11 time-points (moving average) using an application that can handle spreadsheet data.</li>
		<li>Divide F (fluorescence intensities over time) by F0 (fluorescence intensity at the basal level) to calculate the normalized fluorescence intensity. As an example of data visualization, changes in normalized GCaMP6s fluorescence intensity in the pretectum and the ILH are color-coded and mapped on the paramecium trajectories (Figure 4B - D).</li>
	</ol>
	</li>
</ol>
3. Assessment of Behavioral Consequences Following Laser Ablation
<ol>
	<li>Set up a behavioral recording system that consists of a stereoscope equipped with a video camera. Use a camera with an appropriate image acquisition software, commercial or custom-made (Figure 5A).</li>
	<li>Optimize the lighting condition so that the paramecium can be detected by image processing. For example, use a ring white LED light with a spacer (Figure 5B). 
	Note: The distance from, and the angle of, the LED affects the appearance of the sample (i.e., bright in dark background or dark in bright background) and are, therefore, critical for successful image processing. Determine the best height of the spacer (Figure 5C).</li>
	<li>Place a zebrafish larva (recovered from the laser-ablation experiment described in Section 1) in a behavioral recording chamber (which was attached to a glass slide) with the original top seal removed (Figure 5D).</li>
	<li>Collect 50 paramecia from the culture (step 2.3.5) using a micropipette and place them in the recording chamber.</li>
	<li>Place a cover slip on top of the recording chamber. Put a small drop of water on the cover slip before placing on the water surface of the recording chamber to avoid introducing air in the chamber. 
	Note: In this step, some water with paramecia will overflow from the recording chamber. The remaining number of paramecia in the recording chamber will be approximately 30-40.</li>
	<li>Place the recording chamber (containing a larva and paramecia) in the lighting system (Figure 5D).</li>
	<li>Start the time-lapse recording at 10 fps for 11 min (6,600 frames).</li>
	<li>(Optional) For each larva that was tested for behavior, observe the fluorescence of the laser-ablated areas by fluorescent microscopy to confirm the effectiveness of the laser-ablation (i.e., absence, or significantly reduced number of, fluorescent cells in the laser-irradiated area). 
	Note: Even after irradiation, some fluorescent cells can still be observed owing to later onset of Gal4 gene expression in cells that differentiated after ablation27.</li>
	<li>After recording the larva&#39;s behavior, to compensate for the lack of uniformity in the background, perform a pixel-by-pixel division of each frame by an average image in Fiji: click &#39;Process&#39;, &#39;Image Calculator&#39;, &#39;Operation: Divide&#39;, &#39;32-bit(float)result&#39;, and &#39;OK&#39;. To make an average image, click &#39;Image&#39;, &#39;Stacks&#39;, &#39;Z-Project&#39;, &#39;Average intensity&#39;, and &#39;OK&#39; (Figure 5E, F).</li>
	<li>Count the number of paramecia left in the chamber by clicking &#39;Analyze&#39;, &#39;Analyze Particles&#39;, setting an area range that covers all the paramecia, checking the &#39;Show:Masks&#39; checkbox, and clicking &#39;OK&#39;. The &#39;Show:Masks&#39; option will provide an extracted paramecia image (Figure 5G), with a list of the number of particles (paramecia) in each frame.</li>
	<li>Plot the number of paramecia left in the recording chamber over time. Because the estimates of the number of paramecia are noisy, we recommend performing a moving average on each point in a range of 600 frames (1 min) on the spreadsheet.</li>
</ol>

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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BrdU-,+neuroD+(nrd)-+and+Hu-studies+reveal+unusual+non-ventricular+neurogenesis+in+the+postembryonic+zebrafish+forebrain.">BrdU-, neuroD (nrd)- and Hu-studies reveal unusual non-ventricular neurogenesis in the postembryonic zebrafish forebrain.</a> Mech Dev. 117 (1-2), 123-135 (2002).</li><li>Muto, A., 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=Forward+genetic+analysis+of+visual+behavior+in+zebrafish.">Forward genetic analysis of visual behavior in zebrafish.</a> PLoS Genet. 1 (5), 66 (2005).</li><li>Chen, T. W., 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=Ultrasensitive+fluorescent+proteins+for+imaging+neuronal+activity.">Ultrasensitive fluorescent proteins for imaging neuronal activity.</a> Nature. 499 (7458), 295-300 (2013).</li></ol>

The authors have no conflicts of interest to report.

Although the two-photon laser has an excellent spatial resolution to specifically ablate individual neurons, great caution should be taken to avoid any undesired damage on the brain tissue owing to heat. The most important step in the ablation experiment is to determine the optimal amount of laser irradiation. Insufficient irradiation fails to kill the neurons. Too much irradiation will heat-damage the surrounding tissue, which will result in undesired effects. The optimal range of laser irradiation (areas of the ROIs, n...

To understand how behavior arises from neuronal activity in the brain, it is necessary to identify the neural circuits that are involved in the generation of that behavior. At the larval stage, the zebrafish provides an ideal animal model for studying the brain function associated with the behavior because their small, transparent brains make it possible to investigate neuronal activity at a cellular resolution in a broad area of the brain while observing the behavior1. Imaging of neuronal activity in specific neurons has become possible through the invention of genetically encoded calcium (Ca) indicators (GECIs) such as GCaMP2. GCaMP transgenic zebrafish have proven to be useful for associating the functional neural circuit with behavior by conducting Ca imaging in behaving animals3.
While Ca imaging can demonstrate correlations between neuronal activity and behavior, to show causality, suppression of neuronal activity and testing its consequence(s) on behavior are important steps. There are various ways to achieve this: use of genetic mutation that alters specific neural circuits4, expression of neurotoxins in specific neurons5,6, use of optogenetic tools such as halorhodopsin7, and laser ablation of targeted neurons8,9. Laser ablation is particularly suited for eliminating activity in a relatively small number of specific neurons. Irreversible elimination of neuronal activity by killing neurons facilitates assessing behavioral consequences.
One interesting behavior that can be observed at the larval stage in zebrafish is prey capture&#160;(Figure 1A). This visually-guided, goal-directed behavior provides a favorable experimental system for the study of visual acuity10, visuomotor transformation11,12,13, visual perception and recognition of objects14,15,16,17,18, and decision making19. How prey is recognized by predators and how prey detection leads to prey catching behavior has been a central question in neuroethology20. In this paper, we focus on the role of the pretecto-hypothalamic circuit formed by projections of a nucleus in the pretectum (nucleus pretectalis superficialis pars magnocellularis, hereafter, simply noted as the pretectum) to the ILH. Laser-ablation of the pretectum was shown to reduce prey capture activity and abolish neuronal activity in the ILH that is associated with the visual prey perception21. Here, protocols for performing laser ablation and testing its effect using Ca2+ imaging and behavioral recording in zebrafish larvae are described.

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			<td style="width:383px;">low-melting temperature agarose</td>
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			<td>FALCON</td>
			<td>Product#:351007</td>
			<td></td>
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			<td height="40" style="height:40px;">dissecting needle</td>
			<td>AS ONE Corporation</td>
			<td>Cat. No. 2-013-01</td>
			<td>https://keystone-lab.com/en/item/detail/404142</td>
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			<td height="40" style="height:40px;">LSM7MP</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td>two-photon laser scanning microscope</td>
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			<td height="40" style="height:40px;">W Plan-Apochromat 63x/1.0</td>
			<td>Carl Zeiss</td>
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			<td>63X objective lens</td>
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			<td height="40" style="height:40px;">Imager.Z1</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td>an epi-fluorescence microscope</td>
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			<td height="40" style="height:40px;">ZEN</td>
			<td>Carl Zeiss</td>
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			<td>Image acquisition software for confocal microscopes</td>
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			<td height="40" style="height:40px;">Secure-Seal Hybridization Chamber Gasket, 8 chambers, 9 mm diameter x 0.8 mm depth</td>
			<td>Molecular Probes</td>
			<td>Catalogue # S-24732</td>
			<td>Used as a recording chamber in Ca imaging</td>
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			<td>Grace Bio-Labs</td>
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			<td>Hamamatsu Photonics</td>
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			<td style="width:383px;">An software module that enables saving the movie files onto a hard disc drive in a short time</td>
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			<td>Olympus</td>
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			<td>FLIR Systems, Inc.</td>
			<td>Product No. GS3-U3-41C6NIR-C</td>
			<td>CMOS camera</td>
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			<td height="40" style="height:40px;">XIMEA xiQ camera</td>
			<td>XIMEA</td>
			<td>Product No. MQ042RG-CM</td>
			<td>CMOS camera</td>
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		<tr height="40">
			<td height="40" style="height:40px;">a ring LED light</td>
			<td>CCS</td>
			<td>Model: LDR2-100SW2-LA</td>
			<td>White LED</td>
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			<td height="26" style="height:26px;width:316px;"></td>
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			<td height="40" style="height:40px;">Nylon mesh 32&#181;m</td>
			<td>Tokyo Screen</td>
			<td>N-No.380T</td>
			<td>http://www.tokyo-screen.com/cms/sta20347/</td>
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			<td height="26" style="height:26px;">Nylon mesh 13&#181;m</td>
			<td>Tokyo Screen</td>
			<td>N-No. 508T-K</td>
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			<td height="52" style="height:52px;width:316px;">Metal seive 150 micron aperture</td>
			<td>Tokyo Screen</td>
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			<td style="width:383px;">http://www.tokyo-screen.com/cms/sta20341/#ami</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:316px;">Metal seive 75 micron aperture</td>
			<td>Tokyo Screen</td>
			<td></td>
			<td>http://www.tokyo-screen.com/cms/sta20341/#ami</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:316px;">EBIOS</td>
			<td>Asahi Food &#38; Healthcare, Co. Ltd.</td>
			<td></td>
			<td>dry beer yeast</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:316px;">LabVIEW</td>
			<td>National Instruments</td>
			<td></td>
			<td>an integrated development environment for programming</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;width:316px;">Mai-Tai HP</td>
			<td>Spectra Physics&#160;</td>
			<td></td>
			<td>two-photon laser&#160;</td>
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ablation of a neuronal population using a two-photon laser and its assessment using calcium imaging and behavioral recording in zebrafish larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Specific neurons were genetically labeled with either EGFP or GCaMP6s, whose expression were driven in Gal4 lines. A Gal4 line gSAIzGFFM119B was used to label a nucleus in the pretectal area (magnocellular superficial pretectal nucleus), and a subpopulation of olfactory bulb neurons. Another Gal4 line, hspGFFDMC76A, was used to label the ILH. We laser-ablated the pretectal neurons bilaterally (Figure 2A left panel) and also ablated neurons in...

Watch this Scientific Journal Video about Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae at JoVE.com

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

ablation-neuronal-population-using-two-photon-laser-its-assessment

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JoVE Journal

Neuroscience

8.7K Views.  SOKENDAI (The Graduate University for Advanced Studies). Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

Video: Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Prolonged Incubation of Acute Neuronal Tissue for Electrophysiology and Calcium-imaging

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the experimental time, and increases the number of animals that are utilized per study. This limitation specifically impacts protocols such as calcium imaging that require prolonged pre-incubation with bath-applied dyes. Exponential bacterial growth within 3 - 4 h after slicing is tightly correlated with a decrease in tissue health. This study describes a method for&#160;limiting the proliferation of bacteria in acute preparations to maintain viable neuronal tissue for prolonged periods of time (&#62;24 h) without the need for antibiotics, sterile procedures, or tissue culture media containing growth factors. By cycling the extracellular fluid through UV irradiation and keeping the tissue in a custom holding chamber at 15 - 16 &#176;C, the tissue shows no difference in electrophysiological properties, or calcium signaling through intracellular calcium dyes at &#62;24 h postdissection. These methods will not only extend experimental time for those using acute neuronal tissue, but will reduce the number of animals required to complete experimental goals, and will set a gold standard for acute neuronal tissue incubation.

Once removed from the body, neuronal tissue is greatly affected by environmental conditions, leading to eventual degradation of the tissue after 6 - 8 h. Using a unique incubation method, which closely monitors and regulates the extracellular environment of the tissue, tissue viability can be significantly extended for &#62;24 h.

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

Studying Brain Function in Children Using Magnetoencephalography

Magnetoencephalography (MEG) is a non-invasive neuroimaging technique which directly measures magnetic fields produced by the electrical activity of the human brain. MEG is quiet and less likely to induce claustrophobia compared with magnetic resonance imaging (MRI). It is therefore a promising tool for investigating brain function in young children. However, analysis of MEG data from pediatric populations is often complicated by head movement artefacts which arise as a consequence of the requirement for a spatially-fixed sensor array that is not affixed to the child's head. Minimizing head movements during MEG sessions can be particularly challenging as young children are often unable to remain still during experimental tasks. The protocol presented here aims to reduce head movement artefacts during pediatric MEG scanning. Prior to visiting the MEG laboratory, families are provided with resources that explain the MEG system and the experimental procedures in simple, accessible language. An MEG familiarization session is conducted during which children are acquainted with both the researchers and the MEG procedures. They are then trained to keep their head still whilst lying inside an MEG simulator. To help children feel at ease in the novel MEG environment, all of the procedures are explained through the narrative of a space mission. To minimize head movement due to restlessness, children are trained and assessed using fun and engaging experimental paradigms. In addition, children's residual head movement artefacts are compensated for during the data acquisition session using a real-time head movement tracking system. Implementing these child-friendly procedures is important for improving data quality, minimizing participant attrition rates in longitudinal studies, and ensuring that families have a positive research experience.

This article introduces a child-friendly research protocol designed to improve data quality by reducing head movement during pediatric magnetoencephalography (MEG). We familiarize families with the MEG environment, train children to remain still using an MEG simulator, and correct for residual head movement artefacts using a real-time head movement detection system.

Magnetoencephalography (MEG) is a non-invasive neuroimaging technique which directly measures magnetic fields produced by the electrical activity of the ...

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent and cause visual impairment and blindness. Understanding how such diseases progress is critical to formulating new treatments. In vivo&#160;microscopy in animal models of disease is a powerful tool for understanding neurodegeneration and has led to important progress towards treatments of conditions ranging from Alzheimer&#39;s disease to stroke. Given that the retina is the only central nervous system structure inherently accessible by optical approaches, it naturally lends itself towards in vivo imaging. However, the native optics of the lens and cornea present some challenges for effective imaging access.
This protocol outlines methods for in vivo two-photon imaging of cellular cohorts and structures in the mouse retina at cellular resolution, applicable for both acute- and chronic-duration imaging experiments. It presents examples of retinal ganglion cell (RGC), amacrine cell, microglial, and vascular imaging using a suite of labeling techniques including adeno-associated virus (AAV) vectors, transgenic mice, and inorganic dyes. Importantly, these techniques extend to all cell types of the retina, and suggested methods for accessing other cellular populations of interest are described. Also detailed are example strategies for manual image postprocessing for display and quantification. These techniques are directly applicable to studies of retinal function in health and disease.

In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the ...

Evaluation and Manipulation of Neural Activity Using Two-Photon Holographic Microscopy

Recent advances in optical bioimaging and optogenetics have enabled the visualization and manipulation of biological phenomena, including cellular activities, in living animals. In the field of neuroscience, detailed neural activity related to brain functions, such as learning and memory, has now been revealed, and it has become feasible to artificially manipulate this activity to express brain functions. However, the conventional evaluation of neural activity by two-photon Ca2+ imaging has the problem of low temporal resolution. In addition, manipulation of neural activity by conventional optogenetics through the optic fiber can only simultaneously regulate the activity of neurons with the same genetic background, making it difficult to control the activity of individual neurons. To solve this issue, we recently developed a microscope with a high spatiotemporal resolution for biological applications by combining optogenetics with digital holographic technology that can modify femtosecond infrared laser beams. Here, we describe protocols for the visualization, evaluation, and manipulation of neural activity, including the preparation of samples and operation of a two-photon holographic microscope (Figure 1). These protocols provide accurate spatiotemporal information on neural activity, which may be useful for elucidating the pathogenesis of neuropsychiatric disorders that lead to abnormalities in neural activity.

We developed a two-photon holographic microscope that can visualize, assess, and manipulate neural activity using high spatiotemporal resolution, with the aim of elucidating the pathogenesis of neuropsychiatric disorders that are associated with abnormal neural activity.