This work was supported by the Institute of Molecular Biology, Academia Sinica, and National Science and Technology Council, Taiwan. The Machine Shop at the Institute of Physics, Academia Sinica helped to fabricate custom-designed parts. We also want to thank P. Argast (Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland) for the design of the quick-lock mechanism of the head stage.

All animal procedures were approved and carried out in accordance with the guidelines of the Institutional Animal Care and Use Committee of Academia Sinica. Details of the research tools can be found in the Table of Materials.
1. Preparation of recording chamber
<ol>
	<li>Prepare a semi-hexagonal tank, a base plate, and a head stage (Figure 1A; Supplementary Files 1-3). The head stage consists of two metal posts attached to a circular plate. The circular plate contains grooves that can be locked onto the base plate. After being locked together, the head stage and the base plate are laid at the bottom of the semi-hexagonal tank.</li>
	<li>Place the semi-hexagonal tank on the experimental platform of the microscope. The platform should be able to move in the X and Y directions without hitting the limit of the translation stage.</li>
	<li>Aim the 810 nm infrared (IR) light and the camera to the head stage for behavioral recording. Ensure that the field of view of the camera is large enough to fit an adult zebrafish.
	<ol>
		<li>To prevent the two-photon laser and the visual stimulus from interfering with behavioral recording, set up an 875 nm short-pass filter and a 700 nm long-pass filter in front of the camera, respectively.</li>
	</ol>
	</li>
	<li>To present the visual stimulus, attach back-projection films to the inner side of the three walls of the semi-hexagonal tank (Figure 1A).</li>
	<li>Aim the three projectors at the three surfaces of the tank. The three images should be aligned to form a continuous visual scene. To prevent projector lights from contaminating the calcium fluorescence signal, place a neutral density filter and a red color filter (600 nm, long-pass) in front of each projector, and place a band-pass filter (510/80 nm) in front of the photomultiplier tube (PMT).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65526/65526fig1v8.jpg" /> 
Figure 1: Instruments required for head restraint surgery. (A) The quick-lock mechanism between the circular plate of the head stage and the base plate inside the semi-hexagonal tank. Computer-aided design (CAD) files of the custom-made parts can be found in Supplementary Files 1-4. (B) &#937;-shaped head bar for the head restraint. (C) The three-axis micromanipulator used to position the head bar to the attachment site. Inset: orientation of the head bar in the clay. (D) Cannon to hold the fish during surgery. Inset: orientation of fish within the cannon. (E) Fish loading module and the micromanipulator used to load the fish onto the head stage. Inset: orientation of fish within the module. <a href="https://www.jove.com/files/ftp_upload/65526/65526fig1v8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Head restraint surgery
<ol>
	<li>Prepare an &#937;-shaped head bar (Figure 1B; Supplementary File 4) for the zebrafish head restraint. To do this, attach a piece of oil-based modeling clay to a three-axis micromanipulator. Ensure that the clay is solid enough to hold the head bar, but soft enough to detach from the head bar after head restraint.</li>
	<li>Insert an arm of the head bar into the clay and orient the head bar horizontally (Figure 1C). This will ensure that the later attachment of the head bar onto the zebrafish will be level. 
	NOTE: The head bar can be made of titanium (24 mg) or stainless steel (43 mg) and can be reused for multiple experiments. The head bar material does not affect the behavior of adult zebrafish (weight ranges from 300-1,000 mg) under a head restraint.</li>
	<li>Prepare the cannon, a hollow tube for holding the fish body in place during surgery (Figure 1D).</li>
	<li>Prepare 0.03% and 0.01% tricaine methanesulfonate (TMS; see Table of Materials) in fish tank water. This will be used to anesthetize and maintain the fish in a sedated state during surgery.</li>
	<li>Prepare four tissue swabs to remove excess skin and water from the attachment sites on the skull. To prepare each swab, cut a paper towel into a 3 cm square and fold along its diagonal to produce a tube-like structure. Twist the ends of the tube into a fine point. The attachment site is very small, so a fine point allows finer control of the swab.</li>
	<li>Prepare a fish loading module for the micromanipulator (Figure 1E). 
	NOTE: The module consists of two steel posts held together by a right-angle post clamp. One post attaches to the micromanipulator, while the other post holds a rotating post clamp that carries the fish. The module is used to position the fish onto the head stage with fine control. The rotating post clamp serves as a pitch adjuster to change the pitch angle of the fish.</li>
	<li>Anesthetize the fish with 0.03% TMS in fish tank water. Throughout the following steps, use a syringe to deliver 0.01% TMS in fish tank water to the mouth in short pulses every 90 s. Water flow from the perfusion should induce gill movement. This will allow the fish to survive for more than 40 min during surgery. 
	NOTE: Tg[neuroD:GCaMP6f] is used due to its broad expression of the calcium indicator in the forebrain at both the larval and adult stages10. 
	NOTE:&#160;Optionally, use gravity perfusion to provide constant water flow to the mouth during periods in surgery where both hands are occupied.</li>
	<li>Wrap the fish into a capsule (Figure 2A) that holds the fish inside the cannon.
	<ol>
		<li>Wrap the anesthetized fish in a piece of moist paper tissue starting from the tip of the tail to around 1 mm caudal to the gill. Ensure that the wrapping is tight to prevent sliding of the fish out of the tissue.</li>
		<li>Wrap a soft plastic tubing with a longitudinal slit (e.g., a medium-sized heat-shrink tubing cut open) around the paper tissue to cover the fish from the end of the tail to around 2 mm caudal to the gill. The tubing ensures that the fish&#39;s body remains straight throughout the surgery.</li>
		<li>Wrap a paper towel around the tubing to a diameter that is roughly similar to the diameter of the cannon.</li>
		<li>Wrap a soft plastic tube cut from the bulb of a plastic transfer pipette around the paper towel. This ensures that the capsule can be loaded smoothly into the cannon.</li>
		<li>Load the fish into the cannon.</li>
	</ol>
	</li>
	<li>Use a scalpel to remove the skin covering the attachment sites, two triangular areas of the skull rostrolateral to the cerebellum and above the gills (Figure 2B), then remove the skin covering the region between the attachment sites. Use tissue swabs to dry out the attachment sites and clear out any remaining skin.
	<ol>
		<li>Use an adjustable platform to support the head during skin removal, but the platform must not contact the eyes to avoid eye injury during the operation. 
		NOTE: Thorough removal of the skin at the attachment sites is crucial in reducing motion artifacts during imaging.</li>
	</ol>
	</li>
	<li>Apply a drop of tissue glue (see Table of Materials) at the center of each attachment site using a 10 &#181;L pipette tip. 
	NOTE: Tissue glue covers the attachment sites and dries out quickly. The tissue glue provides a bonding interface between the skull and the dental cement used to glue the head bar, and prevents water from the gills from reaching the attachment site.</li>
	<li>Put the fish body in an upright position and position the head bar slightly above and caudal to the attachment sites in preparation for head bar gluing.</li>
	<li>Prepare dental cement (see Table of Materials) and use it immediately to glue the head bar to the skull (Figure 2C). The procedure is time sensitive and must be done within 45 s.
	<ol>
		<li>Mix one small spoon of polymer with four drops of quick monomer and one drop of catalyst V. Stir the mixture evenly for 15 s.</li>
		<li>Use a micropipette and a 10 &#181;L pipette tip to apply the mixture onto the attachment sites and the region between the sites. Avoid covering the gills and the eyes.</li>
		<li>Gently press the head bar onto the cement using the micromanipulator. Cover the head bar with the remaining cement.</li>
		<li>Wait for 12 min to allow curing of the dental cement. Avoid water contact with the cement.</li>
	</ol>
	</li>
	<li>To improve optical access to the forebrain for calcium imaging, remove the skin above the telencephalon. Skin removal can be done in 3 min with five cuts using a scalpel (Figure 2D). Make sure that lipid droplets adhering to the surface of the skulls are removed.</li>
	<li>After curing, remove the clay from the head bar.</li>
	<li>Transfer the entire capsule from the cannon to the pitch adjuster in the fish loading module of the micromanipulator.</li>
	<li>Use the micromanipulator to position the fish. The head bar should be positioned on top of the metal posts of the head stage. Slightly increase the pitch angle of the fish so that the surface of the forebrain can be better aligned to the optical plane during two-photon imaging.</li>
	<li>Glue the head bar to the metal posts with ultraviolet (UV)-curable glue. A 15 s UV exposure is sufficient for curing.</li>
	<li>Pull out the fish from the capsule and lock the head stage onto the base plate within the semi-hexagonal tank.</li>
	<li>Immerse the animal in fish tank water. The fish should start breathing and recover from anesthesia within 1-2 min. 
	NOTE: Optionally, use syringe perfusion to deliver fresh water to the mouth.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65526/65526fig02.jpg" /> 
 
Figure 2: Key steps during head restraint surgery. (A) Composition of the capsule inside the cannon. (B) Attachment sites on the skull (red). Red arrows specify blood vessel sites. (C) Top: head bar attached to the fish skull. Bottom: fish loaded onto the head stage. (D) Cuts needed to remove the skin above the forebrain. Numbers denote the cutting sequence. Avoid skin removal at the marked site (arrowhead) to prevent bleeding of the animal. <a href="https://www.jove.com/files/ftp_upload/65526/65526fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Two-photon imaging
<ol>
	<li>Turn on the laser and wait for 30 min for the power to stabilize. Set the wavelength to 920 nm and power to around 50 mW at the sample. 
	NOTE: The laser beam controlled by the resonant scanner moves slowly at the turnaround points of the scan path. To prevent tissue damage, a Pockels cell is used to reduce laser power at the turnaround points.</li>
	<li>Place the recording chamber containing the head-restrained zebrafish on the experimental platform (Figure 3A). The platform should be able to move in the X and Y directions without hitting the limit of the translation stage.</li>
	<li>Place the objective lens as close as possible to the forebrain surface. The objective lens should be aimed at the front edge of the pupil (Figure 3B).</li>
	<li>Open the laser shutter and enable the PMT. Gradually elevate the objective lens (~1 mm) until the dorsal forebrain is revealed in the fluorescence image (Figure 3C).</li>
	<li>To increase the number of neurons recorded, use a piezo actuator to acquire images at multiple depths (six image planes, 15 &#181;m apart). This will increase the data yield at the expense of the frame rate. Enable fast Z-axis scanning to control the piezo actuator (uniform mode, number of slices = 6, step size = 15 &#181;m, waveform = sawtooth, flyback time = 4 ms, actuator lag = 8 ms).</li>
	<li>For behavior recording, turn on the 810 nm infrared (IR) lights on both sides of the tank. Adjust the angle to illuminate the entire body, which should be clearly visible in the camera.</li>
	<li>Turn on the projectors.</li>
	<li>Start recording data.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65526/65526fig3v2.jpg" /> 
Figure 3: Setup to perform calcium imaging, behavior recording, and visual stimulus display. (A) Three projectors present a visual stimulus onto the walls of the semi-hexagonal tank. IR lights on the side are used to illuminate the body of the zebrafish. (B) Positioning of the objective lens. Left: front view. Right: side view. The distance between the 16x objective lens and the targeted brain region is around 2.5 mm. (C) Example two-photon image. Left: maximum projection of the entire dorsal forebrain in Tg[neuroD:GCaMP6f]. Right: zoomed-in image to reveal neurons across multiple brain regions. Inset: a higher magnification from a different brain region. Images are averages of 10 s of data recorded at 5Hz. <a href="https://www.jove.com/files/ftp_upload/65526/65526fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Behavior Tracking and Neural Activity Pattern in the Zebrafish Forebrain

<ol>
	<li>Grienberger, C., Konnerth, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+calcium+in+neurons.">Imaging calcium in neurons.</a> Neuron. 73 (5), 862-885 (2012).</li><li>Chow, D. M., 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=Deep+three-photon+imaging+of+the+brain+in+intact+adult+zebrafish.">Deep three-photon imaging of the brain in intact adult zebrafish.</a> Nature Methods. 17 (6), 605-608 (2020).</li><li>Mittmann, 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=Two-photon+calcium+imaging+of+evoked+activity+from+L5+somatosensory+neurons+in+vivo.">Two-photon calcium imaging of evoked activity from L5 somatosensory neurons in vivo.</a> Nature Neuroscience. 14 (8), 1089-1093 (2011).</li><li>Friedrich, R. W., Jacobson, G. A., Zhu, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circuit+neuroscience+in+zebrafish.">Circuit neuroscience in zebrafish.</a> Current Biology. 20 (8), R371-R381 (2010).</li><li>Kappel, J. M., 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=Visual+recognition+of+social+signals+by+a+tectothalamic+neural+circuit.">Visual recognition of social signals by a tectothalamic neural circuit.</a> Nature. 608 (7921), 146-152 (2022).</li><li>Bartoszek, E. M., 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=Ongoing+habenular+activity+is+driven+by+forebrain+networks+and+modulated+by+olfactory+stimuli.">Ongoing habenular activity is driven by forebrain networks and modulated by olfactory stimuli.</a> Current Biology. 31 (17), 3861-3874 (2021).</li><li>Valente, A., Huang, K. H., Portugues, R., Engert, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ontogeny+of+classical+and+operant+learning+behaviors+in+zebrafish.">Ontogeny of classical and operant learning behaviors in zebrafish.</a> Learning &amp; Memory. 19 (4), 170-177 (2012).</li><li>Buske, C., Gerlai, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maturation+of+shoaling+behavior+is+accompanied+by+changes+in+the+dopaminergic+and+serotoninergic+systems+in+zebrafish.">Maturation of shoaling behavior is accompanied by changes in the dopaminergic and serotoninergic systems in zebrafish.</a> Developmental Psychobiology. 54 (1), 28-35 (2012).</li><li>Huang, K. H., 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=A+virtual+reality+system+to+analyze+neural+activity+and+behavior+in+adult+zebrafish.">A virtual reality system to analyze neural activity and behavior in adult zebrafish.</a> Nature Methods. 17 (3), 343-351 (2020).</li><li>Rupprecht, P., Prendergast, A., Wyart, C., Friedrich, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Remote+z-scanning+with+a+macroscopic+voice+coil+motor+for+fast+3D+multiphoton+laser+scanning+microscopy.">Remote z-scanning with a macroscopic voice coil motor for fast 3D multiphoton laser scanning microscopy.</a> Biomedical Optics Express. 7 (5), 1656-1671 (2016).</li><li>Papadopoulos, I. N., Jouhanneau, J. -. S., Poulet, J. F. A., Judkewitz, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scattering+compensation+by+focus+scanning+holographic+aberration+probing+(F-SHARP).">Scattering compensation by focus scanning holographic aberration probing (F-SHARP).</a> Nature Photonics. 11 (2), 116-123 (2017).</li><li>Torigoe, M., 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=Zebrafish+capable+of+generating+future+state+prediction+error+show+improved+active+avoidance+behavior+in+virtual+reality.">Zebrafish capable of generating future state prediction error show improved active avoidance behavior in virtual reality.</a> Nature Communications. 12 (1), 5712 (2021).</li></ol>

The authors declare that they have no competing financial interests.

Here, we describe a detailed protocol to restrain the head of adult zebrafish for two-photon calcium imaging. There are two critical steps to achieve a head restraint that is stable enough for laser scanning imaging. First, the head bar has to be glued to the specific attachment sites of the skulls. Other parts of the skull&#160;are often too thin to provide mechanical stability and may even be fractured during strong body movements. Second, the skin above the attachment sites has to be thoroughly removed. Residual water...

Calcium fluorescence imaging with genetically encoded indicators or synthetic dyes has been a powerful method of measuring neuronal activity in behaving animals, including non-human primates, rodents, birds, and insects1. The activity of hundreds of cells, up to approximately 800 &#181;m below the brain surface, can be measured simultaneously using multi-photon imaging2,3. The activity of specific cell types can also be measured by expressing calcium indicators in genetically defined neuronal populations. Application of the imaging method for small vertebrate models opens up new possibilities in the field of neuronal computation across brain regions.
Zebrafish are a widely used model system in neuroscience research. Larval zebrafish at around 6 days post-fertilization have been used for calcium imaging due to their miniature brain and transparent body4. Juvenile zebrafish (3-4 weeks old) are also used for studying the neural mechanisms underlying sensorimotor pathways5,6. However, the maximal performance level for complex behaviors, including associative learning and social behaviors, is reached at an older age7,8. Thus, a reliable protocol is required to study multiple cognitive functions in the brains of adult zebrafish using imaging methods. While zebrafish larva and juvenile zebrafish can be embedded in agarose for in vivo imaging, adult zebrafish at 2 months or older suffer from hypoxia in such conditions and are physically too strong to be restrained by agarose. Therefore, a surgical procedure is required to stabilize the brain and enable the animal to breathe freely through the gills. 
Here, we describe a head restraint protocol that involves a novel design of a single head bar. The reduced surgery time of 25 min is twice as fast as the previous method9. We also describe the design of the recording chamber (semi-hexagonal tank), the head stage and a quick-lock mechanism to combine the two parts9. Finally, the setup to present an immersive visual stimulus to study visually triggered brain activity and behaviors is also described. Overall, the procedures described here can be used to perform two-photon calcium imaging in genetically defined cell populations in a head-restrained adult zebrafish, enabling the investigation of brain activities during various behavioral paradigms.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acquisition card</td>
			<td>MBF Bioscience</td>
			<td>Vidrio vDAQ</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Back-projection film</td>
			<td>Kimoto</td>
			<td>Diland screen - GSK</td>
			<td>present visual stimulus</td>
		</tr>
		<tr>
			<td>Band-pass filter (510/80 nm)</td>
			<td>Chroma</td>
			<td>ET510/80m</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Base plate for the semi-hexagonal tank</td>
			<td>custom made</td>
			<td>see supplemental files</td>
			<td>recording chamber</td>
		</tr>
		<tr>
			<td>Camera filter (&#60;875 nm)</td>
			<td>Edmund optics</td>
			<td>#86-106</td>
			<td>Behavior recording</td>
		</tr>
		<tr>
			<td>Camera filter (&#62;700 nm)</td>
			<td>Edmund optics</td>
			<td>#43-949</td>
			<td>Behavior recording</td>
		</tr>
		<tr>
			<td>Camera lens</td>
			<td>Thorlabs</td>
			<td>MVL50M23</td>
			<td>Behavior recording</td>
		</tr>
		<tr>
			<td>Chameleon Vision-S</td>
			<td>Coherent</td>
			<td>Vision-S</td>
			<td>Laser</td>
		</tr>
		<tr>
			<td>Circular plate for the head stage</td>
			<td>custom made</td>
			<td>see supplemental files</td>
			<td>recording chamber</td>
		</tr>
		<tr>
			<td>Controller for piezo actuator</td>
			<td>Physik Instrumente&#160;</td>
			<td>E-665. CR</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Current amplifier</td>
			<td>Thorlabs</td>
			<td>TIA60</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Elitedent Q-6</td>
			<td>Rolence Enterprise</td>
			<td>Q-6</td>
			<td>Surgery: UV lamp</td>
		</tr>
		<tr>
			<td>Emission Filter 510/80 nm</td>
			<td>Chroma</td>
			<td>ET510/80m</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Head bar</td>
			<td>custom made</td>
			<td>see supplemental files</td>
			<td>recording chamber</td>
		</tr>
		<tr>
			<td>Infrared light</td>
			<td>Thorlabs</td>
			<td>M810L3</td>
			<td>Behavior recording</td>
		</tr>
		<tr>
			<td>LED projector</td>
			<td>AAXA</td>
			<td>P2B LED Pico Projector</td>
			<td>present visual stimulus</td>
		</tr>
		<tr>
			<td>Moist paper tissue (Kimwipe)</td>
			<td>Kimtech Science</td>
			<td>34155</td>
			<td>Surgery: moist paper tissue</td>
		</tr>
		<tr>
			<td>Motorized XY sample stage</td>
			<td>Zaber</td>
			<td>X-LRM050</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Neutral Density Filters (50% Transmission)</td>
			<td>Thorlabs</td>
			<td>NE203B</td>
			<td>present visual stimulus</td>
		</tr>
		<tr>
			<td>&#216;1/2&#34; Post Holder</td>
			<td>ThorLabs</td>
			<td>PH1.5V</td>
			<td>Surgery: hollow tube for cannon</td>
		</tr>
		<tr>
			<td>&#216;1/2&#34; Stainless Steel Optical Post</td>
			<td>ThorLabs</td>
			<td>TR150/M</td>
			<td>Surgery: fish loading module</td>
		</tr>
		<tr>
			<td>Objective lens 16x, 0.8NA</td>
			<td>Nikon</td>
			<td>CF175</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Oil-based modeling clay</td>
			<td>Ly Hsin Clay</td>
			<td>C4086</td>
			<td>Surgery: head bar holder</td>
		</tr>
		<tr>
			<td>Optical adhesive</td>
			<td>Norland Products</td>
			<td>NOA68</td>
			<td>Surgery: UV curable glue</td>
		</tr>
		<tr>
			<td>Photomultiplier tube</td>
			<td>Hamamatsu</td>
			<td>H11706P-40</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Piezo actuator</td>
			<td>Physik Instrumente&#160;</td>
			<td>P-725.4CA PIFOC</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Pockels Cell</td>
			<td>Conoptics</td>
			<td>M350-80-LA-BK-02</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Red Wratten filter (&#62; 600 nm)</td>
			<td>Edmund optics</td>
			<td>#53-699</td>
			<td>present visual stimulus</td>
		</tr>
		<tr>
			<td>Resonant-Galvo Scan System</td>
			<td>INSS</td>
			<td>RGE-02</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Right-Angle Clamp for &#216;1/2&#34; Post</td>
			<td>ThorLabs</td>
			<td>RA90/M</td>
			<td>Surgery: fish loading module</td>
		</tr>
		<tr>
			<td>Rotating Clamp for &#216;1/2&#34; Post</td>
			<td>ThorLabs</td>
			<td>SWC/M</td>
			<td>Surgery: fish loading module</td>
		</tr>
		<tr>
			<td>ScanImage</td>
			<td>MBF Bioscience</td>
			<td>Basic version</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Semi-hexagonal tank</td>
			<td>custom made</td>
			<td>see supplemental files</td>
			<td>recording chamber</td>
		</tr>
		<tr>
			<td>Super-Bond C&#38;B Kit</td>
			<td>Sun Medical Co.</td>
			<td>Super-Bond C&#38;B</td>
			<td>Surgery: dental cement</td>
		</tr>
		<tr>
			<td>Tricaine methanesulfonate</td>
			<td>Sigma Aldrich</td>
			<td>E10521</td>
			<td>Surgery: anesthetic</td>
		</tr>
		<tr>
			<td>USB Camera</td>
			<td>FLIR</td>
			<td>BFS-U3-13Y3M-C</td>
			<td>Behavior recording</td>
		</tr>
		<tr>
			<td>Vetbond</td>
			<td>3M</td>
			<td>1469SB</td>
			<td>Surgery: tissue glue</td>
		</tr>
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two-photon calcium imaging of forebrain activity in behaving adult zebrafish

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used for measuring activities across brain regions through optical imaging methods. However, reports on the recording of brain activity in behaving adult zebrafish have been scarce. The present study describes procedures to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish. We focus on steps to restrain adult zebrafish from moving their heads, which provides stability that enables laser scanning imaging of the brain activity. The head-restrained animals can freely move their body parts and breathe without aids. The procedure aims to shorten the time of head restraint surgery, minimize brain motion, and maximize the number of neurons recorded. A setup for presenting an immersive visual environment during calcium imaging is also described here, which can be used to study neural correlates underlying visually triggered behaviors.

The protocol consists of two parts: head restraint surgery and two-photon calcium imaging of neuronal activities in the forebrain. The success of surgery is defined by the survival of the animal and the stability of the head restraint. The survival rate can be greatly improved by frequent perfusion of 0.01% TMS solution through the mouth during surgery. Fish should recover from anesthesia and breathe actively within 1-2 min after being immersed into fish tank water. Two-photon calcium imaging enables activity recording o...

Watch this Scientific Journal Video about Advancements in Adult Zebrafish Brain Research at JoVE.com

Author Spotlight: Advancements in Adult Zebrafish Brain Research 

Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Two-Photon Calcium Imaging of Forebrain Activity in Behaving Adult Zebrafish

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used ...

two-photon-calcium-imaging-forebrain-activity-behaving-adult

Research

JoVE Journal

Neuroscience

2.3K Views.  Academia Sinica. Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Video: Author Spotlight: Advancements in Adult Zebrafish Brain Research 

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

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 ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

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+ ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

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.

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 ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...