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/pubmed?term=((Imaging+calcium+in+neurons%5BTitle%5D)+AND+%22Neuron%22%5BJournal%5D)">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/pubmed?term=((Deep+three-photon+imaging+of+the+brain+in+intact+adult+zebrafish%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">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/pubmed?term=((Two-photon+calcium+imaging+of+evoked+activity+from+L5+somatosensory+neurons+in+vivo%5BTitle%5D)+AND+%22Nature+Neuroscience%22%5BJournal%5D)">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/pubmed?term=((Circuit+neuroscience+in+zebrafish%5BTitle%5D)+AND+%22Current+Biology%22%5BJournal%5D)">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/pubmed?term=((Visual+recognition+of+social+signals+by+a+tectothalamic+neural+circuit%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">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/pubmed?term=((Ongoing+habenular+activity+is+driven+by+forebrain+networks+and+modulated+by+olfactory+stimuli%5BTitle%5D)+AND+%22Current+Biology%22%5BJournal%5D)">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/pubmed?term=((Ontogeny+of+classical+and+operant+learning+behaviors+in+zebrafish%5BTitle%5D)+AND+%22Learning+%26+Memory%22%5BJournal%5D)">Ontogeny of classical and operant learning behaviors in zebrafish.</a> Learning & Memory. 19 (4), 170-177 (2012).</li>
<li>Buske, C., Gerlai, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Maturation+of+shoaling+behavior+is+accompanied+by+changes+in+the+dopaminergic+and+serotoninergic+systems+in+zebrafish%5BTitle%5D)+AND+%22Developmental+Psychobiology%22%5BJournal%5D)">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/pubmed?term=((A+virtual+reality+system+to+analyze+neural+activity+and+behavior+in+adult+zebrafish%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">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/pubmed?term=((Remote+z-scanning+with+a+macroscopic+voice+coil+motor+for+fast+3D+multiphoton+laser+scanning+microscopy%5BTitle%5D)+AND+%22Biomedical+Optics+Express%22%5BJournal%5D)">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/pubmed?term=((Scattering+compensation+by+focus+scanning+holographic+aberration+probing+%28F-SHARP%29%5BTitle%5D)+AND+%22Nature+Photonics%22%5BJournal%5D)">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/pubmed?term=((Zebrafish+capable+of+generating+future+state+prediction+error+show+improved+active+avoidance+behavior+in+virtual+reality%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">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><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 (&amp;lt;875 nm)</td><td>Edmund optics</td><td>#86-106</td><td>Behavior recording</td></tr><tr><td>Camera filter (&gt;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&amp;nbsp;</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>&amp;Oslash;1/2&quot; Post Holder</td><td>ThorLabs</td><td>PH1.5V</td><td>Surgery: hollow tube for cannon</td></tr><tr><td>&amp;Oslash;1/2&quot; 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&amp;nbsp;</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 (&gt; 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 &amp;Oslash;1/2&quot; Post</td><td>ThorLabs</td><td>RA90/M</td><td>Surgery: fish loading module</td></tr><tr><td>Rotating Clamp for &amp;Oslash;1/2&quot; 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&amp;amp;B Kit</td><td>Sun Medical Co.</td><td>Super-Bond C&amp;amp;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></tbody></table>

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 ...

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Neuroscience

2.7K 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 

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

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

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

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

Biology

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 ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

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

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

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

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

Developmental Biology

Simultaneous Calcium Imaging and Glucose Stimulation in Living Zebrafish to Investigate In Vivo &#946;-Cell Function

The pancreatic &#946;-cells sustain systemic glucose homeostasis by producing and secreting insulin according to the blood glucose levels. Defects in &#946;-cell function are associated with hyperglycemia that can lead to diabetes. During the process of insulin secretion, &#946;-cells experience an influx of Ca2+. Thus, imaging the glucose-stimulated Ca2+ influx using genetically encoded calcium indicators (GECIs) provides an avenue to studying &#946;-cell function. Previously, studies showed that isolated zebrafish islets expressing GCaMP6s exhibit significant Ca2+ activity upon stimulation with defined glucose concentrations. However, it is paramount to study how &#946;-cells respond to glucose not in isolation, but in their native environment, where they are systemically connected, vascularized, and densely innervated. To this end, the study leveraged the optical transparency of the zebrafish larvae at early stages of development to illuminate &#946;-cell activity in vivo. Here, a detailed protocol for Ca2+ imaging and glucose stimulation to investigate &#946;-cell function in vivo is presented. This technique allows to monitor the coordinated Ca2+ dynamics in &#946;-cells with single-cell resolution. Additionally, this method can be applied to work with any injectable solution such as small molecules or peptides. Altogether, the protocol illustrates the potential of the zebrafish model to investigate islet coordination in vivo and to characterize how environmental and genetic components might affect &#946;-cell function.

Here, the study presents a protocol for calcium imaging and glucose stimulation of the pancreatic &#946;-cells of the zebrafish in vivo.

The pancreatic &#946;-cells sustain systemic glucose homeostasis by producing and secreting insulin according to the blood glucose levels. Defects in ...

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 Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Deep-Tissue Three-Photon Fluorescence Microscopy in Intact Mouse and Zebrafish Brain

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo imaging with subcellular resolution. 3PM has two major advantages for deep-tissue imaging over 2PM that has been widely used in biology laboratories: (i) longer attenuation length in scattering tissues by employing ~1,300 nm or ~1,700 nm excitation laser; (ii) less background fluorescence generation due to higher-order nonlinear excitation. As a result, 3PM allows high-contrast structural and functional imaging deep within scattering tissues such as intact mouse brain from the cortical layers to the hippocampus and the entire forebrain of adult zebrafish.
Today, laser sources suitable for 3PM are commercially available, enabling the conversion of an existing two-photon (2P) imaging system to a three-photon (3P) system. Additionally, multiple commercial 3P microscopes are available, which makes this technique readily available to biology research laboratories. This paper shows the optimization of a typical 3PM setup, particularly targeting biology groups that already have a 2P setup, and demonstrates intravital 3D imaging in intact mouse and adult zebrafish brains. This protocol covers the full experimental procedure of 3P imaging, including microscope alignment, prechirping of ~1,300 and ~1,700 nm laser pulses, animal preparation, and intravital 3P fluorescence imaging deep in adult zebrafish and mouse brains.

Three-photon microscopy enables high-contrast fluorescence imaging deep in living biological tissues, such as mouse and zebrafish brains, with high spatiotemporal resolution.

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo ...

In Vivo Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...

In Vivo Confocal Fluorescence Imaging of Neural Activity Induced by Sensory Stimulation in Partially Restrained Larval Zebrafish

Zebrafish larvae are a promising vertebrate model system for studying the neural mechanisms of behavior. Their translucence and relatively simple neural circuitry facilitate the use of optogenetic techniques in cellular analyses of behavior. Fluorescent indicators of in vivo neural activity, such as GCaMP6s, have been widely used to study the neural activity associated with simple behaviors in larval zebrafish. Here, we present a protocol for detecting sensory-induced activity in semi-restrained zebrafish larvae using the transgenic line Tg(elav3:GCaMP6s). In particular, we use the chemical agent allyl isothiocyanate to induce a robust, reproducible fluorescent response in a brain region at the border of the hindbrain and spinal cord. We discuss the potential uses of GCaMP6s for optical monitoring of neural activity during a range of behavioral paradigms and the limitations of this technique. Our protocol outlines an accessible approach for monitoring dynamic, behavior-related in vivo neural activity in the larval zebrafish brain.

Here, we present a detailed protocol to examine neural activity in brain regions of transgenic zebrafish that express GCaMP calcium indicators using confocal microscopy.

Zebrafish larvae are a promising vertebrate model system for studying the neural mechanisms of behavior. Their translucence and relatively simple neural ...