We thank Fumi Kubo, Tod Thiele and HerwigBaier (UCSF/Max Planck Institute) for advice on behavior experiments and DPSS laser set up; Heesoo Kim and Chiara Cerri from the MBL Neurobiology Course for assisting in ChEF-tdTomato experiments; PetronellaKettunen (University of Gothenburg)for initial collaboration on optogenetic experiments; BaljitKhakh, Eric Hudson, Mike Baca and John Milligan (UCLA) for technical advice; and Roger Tsien (UCSD) for the ChEF-tdTomato construct. This work was supported by an NRSA (5F31NS064817) award to AMSP and a grant from the NSF (RIG:0819010) to AS.

Prepare the following ahead of time.
1. Prepare Optic Cable
<ol>
 <li>Create a storage unit for the optic cable by melting the tapered neck of a glass Pasteur pipette over a Bunsen burner to create a ~150 &deg; angle.</li>
 <li>Using a wire cutter or a razor blade, carefully cut the optic cable into two pieces. Each piece should have one end with a FC/PC adaptor and one exposed end. Store one piece as a reserve cable.</li>
 <li>Strip the optic cable down to the cladding by removing the fiber jacket and strengthening fibers (Figure 1A) from 5.0 cm of cut end of the cable.</li>
 <li>Insert optic cable into prepared Pasteur pipette. Make sure cable can easily move in and out of the pipette tip.</li>
 <li>With optic cable protruding from tip of Pasteur pipette, carefully cut and remove fiber cladding from around glass fiber, ~2 mm from cut end.</li>
 <li>Using a diamond pen/glass cutter, nick the glass fiber and break off the end to create a clean cut/surface at the end of the fiber (Figure 1B).</li>
 <li>Retract optic cable into Pasteur pipette for storage. Repeat step (1.6) if tip of optic fiber gets chipped or breaks unevenly.</li>
 <li>Position optic cable in Pasteur pipette using a micromanipulator (Figure 1C).</li>
</ol>
Procedures 2-8 describe a method for injecting transgenes into embryos generally applicable to many zebrafish experiments. Variations on this method, like those described in other JoVE videos 6, 7, 8, 9, 22 are equally effective.
2. Pull Injection Needles
<ol>
 <li>Using a needle puller, pull borosilicate glass tubing into two injection needles with a gradually tapered tip. Puller settings will vary. (On a Sutter Instruments needle puller we use settings: P = 500, Heat = 720, Pull = 50, Velocity = 70 and Time = 150). Every needle puller is different, so optimize puller settings empirically. For more tapered needles, increase the Heat and/or Pull. For less tapered needles, increase Time and/or decrease Pull.</li>
 <li>Store needles in a secure container (i.e. a Petri dish with rolled tape, adhesive side out).</li>
</ol>
3. Pour Injection Molds
<ol>
 <li>Melt 0.5 g agarose in 30 ml embryo/blue water, until agarose has completely dissolved.</li>
 <li>Pour into bottom half of a Petri dish.</li>
 <li>Place rectangular mold with mounting wells (Figure 2) into agarose, being careful to limit bubble formation around the wells.</li>
 <li>Allow agarose to solidify.</li>
 <li>Remove mold and fill Petri dish with blue/embryo water.</li>
 <li>Store upright, filled with clean water, at room temp for same day use or at 4 &deg;C for future use.</li>
</ol>
4. Make Plasmid DNA Mix for Injections
<ol>
 <li>Dilute plasmid DNA to a concentration of 50 ng/&mu;l with 1:10 phenol red in ddH2O. For example:
 <table border="1">
 <tbody>
 <tr>
 <td>1.0 ml</td>
 <td>plasmid DNA (250 ng/ml)</td>
 </tr>
 <tr>
 <td>0.5 ml</td>
 <td>phenol red</td>
 </tr>
 <tr>
 <td>3.5 ml</td>
 <td>ddH2O</td>
 </tr>
 </tbody>
 </table>
 </li>
 <li>DNA mix can be kept at room temperature if using immediately or stored at 4 &deg;C for several days.</li>
</ol>
5. Set up Mating Pairs
This should be done the evening before you plan to do injections.
<ol>
 <li>Fill breeding tanks with system water and place male and female fish together. If injections cannot be performed as soon as lights turn on in the facility, separate male and female fish with a divider.</li>
</ol>
6. Prepare for Injections (Can be done while waiting for embryos)
<ol>
 <li>Turn on pressure injector rig. Make sure the system is set to pulse. Start with the PULSE DURATION set to 1.</li>
 <li>Turn on the AIR valve and adjust pressure injector to ~20 psi.</li>
 <li>Using a dissecting scope at the highest magnification, break tip of needle with forceps or a poker to create a ~2 &mu;m opening (Figure 3, Movie 1).</li>
 <li>Fill needles with DNA mix by: 1. Placing the needle, tip side up, into the DNA mix and letting the needle fill by capillary action or 2. Using a long-reach tip to pipette 1-2 &mu;l of the DNA mix directly into the needle.</li>
 <li>Place filled needle in a safe place until ready to inject.</li>
</ol>
7. Collect Embryos
<ol>
 <li>After facility lights have turned on, remove dividers (if applicable). Collect embryos with a strainer/small sieve and transfer them to a Petri dish with fresh blue/embryo water.</li>
</ol>
8. Inject Embryos at the 1-2 Cell Stage
<ol>
 <li>Transfer harvested embryos to injection molds using a plastic pipette.</li>
 <li>Using a dissecting microscope, gently push embryos into wells with forceps or a blunt poker (Figure 4, Movie 2).</li>
 <li>Place loaded needle into a micromanipulator and position over embryos.</li>
 <li>Calibrate injection volume by adjusting the PULSE DURATION in 1-step increments until you achieve the desired volume (~1 nl). This could be calibrated with a micrometer slide, as described in a previous JoVE videos 8, 22, but for this experiment precision is not necessary, since a wide range of injection volumes will result in adequate expression.</li>
 <li>Inject ~1 nl DNA mix directly into the cell and repeat for all embryos. DNA can also be injected into the yolk, but this tends to be less efficient (Figure 5, Movie 3).</li>
 <li>Remove injected embryos from mold using a gentle stream of blue/embryo water.</li>
 <li>Store injected embryos at 28.5 &deg;C in the dark.</li>
 <li>Remove unfertilized, damaged or dead embryos periodically.</li>
 <li>Treat embryos with PTU between 18-24 hpf to prevent pigmentation.</li>
</ol>
9. Screen for Transgene Expression
<ol>
 <li>Manually dechorionate 24-48 hpf embryos using forceps.</li>
 <li>Anesthetize larvae using 0.02% tricaine.</li>
 <li>Using a fluorescent dissecting microscope, identify larvae with RB or trigeminal neuron expression and transfer these to a new dish with fresh PTU blue/embryo water. Embryos with sparse expression in easily identifiable cells are optimal, but individual neurons will be targeted for activation with a fiber optic cable so a wide range of expression density is acceptable.</li>
 <li>Store embryos at 28.5 &deg;C in the dark until desired experimental stage.</li>
</ol>
10. Mount Larvae for Behavior Experiments
<ol>
 <li>Make 1.5% low melt agarose in ddH2O and store in a 42 &deg;C heat block to prevent it from solidifying.</li>
 <li>Using a glass Pasteur pipette, transfer one of the pre-screened larvae into a tube of 1.5% low melt agarose with as little blue/embryo water as possible.</li>
 <li>Transfer larvae in a drop of agarose onto a small Petri dish.</li>
 <li>Under a dissecting microscope, position larva dorsal up.</li>
 <li>When agarose has solidified, cut away the agarose with a thin razor blade (#11 scalpel), leaving a wedge of agar around the entire larva.</li>
 <li>Make two diagonal cuts at either side of the yolk; take care not to nick the larva.</li>
 <li>Fill the area surrounding the agarose with embryo/blue water.</li>
 <li>Pull agarose away from the trunk and tail of the larva (Figure 6).</li>
</ol>
11. Prepare High-speed Camera and Imaging Software
<ol>
 <li>Mount high-speed camera onto dissecting scope.</li>
 <li>Connect camera to computer.</li>
 <li>Turn on computer.</li>
 <li>Turn on high-speed camera.</li>
 <li>Open video/imaging software (We use AOS imaging software and will describe procedures for using it here, but other imaging software is equally acceptable).</li>
 <li>Adjust camera settings accordingly (i.e. 1,000 frames per sec (fps), 50% trigger buffer or other preferred settings).</li>
 <li>Start recording.</li>
</ol>
12. Activate Single Neurons Using a 473 nm Laser
<ol>
 <li>Attach stimulator, laser and optic cable.</li>
 <li>Turn on stimulator.</li>
 <li>Set stimulator to a maximum of 5 Volts and a pulse duration of 5 msec.</li>
 <li>Turn on laser according to manufacturer's instructions.</li>
 <li>Use a dissecting microscope to position the tip of the optic cable near cell body of a neuron with ChEF-tdTomato expression (Figure 6).</li>
 <li>Deliver pulse of blue light to activate sensory neuron.</li>
 <li>Record behavior using a high-speed camera set at 500 or 1,000 frames per sec.</li>
 <li>Repeat experiment as desired, waiting 1 min between each activation to avoid habituation. (We record a minimum of three responses for each neuron).</li>
 <li>To release larvae, pry apart agarose with forceps, taking care not to injure the animal. This animal can be allowed to develop further and the procedure can be repeated at an older stage to characterize development of the behavior. The embryo can also be remounted for high-resolution confocal imaging of the activated cell to correlate behavior with cellular structure, as described below.</li>
 <li>Transfer larva to culture plate with fresh blue/embryo water. We use a 24-well plate to keep track of individual larvae.</li>
</ol>
13. Image Neuron(s) with a Confocal Microscope
<ol>
 <li>Anesthetize larvae with 0.02% tricaine.</li>
 <li>Mount larvae, dorsal side up, in 1.2% low melt agarose or in a dorsal mold.</li>
 <li>Image ChEF-tdTomato neurons with a 543 nm laser and appropriate filter and objective. We use a 20x objective.</li>
 <li>Remove larvae from agarose and return to individual well with blue/embryo water.</li>
 <li>Store at 28.5 &deg;C in the dark for further analysis.</li>
</ol>

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The authors declare that they have no competing financial interest.

We have described an approach for optogenetic activation of single RB neurons in live zebrafish. Our method employs transient transgenesis to express a fluorescently tagged channelrhodopsin variant, ChEF-tdTomato13, in specific somatosensory neurons. This approach could easily be adapted for use in other larval zebrafish cell populations.
Using this approach we consistently elicited behavioral responses from 34-48 hpf larvae expressing ChEF-tdTomato. Using a 5 msec pulse of bl...

The development of optogenetic methods for promoting or inhibiting neuronal excitability with defined wavelengths of light has made it possible to study the function of distinct populations of neurons in neural circuits controlling behavior 1, 19, 21. This technique is often used to activate groups of neurons, but it can also be used to activate individual neurons. Zebrafish larvae are particularly amenable to these methods since they are translucent, their nervous system develops quickly, and creating transgenic animals is fast and routine. However, significant technical hurdles must be overcome to reliably achieve single neuron activation.
To optimize a procedure for optogenetic activation of single zebrafish neurons, we focused on somatosensory neurons. Zebrafish larvae detect a variety of somatosensory stimuli using two populations of neurons: trigeminal neurons, which innervate the head, and Rohon-Beard (RB) neurons, which innervate the rest of the body. Each trigeminal and RB neuron projects a peripheral axon that branches extensively in the skin to detect stimuli and a central axon that connects to downstream neural circuits. Animals respond to touch as early as 21 hr post-fertilization (hpf), indicating that coherent somatosensory circuits have formed 5, 18. During larval development at least some trigeminal and RB neurons synapse onto the Mauthner cell to activate classic escape responses, but accumulating evidence suggests that there are multiple classes of somatosensory neurons with different patterns of connectivity that may elicit variations on the escape behavior 2, 4, 10, 12, 14, 15, 16, 17. Our motivation for developing this method was to characterize the behavioral function of different classes of somatosensory neurons, but this approach could in principle be used to study the function of almost any neuron or population of neurons in larval zebrafish.
Douglass et al. previously described a method for activating Channelrhodopsin-2-expressing somatosensory neurons with blue light, eliciting escape behavior 3. Their approach used an enhancer element from the isl1 gene to drive expression of ChR2-EYFP in somatosensory neurons. This transgene, however, was reported to display relatively weak fluorescence, requiring co-injection of a second reporter, UAS::GFP, to allow visualization of cells expressing ChR2-EYFP. This approach was used to elicit behavior responses between 24-48 hpf, but could never elicit a response past 72 hpf. Thus, while this method works for studying neural circuitry at very early larval stages (24-48 hpf), it is inadequate for characterizing neural circuits and behavioral responses in older larvae, when more diverse behavioral responses are apparent and neural circuits are more mature.
We sought to improve the sensitivity of this technique in order to characterize the function of subpopulations of larval RB neurons. To improve expression we used a somatosensory-specific enhancer (CREST3) 20 to drive expression of LexA-VP16 and a stretch of LexA operator sequences (4xLexAop) 11 to amplify the expression of a fluorescently tagged light-activated channel. This configuration amplified expression of the channel, eliminating the need for co-expressing a second reporter and allowing us to directly determine the relative abundance of the channel in each neuron. Using the LexA/LexAop sequence had the additional advantage of allowing us to introduce the transgene into zebrafish reporter lines that use the Gal4/UAS system. Transient expression of this transgene resulted in varying levels of expression, but was usually robust enough to visualize both the cell body and axonal projections of individual neurons over several days. To optimize sensitivity to light we used the light activated channel ChEF, a channelrhodopsin variant consisting of a chimera of channelopsin-1 (Chop1) and channelopsin-2 (Chop2) with a crossover site at helix loop E-F 13. This channel is activated at the same wavelength as ChR2, but requires lower light intensity for activation, making it more sensitive than other commonly used channels, including ChR2. The ChEF protein was fused to the red fluorescent protein, tdTomato, enabling us to screen for protein expression without activating the channel. As a light source, we used a diode pumped solid-state (DPSS) laser coupled to a fiber optic cable to deliver a precise, high-powered pulse of blue light to a specific region of the larvae. This allowed us to focus laser light on individual neurons, eliminating the need for finding rare transgenic animals expressing the channel in a single neuron. Using this approach, we were able to activate single RB neurons, record behavioral responses with a high-speed video camera, and image the activated neurons at high resolution with confocal microscopy.

<table border="1">
<tbody>
 <tr>
 <td>Name of 			Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Materials</td>
 </tr>
 <tr>
 <td>Glass 			Pasteur pipette</td>
 <td>Fisher</td>
 <td>1367820B</td>
 <td>or 			equivalent (10-15 mm diameter)</td>
 </tr>
 <tr>
 <td>200 			&mu;m 			optic fiber</td>
 <td>ThorLabs</td>
 <td>AFS200/220Y-CUSTOM</td>
 <td>Patch Cord, Length: 3 m, End A: FC/PC, End B: FC/PC, Jacket: FT030</td>
 </tr>
 <tr>
 <td>50 			&mu;m 			optic fiber</td>
 <td>ThorLabs</td>
 <td>AFS50/125Y-CUSTOM</td>
 <td>Patch Cord, Length: 3 m, End A: FC/PC, End B: FC/PC, Jacket: FT030</td>
 </tr>
 <tr>
 <td>Adjustable 			Stripping Tool</td>
 <td>ThorLabs</td>
 <td>AFS900</td>
 <td>or 			Three-Hole Stripping Tool (FTS4)</td>
 </tr>
 <tr>
 <td>Diamond 			Wedge scribe</td>
 <td>ThorLabs</td>
 <td>S90W</td>
 <td></td>
 </tr>
 <tr>
 <td>Flaming/Brown Micropipette 			Puller</td>
 <td>Sutter 			Instruments</td>
 <td>P-97</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Borosilicate 			glass tubing with filament</td>
 <td>Sutter 			Instruments</td>
 <td>BF-100-78-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Injection 			mold</td>
 <td>n/a</td>
 <td>n/a</td>
 <td>Figure 			5</td>
 </tr>
 <tr>
 <td>1.5 			ml centrifuge tubes</td>
 <td>Any</td>
 <td>Any</td>
 <td></td>
 </tr>
 <tr>
 <td>Petri 			dish (100x15 mm)</td>
 <td>Any</td>
 <td>Any</td>
 <td></td>
 </tr>
 <tr>
 <td>Petri 			dish (60x15 mm)</td>
 <td>Any</td>
 <td>Any</td>
 <td></td>
 </tr>
 <tr>
 <td>Pressure 			injector</td>
 <td>ASI</td>
 <td>MPPI-3</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Micromanipulator 			and metal stand</td>
 <td>Narashige</td>
 <td>M152</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Disposable 			plastic pipettes</td>
 <td>Fisherbrand</td>
 <td>13-711-7</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Poker 			(Pin holder and Insect pin)</td>
 <td>Fine 			Science Tools, Inc.</td>
 <td>26018-17 			and 26000-70</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>Fine 			Science Tools, Inc.</td>
 <td>11255-20</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Microloader 			pipette tips</td>
 <td>Eppendorf</td>
 <td>9300001007</td>
 <td></td>
 </tr>
 <tr>
 <td>28.5 &deg;C 			incubator</td>
 <td>any</td>
 <td>any</td>
 <td></td>
 </tr>
 <tr>
 <td>42 &deg;C 			heat block</td>
 <td>Any</td>
 <td>Any</td>
 <td></td>
 </tr>
 <tr>
 <td>Non-Sterile 			scalpel blades #11</td>
 <td>Fine 			Scientific Tools, Inc.</td>
 <td>10011-00</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Dissecting 			scope</td>
 <td>Zeiss</td>
 <td>Stemi</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Fluorescent 			dissecting scope with standard filter</td>
 <td>Any</td>
 <td>any</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Confocal 			microscope </td>
 <td>Zeiss</td>
 <td>LSM 			510 or 710</td>
 <td>or 			equivalent with lasers for GFP and RFP, and 10x, 20x and 40x 			objectives </td>
 </tr>
 <tr>
 <td>High 			speed camera</td>
 <td>AOS 			Technologies, Inc.</td>
 <td>X-PRI 			(130025-10)</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>473 			nm portable laser</td>
 <td>Crystal 			lasers</td>
 <td>CL-473-050</td>
 <td>or 			higher power, with TTL option</td>
 </tr>
 <tr>
 <td>S48 			Stimulator</td>
 <td>Astro-Med, 			Inc. Grass Instrument division</td>
 <td>S48K</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>FC/PC 			to FC/PC mating sleeve</td>
 <td>ThorLabs</td>
 <td>ADAFC1</td>
 <td>May 			need for optic cable connection</td>
 </tr>
 <tr>
 <td>Laser 			Safety Glasses</td>
 <td>ThorLabs</td>
 <td>LG10</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>24 			culture 			plates</td>
 <td>Genesee</td>
 <td>25-102</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Single 			depression slides</td>
 <td>Fisher</td>
 <td>S175201</td>
 <td>Or 			equivalent</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Instant 			ocean</td>
 <td>Aquatic 			Ecosystems</td>
 <td>IS50</td>
 <td></td>
 </tr>
 <tr>
 <td>Methylene 			blue</td>
 <td>Fisher</td>
 <td>S71325</td>
 <td></td>
 </tr>
 <tr>
 <td>Phenol 			red</td>
 <td>Sigma</td>
 <td>P4758</td>
 <td></td>
 </tr>
 <tr>
 <td>Agarose</td>
 <td>EMD</td>
 <td>2125</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>Low 			Melt agarose</td>
 <td>Sigma</td>
 <td>A9045</td>
 <td>or 			equivalent</td>
 </tr>
 <tr>
 <td>PTU</td>
 <td>Sigma</td>
 <td>P7629</td>
 <td></td>
 </tr>
 <tr>
 <td>Tricaine</td>
 <td>Sigma</td>
 <td>A5040</td>
 <td></td>
 </tr>
 <tr>
			<td>blue/embryo water</td>
			<td></td>
			<td></td>
			<td>10 L ddH2O 
			0.6 g Instant Ocean 
			6 drops methylene blue</td>
		</tr>
		<tr>
			<td>phenol red </td>
			<td></td>
			<td></td>
			<td>(5 mg/ml in 0.2 M KCl)</td>
		</tr>
		<tr>
			<td>100x PTU</td>
			<td></td>
			<td></td>
			<td>0.150 g PTU 
			50 ml ddH2O 
			dissolve at 70 &deg;C, shake often 
			aliquot and store at -20 &deg;C</td>
		</tr>
		<tr>
			<td>1x PTU</td>
			<td></td>
			<td></td>
			<td>1 ml 100x PTU 
			99 ml blue/fish water</td>
		</tr>
		<tr>
			<td>Tricaine stock solution</td>
			<td></td>
			<td></td>
			<td>400 mg tricaine 
			97.9 ddH2O</td>
		</tr>
		<tr>
			<td>~2.1 ml 1M Tris, pH9.0</td>
			<td></td>
			<td></td>
			<td>adjust pH to ~7.0 
			store in 4 &deg;C or -20 &deg;C for long term storage</td>
		</tr>
 </tbody>
</table>

optogenetic activation of zebrafish somatosensory neurons using chef-tdtomato

Larval zebrafish are emerging as a model for describing the development and function of simple neural circuits. Due to their external fertilization, rapid development, and translucency, zebrafish are particularly well suited for optogenetic approaches to investigate neural circuit function. In this approach, light-sensitive ion channels are expressed in specific neurons, enabling the experimenter to activate or inhibit them at will and thus assess their contribution to specific behaviors. Applying these methods in larval zebrafish is conceptually simple but requires the optimization of technical details. Here we demonstrate a procedure for expressing a channelrhodopsin variant in larval zebrafish somatosensory neurons, photo-activating single cells, and recording the resulting behaviors. By introducing a few modifications to previously established methods, this approach could be used to elicit behavioral responses from single neurons activated up to at least 4 days post-fertilization (dpf). Specifically, we created a transgene using a somatosensory neuron enhancer, CREST3, to drive the expression of the tagged channelrhodopsin variant, ChEF-tdTomato. Injecting this transgene into 1-cell stage embryos results in mosaic expression in somatosensory neurons, which can be imaged with confocal microscopy. Illuminating identified cells in these animals with light from a 473 nm DPSS laser, guided through a fiber optic cable, elicits behaviors that can be recorded with a high-speed video camera and analyzed quantitatively. This technique could be adapted to study behaviors elicited by activating any zebrafish neuron. Combining this approach with genetic or pharmacological perturbations will be a powerful way to investigate circuit formation and function.

<img src="/files/ftp_upload/50184/50184fig1.jpg" alt="Figure 1"/> 
 Figure 1. Optic cable set up. (A) Layers of a fiber optic cable. (B) Stripped fiber optic cable in a Pasteur pipette. (C) Fiber optic cable in Pasteur pipette positioned using a micromanipulator.
<img src="/files/ftp_upload/50184/50184fig2.jpg" alt="Figure 2"/> 
 Figure 2. Injection mold template.
<p c...

Watch this Scientific Journal Video about Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato at JoVE.com

Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Optogenetic techniques have made it possible to study the contribution of specific neurons to behavior. We describe a method in larval zebrafish for activating single somatosensory neurons expressing a channelrhodopsin variant (ChEF) with a diode-pumped solid state (DPSS) laser and recording the elicited behaviors with a high-speed video camera.

Larval  zebrafish are emerging as a model for describing the development and function  of simple neural circuits. Due to their  external fertilization, ...

optogenetic-activation-zebrafish-somatosensory-neurons-using-chef

Research

JoVE Journal

Neuroscience

14.7K visualizações.  University of California, Los Angeles. Optogenética técnicas tornaram possível estudar a contribuição dos neurónios específicos para o comportamento. Nós descrevemos um método em zebrafish larval para ativar neurônios individuais somatossensoriais expressando uma variante channelrodopsina (chef) com um diodo-bombeado do laser de estado sólido (DPSS) e registrando os comportamentos eliciados com uma câmera de vídeo de alta velocidade.

Vídeo: Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific regions of developing embryos, including those of C. elegans, chick, Xenopus, zebrafish, and mouse 1. In zebrafish, in vivo electroporation has been shown to have excellent spatial and temporal resolution for the delivery of these reagents 2-7. The temporal resolution of this method is important because it allows for incorporation of these reagents at specific stages in development. Furthermore, because expression from electroporated vectors occurs within 6 hours 7, this method is more timely than transgenic approaches. While the spatial resolution can be extremely precise when targeting a single cell 2, 6, it is often preferable to incorporate reagents into a specific cell population within a tissue or structure. When targeting multiple cells, in vivo electroporation is efficient for delivery to a specific region of the embryo; however, particularly within the developing nervous system, it is difficult to target specific cell types solely through spatially discrete electroporation. Alternatively, enhancer trap transgenic lines offer excellent cell type-specific expression of transgenes 8. Here we describe an approach that combines transgenic Gal4-based enhancer trap lines 8 with spatially discrete in vivo electroporation 7, 9 to specifically target developing neurons of the zebrafish olfactory bulb. The Et(zic4:Gal4TA4,UAS:mCherry)hzm5 (formerly GA80_9) enhancer trap line previously described 8, displays targeted transgenic expression of mCherry mediated by a zebrafish optimized Gal4 (KalTA4) transcriptional activator in multiple regions of the developing brain including hindbrain, cerebellum, forebrain, and the olfactory bulb. To target GFP expression specifically to the olfactory bulb, a plasmid with the coding sequence of GFP under control of multiple Gal4 binding sites (UAS) was electroporated into the anterior end of the forebrain at 24-28 hours post-fertilization (hpf). Although this method incorporates plasmid DNA into multiple regions of the forebrain, GFP expression is only induced in cells transgenically expressing the KalTA4 transcription factor. Thus, by using the GA080_9 transgenic line, this approach led to GFP expression exclusively in the developing olfactory bulb. GFP expressing cells targeted through this approach showed typical axonal projections, as previously described for mitral cells of the olfactory bulb 10. This method could also be used for targeted delivery of other reagents including short-hairpin RNA interference expression plasmids, which would provide a method for spatially and temporally discrete loss-of-function analysis.

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

The temporal and spatial resolution of genetic manipulations determines the spectrum of biological phenomena that they can perturb. Here we use temporally and spatially discrete in vivo electroporation, combined with transgenic lines of zebrafish, to induce expression of a GFP transgene specifically in neurons of the developing olfactory bulb.

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific ...

Using Affordable LED Arrays for Photo-Stimulation of Neurons

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single cells in response to electrical or pharmacological manipulations1,2. With the invention of methods to optically control genetically targeted neurons (optogenetics), researchers now have an unprecedented level of control over specific groups of neurons in the standard slice preparation. In particular, photosensitive channelrhodopsin-2 (ChR2) allows researchers to activate neurons with light3,4. By combining careful calibration of LED-based photostimulation of ChR2 with standard slice electrophysiology, we are able to probe with greater detail the role of adult-born interneurons in the olfactory bulb, the first central relay of the olfactory system. Using viral expression of ChR2-YFP specifically in adult-born neurons, we can selectively control young adult-born neurons in a milieu of older and mature neurons. Our optical control uses a simple and inexpensive LED system, and we show how this system can be calibrated to understand how much light is needed to evoke spiking activity in single neurons. Hence, brief flashes of blue light can remotely control the firing pattern of ChR2-transduced newborn cells.

Adult-born neurons expressing ChR2 can be manipulated in slice electrophysiological preparations in order to examine their contribution towards the function of olfactory neural circuits.

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single ...

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Somatosensory Event-related Potentials from Orofacial Skin Stretch Stimulation

Cortical processing associated with orofacial somatosensory function in speech has received limited experimental attention due to the difficulty of providing precise and controlled stimulation. This article introduces a technique for recording somatosensory event-related potentials (ERP) that uses a novel mechanical stimulation method involving skin deformation using a robotic device. Controlled deformation of the facial skin is used to modulate kinesthetic inputs through excitation of cutaneous mechanoreceptors. By combining somatosensory stimulation with electroencephalographic recording, somatosensory evoked responses can be successfully measured at the level of the cortex. Somatosensory stimulation can be combined with the stimulation of other sensory modalities to assess multisensory interactions. For speech, orofacial stimulation is combined with speech sound stimulation to assess the contribution of multi-sensory processing including the effects of timing differences. The ability to precisely control orofacial somatosensory stimulation during speech perception and speech production with ERP recording is an important tool that provides new insight into the neural organization and neural representations for speech.

This paper introduces a method for obtaining somatosensory event-related potentials following orofacial skin stretch stimulation. The current method can be used to evaluate the contribution of somatosensory afferents to both speech production and speech perception.

Cortical processing associated with orofacial somatosensory function in speech has received limited experimental attention due to the difficulty of ...

M&#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after severe damage. Investigating the regeneration pathways in zebrafish might bring new insights to develop innovative strategies for the treatment of retinal degenerative diseases in mammals. Herein, we focused on the induction of a focal lesion to the outer retina in adult zebrafish by means of a 532 nm diode laser. A localized injury allows investigating biological processes that take place during retinal degeneration and regeneration directly at the area of damage. Using non-invasive optical coherence tomography (OCT), we were able to define the location of the damaged area and monitor subsequent regeneration in vivo. Indeed, OCT imaging produces high-resolution, cross-sectional images of the zebrafish retina, providing information which was previously only available with histological analyses. In order to confirm the data from real-time OCT, histological sections were performed and regenerative response after the induction of the retinal injury was investigated by immunohistochemistry.

M&amp;#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

The zebrafish is a popular animal model to study mechanisms of retinal degeneration/regeneration in vertebrates. This protocol describes a method to induce localized injury disrupting the outer retina with minimal damage to the inner retina. Subsequently, we monitor in vivo the retinal morphology and the M&#252;ller glia response throughout retinal regeneration.

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a ...

Isolation and RNA Extraction of Neurons, Macrophages and Microglia from Larval Zebrafish Brains

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is important to isolate these cells without changing their gene expression profile. The zebrafish model provides a large number of transgenic fish lines in which specific cell types are labelled; for example neurons in the NBT:DsRed line or macrophages/microglia in the mpeg1:eGFP line. Furthermore, antibodies have been developed to stain specific cells, such as microglia with the 4C4 antibody.
Here, we describe the isolation of neurons, macrophages and microglia from larval zebrafish brains. Central to this protocol is the avoidance of an enzymatic tissue digestion at 37 &#176;C, which could modify cellular profiles. Instead a mechanical system of tissue homogenization at 4 &#176;C is used. This protocol entails homogenization of brains into cell suspension, their immuno-staining and the isolation of neurons, macrophages and microglia by FACS. Afterwards, we extracted RNA from those cells and evaluated their quality/quantity. We managed to obtain RNA of high quality (RNA Integrity Number (RIN) &#62; 7) to perform qPCR on macrophages/microglia and neurons, and transcriptomic analysis on microglia. This approach enables a better characterization of these cells, as well as a clearer understanding about their role in development and pathologies.

We present a protocol to isolate neurons, macrophages and microglia from larval zebrafish brains under physiological and pathological conditions. Upon isolation, RNA is extracted from these cells to analyze their gene expression profile. This protocol allows for the collection of high-quality RNA for performing downstream analysis like qPCR and transcriptomics.

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is ...