Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Prepare the following ahead of time.

1. Prepare Optic Cable

1. 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 ° angle.
2. 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.
3. 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.
4. Insert optic cable into prepared Pasteur pipette. Make sure cable can easily move in and out of the pipette tip.
5. With optic cable protruding from tip of Pasteur pipette, carefully cut and remove fiber cladding from around glass fiber, ~2 mm from cut end.
6. 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).
7. Retract optic cable into Pasteur pipette for storage. Repeat step (1.6) if tip of optic fiber gets chipped or breaks unevenly.
8. Position optic cable in Pasteur pipette using a micromanipulator (Figure 1C).

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

1. 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.
2. Store needles in a secure container (i.e. a Petri dish with rolled tape, adhesive side out).

3. Pour Injection Molds

1. Melt 0.5 g agarose in 30 ml embryo/blue water, until agarose has completely dissolved.
2. Pour into bottom half of a Petri dish.
3. Place rectangular mold with mounting wells (Figure 2) into agarose, being careful to limit bubble formation around the wells.
4. Allow agarose to solidify.
5. Remove mold and fill Petri dish with blue/embryo water.
6. Store upright, filled with clean water, at room temp for same day use or at 4 °C for future use.

4. Make Plasmid DNA Mix for Injections

1. Dilute plasmid DNA to a concentration of 50 ng/μl with 1:10 phenol red in ddH₂O. For example:

   1.0 ml	plasmid DNA (250 ng/ml)
   0.5 ml	phenol red
   3.5 ml	ddH2O

2. DNA mix can be kept at room temperature if using immediately or stored at 4 °C for several days.

5. Set up Mating Pairs

This should be done the evening before you plan to do injections.

1. 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.

6. Prepare for Injections (Can be done while waiting for embryos)

1. Turn on pressure injector rig. Make sure the system is set to pulse. Start with the PULSE DURATION set to 1.
2. Turn on the AIR valve and adjust pressure injector to ~20 psi.
3. Using a dissecting scope at the highest magnification, break tip of needle with forceps or a poker to create a ~2 μm opening (Figure 3, Movie 1).
4. 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 μl of the DNA mix directly into the needle.
5. Place filled needle in a safe place until ready to inject.

7. Collect Embryos

1. 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.

8. Inject Embryos at the 1-2 Cell Stage

1. Transfer harvested embryos to injection molds using a plastic pipette.
2. Using a dissecting microscope, gently push embryos into wells with forceps or a blunt poker (Figure 4, Movie 2).
3. Place loaded needle into a micromanipulator and position over embryos.
4. 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.
5. 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).
6. Remove injected embryos from mold using a gentle stream of blue/embryo water.
7. Store injected embryos at 28.5 °C in the dark.
8. Remove unfertilized, damaged or dead embryos periodically.
9. Treat embryos with PTU between 18-24 hpf to prevent pigmentation.

9. Screen for Transgene Expression

1. Manually dechorionate 24-48 hpf embryos using forceps.
2. Anesthetize larvae using 0.02% tricaine.
3. 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.
4. Store embryos at 28.5 °C in the dark until desired experimental stage.

10. Mount Larvae for Behavior Experiments

1. Make 1.5% low melt agarose in ddH₂O and store in a 42 °C heat block to prevent it from solidifying.
2. 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.
3. Transfer larvae in a drop of agarose onto a small Petri dish.
4. Under a dissecting microscope, position larva dorsal up.
5. When agarose has solidified, cut away the agarose with a thin razor blade (#11 scalpel), leaving a wedge of agar around the entire larva.
6. Make two diagonal cuts at either side of the yolk; take care not to nick the larva.
7. Fill the area surrounding the agarose with embryo/blue water.
8. Pull agarose away from the trunk and tail of the larva (Figure 6).

11. Prepare High-speed Camera and Imaging Software

1. Mount high-speed camera onto dissecting scope.
2. Connect camera to computer.
3. Turn on computer.
4. Turn on high-speed camera.
5. Open video/imaging software (We use AOS imaging software and will describe procedures for using it here, but other imaging software is equally acceptable).
6. Adjust camera settings accordingly (i.e. 1,000 frames per sec (fps), 50% trigger buffer or other preferred settings).
7. Start recording.

12. Activate Single Neurons Using a 473 nm Laser

1. Attach stimulator, laser and optic cable.
2. Turn on stimulator.
3. Set stimulator to a maximum of 5 Volts and a pulse duration of 5 msec.
4. Turn on laser according to manufacturer’s instructions.
5. Use a dissecting microscope to position the tip of the optic cable near cell body of a neuron with ChEF-tdTomato expression (Figure 6).
6. Deliver pulse of blue light to activate sensory neuron.
7. Record behavior using a high-speed camera set at 500 or 1,000 frames per sec.
8. Repeat experiment as desired, waiting 1 min between each activation to avoid habituation. (We record a minimum of three responses for each neuron).
9. 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.
10. Transfer larva to culture plate with fresh blue/embryo water. We use a 24-well plate to keep track of individual larvae.

13. Image Neuron(s) with a Confocal Microscope

1. Anesthetize larvae with 0.02% tricaine.
2. Mount larvae, dorsal side up, in 1.2% low melt agarose or in a dorsal mold.
3. Image ChEF-tdTomato neurons with a 543 nm laser and appropriate filter and objective. We use a 20x objective.
4. Remove larvae from agarose and return to individual well with blue/embryo water.
5. Store at 28.5 °C in the dark for further analysis.