1. Preparing for microinjection
- Prepare for the injection by pulling glass capillaries using a micropipette puller (see Table of Materials), following the five-step protocol for glass capillary tube pulling (Table 1).
- Open the tip of the needle (thin wall glass capillaries [3.5 in] with filament, outer diameter [OD] 1.14 mm) to the appropriate size at an angle using forceps (Figure 1A).
- Fill the needle with 0.1 mL of mineral oil to ensure that there are no bubbles.
- Remove the screw cap of the steel needle of the microinjection apparatus. Align the glass needle hole with the steel needle and then tighten the screw cap. Mount the loaded needle microinjection apparatus in a micromanipulator (see Table of Materials).
- Adjust the position of the microinjection apparatus so that the micromanipulator can flexibly move the apparatus under the microscope. Discharge a certain volume of paraffin oil to make the steel needle enter the capillary glass tube.
- Drop the injection solution (such as 1× phosphate-buffered saline (PBS) or 5 mg/ mL LPS) on a glass slide sterilized with 70% ethanol and adjust the microinjection apparatus so that the tip is inserted into the liquid drop. Load ~2 µL of injection solution into the needle.
- Set up the micromanipulator (fine adjustment settings of up, down, left, and right) so that the needle tip of the microinjection apparatus is in the same field of vision as the larvae on high magnification (Figure 1B).
2. Mounting zebrafish for microinjections
NOTE: Zebrafish brain development occurs within 3 days post-fertilization (dpf) and matures at 5 dpf with a well-developed central nervous system. Therefore, 5 dpf larvae are already suitable for studying LPS-mediated neuronal damage as well as behavioral and inflammatory responses.
- Inject the zebrafish larvae within approximately 30 min of reaching the stage of interest, to maintain the consistency of injection timing.
- To anesthetize the zebrafish larvae, combine tricaine with clean fish tank water (final concentration of 0.02% w/v tricaine).
- Melt a solution of 2% agarose in double-distilled water (ddH2O) using a microwave. Pour the molten agarose into a plastic dish. The 2% agarose-coated plastic dish can be stored at 4 °C for up to 2 weeks.
- Use a plastic transfer pipette to transport anesthetized larvae to the center of the 2% agarose-coated plastic dish. Orient the mounted larvae with the brain side up for needle access.
3. Injecting the brain ventricle
- Adjust the magnification of the microscope so that the brain ventricular structure of zebrafish is clearly displayed in the field of vision (Figure 1B).
- Place the needle carefully above the brain tectum (Figure 1C).
- Puncture the skin of the zebrafish brain with the needle tip slowly using the micromanipulator.
- Press the foot pedal to eject 1 nL of 1x PBS or different concentrations of LPS (1.0, 2.5, and 5.0 mg/mL) (see Table of Materials). Successful ventricle injection images at which the brain ventricle was injected with 1 nL of 1% Evans blue dye (diluted in PBS) are shown as an example (Figure 1D). Transfer the larvae to clean egg water (E3) medium immediately after the injection. After 24 h, collect the larvae for microscopic imaging, locomotive behavioral assay, and determination of other indicators.
