电机神经切断和时间推移成像胶质细胞行为在现场斑马鱼

The overall goal of this procedure is to time-lapse image glial cell behavior following peripheral nerve injury in live zebrafish larvae. This is accomplished by first mounting anesthetized zebrafish larvae in low melting point aros on glass bottomed petri dishes and positioning them with a dissecting needle. Next, the mounted larvae is placed on the confocal microscope.

A nerve is selected for ablation and an image acquired of the axons and glial cells of the uninjured nerve. Then ROIs are selected along the nerve and the laser is fired to ablate the selected areas creating a nerve transection. Finally, the axons and glial cells are time-lapse imaged following the nerve transection.

Ultimately, results can be obtained that show dynamic glial cell behavior during nerve degeneration and regeneration through time-lapse confocal microscopy. This method can help answer key questions in the fields of regeneration and glial cell biology, such as how do glial cells respond to nerve injury? And how do distinct glial subtypes coordinate their behavior during degeneration and regeneration?

After crossing adult zebrafish containing transgenes that fluorescently label motor neurons and glial cell types of interest and incubating the embryos to 24 hours post fertilization or HPF, remove the egg water and replace it with one pH tooth threa or PTU in egg water before returning the embryos to the incubator. From this time point until about 96 HPF, use a fluorescent dissecting scope to screen embryos for the presence of the desired fluorescent gene. Transfer positive embryos into fresh PTU egg water and return them to the incubator.

When the larvae reached six days post fertilization or DPF, select a few embryos for mounting and transfer them to a smaller dish. Remove the water from the dish and immediately replace it with 0.02%trica In PTU egg water, incubate the larvae in anesthetic approximately five minutes. Place an Eloqua of 0.8%low melt agros previously prepared according to the text protocol into a beaker with tap water and a microwave.

For 30 seconds or until the agros is melted, allow the beaker to cool until the tube of agros feels lukewarm to the touch. Next, transfer an anesthetized larvae to a 35 millimeter glass bottom dish. Remove any water in the dish.

Then immediately cover the larvae with enough warm aros to fill the glass bottomed portion of the dish. As the agros hardens, use a dissecting needle to position the larvae on its side. Then tilt it slightly on its back, making sure that the mounted larvae is touching the glass on the bottom of the dish, which is necessary for injury and imaging.

Use the needle to maintain the larvae in position until the agro solidifies and the larvae is immobilized. Then slowly pipette enough trica water into the dish to completely cover the agros and larvae. Turn on all confocal microscope instrumentation, appropriate diode lasers for exciting zebrafish transgenes, and the nitrogen pumped dye laser.

Be sure the blank beam splitter is in place and the 435 nanometer coer and dye cell is in place. Open the laser attenuator fully and then open the imaging software with the 63 x 1.2 na water immersion lens and a glass slide with one mirrored side. Use brightfield illumination to find and focus on a scratch or etch in the mirror.

Using imaging software view and focus the etchings on the computer screen from the window that controls the laser calibration and power settings. Set the number of pulses to one and the attenuation to 3%transmission from the main toolbar. Select the ellipse tool and click the image on the computer screen.

To create a single circular ROI repeat the process three more times until there are four circular RI spaced randomly over the image. Next, fire the laser. If the laser is working and calibrated properly, this will create four small spot etchings, one within each circular ROI Then remove the glass slide from the microscope stage and clean the objective.

Replace the blank beam splitter with the 100%ILL beam splitter to transect a motor nerve using laser ablation. Apply a small drop of water immersion medium to the objective and place the dish with mounted larva on the appropriate stage using clips to stabilize it using the eyepiece and widefield illumination. Focus on the larvae and locate the motor neurons.

Scan the nerves in hemi segments 10 to 20 and select a motor nerve for transection. Next, bring up a live view of the nerve on the computer screen. Select Z planes and acquire an image of the axons and glial cells of the uninjured nerve.

Then prepare time-lapse imaging settings to capture Z projections of all cell types in five to 30 minute intervals. Depending on the experiment, remove the 100%ILL beam splitter and replace with the blank. Return to the live view of the nerve and use the appropriate fluorescent channel to view the axons that will be transected.

Using the ellipse tool, create a thin elliptical ROI in the area to be ablated. Then create smaller ROIs within the selected region in the window that controls the laser settings. Set the number of pulses to two and the attenuation plate to 18%transmission.

Then fire the laser within the selected ROIs. Wait 10 or more seconds and check the ablated area again for fluorescence. Be aware that an ROI may initially appear ablated when it is actually photo bleached and the ROIs may need to be modified slightly during the course of ablation to achieve a complete transection.

If fluorescence returns, increase the laser power and fire the laser. Again, repeat this until fluorescence disappears within the R OIS and does not return within 10 seconds. Once the ablation is completed, begin time-lapse imaging when the time-lapse is complete.

Use imaging software to compile data and create color composite Z projections for each time point. Create a quick time movie to analyze the behavior of axons glial cells simultaneously. The assay described here can be used to assess the response of glial cells and other nerve associated cell populations to axonal injury in vivo.

Shown here is an example of a nerve injury created using this method and the response of surrounding glial cells. The experiment was performed in six DPF transgenic zebra fish that expressed a membrane targeted EGFP in perineural ggl, and cytosolic DS red in motor neurons. The injury was made along the rostral projection of a trunk motor nerve that was then time-lapse imaged in both the EGFP and DS red channels.

This allowed simultaneous visualization of axon and glial cell behaviors immediately following the injury. These images are static time points taken from the time-lapse movie. The dotted ellipse shows the ROI that was ablated using the laser at one minute.

Post transection or MPT, the ablated area lacked fluorescence and the injury zone measured approximately 3.5 micrometers from the proximal to distal stump. The success of a transection can be confirmed by imaging the distal nerve stump and looking for signs of wallerian degeneration, including distal axon fragmentation and rapid clearance. The absence of axonal fluorescence along the distal stump at 120 MPT indicates these axons have indeed undergone wallerian degeneration and the transaction was successful.

Adjusting the laser power to an ideal setting is critical when performing laser ablation experiments. Ideal laser power settings will cleanly ablate the nerve only within the selected ROI and laser power settings that are either too low or too high will yield suboptimal results seen here is an injury that was performed with the laser power that was too low. Fluorescence remained within the ROI after firing the laser resulting in an incomplete transection.

On the other hand, as demonstrated in this figure, a laser power that was too high produced an extremely large ablation. After watching this video, you should have a good understanding of how to mount zebrafish for laser transection experiments. Create a transection using a nitrogen pump dye laser, and assess the response of surrounding cells using time-lapse confocal imaging.