To cut axons using In vivo two photon imaging position anesthetized zebrafish embryos within a drop of aros as it hardens in a specialized chamber consisting of a TEFL laundering and cover slip. A glass light is placed on top of the chamber, which is then flipped upside down prior to the ex otomy. Take a before image of the axon you wish to cut at 40 x zoom into 70 x on the region you wish to exo mize.
And with the laser off, turn up the laser power. Briefly turn on the laser. In the after image of the axon, you will see scattered axonal debris if the ex otomy is successful.
Hi, I'm Georgia O'Brien. Hi, I'm Sandra Rigo. And I'm Shauna Martin.
And we're from the laboratory of Alvarez TI in the Department of Molecular Cell and developmental biology at the University of California Los Angeles. Today we'll show you a procedure for two photon ex sodomy and time lapse confocal imaging in live zebrafish embryos. We use this procedure in our laboratory to study degeneration and regeneration of peripheral somatosensory axons.
So let's get started. To begin, we'll prepare a 1%low melt agro solution for embedding. We dissolve the agros in double distal water by heating in a microwave before Alec watering into small tubes and storing in a heating block at 42 degrees Celsius.
First embryos can be placed into a 5%PTU ringer solution at 22 to 24 hours post fertilization to inhibit the formation of pigment, which is brightly, autofluorescent, and will obscure axons on the two photon microscope. Next, we select embryos for imaging and remove their choon by gently pulling them apart with forceps. Once the embryos are selected, they're anesthetize by adding about 0.02%trica to the PTU ringers.
Before proceeding, make sure animals are not responsive to touch. Prepare the long-term imaging chamber by applying vacuum grease to one side of a glass or Teflon ring and fixing it onto a glass cover slip. Taking care not to transfer much of the PTU ringers.
Transfer one embryo into the 42 degree agro solution with a glass pipette. Then transfer the embryo with one drop of agros onto a prepared cover slip. Once transferred, position the embryo as desired while the agros hardens.
Keep in mind that the imaging chamber will be flipped when you're done so that the copper slip will be the top surface. If you have a motorized stage and can image multiple locations at once, repeat these steps for each embryo. Allow the agro roses to completely solidify.
Then fill the ring with 0.02%tri canine PTU ringers. Finally, apply vacuum grease to the other side of the rings. Then cover them with a glass light.
Flip the seal chambers over. Now we are ready for two four ton exo tomy imaging. To prepare for imaging, place the mounted embryos on a slide holder under the microscope.
Focus on one embryo with a 40 x 0.8 numerical aperture water objective. Then turn on the laser. Set the laser to a wavelength of 910 nanometers and a power of 30 milliwatts.
Open the imaging software. We use scan image software developed in corals for Boar's Laboratory. At this power, one can scan the field for a candidate axon that will be severed.
Once found, take an image of the target axon. Mark the first and last Z positions, acquire the image and make a maximum projection of the Z stacks. Next zoom in 70 x on the branch of the axon you wish to ex itemize.
Turn up the power to 180 milliwatts and set the number of Z slices to one. Initiate a single scan with a laser by pressing grab. This should be sufficient to sever the axon, however it is necessary to optimize this procedure for your microscope and experimental goal.
Finally, zoom out. Reduce the power to 30 milliwatts and take an image of our newly severed axon. If a custom built two photon scope is not available in your laboratory, the Zes five 10 confocal two photon microscope can also be used to sever axons.
We begin by placing the mounted embryo onto the stage and bringing it into focus using a 25 x water objective. Next, turn on the two photon in Argonne lasers in a multi-track setting so that it is possible to switch from one to the other. Although both lasers are used to detect GFP, the two photon emission is visualized with red and the argonne laser emission with green.
In order to differentiate the two, use the Argonne laser to identify an axon to endure under Z settings. Mark the first and last optical sections. Take a confocal image and create a maximum projection of the Zack.
Turn off the argon laser and turn on the two photon laser scan at an intensity of about 9%transmission. To make sure that the axon is still in focus, click the stop button so that the crop tool will be available. Use crop to zoom in on the area of interest.
Usually we zoom to about 70 x. Choose the region of the axon to be injured and bring this region into focus. The zoom can be checked under the mode tab.
Next, under the channels tab, change the intensity of the two photon from about 9%transmission to 15 to 30%transmission. To activate the new settings, click on the fast XY button and then click stop quickly afterward. To avoid excess damage, the axon should be seen as scattered debris if the procedure worked.
Finally, to ensure that the axon was indeed damaged, switch back to the 4 88 nanometer argonne laser. Take another confocal image and create a maximum projection of the Zacks. Now that we've shown you two ways to sever axons using two photon laser microscopy, we'll look at confocal TimeLapse imaging of regeneration using the zes LSM five 10.
To prepare time lapse imaging, be sure to turn on your heated stage at least 30 minutes before setting up your time-lapse movie. Place the mounted embryos on a slide holder under the microscope. Focus on one embryo with the desired air objective.
We use a 20 x 0.5 numerical aperture Objective, open the Zeiss LSM five 10 imaging software and then turn on the appropriate lasers and set up the desired configuration. Open the scan stage and multi-time windows. If you have multi-time software, define the position and configuration of your first embryo in the multi-time window.
Activate fast scan on the scan window, then move the stage to the desired XY position. Mark the first and last Z positions in the scan window. Press stop and then mid also in the scan window, wait for the scan of the middle Z slice to finish.
Then press mark position in the stage window. If you are only imaging one embryo, click on the single location tab in the multi-time window. Click on replace X, y, Z.Then click on save configuration.
Agree when asked to override the previous configuration. Now you can set up the parameters for image acquisition in the multi lo window. If you have a mechanized stage, you can set up multiple locations to image multiple embryos.
In this case, you should select the multiple locations tab before defining the X, Y, Z and configuration for your first location. Also, be sure that the pull down menu just below the multiple locations tab is on location one and that the pull down menu in the configuration section says multi lo one. Before you click on save configuration, define the position and configuration of your remaining embryos in the multi-time window.
Locate your next embryo press fast scan, then move the stage into the desired XY position and mark your first and last Z positions in the scan window. Press stop. Then mid also in the scan window, wait for the scan of the middle Z slice to finish.
Then press mark position In the stage window, click on replace X, Y, Z.Check that the pull-down menu just below the multiple locations tab is on location two and that the pull down menu in the configuration section says multi lobe two. Then click on save configuration.Agree. When asked to overwrite the previous configuration, repeat this process for the remaining embryos.
Set up the parameters for image acquisition in the multi lo window. Choose the GR GL option in the list of block section. Then enter the desired number of group repetitions.
This is the number of times the confocal will acquire an image at each location. Enter the desired weight interval in the parameter section. This is the amount of time from the start of one imaging repetition to the start of the next.
Select Zack XY in the configuration section. Enter your base file name in the bottom section. Click on select image db and choose your database.
Click on the options button, then select your MDB folder to save temp files too. Check keep final image open. Save final image, middle of the Zack and select weight interval.
Click okay to close the options window, then click on start time. Leave the multi-time window open for the duration of imaging. Now it's time to analyze your data.
When the time-lapse imaging has finished. The multi-time software will compile all of your temporary files into a summary file for each location. If you want to stop the movie before it is complete, be sure to press finish instead of stop so that a summary file will be created.
The temp files and summary files should be automatically saved to the designated MDB folder. Here is a representative example of a time lapse confocal movie following ex otomy at 54 hours post fertilization. The first frame shows a dorsal view of the axon before ex otomy with a yellow arrow pointing to the ex otomy site the following time.
Lapse movie begins within one hour after ex otomy and lasts 12 hours. One confocal stack was collected every 20 minutes. This lapse confocal movie is recorded using the same parameters at 78 hours post fertilization.
The first frame shows a lateral view of the axon before ex sodomy with a yellow arrow pointing to the ex sodomy site. We've just shown you how to generate precise tissue damage and visualize live zebrafish embryos using two photon sodomy and time-lapse confocal imaging. When doing this procedure, it is important to optimize the amount of power used for two photon laser ex sodomy to avoid causing excess damage.
So that's it. Thanks for watching and good luck with your Experiment.
