The overall goal of this procedure is to reconstruct and visualize an entire mouse brain with micron scale resolution. This is accomplished by first dehydrating, a fixed mouse brain integrated tetrahedran series. The second step is to clear the sample in Benzyl Ether.
Next, the brain is imaged with a confocal light sheet microscope. The final step is to use software to align and stitch together the acquired image stacks. Ultimately, a multi resolution visualization tool can be used to navigate the reconstructed image volume.
The main advantage of this technique of other methods like confocal or to photo microscopy is that the complete volume of a mouse brain can be image in just a few days. This method can help answer key questions in the neuroscience field, such as the distribution of different neuronal types throughout the whole brain. Does This metal cap provide insight into fine brainin autonomy?
It can be applied also to mouse embryos or foot flies, demonstrating the procedure will be reini. A grad student from my lab. To begin this protocol perform standard fixing of mouse brains via trans cardio perfusion and postfix overnight before performing dehydration and clearing.
Then since the dehydration solvent, THF strongly quenches XFP fluorescence perform filtration to remove peroxides with basic activated aluminum oxide using a chromatography column, avoid cracking of the column, which would reduce peroxide removal at a stabilizer to the final solution. Even if the THF was already stabilized before filtration, the stabilizer will be retained by the aluminum oxide. Note that THF without stabilizer may develop large amounts of peroxides and can be explosive and dangerous.
Next, dehydrate the whole mouse brain integrated series of THF in pure water. Then place the vial with the sample on a rotating wheel to avoid light exposure. Wrap the vial with an aluminum foil.
Finally, filter the clearing solvent with aluminum oxide using a filtering funnel. Then clear the sample in 100%DBE, changing the solution after three and six hours when the brain appears transparent. Typically one hour after the second DBE change mount it on a tipped imaging plate by one of the methods seen here.
Direct mounting mounting via agros disc or mounting via an agro speaker for direct mounting, gently plunge the sample onto one of the tips to use the aros disc plunge. A disc made of 6%aros gel onto the tips. Then gently fix the sample on the agros disc with acrylic glue and wait about one minute for curing to mount.
Using the agros beaker plunge, a beaker made of 3%agros gel onto the tips. Then gently placed the sample onto the bottom of the beaker. After mounting the specimen following one of these methods, position the imaging plate inside the specimen chamber using tweezers.
Then fill the chamber with clearing solution. Next, prepare for tomography. First, remove the slit to collect as much fluorescence as possible, and then illuminate the sample with low laser power.
To prevent photobleaching, move the sample in the three x, y, and Z directions using the software driven micro positioning system. For each direction, identify the minimum and maximum coordinates for which the sample is illuminated while within the field of view of the camera. Then insert the determined coordinates as parameters for the pre tomography subroutine.
Also specify the distance between adjacent stacks to be used in the tomography and the pre tomography sampling step in the Z direction. Finally, start the pre tomography, which will collect the images in every stack with the Z sampling parameters as specified in the previous step. To begin acquisition, first, move the sample outside of the light sheet using the micro positioning system.
Then increase the laser power and stop motion of all the scanning systems in order to illuminate only a fixed line at the center of the field of view. Next, mount the slit and align it using the autofluorescent signal from the clearing solution. You should now clearly see the slit line in the center of the field of view.
Now reactivate the scanning system. Reduce laser power and move the sample inside the light sheet using the micro positioning system. Note that the laser power used with the slit will be higher than that used without it.
Since the spatial filter blocks a non negligible fraction of light, adjust the amplitude and offset of the scanning and d scanning systems with the control software until the images look clear and bright. Then set the appropriate Z step for the experiment and run the tomography acquisition software. The volume will be imaged in many parallel, partially overlapping stacks, and the images will be saved in a hierarchical folder structure.
In order for the stitching software to work with a complete cubic volume, complete the acquired volume with synthetic black images, meaning images of the same type and dimensions of the collected ones, but with zero intensity using automated software. Next, launch the VA A 3D software with plugins, Terra, Stitcher, and tarly installed and load the Terra Stitcher plugin. Select the directory containing the imaged volume and indicate the relative orientations of the axes with respect to a reference right-handed coordinate system and the voxel size.
Now launch the first part of the stitching. The software will calculate the relative displacement between pairs of stacks and find an overall optimal placement for all the stacks together select to save the stitched volume in single resolution or in multi resolution format. The ladder enables for multi resolution visualization with the tarly plugin.
In both cases, if the higher resolution image is larger than a few gigabytes, also select the multi-stack save modality and specify the size of individual sub stacks. This will enable efficient access to the stored data. Now launch the second part of the stitching.
The software will merge the aligned stacks and save them in either single or multi-stack mode. When completed, close the Terra Stitcher plugin. Next, load the terra fly plugin.
Select the folder with the multi-stack multi resolution volume and indicate foxhole size and axis orientations. The volume will be loaded at the minimum resolution. To zoom to a higher resolution, simply use the mouse scroll.
Alternatively, one can double click on a specific point of the image or select to zoom in around an existing landmark or directly specify the coordinates of the volume of interest. To zoom back to lower resolution, simply use the mouse scroll. The described protocol can be used to reconstruct with micron scale resolution either entire mouse brains or excised parts without the need for physical sectioning.
As a representative result, the whole cerebellum of an L seven GFP mouse postnatal day 10 is shown here in this animal. All Perkin gene neurons are labeled with EGFP. After zooming in the typical lamellar structure of the cerebellar cortex can be seen further zooming in allows clearly distinguishing each burkini cell soma.
The described protocol can thus be used to screen neuronal spatial organization in various neurodevelopment studies. As a second representative result, here we can see images from the UN sectioned brain of an adult thigh, one GFPM mouse. In this transgenic animal, EGFP is expressed in a random sparse neuronal subset.
The right half of the brain is shown here. As we zoom in further and further neuronal processes become distinguishable. This result demonstrates that the described protocol allows micron scale resolution imaging in entire adult mouse brains opening the possibility of studying whole brain anatomy at cellular resolution.
In mouse models of neurodegenerative disease Once mastered, this technique can be done in three to four days if it's performed properly. While attempting this procedure, it's important to remember to properly filter the dehydration and cleaning solution in order to preserve GFP fluorescence. The multi resolution visualization tool presented here can also be used to explore very large data sets obtained with other techniques such as micro optical section in tomography.
After watching this video, you should have a good understanding of how to date, rate and clear fixed mouse brain. Image them with confocal sheet microscopy and visualize the reconstructed volume with the VR 3D using the Tely plugin.
