Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

The overall goal of this protocol is to demonstrate modifications to the original CLARITY protocol, that yield more consistent and cost effective results. This method can help answer key questions in neuroscience, such as 3D morphology, and connectivity in the central nervous system. The main advantage of this technique is that it improves the consistency of optical clearing and microscopy in the mouse central nervous system.

In a cost effective and reproduceable manner. Visual demonstration of this method is critical. As the imaging chamber construction is complicated and not readily obvious from written description.

Obtain paraformaldehyde fixed brain and spinal cord tissue from mice expressing fluorescent proteins. Hemisect the brain by placing it in a plastic mouse brain matrix and cutting along the midline with a matrix blade. Add each hemisphere and spinal cord to separate 50 milliliter centrifuge tubes.

Each containing 40 milliliters of hydrogel solution. Cover the tubes with aluminum foil to protect the fluorophores. And incubate the tubes containing samples and hydrogel at four degrees celsius with gentle shaking for seven days.

Following the incubation period discard 25 milliliters of hydrogel leaving the sample and approximately 25 milliliters of hydrogel in the centrifuge tube. Next deoxygenate the sample by removing the cap, placing the centrifuge tube in a desiccator jar, and connecting the jar to a vacuum. Apply the vacuum for at least 10 minutes.

Gently shake to dislodge the bubbles of emitted oxygen. After 10 minutes, replace the vacuum in the desiccator jar with 100%nitrogen gas over two to three minutes. Once the pressure equalizes and the top of the desiccator jar can be opened, quickly cap the tube to prevent the reintroduction of oxygen.

Next polymerize the hydrogel by placing the tube with sample in a 37 degrees celsius water bath for three hours until polymerization is complete. After three hours the polymerized hydrogel will take on a gel like consistency. Use a spatula to remover the polymerized sample from the centrifuge tube then cut away the excess hydrogel from the sample.

Lastly, remove the remaining hydrogel by placing the sample on a lint free wipe, and gently rubbing and rolling the tissue on it, leaving behind only a thin layer of polymerized hydrogel on the sample. To begin the process of lipid removal, transfer the polymerized sample to a clean 50 milliliter centrifuge tube with 45 milliliters of clearing buffer, and incubate at 45 degrees celsius with gentle shaking for seven days. Change the buffer daily during this initial seven day period by pouring off the used clearing buffer into a chemical waste container.

And adding 45 milliliters of fresh clearing buffer. Once all lipids are solubilized and the tissue reaches transparency transfer the sample to a new 50 milliliter centrifuge tube filled with 45 milliliters of PBST, and wash four times at 40 degrees celsius with gentle shaking over the next 24 hours, to remove residual SDS. After the final PBST wash, transfer the sample to a clean 15 milliliter centrifuge tube filled with one microgram per milliliter DAPI and 1xPBST.

Wrap with foil and incubate at room temperature for 24 hours. The next day wash three times in PBST at room temperature for at least six hours per wash. And then proceed to refractive index matching.

To begin this part of the procedure, transfer the sample to a clean 15 milliliter centrifuge tube filled with 2-2 thiodiethanol and 1xPBS at PH 7.5 and incubate as shown on screen. Towards the end of the second incubation, create and imaging chamber by rolling a piece of reusable adhesive into a cylinder with uniform diameter just slightly greater than the thickness of the sample. Lay the cylinder in an open ended circle on a 40 millimeter glass bottom dish.

Then use a P1000 pipette tip to create a seal between the reusable adhesive and glass by rolling it around the outer rim of the cylinder. Pipette approximately 100 microliters of 63%TDE onto the glass dish and spread it around to cover the glass surface within the circle of reusable adhesive. Then use a spatula to carefully place the sample in the middle of the circle taking care not to form bubbles.

Next, pipette another 100 microliters of 63%TDE directly onto the sample, and immediately place a 40 millimeter circular cover glass over the reusable adhesive. Use the index, middle, and ring finger of both hands to carefully press around the perimeter of the circle until the cover glass makes contact with the sample. Now, slowly bring the glass bottom dish to an upright position resting against a surface, then add 63%TDE with a pipette until the solution reaches the small opening at the top.

Use a lint free wipe to dry off the pipette tip so that the solution does not drip on the glass. Lastly, add silicon elastomer to the small opening to enclose the circle and create a closed chamber. Wait five minutes to dry, and then proceed to imaging.

Place the imaging chamber with the sample inside on the stage of a laser scanning confocal microscope. Open the imaging software, to turn on the argon excitation laser, click the configuration tab at the top of the program, then the laser icon. Check the box next to argon to power the laser and move the slide scale to 30%Return to the acquire tab at the top.

In the beam path settings box, go to the load-save setting box, and select the drop down menu to select the appropriate wavelength channel to emit YFP. In the channel settings box, select the appropriate objectives for imaging by clicking the red hyperlink labeled objective current object. Use objectives with a working distance greater than the thickness of the tissue.

And a large numerical aperture to maximize the inplane image resolution, and to minimize optical section thickness. On the left column of drop down menus open the XY resolution window to set the acquisition matrix, called format to 1024x1024 or a value appropriate for the lateral resolution limit of the objective. Set the zoom factor as low as possible.

1.7 is used here. In the same window set eighth line average and one frame average parameters by dropping down the individually labeled menus accordingly. Locate the sample using the stage controls.

Once a representative field is visible, set the gain to between 760 to 780, and the offset to between 1.0 and 1.5 for YFP. Select tile scan. And then set the upper and lower limits of the Z stack to be acquired.

Set the thickness of the optical section based on the objectives optical section resolution limit and then start the acquisition. This image shows YFP positive neurons in the cerebral cortex and hippocampus imaged as 5x magnification and reconstructed in 3D. The scale bar represents one millimeter.

This maximum intensity projection of a 10x image stack of the cerebral cortex demonsrates the dendritic aborization of cortical layer five pyramidal neurons. Here the scale bar represents 100 microns. Here THY-1YFP expression is shown in an intact cervical and thoracic spinal cord, imaged at 5x magnification, and reconstructed in 3D.

The scale bar represents two millimeters. Finally this maximum intensity projection of a 10x imaged stack of the spinal cord, demonstrates individual axons, scale bars represent 100 microns. If preformed properly this technique can be done in two to four weeks for whole spinal cords, and four to six weeks for hemisected brains.

Following this procedure, other methods such as immunohistic chemistry can be preformed in order to answer questions that require more complex biochemical phenotyping. After watching this video you should have a good understanding of how to optically clear and image a mouse central nervous system. Using passive clarity, and laser confocal microscopy.