The goal of this technology is to provide histological tools that integrate complex model systems in a high-throughput, observer independent, and fully digital manner that is both cost-effective and meaningful to musculoskeletal biologists. Estology, while critical to most biological researchers, is often labor-intensive and non-quantitative. Therefore, there is a need for histological techniques that provide efficient, high-throughput, and automated analysis.
With our cryohistology platform, the researcher can associate multiple molecro signals to specific cells within a given specimen by imaging and staining the same slide multiple times in high-throughput manner. Demonstrating the procedure will be Doctor Xi Jiang and Li Chen, research assistants in my laboratory, who have pioneered the development and the execution of these techniques. To begin this procedure, remove the specimen from formalin, dissect away any excess tissue, and transfer to 30%sucrose, made in 1x PBS.
Incubate the specimens in sucrose for 12 to 24 hours at four degrees Celsius. Next, fill a cryomold with cryoembedding medium and place the specimens in the mold. Then, align the samples in the mold such that the cutting plain for the region of interest is parallel to the bottom of the mold.
After that, place the cryomold onto a piece of dry ice until a thin layer of cryoembedding medium freezes, locking the spicimens into place. Then, fill the remaining volume of the cryomold with cryoembedding medium, while maintaining the cryomold on the dry ice pellet. Following that, place the cryomold in a container containing 2-methylbutane pre-chilled by dry ice.
Once the specimens are completely frozen, remove the cryomold and shake off excess 2-methylbutane. Wrap the cryomold in cellophane and store it at minus 20 degrees Celsius, or minus 80 degrees Celsius. In this procedure, transfer the specimen block from the freezer to a cryostat set at minus 20 to minus 25 degrees Celsius.
Next, remove the block from the cryomold and trim away excess cryoembedding medium with a razor blade. Then, add some cryoembedding medium onto the specimen disk, and align the block such that the surface of the specimen disk is parallel to the bottom surface of the block. Wait for the cryoembedding medium to freeze.
Afterward, place the specimen disk onto the specimen head and adjust the head such that the blade cuts parallel to the surface of the specimen block. Next, cut pieces of cryotape to cover the region of interest and pre-chill them in the cryostat. Afterward, trim the block down to a level within the region of interest, continuing to make adjustments as needed.
Once the region of interest has been reached, and a section can be made, brush off any shavings from the surface of the block. Next, remove the non-adherent backing from the cryotape by grasping the tape by the non-sticky silver gold tabs. Then place the tape onto the block with the sticky side down.
Apply pressure to the cryotape using the roller, and make a section using either the automatic motor or manual cutting wheel of the cryostat. Following this, place the section tissue side up on a plastic microscope slide within the cryostat. Remove the plastic slide from the cryostat and allow the cryoembedding medium to melt, such that the section adheres to the slide's surface.
Repeat sectioning for the serial sections or other regions of interest. Once finished, place the sections in a slide box and store them at four, minus 20, or minus 80 degrees Celsius, depending on the length of storage required and the downstream response measures. To perform the UV curable adhesive method, first label the microscope slide.
Apply a drop of UV activated optical adhesive to two glass slides, and use the edge of a slide to spread a thin layer of adhesive across the surface of the glass slides. Then, cut off the silver gold tab and lay the taped section tissue side up, on the adhesive layer of the first slide. Apply the section in a rolling motion from one edge to the other in order to minimize the formation of entrapped bubbles underneath the tape.
Afterward, remove the taped section from the first slide, then place it on the adhesive layer of the second slide, and take care to avoid entrapment of bubbles. Once the appropriate number of sections for the given experiment is placed on each slide, wipe off the excess adhesive from the surrounding areas. Next, inspect the sections closely for bubbles or fibers underneath the tape.
Then place the microscope slides under the UV black light for five to ten minutes to cross-link the adhesive layer. In this step, prepare the reference marker solution by dissolving 50 microliters of green, and 50 microliters of red microspheres in 100 microliters of water. Store the solution in the dark at four degrees Celsius.
Then dip a pipette tip into the microsphere solution, and dab the microsphere solution onto each section, adjacent to the region of interest. Next, dry the slides at room temperature and protect them from light by covering them with foil for 30 minutes, if processing the slides on that day. If not, place the slides in a slide box at four degrees Celsius.
After staining the sections, mount the cover slips on the slides using acquiesce mounting medium. To perform imaging, load the slides into the microscope trays and insert the trays into the tray stack of the slide scanning microscope. Then, click on the profile list in the microscope software to load a profile for each slide with the appropriate exposure time for each flurophore.
The, click on the Start Preview Scan button to take a preview image of each slide. The software will apply a file name automatically by reading the bar codes placed on the slide labels. Alternatively, a file name can be manually assigned to each slide.
Set the regions of interest by entering the tissue detection wizard, drawing each region using the draw tool, making any necessary adjustments, and saving them for subsequent rounds of imaging. Once each slide is set up, start the scan process by clicking the Start Scan button. Following each round of imaging, submerge the slides in a coplin jar filled with one 1x PBS until the cover slips fall off from the slides, before the next staining step.
After all of the imaging rounds have been completed, export the images from each individual channel as TIF or JPG files for image assembly and analysis. In this figure, a section from a three month old mouse femur was stained with calcein blue and imaged for accumulated mineral, seen in white, and mineralization labels, seen in green and red in the first round of imaging. Slides were then imaged for trap activity, seen in yellow in the second round, and AP activity, seen in red in the third round of imaging.
Finally, the slides were stained with toluidine blue in the fourth round. The reference markers were imaged during every imaging round including the chromogenic round, and were aligned using the custom software. Once mastered, this technique can be done in less time than it takes to decalcify a typical specimen.
After watching this video, you should have a good understanding of how to produce high quality high content images of mineralized tissues, which we hope can be more broadly used by the musculoskeletal community.
