Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

This method will enable you to generate three-dimensional histological images from small millimeter scale samples, including small model organisms. The main advantage of this technique is that the embedded samples are very stable, enabling you to keep the samples for long periods of time and to image them multiple times, say with different imaging methods over years. The output of this method will be highly useful for high throughput model system phenomics, toxicology, and potentially even human tissue diagnostics.

Generally individuals new to this method will struggle because the samples are relatively small and difficult to orient or maneuver. Visual demonstration is critical because attention is required to avoid sample damage. For juvenile or older zebrafish, starve them for at least 24 hours in order to reduce the volume of gut content prior to fixation.

Move 10%neutral buffer formalin or NBF and 2x Tricaine-S pre-chilled to four degrees celsius onto ice. Double the fish water volume with chilled 2x MS-222 for rapid and humane euthanasia. Sixty seconds after the fish have stopped moving, after humane euthanasia, place the fish in chilled 10%NBF solution in flat-bottomed glass vials at room temperature, and fix them overnight.

On the second day, rinse the fixed fish specimens three times in 1x PBS pH 7.4 for 10 minutes each. After washing, submerge the samples in 35%ethanol and incubate for 20 minutes at room temperature with gentle agitation. Then repeat this step with 50%ethanol.

Incubate the samples in freshly prepared 0.3%phosphotungsten acid or PTA solution overnight at room temperature with gentle agitation to stain them. Start the third day by preparing a one-to-one volume-by-volume mixture of 100%ethanol and LR white acrylic resin. Rinse the samples three times in 70%ethanol for 10 minutes each, then incubate the samples in 90%ethanol at room temperature for 30 minutes with gentle agitation.

Repeat the incubation under the same conditions but 95%ethanol and then two more times with 100%ethanol. At the end, submerge the samples in one-to-one ethanol and LR white acrylic resin mixture, and incubate at room temperature overnight with gentle agitation. On the fourth day, remove the mixture from the vial, and add 100%LR white resin to submerge the samples.

Incubate for two hours at room temperature with gentle agitation. After two hours, replace the resin with fresh 100%LR white resin, and incubate for one hour at room temperature with gentle agitation. To assemble the embedding apparatus, use the poly MI tubing with a diameter of at least 0.1 millimeters larger than the diameter of the specimen, and cut it to a standard length of 30 millimeters.

Then attach a P1000 micro-pipe head to the white end of the embedding adapter and insert the poly MI tubing to its narrow end. After the samples have incubated for one hour in 100%LR white resin, transfer them to a small weigh boat and fully submerge them in 100%LR white resin. Aim the tubing of the embedding apparatus at the wide end of the fixed sample, or towards the head, and pipet the specimen slowly into the tubing.

Position the sample in the middle of the tubing, making sure that the tubing above and below the specimen is filled with resin. Immediately after, to seal the open end of the tubing, flatten a piece of oil-based soft modeling clay into a sheet about one millimeter thick and stabilize the tubing between index and middle fingers. Then slowly press the clay against the end of the tubing with thumb, and remove any excess clay.

Remove the embedding apparatus from the micro-pipe head. Pull out the tubing by gentle rotation. While holding a finger against the sealed end to prevent the ejection of the clay, seal the other end by slowly pushing the unsealed end of the tubing into the clay.

Place the tubing horizontally on a tube rack, and allow the resin to polymerize for 24 hours at 65 degrees celsius in the oven. At the end of the following day, collect the samples for image acquisition. A successfully embedded zebrafish larva has no air bubbles trapped.

If air is trapped during the embedding process, it can move toward the specimen if the sample is not placed horizontally during the polymerization, which can degrade the image quality. This protocol was successful in embedding various model organisms. Such as zebrafish, Drosophila, Daphnia, and mouse embryo.

The successful embedding can be shown with reconstructed, 3D renderings, imaged by micro-CT scan for zebrafish, Daphnia, and Drosophila. Embedding resin can be seen in all scans outside of the sample, but does not interfere with the sample itself. These specimens can be stored for an extended period of time and reused for multiple imaging sessions, as shown with zebrafish larva imaged in 2011.

And then again in 2013. Anatomical features are preserved after the storage, as seen on close-ups of muscles from both scans. Furthermore, intensity profiles normalized to their corresponding averages along segmented lines, shows spatially matching local peaks in both scans.

Average intensity values for selected regions in both scans were divided by an average for background and used to generate signal-to-noise ratios, which corresponded well between the scans. While attempting this method, it's important to work gently to avoid sample damage and to work carefully to prevent introduction of air bubbles. Following this procedure, other methods such as histology or electron microscopy can be used in order to study patterns of protein expression, or achieve higher resolution of regions of interest in 2D.

Our hope is that this publication will help you to access the power of high-resolution tissue micro-CT.