Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Here, we describe computational tools and methods that allow visualization and analysis of three and four-dimensional image data of mouse embryos in the context of axial elongation and segmentation, obtained by in toto optical projection tomography, and by live imaging and whole-mount immunofluorescence staining using multiphoton microscopy.

The organogenesis stages in the mouse embryos remain poorly studied partially due to the difficult visualization of the embryo complex structures. This protocol describes techniques that allow 3D and 4D visualization and analysis of mouse embryos during axial extension and segmentation. The main advantage of this technique is that it allow researchers to study and show the complex structures of the mouse embryo as 3D dynamic objects.

This protocol contains techniques that can be used in the study of congenital malformations affecting the vertebra and the spinal cord of mammals, such as scoliosis. These techniques can also be used with in vitro model systems, thus allowing the study of human embryonic development. To prepare mouse embryos between embryonic day 8.25 and embryonic day 10.5 for live imaging, dissect the embryos in pre-warmed M2 medium at 37 degrees Celsius.

Gently remove the yolk sac using clean forceps and wash the embryo once with fresh M2 medium to remove blood and debris produced during dissection. During the live imaging procedure, incubate embryos in low glucose DMEM medium supplemented with 10%HyClone defined FBS, two millimolar L-glutamine, and 1%penicillin streptomycin in a 37 degree Celsius heated chamber and a 65%oxygen with 5%carbon dioxide environment. To prepare similar stage mouse embryos for immunofluorescence microscopy, instead of using heated M2, dissect the embryos in cold PBS and fix in 4%paraformaldehyde.

After performing the whole mount immunofluorescence staining protocol as described in the text manuscript, tissue clearance can be achieved using RapiClear. First, place the embryos in the center of a 75 by 25 millimeter depression concave glass slide, remove all the PBST in the microscope depression slide and add 200 microliters of clearing solution, then cover the slide preparation with a box. After 10 minutes protected from the light, when embryos start to become transparent, replace the clearing solution and wait an additional 20 minutes.

Top the embryo with a coverslip starting from one side and gently moving towards the other. The sample is now ready for imaging. For clearing using methyl salicylate or BABB, ensure that the embryos are completely dehydrated in 100%methanol and then start the series of BABB and methanol with 20%successive increases in concentration and a 20-minute incubation for each step.

When the BABB solution reaches 100%make two additional changes for a fresh solution. Prepare a 20 by 60 millimeter 1.5 cover glass slide to mount the embryos to avoid compressing the sample. Add spacers made from thin metal washers.

Transfer the cleared embryos to the microscope slides using a toothpick, a fine cotton tip, or a Pasteur pipette. Then add a drop of mounting medium, cover with another similar cover glass and seal with melted paraffin to better stabilize the preparation to avoid bubbles. Place the coverslip on one side and then gradually and gently slide it sideways, adding more medium if necessary.

The sample is now ready to be imaged in the microscope. Use Fiji/ImageJ to reposition the embryo into a standardized anterior-posterior and dorsal-ventral axis position after imaging. Reduce the dataset size in Fiji's image, then scale and insert the X, Y, and Z scale values necessary to reduce the dataset to less than 200 megabytes.

Make sure the create new window option is ticked. Then go to plugins and open 3D Viewer. Inside the 3D Viewer window, click over the embryo to select it, causing a red 3D box to appear.

When using this mode, control the rotation of the dataset with a mouse. After ensuring the perfect position of the embryo, select edit, transform the image, and export transformed image in the 3D Viewer window. Inspect the different orthogonal planes, then go to image, then stack and select orthogonal views.

When the embryo is correctly positioned, select the 3D Viewer window menu, then edit, transform, and save transformation matrix to a text file with a mat extension. Open the text file using Fiji/ImageJ, remove the first two lines and re-save. This creates a transformation matrix file compatible with the TransformJ plugin.

Switch to the full resolution dataset and perform the operation plugins TransformJ and TransformJ affine. In the new window, browse to search for the matrix file previously saved, select the cubic B-spline interpolation and resample isotropically, then click OK.After properly repositioning the embryo, trim most of the created empty space. To perform a full Z projection image, click on stacks, Z Project, and choose maximum intensity.

Then draw the minimum ROI that contains the whole embryo in X and Y.Switch to the original dataset image windows by clicking on edit, selection, restore selection, and crop by selecting image. Trim the slices in the beginning and end that do not intersect embryo tissues by selecting image and opening stacks, click on tools, select the slice remover, and specify the first and last slice to remove, making sure to change the increment to one. If desired, perform further analysis and 3D reconstructions using the instructions in the text manuscript.

To perform 3D analysis of more developed embryos, dissect embryonic day 18.5 fetuses in cold PBS, remove all extra embryonic membranes, then wash the fetuses several times in fresh PBS to remove blood and debris produced during the dissection procedure. Fix fetuses in 4%PFA made in PBS at four degrees Celsius for five to seven days. After washing the embryo several times in PBS, dehydrate the fetuses by incubating in 10%10 successive increases of methanol solutions until 100%methanol is reached.

Perform each incubation for 25 minutes on a shaker at room temperature. Next, incubate fetuses separately on a shaker first for one day in 5%hydrogen peroxide in methanol and then in 10%hydrogen peroxide in methanol for up to three days until the embryos lose all natural pigmentation. Gradually rehydrate the embryos in a reverse methanol series in demineralized water with each incubation for 20 minutes on a shaker at room temperature, then wash the embryos three times for 30 minutes per wash in demineralized water.

To embed fetuses in 1%agarose blocks, fill a 50 milliliter plastic tube or syringe with melted agarose, then place the fetus in the agarose in a vertical position and maintain this position with forceps during solidification of the agarose. Place the mold with the block at four degrees Celsius for 30 minutes for the agarose to fully jellify, then remove the agarose block with the fetus from the mold and place it in a container with demoralized water. Perform fetus dehydration gradually with 10%increases in methanol concentration diluted in demineralized water or PBS, keeping the sample in methanol concentration for at least 45 minutes at room temperature on a shaker until 100%methanol is reached.

Clear the fetuses by moving them through a series BABB solution and methanol for 2.5 hours in each concentration with 25%increases in BABB concentration until reaching 100%BABB. Replace the BABB solution with a fresh one every day until the fetus is completely transparent. This process might take three to five days.

Use glue to attach the cleared agarose block to the motor axis of the OPT scanner, then adjust the optics to obtain an image of the whole fetus and proceed to acquire a full projection dataset. The 4D live imaging technique enables the dynamic analysis of LuVeLu reporter expression and embryonic day 8.5 Snai1-conditional knockout embryos. Along with the normal LuVeLu signal on the presomitic mesoderm, Snai1-conditional knockout embryos display LuVeLu expression in the ectopic bulge that arises from the primitive streak.

Whole mount immunofluorescence assays allow the detection of potential NMPs and key regulators of mesoderm differentiation. White and yellow arrows highlight differences in the mutant and wild type embryos. 3D renderization of whole mount immunostainings enable a better comprehension of the tissue or structure in the study.

In this case, the location of potential NMPs in the tail bud was investigated. 3D tissue reconstructions are also important to understand and characterize morphological defects in mouse embryos. Interactive visualization of 3D reconstruction can also be built in user-friendly formats.

For example, in a PDF file. These 3D embryo caudal structures can be used to illustrate the power of 3D models to a more general audience and to help in the study in teaching a vertebrate axial elongation and segmentation. Finally, 3D in toto visualization of more developed mouse embryos is possible using the OTP microscopy technique.

Animated renderings are shown here, highlighting key slices of embryonic day 18.5 fetuses. The most important thing to remember is that the result must remain a faithful representation of what is observed in the embryo. Researchers can use these methods to go beyond presenting just static to the images, but also include videos, three-dimensional reconstructions, and even interactive 3D models of their data.