So using this protocol, we can visualize cytonemes in fixed mouse tissue throughout the entire embryo with standard light microscopy. Using this technique, we can probe for endogenous signaling proteins that travel along cytonemes, rather than rely on genetically manipulated mouse models that use fluorescently-tagged proteins. While attempting this protocol, it is important to gently handle tissue sections for the optimal preservation of cytonemes.
Demonstrating this procedure will be Miriam Dillard, a lead researcher in my group, and Christina Daly, a St.Jude graduate student. To begin, use dissecting scissors and forceps to make a Y incision to the peritoneal cavity. Excise the uterus containing E 9.5 embryo.
Dissect the embryos in complete DMEM growth medium. Use forceps to remove the yolk SAC, placenta, and surrounding membranes. Rinse the isolated embryos in HBSS to remove any residual amniotic tissue and blood.
Next, prepare the fixative by adding paraformaldehyde to HBSS for a working concentration of 4%paraformaldehyde. Add one milliliter of this solution to each well of a 24-well plate. Place the embryos into individual wells.
Incubate the embryos for 45 minutes with gentle agitation on a rocker. After incubation, remove the fixative and wash the embryos thrice, for 30 minutes in PBS with added calcium, magnesium, and 0.1%Triton. Then incubate the embryos in blocking solution with gentle agitation twice, for one hour each.
After the second incubation, rinse the embryos using fresh blocking solution. In the meantime, prepare the primary antibody solution by diluting the antibodies in supplemented PBS. After the rinse is complete, remove the blocking solution and add one milliliter of primary antibody solution to each well.
Incubate the plate at four degrees Celsius with gentle rotation for three days. Following the primary antibody incubation, wash the embryos with supplemented PBS five times for one hour on a rocker at 20 RPM. Then add one milliliter of secondary antibody solution to each well.
Incubate the plate with gentle rocking at four degrees Celsius in the dark for three days. Remove the secondary antibody solution and wash the embryos thrice for 30 minutes in supplemented PBS. Prepare a 4%solution by weight of low melting point agarose in PBS with calcium and magnesium.
Simultaneously, place a 12-well plate in the 55 degrees Celsius bead bath and add three milliliters of agarose to each well. Next, place the plate on a bench top and transfer the embryos into individual wells using a perforated spoon. Use pipette tips to gently embed and orient the embryo such that it is centered within the solution.
Once the embryos are oriented, place the plate at minus 20 degrees Celsius for 10 minutes for solidification. Then using a scalpel, remove the entire agarose block from the well. Cut a rectangular block around the embryo, leaving approximately 0.3 centimeters of the block on each side.
Leave an extra length of the block along the caudal end of the embryo. To mount the embryo on the vibratome, first apply a strip of tape to the specimen holder. Orient the embryo in an upright position in the upper section of the block and super glue the agarose block to the tape, such that the blade will generate axial sections in an anterior to posterior sequence.
Next, fill the vibratome chamber with cold HBSS to fully immerse the sample and then surround the chamber with ice. Set the parameters on the vibratome and perform serial axial sectioning of the embryo. Use forceps to transfer individual sections to a separate dish filled with HBSS.
Remember to use forceps for holding only the agarose to avoid tissue damage and destruction of cytonemes. To perform F-actin staining, remove HBSS and incubate the sections for 40 minutes with ActinRed and DAPI solution in supplemented PBS. After incubation, wash sections thrice for 20 minutes in supplemented PBS.
With a hydrophobic marker pen, draw a barrier around the edges of a charged microscope slide and add a small volume of HBSS to the fill area. Then, use forceps to transfer sections to the slide. Remove excess agarose using forceps.
Once all the sections have been transferred to the slide, remove excess liquid by pipetting and using the corner of an absorbent towelette. Then add several drops of mounting medium to the slide. Mount the cover slip by gently placing it on the slide.
For analysis, perform imaging of the tissue sections for a minimum of three embryos per genotype on a confocal or any high-resolution microscope. Correctly-oriented sections prepared using this protocol are shown here. Compared to vibratome sectioning, cryostat sectioning of the tissue did not preserve cellular extensions.
A few GFP-positive membrane fragments were detected between cells of the notochord and neural tube, and between adjacent neural tube cells in cryostat sections. However, F-actin staining of cellular extensions in the mesenchymal cells surrounding the neural tube was impaired in cryostat sections. Vibratome sectioning ensured minimal disruption of whole embryo and individual tissue sections.
Optimally-handled sections allowed the detection of ctyonemes between adjacently localized floor plate neural epithelial cells and mesenchymal cells. Folded or buckled sections were evident by a large separation between the notochord and the ventral floor plate of the neural tube. And a loss of cellular membrane extensions migrating between epithelial cells.
F-actin and DAPI-stained sections had a consistent spacing of mesenchymal cells and ctyonemes surrounding the neural tube and notochord. Minor distortions to the sections may have caused fragmentation of actin-based extensions and large gaps to form between cells, underscoring the need for delicate handling. After embryo sectioning, it's imperative to minimize any bending or folding of tissue sections.
to prevent the breaking of cytonemes. Using this technique, we can directly visualize how signaling molecules, like morphogens are spread across tissues. This is giving us new insights into how tissues and organs are patterned.
