Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

This protocol can be implemented in most esophagal developmental biology labs, allowing researchers who do not have access to super resolution microscopes to produce super resolution images. The main advantage of this technique is that it allows researchers to bypass the diffraction limit of conventional confocal microscopy without requiring a super resolution microscope by expanding the sample itself. This protocol will be beneficial for those studying how protein localization affects cell morphology or function in intact embryos, particularly when those proteins are organized into complex subcellular structures or networks.

The most common problem people encounter is losing embryos throughout the protocol. It is advisable to apply multiple coats of Poly-L-Lysine to adhere the embryos firmly. After the embryos, take a six centimeter plastic Petri dish base filled half with 3%agar.

Score a five by three centimeter rectangle in the agar with a razor blade or scalpel. Remove the agar slab using a small lab spatula. Then invert the base of the Petri dish, set it on the bench, and place the agar slab on top of the inverted dish.

Remove the lid from the Petri dish and ensure it is dry. Using gloves, affix a piece of double-sided tape to the inside of the lid. Take the vials containing the coated and fixed embryos out of the shaker and place them vertically on the workbench.

Allow the organic and aqueous phases to separate. Properly Fixed embryos should remain at their interface. Completely remove the bottom aqueous phase using a Pasteur pipette and a P200 pipette.

Use a glass Pasteur pipette fitted with a latex bulb to transfer the fixed embryos onto an agar slab in multiple small batches so that they don't adhere to the inside of the pipette. Once the embryos are on the agar slab, use a P200 pipetter to remove any remaining heptane near the embryos. Now, drop the double-sided tape lid onto the agar slab from a height of two centimeters to adhere the embryos to the tape.

Gently remove the lid from the agar slab, place it upside down on the bench, and add enough PBS Tween to cover the embryos in the lid. To collect the desired embryos, use a stereo dissecting microscope at 100x magnification with indirect lighting. Prick the vitelline membrane near the anterior or posterior end of the embryo with a fine glass needle to deflate it and release pressure.

Use fine forceps or a metal probe to gently push the embryo out through the hole with the vitelline membrane still adhered to the double-sided tape. Leave undesired embryos on the tape. Periodically use a glass Pasteur pipette to collect any floating devitellinized embryo and move them to a 1.5 milliliter microfuge tube.

Prepare the PDMS solution in a 50 milliliter conical tube and create a balanced tube by adding an appropriate amount of water to a second 50 milliliter conical tube. Centrifuge the PDMS solution at 500G for three minutes at 15 degrees Celsius. Then pour it into a 10 centimeter Petri dish to a depth of one millimeter.

Allow the PDMS solution to solidify overnight at 55 degrees Celsius. Once the PDMS slab is solidified, score square areas slightly smaller than a 22 by 22 millimeter cover slip using a scalpel. Inside each square, score and remove an eight millimeter wide square well.

transfer each square PDMS well onto a 22 by 22 millimeter cover slip and firmly adhere it. To adhere the embryos to the cover slips, apply enough 0.1%Poly-L-Lysine to cover the cover slip surface inside each well and place them in a 55 degree Celsius incubator to dry. Briefly rinse the embryos and PBS to remove the Tween detergent.

Then transfer more than 10 embryos into each Poly-L-Lysine coated well. Allow the embryos to settle to the bottom of the wells. Remove the excess liquid from the adhered embryos using a Pasteur pipette and immediately proceed to the next step.

While the embryos sit in the monomer solution, prepare a gelation solution to cover the PDMS wells. Dilute the catalytic oxidant freshly from the powder. Combine 3, 920 microliters of monomer solution with 60 microliters of 10%TEMED and 20 microliters of 1%TEMPO.

Split the gelation solution in 125 microliter aliquots into multiple PCR tubes. Remove the monomer solution from the PDMS wells using a vacuum or pipetter while being careful not to disrupt the embryos. Add five microliters of APS to one of the PCR tubes containing gelation solution to initiate polymerization.

Quickly distribute the polymerizing gelation solution amongst the three wells. Repeat this until all the wells and embryos are covered. Let the samples gel for 1.5 to 2.5 hours at 37 degrees Celsius.

Thicker hydrogels will take longer to complete polymerization and solidify. Agitate the hydrogels often to monitor the polymerization. Once solidified, hydrogels will not wiggle.

After gelation, peel the PDMS wells from the cover slip without disturbing the hydrogels. Transfer the hydrogels individually into the wells of a six well plate. The hydrogels may slightly expand during digestion.

Cover the gels completely with digestion buffer. 30 milliliters of digestion buffer is sufficient to cover them in a six well plate and incubate them for one hour at 37 degrees Celsius. After digestion, move the hydrogels into a six centimeter Petri dish and fill it with deionized water to expand.

The hydrogels may detach from the cover slips at this point and expand four folds in linear dimension. Using a Pasteur pipette, remove as much excess water from the Petri dish as possible to minimize the movement of the gels when handled. Maneuver the expanded gels with the embryos on the bottom surface onto a large cover slip for imaging.

Mount each cover slip with gel over the objective of an inverted laser scanning confocal microscope. After locating the properly staged and oriented specimens switched to a high magnification oil or water immersion objective to image at high resolution. Measurement of embryo length along the head to tail axis under a 10x objective showed that unexpanded embryos spanned approximately one half of a field of view, whereas the expanded embryos spanned approximately two full fields of view.

The average head to tail length of the unexpanded control embryos was 398.8 micrometers. For experiments one, two, and three, the average embryo lengths were expansion factors of 4.0 fold, 4.7 fold, and 4.9 fold respectively. In the control sample, the cells in the maxillary segment had an average width of 4.76 micrometers.

In the expanded samples, the cells in the maxillary segment had an average width of 19.10 micrometers, representing a 4.0 fold expansion. The actomyosin cytoskeleton was imaged in unexpanded control versus expanded embryos, undergoing convergent extension. They appeared as a single line where neighboring cells met.

By contrast, in expanded stage seven embryos, parallel lines of myosin two could be observed at cell cell junctions, representing cortical protein pools in adjacent cells. In the unexpanded stage six embryos, the mitochondria labeled with streptavidin appeared as a heterogeneous cytoplasmic haze without any clear subcellular details. However, in the expanded embryos, numerous puncta were resolvable within the cytoplasm, likely representing fragmented mitochondria or portions of the mitochondrial network.

One important thing to remember when attempting this procedure is to protect the embryos from excessive light exposure after the secondary antibody incubation.