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

This protocol follows the University of Arkansas (UARK) guidelines for research on invertebrate animals, such as Drosophila melanogaster, and was approved by the UARK Institutional Biosafety Committee (protocol #20001).

1. Drosophila embryo fixation and devitellinization

NOTE: Step 1 describes a procedure (hand-peeling) for the manual removal of the vitelline membrane, a transparent impermeable membrane that surrounds the embryo. Importantly, hand-peeling allows for the selection of properly staged embryos at the start of the ExM protocol, thus greatly enhancing the likelihood of obtaining embryos in a useable orientation at the end of the ExM protocol. However, this ExM protocol is completely compatible with bulk embryo collection and standard procedures for methanol-based removal of the vitelline membrane, in which case one can skip directly to step 2 (immunofluorescence labeling).

1. Prepare or purchase a number of fine glass needles. The actual dimensions of the needle tip are not critical, but ensure the needles are rigid and sharp enough to pierce the vitelline membranes of fixed embryos. Make needles from glass capillary tubes (1 mm outer diameter, 0.75 mm inner diameter) using a micropipet puller, as one would prepare needles for embryo microinjections¹⁸; alternatively, purchase pre-pulled needles.
2. Collect embryos using standard Drosophila techniques¹⁹ by placing >100 adult Drosophila in a vented plastic cup sealed with a fruit-juice/agar plate²⁰. Use timed collection windows to enrich for embryos of the proper stage¹⁶. For example, to enrich for the gastrulation (stage 6) and convergent-extension (stage 7) stages, change the fruit-juice plate, collect the embryos for 2 h at 25 °C, then remove the plate, and age it for a further 2 h at 25 °C to obtain embryos that are ~2-4 h old.
3. To remove the eggshell-like chorion from the embryos, cover the surface of the fruit-juice plates with 50% bleach (Table 1), release the embryos from the surface of the agar by agitating them with a small paintbrush, and wait 3 min for the chorion to dissolve.
4. Transfer the dechorionated embryos using a paintbrush to a 30 mL scintillation vial containing 4 mL of heptane (organic top phase) and 4 mL of fixation buffer (aqueous bottom phase; Table 1). Freshly dilute the formaldehyde from a freshly prepared or recently opened stock, and mix with 10x PBS and deionized water immediately before adding the embryos.
   NOTE: Prepare the formaldehyde from paraformaldehyde power or 16% EM-grade formaldehyde in glass ampules. Stocks of concentrated formaldehyde (e.g., 37% formaldehyde) can be used, although the results may be less consistent.
5. The embryos will accumulate at the interface between the organic and aqueous phases. Add as many embryos as will form a single layer at the interface. If too many embryos are added to a vial, they will not fix as well.
6. Using strong tape, immobilize the scintillation vials on their sides on a tabletop shaker, and agitate them for 20 min at 220 rpm. For optimal fixation, maintain a vigorous emulsion between the organic and aqueous phases during the entire fixation.
7. During the fixation time, prepare one of the following for each of the samples.
   1. Take a plastic 6 cm Petri dish base filled halfway with 3% agar and score a ~5 cm x 3 cm rectangle in the agar with a razor blade or scalpel. Fruit juice/agar plates can also be used for this purpose.
   2. Using a small lab spatula, remove the agar slab. Invert the base of the Petri dish, and set it on the bench. Place the agar slab on top of the inverted dish (Figure 2A).
   3. Take the lid of the Petri dish and make sure it is dry. Wearing gloves, place a piece of double-sided tape inside of the lid (the piece of tape should be slightly larger than the agar slab; Figure 2B).
8. Remove the vials from the shaker, set them upright on the bench, and allow the organic and aqueous phases to separate. Properly fixed embryos will remain at the interface between the two phases.
9. Transfer the fixed embryos onto the agar slab using a glass Pasteur pipet fitted with a latex bulb. To prevent the embryos from adhering to the inside of the pipet, try to keep the embryos within the narrow neck of the pipet, and transfer the embryos in multiple small batches rather than all at once. Once all the embryos are on the agar slab, remove most of the residual heptane from around the embryos using a P200 pipettor. Perform this step as quickly as possible (<3 min) to avoid drying out the fixed embryos, which can negatively affect the morphology.
10. From a height of ~2 cm, drop the lid with the double-sided tape onto the agar slab to adhere the embryos to the tape (Figure 2C). Gently remove the lid from the agar slab, place it upside down on the bench, and then add enough PBS-Tween (Table 1) to cover the embryos in the lid.
11. Using a stereo dissecting microscope at approximately 100x magnification with indirect lighting, identify properly staged embryos using morphological markers. For stage 6 embryos, use markers like a visible cephalic furrow and invaginated mesoderm; for stage 7 embryos, use markers like an extended germband; for stage 11 embryos, use markers like a fully extended germband and visible segmentation along the head-to-tail axis¹⁶.
    1. To collect the desired embryos, first prick the vitelline membrane (a transparent oval membrane around the embryo) near the anterior or posterior end of the embryo with a fine glass needle; the membrane will deflate a bit as the pressure releases. Then, using fine forceps or a metal probe, gently push the embryo on the other end through the hole; the vitelline membrane will remain adhered to the double-sided tape. Leave undesired embryos adhered to the tape.
12. Periodically collect the floating devitellinized embryos with a glass Pasteur pipet, and move them to a 1.5 mL microfuge tube.
13. At this point, perform one of the following steps.
    1. Continue directly to the immunofluorescence labeling steps. The devitellinized embryos can go directly into blocking solution (step 2.2). Do not allow the embryos to remain in PBS-Tween or blocking solution (Table 1) for longer than 16 h before proceeding to the next step.
    2. Move the embryos into methanol for storage. Remove as much of the PBS-Tween as possible, and then add 1 mL of methanol. Once the embryos have settled, remove as much methanol as possible, and add 1 mL of fresh methanol. Store the embryos at −20 °C indefinitely. Methanol storage also allows for the pooling of embryos from multiple collections.

2. Immunofluorescence labeling

NOTE: Aside from the antibody incubation steps, exact liquid amounts and times are not critical in this section. To perform a rinse or wash, allow the embryos to settle to the bottom of the tube, remove as much liquid as possible without sucking up embryos, and then add ~1 mL of new liquid; use a glass Pasteur pipet fitted with a latex bulb for optimal clarity and control. For the rinse step, the embryos are not rocked, just allowed to settle; for the wash step, the embryos are rocked on a nutator for the indicated amount of time and then allowed to settle.

1. If embryos were not stored in methanol, proceed to step 2.2. If the embryos were stored in methanol, rinse twice with PBS-Tween, and then wash for 20 min twice with PBS-Tween.
2. Wash the embryos for 30-60 min in 1 mL of blocking solution.
3. Incubate the embryos with primary antibodies diluted in antibody solution (Table 1) for 2 h at room temperature or preferably overnight at 4 °C. Perform this step in as small a volume as possible (50-300 µL) to conserve the primary antibodies; rocking on a nutator is not strictly required.
   1. Increase the amount of primary antibody used in a typical immunofluorescence experiment by at least 50% for ExM. Use the following primary antibody concentrations: 1:200 for anti-Par-3 guinea pig polyclonal²¹ and 1:100 for anti-GFP rabbit polyclonal.
4. Remove the primary antibody solution (save at 4 °C if desired), rinse twice with PBS-Tween, and then wash for 15 min four times with PBS-Tween.
5. Incubate the embryos with fluorescent secondary antibodies in a final volume of 300 µL (diluted in antibody solution) for 1 h at room temperature on a nutator. Fluorescently labeled streptavidin can be added during this step. From this step onward, protect the embryos from excessive and prolonged light exposure when possible, for example by covering the tubes with an opaque box lid or keeping the samples in a drawer.
   1. Use the following concentrations: 1:500 for anti-rabbit IgG goat polyclonal fused to Alexa Fluor 488; 1:500 for anti-guinea pig IgG goat polyclonal fused to Alexa Fluor 568; and 1:1000 for streptavidin-Alexa Fluor 488.
6. Remove and dispose of the secondary antibody solution. Rinse the embryos twice with PBS-Tween, and wash for 15 min four times with PBS-Tween.
7. At this point, the embryos can be stored at 4 °C in the dark but process the samples as quickly as possible (<24 h).

3. Preparing PDMS wells

NOTE: The PDMS wells can be made up to 2 weeks in advance.

1. Set an incubator or hot plate to 55 °C, and set a centrifuge that can spin conical tubes to 15 °C.
2. To prepare the PDMS solution (Table 1), place a 50 mL conical tube in a secondary container on a scale, and add 10 g of silicone elastomer base to the tube using a syringe. Then, add 1 g of silicone elastomer curing agent, and invert the tube several times to mix.
3. Create a balance tube by adding an appropriate amount of water to a second 50 mL conical tube. Centrifuge the PDMS solution at 500 x g for 3 min at 15 °C, and then pour it into a 10 cm Petri dish to a depth of ~1 mm. If necessary, remove bubbles by blowing gently on the solution with an air hose. Allow the PDMS solution to solidify overnight at 55 °C.
4. Once the PDMS slab is solidified, using a scalpel, score square areas that are slightly smaller than a 22 mm x 22 mm coverslip. Inside each square, score and remove an ~8 mm-wide square well.
5. Transfer each square PDMS well onto a 22 mm x 22 mm coverslip, and firmly adhere it (Figure 2D). Preparing six or more coverslips should yield a good number of expanded embryos to image.

4. Adhering the embryos to the coverslips

1. Apply enough 0.1% poly-L-lysine to cover the coverslip surface inside of each well (~50 µL), and place them in a 55 °C incubator to air dry. Repeat this step to increase adhesiveness.
2. Briefly rinse the embryos once in 1x PBS to remove the Tween detergent, and then transfer >10 embryos into each of the poly-L-lysine-coated wells.
3. Allow the embryos to settle to the bottom of the wells. Remove the excess liquid from the adhered embryos using a Pasteur pipet. Immediately proceed to the next step.

5. Activation and gelation

NOTE: Activation refers to the addition of MA-NHS to the embryos, which will modify the sample proteins and antibodies so they can bind to the hydrogel. Gelation refers to the generation of a hydrogel in and around the embryos in each well. During gelation, the embryos are permeated with a monomer solution and then treated with a gelation solution to form the hydrogel.

1. Activate the embryos for 1 h at room temperature by filling the wells with activation solution (1 mM MA-NHS freshly diluted in 1x PBS; Table 1). Change this solution approximately every 10 min over the course of 1 h.
2. Rinse the embryos with 1x PBS three times. Incubate the embryos in monomer solution (Table 1) for 45 min at 4 °C.
3. While the embryos are sitting in the monomer solution, prepare the gelation solution (Table 1). Preparing ~2 mL of gelation solution is enough to cover the PDMS wells from an entire 10 cm Petri dish. Be sure to add the APS last, as it will initiate polymerization and begin the gelation.
   1. Dilute the catalytic oxidant freshly from powder (e.g., 1% TEMPO w/v in water). Combine 1,960 µL of monomer solution with 30 µL of 10% TEMED and 10 µL of 1% TEMPO.
   2. To avoid polymerizing the entire batch of gelation solution at once, work in small batches. Split the gelation solution (without APS) into 125 µL aliquots between the eight tubes of a PCR strip.
   3. Remove the monomer solution from the three PDMS wells using a vacuum while being careful not to disrupt the embryos. Add 5 µL of APS to one of the PCR tubes containing gelation solution to initiate polymerization. Quickly distribute the polymerizing gelation solution amongst the wells (~40 µL per well). Repeat this until all of the wells and embryos are covered.
4. Let the samples gel for 1.5-2.5 h at 37 °C. Agitate the hydrogels every so often to monitor the polymerization. Solidified hydrogels will not wiggle. Thicker hydrogels will take longer to complete polymerization and solidify.

6. Digestion and expansion

NOTE: Thicker and larger gels will take longer to expand, and the center of the gels may take several hours to completely expand; this can be sped up by trimming the edges of the gel. As the gels expand, their refractive index will become nearly identical to that of water, and they will become very hard to see.

1. After gelation is complete, peel the PDMS wells from the coverslip while trying not to disturb the hydrogels. Cut away excess hydrogel material, if desired.
2. Transfer the hydrogels (still attached to the coverslips) individually into the wells of a six-well plate (Figure 2E). Note that the hydrogels may slightly expand during digestion.
3. Cover the gels completely with digestion buffer (Table 1) for 1 h at 37 °C. In general, 30 mL of digestion buffer is sufficient to cover the gels in a 6-well plate.
4. After digestion, transfer each hydrogel individually into a 6 cm Petri dish by sliding them off the coverslip. It may be necessary to use a second coverslip to dislodge the gel. Fill each Petri dish with deionized water to expand the gel. Change the water three to four times over the course of 1-2 h until the gels are fully expanded (expect an approximate four-fold increase in width).

7. Mounting and imaging

NOTE: Expanded hydrogels are composed almost entirely of water, making them nearly transparent and extremely fragile. The gels can be manipulated using long coverslips to move them around and pick them up. Mount and image only one or two gels at a time, as gels will gradually release water and begin to slide around the coverslip.

1. Using a Pasteur pipet, remove as much excess water from the Petri dish as possible to minimize the gels moving around when handled.
2. Maneuver each expanded gel, with the embryos on the bottom surface, onto a large coverslip (e.g., 24 mm x 40 mm) for imaging.
3. Mount each coverslip with gel over the objective of an inverted laser-scanning confocal microscope. Locate properly staged and oriented specimens using the epifluorescence or brightfield microscopy modes on the microscope with low-magnification (5x or 10x) or medium-magnification (20x) air objectives.
4. To image at high resolution, switch to a high-magnification (60x, 63x, or 100x) oil- or water-immersion objective. The surface of the embryos must be within the focusing range of the objective (<300 µm from the coverslip for a 63x objective) in order to be imaged.
5. Collect data from samples using the laser-scanning confocal mode on the microscope. Be sure to collect non-saturated images with good dynamic range, and use an appropriate number of pixels per image to capture the maximal possible information about the sample²³.

Figure 2: Manual devitellinization and working with hydrogels. (A) Cutting an agar slab from an agar/fruit-juice plate. (B) Placing double-sided tape inside the lid of a 6 cm Petri dish. (C) Adhering embryos to the taped lid. (D) A PDMS slab with a square well adhered to a 22 mm x 22 mm coverslip. (E) Coverslip with a PDMS well inside a 6-well plate. Please click here to view a larger version of this figure.