Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

1. Prepare embryo collection cups and apple juice agar plates

1. Five days prior to the injection experiment, construct²⁸ or procure at least two small embryo collection cups. Make fresh 60 mm apple juice agar plates to be used with small collection cups²⁸. Store plates in plastic boxes covered with damp paper towels at 4 °C.
   NOTE: Small embryo collection cups, populated with fly numbers as described in step 1.3, will provide sufficient embryo numbers per experiment, while also ensuring that embryo handling and injection can be done in a short enough time to allow imaging of early developmental stages.
2. Warm apple juice plates to 18 °C and add a dab of yeast paste to the center of the plate. Yeast paste is a simple paste of active yeast and distilled water.
3. To promote the most generous egg laying, set up collection cups with flies 2 days prior to the experiment. Add at least 100 females and 50 male flies to the collection cups, and top with a prepared apple juice plate (Figure 1, step 1). On the days leading up to the injection experiment change the apple juice plates at least twice each day, once in the morning and once in the evening.
   NOTE: The best injection and imaging results are obtained when embryo collection cups are kept at 18 °C with a 12 h light on/light off cycle.

2. Prepare a working stock solution of G-actin^Red for microinjection

NOTE: This preparation only makes 2 μL of a 5 mg/mL working stock of G-actin^Red, so if users are unaccustomed to the microinjection technique, it is advantageous to skip to step 3 and practice the microinjections with a neutral pH buffer to conserve precious working stock. The 10 μg stock of G-actin^Red from the vendor can be stored in its original packaging in a 16 oz screw top jar with ~500 g of desiccant at 4 °C for up to 6 months.

1. Prepare a G-buffer stock solution in advance: 5 mM Tris-HCl, 0.2 mM CaCl₂, pH 8.0. Filter and store at room temperature.
2. On the day of injections, prepare 1 mL of a G-buffer working solution in a fresh snap cap microcentrifuge tube on ice using the G-buffer stock from step 2.1, supplemented with the following at the indicated final concentrations: 1 mM dithiothreitol (DTT) and 0.2 mM ATP, pH 8.0.
   NOTE: The 1 mL volume of G-buffer working solution is more than what is needed for an experiment but simplifies the preparation. Excess can be discarded after the experiment or users can scale down according to their preference.
   1. Keeping the 10 μg G-actin^Red stock on ice, first add 1 μL of filtered, distilled water to the top of the pink droplet of G-actin^Red inside the tube.
   2. Next, add 1 μL of the cold, freshly prepared G-buffer working solution from step 2.2. Pipet up and down ~20 times to mix well with the pipette volume set to 1 μL. The G-actin^Red stock will now be at the final dilution of 5 mg/mL.
3. Incubate the prepared G-actin^Red for 30 min on ice undisturbed.
4. Centrifuge the prepared G-actin^Red at 16,000 x g for 20 min at 4 °C in a microcentrifuge to remove any precipitate.
5. Carefully pipet 1.5 μL of the supernatant into a fresh snap cap microcentrifuge tube on ice, avoiding the dark pink pellet.
6. Store the prepared G-actin^Red supernatant on ice for up to 6 h until ready to load into a microneedle.
   NOTE: Microneedles can be pulled from capillary tubes in advance on a micropipette puller, then stored at room temperature on a strip of modeling clay in a 100 x 20 mm Petri dish. Suggested parameters for microneedle pulling can be found in the Pipette Cookbook²⁹.

3. Collect embryos and mount for injection

1. Allow flies to lay embryos for 30 min on apple juice plates with yeast paste at 18 °C.
2. While flies are laying, pre-warm an apple juice agar plate without yeast paste to room temperature, cut out a 4 cm x 1 cm rectangular wedge of apple juice agar with a razor blade, and place on a 25 mm x 75 mm glass slide.
   1. Harvest plate from the 30 minute collection and dechorionate embryos by pouring fresh bleach, diluted 1:1 with distilled water, onto the plate and swirling the plate for 1 min, as described in²⁸ (Figure 1, step 2).
      NOTE: Different brands of bleach are sold at different concentrations. The bleach used here is 6% sodium hypochlorite from the bottle and is diluted to a final concentration of 3% sodium hypochlorite. Other bleach brands at slightly lower concentrations will work equally well.
   2. Pour the bleach and dechorionated embryos into a collection basket (a 70 μm cell strainer) and rinse the plate twice with distilled water from a squirt bottle, adding these washes to the collection basket.
   3. Vigorously rinse the dechorionated embryos in the collection basket with distilled water until no yeast clumps are visible and the basket leaves no pink marks from excess bleach when blotted on a paper towel.
3. Using a paintbrush with bristles dampened with distilled water, transfer dechorionated and washed embryos from the collection basket onto the prepared apple juice agar wedge on the glass slide.
4. Use a pair of fine tip tweezers or a dissecting needle to arrange ten embryos in a straight line along the long axis of the rectangular agar wedge (Figure 1, step 3). Arrange the embryos head-to-tail, such that their anterior pole is facing to the right and dorsal side is facing the researcher (Figure 1, step 3, magnified).
5. Cut off 0.5 cm of the end of a P200 pipette tip with a razor blade and dip into the “embryo glue” (described in²⁸). Generously coat a region 5 mm in width (Figure 1, step 4) along the long edge of a 24 mm x 50 mm rectangular coverslip, and let dry, glue side up. Drying will take ~30 s and is complete once the entire glue-coated region appears matte rather than wet or shiny.
   NOTE: Prepare “embryo glue” at least 48 h in advance. Add n-Heptane to strips of double-sided tape in a scintillation vial as described in²⁸.
6. Once the “embryo glue” has dried, gently place the coverslip glue side down on top of the row of aligned embryos on the agar, leaving 2-3 mm of space between the edge of the coverslip and the row of embryos.
   NOTE: Sticking the embryos too close to the edge of the coverslip may lead to the embryos drying out too much during the course of the experiment.
7. Flip the coverslip over so that the embryos are now facing up. They should be stuck in a line along one long edge of the coverslip, and their ventral region facing the closest edge of the coverslip (Figure 1, step 4, magnified).
8. Desiccate the embryos by placing the coverslip with embryos gently on top of 150 g of fresh blue desiccant stored in a 16 oz screw top jar. Tightly screw on the lid and incubate for 8-10 min (Figure 1, step 5).
9. After desiccation, remove the coverslip from the desiccant jar and tape each short side of the coverslip to a microscope slide, embryo side up, with two 4 cm² pieces of double-sided tape so that the embryo coverslip will fit onto the injection stage (Figure 1, step 6).
10. Add 2-3 drops of Halocarbon 27 oil with a Pasteur pipette to cover the aligned embryos and protect them from further dehydration (Figure 1, step 6).

4. Inject and heat stress embryos to promote actin rod formation

NOTE: All injections are done in a temperature-controlled room at 18 °C.

1. Prepare humid incubation chambers from a glass Petri dish at least 100 mm x 20 mm in size and line the chamber with twists of lab tissue wipers dampened with distilled water (Figure 1, step 8). Pre-warm the incubation chambers at 32 °C or the desired incubation temperature prior to injecting embryos.
2. Open the airflow valve for the microinjector and turn on the microinjector (compressed air or house air with a pressure of at least 90 psi is suitable).
3. While embryos are desiccating, backload the previously prepared G-actin^Red supernatant into the microneedle using a micro loader tip. Set the pipette to draw up 1-1.5 μL.
   NOTE: Because of the viscosity of the actin, loading volumes may not be accurate and there may be enough actin left to load at least one to two more microneedles. Up to 60 embryos can be injected per loaded microneedle if the microneedle is calibrated properly and does not become clogged during the course of the experiment.
4. Attach the microneedle to the needle holder and tighten the screw. Connect the air tube to the microinjector and ensure that the backflow pressure on the microneedle equilibrates to 30 hPa.
5. Calibrate the microinjector settings to expel a 100 μm diameter bubble of G-actin^Red (~500 pL) on a slide micrometer. Rotate the pressure knob (500-1500 hPa) and injection pulse time knob (0.1-0.5 s) on the microinjector to get the right bubble size. Adjust these settings each time a new microneedle is loaded to account for variability in actin viscosity and microneedle tip size.
   NOTE: The prepared G-actin^Red is viscous and there may be air in the tip of the microneedle that should be expelled before injecting embryos. If the G-actin^Red does not readily expel from the tip of the microneedle, gently break the microneedle tip against the edge of the slide micrometer.
6. Place the slide with mounted embryos onto the microscope stage.
   NOTE: Every injection set up will be different, so researchers will have to adjust their injection method accordingly. Here the embryos are moved with respect to a stationary microneedle, injecting each embryo by running the embryo into the microneedle.
7. Adjust the micromanipulator stage and focus of the 10x objective on the light microscope so that the embryos are visible. The embryos are in the correct focal plane when the outlines of the vitelline membrane are sharpest and the embryo appears largest. Choose embryos to inject that are in the correct developmental stage, so that by the time the post-injection incubation is complete, most of the clutch reaches the desired developmental stage (e.g., inject embryos at Bownes’ stage 2-3³⁰ in order to observe rods at cellularization after heat stress at 32 °C).
8. Use the microneedle controls to bring the needle into the same focal plane as the embryos.
   NOTE: If the microneedle catches on the coverslip while moving the stage or microneedle, then the microneedle is too close to the slide and is not in the correct focal plane. The microneedle should be parallel to the coverslip, and not at a significant angle (Figure 1, step 7).
9. Insert the microneedle into the embryo so that it hits the embryo in the middle of its ventral region, at the embryo “equator”. Trigger injection with the foot pedal or “inject” button when the microneedle tip is visible inside the middle of the embryo (Figure 1, step 7).
10. Inject the G-actin^Red once and slowly remove the microneedle. Move the stage and repeat for each embryo of the appropriate developmental stage.
    NOTE: Expansion of the embryo is normal as the G-actin^Red is injected and a bit of cytoplasm may leak out of the embryo.
11. After all the embryos have been injected, place the slide with the embryos in the prepared humid incubation chamber and close the lid (Figure 1, step 8). If trying to obtain embryos that reach cellularization, heat stress the embryos at 32 °C for 60-75 min in the humid incubation chamber.
    NOTE: Incubation times are noted that allow visualization of rods in cellularizing heat stressed embryos. The incubation time will be longer for non heat stress control embryos because development will be slower at a lower temperature³¹. These control embryos can be incubated in humid chambers at temperatures such as 18 °C or 25 °C, depending on design of the specific experiment and the question to be asked. The minimum incubation time necessary for the G-actin^Red to diffuse throughout the embryo is 30 min.

5. Image actin rods in heat stressed embryos by confocal microscopy

1. While the embryos are being heat stressed, turn on the confocal microscope and select the 561 nm laser channel.
2. Move the objective lens (25x, 40x or 63x recommended) to the working position.
3. If imaging heat stressed embryos, then set the heated stage incubator to achieve an internal temperature of 32 °C. A point-and-shoot or infrared thermometer can be used to check the temperature at or near the objective.
4. Remove the slide with injected embryos from the humid incubation chamber after incubation is complete (Figure 1, step 9).
5. Working quickly, gently pry off the double-sided tape pieces that were used to adhere the coverslip with mounted embryos to the slide (Figure 1, step 9).
   CAUTION: Be gentle during these steps as coverslips can easily shatter if too much force is applied.
6. Stick two 2.5 cm long pieces of double-sided tape together and cut the tape in half lengthwise to make two strips, 2.5 x 0.5 cm long (Figure 1, step 10).
7. Stick two-thirds of the length of each tape strip onto the first coverslip, flanking each side of the embryos in Halocarbon 27 oil (Figure 1, step 10, orange), leaving one-third of the tape strips hanging off the edge of the first coverslip where the embryos are stuck. Use gloved hands and be careful not to touch the embryos during this step.
8. Gently place a second rectangular coverslip on top of the tape strips to sandwich the embryos between the coverslips (Figure 1, step 10, blue). Align the 25 mm edges but keep the 50 mm edges offset from one another by 1 cm in width.
   NOTE: This second coverslip’s full surface will become the new imaging surface that will face the objective lens, so take care not to get fingerprints or Halocarbon 27 oil on this second coverslip surface. The offset is necessary so that extra Halocarbon 27 oil can be added to evenly immerse the embryos. If needed, add Halocarbon 27 oil at the seam where the two coverslips meet at the top of the sandwich and it will coat the embryos by capillary action (see dashed lines in Figure 1, step 10).
9. Gently tap down on the areas of the coverslip that are directly on top of the tape strips with the blunt side of a razorblade to get the coverslip to adhere to the tape.
10. Flip the coverslip sandwich over and place on a lab tissue wiper to keep the imaging surface clean and carry to the confocal microscope (Figure 1, step 11). Ensure that the tape is completely stuck to both coverslips before imaging.
11. Confirm that the heated stage is at temperature and if using an inverted microscope, add immersion liquid onto the selected objective lens.
12. Place the coverslip sandwich onto the stage carefully to ensure that the new imaging surface (2^nd coverslip) is the one touching the immersion liquid (Figure 1, step 11).
    NOTE: If needed, adhere the coverslip sandwich to the stage with two small pieces of double-sided tape to prevent unnecessary movement during imaging if the coverslips do not fit well in the heated stage.
13. Focus on an embryo that is in cellularization (Bownes stage 4a³⁰) or desired developmental stage using either transmitted light or fluorescence.
14. Once an embryo has been brought into focus, switch to the laser acquisition mode on the confocal microscope and adjust laser power and gain, frame size, tiling, and projection settings as desired.
15. Take surface-view images through the focal planes of the embryo’s nuclei to find intranuclear actin rods. Rods should appear in multiple orientations as bright streaks or dots inside the comparatively dark nuclei (Figure 2A, 2C).

6. Alternative imaging experiments

1. Perform FRAP to investigate actin turnover along the length of intranuclear actin rods or in cytoplasmic actin structures, such as the tips of the plasma membrane furrows during cellularization.
2. In the imaging software, choose a rectangular region around furrow tips or actin rods in which to acquire the experiment.
3. Choose a small square region of the center of an actin rod within the rectangular acquisition region to bleach. Ensure that the tips of the actin rod are visible and do not get bleached during the course of the experiment to allow for accurate tracking of the rod inside the nucleus.
4. Set the bleach laser to iterate 50x and set the bleach laser power to maximum.
5. Choose a time course to acquire images every second for a total up to 120 s at maximum pixel dwell speed and set the laser to bleach after the first two seconds of image acquisition.
6. Quantify the data to determine the half-time of fluorescence recovery¹.