Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

1. Prepare necessary materials

NOTE: These preparations are best done days or weeks ahead of time.

1. Prepare injection needles.
   NOTE: Needles can be stored indefinitely in a covered container. Needles must be fine enough to deliver ~700 fL while being strong enough to pierce the vitelline membrane.
   1. Place a capillary into the capillary holder on the needle puller. Secure the capillary with the wingnuts. Align the center of the capillary with the heating element so that two symmetric needles are generated.
   2. Choose appropriate settings on the needle puller according to the manufacturer's instructions.
   3. Perform quality control using a dissecting scope.
      NOTE: Needle tips should be as fine as possible. Cracked, jagged, or large-bore needle tips are discarded.
   4. Prepare batches of 10 needles (5 capillaries worth) at a time.
   5. Place a deformable putty, which has been rolled up into a cylinder, at the center of the bottom of a container with a lid. Carefully press the needles into the putty such that it holds them horizontally with the needle tips safely suspended in the air to prevent damage. Cover with the lid, and store until use.
2. Prepare egg collection plates. Store at 4 °C.
   NOTE: Egg collection plates are small agar plates supplemented with fruit juice to encourage egg-laying. How to prepare them is described in many protocols, including¹³.
3. Prepare heptane glue.
   NOTE: This glue is extracted from double-sided tape and is used to securely attach the embryos to a coverslip. It stops the embryo from floating out of focus during injection and imaging. The glue can be prepared days or weeks ahead of time.
   1. Ball up double-sided tape to a size bigger than a golf ball and place it at the bottom of a ~50 mL glass container with a tightly sealing lid. Pack tape tightly so enough adhesive will be present.
   2. In a fume hood, fill the glass container with heptane to cover all the tape. Keep the container tightly closed when not in use.
      CAUTION: Heptane is flammable as a liquid and vapor. It can cause eye, skin, and respiratory tract irritation. Breathing vapors may cause drowsiness, dizziness, and lung damage. Keep heptane away from sources of ignition and store it in a well-ventilated area meant for flammables and away from incompatible substances.
   3. Let the heptane glue sit overnight or longer. For best results, place the container on a shaker or another agitator overnight to help with dissolving the adhesive.
      NOTE: The tape itself will remain, but the adhesive will be dissolved in the heptane. In step 5.1.2, the glue is applied to a coverslip; heptane evaporates, leaving the glue behind.
   4. Test the heptane glue preparation by placing a drop of it on a glass slide and making sure a sticky residue remains once the heptane has evaporated. If adhesive residue is not visible afterward, add more tape to the glass container and repeat the previous step.
      NOTE: Once prepared, the glue can be used for months or even years.
4. Prepare a 1 mg/mL (3.8 mM) BODIPY493/503 stock solution by diluting commercially obtained preparation in pure anhydrous DMSO. Keep it covered to protect it from light and water. Store indefinitely at -20 °C or short term at 4 °C.

2. Prepare egg collection cages

NOTE: Do this at least a day (preferably 2 or 3 days) before the planned injection(s).

1. Prepare yeast paste.
   NOTE: Yeast paste supplements protein and other vital nutrients not provided in the apple juice plates. It promotes egg production and egg-laying.
   1. Add 2-10 g of dry baker's yeast, and then 1 mL of tap water to a small beaker. Mix using a spatula.
   2. Keep adding tap water in 1 mL increments until the desired, tooth-paste-like consistency is reached.
   3. Cover the mixture and store it at 4 °C.
2. Set up fly cages for egg collection.
   NOTE: Male and female flies of the desired genotype are required: 10-50 females less than 2 weeks old and a similar number of males. A number of different options for fly cages are available, including homemade options^14,13. It is important that the size of the apple juice plates is matched to the size of the fly cages.
   1. Place a small smear of yeast paste on an apple juice plate. Keep it covered and let it come to room temperature because flies will not lay eggs on the plate if it is too cold.
   2. Transfer flies to an embryo collection cage, seal with the yeasted apple juice plate, and secure the plate to the embryo collection cage.
   3. Allow newly transferred flies to acclimate to the cage for 1-2 days, replacing the plate with a fresh one daily. If all the yeast has been eaten by the next day, increase the amount of yeast paste for future collections.
      ​NOTE: This is an important step to increase egg yield as well-fed females lay more eggs.

3. On the day of injection, prepare the injection solution and load it into the needle

1. Use the stock solution of BODIPY 493/503 (3.8 mM in DMSO) as the injection solution.
2. Load a single needle with 1 µL of the injection solution using needle loading tips.
   NOTE: Strive to get the liquid all the way to the tip without trapping air bubbles. Be ready to quickly replace the loaded needle in case of accidental breakage.
   1. Attach a loading tip to a micropipette and draw 1 µL of the injection solution into the tip. Using gloves, hold the needle in one hand with its tip pointed away to minimize the risk of breakage.
   2. Carefully insert the loading tip into the needle and push it close to the needle tip. Dispense the liquid near the needle tip. After removing the pipette, hold the needle vertically with its tip down until the liquid flows to the tip.
3. Store the loaded needles in a separate container with putty as above (see step 1.1.5).
   ​NOTE: Needles should be prepared in advance (up to several hours) of injection but should not be used after more than a day.
4. Keep needles out of ambient light to prevent bleaching of the dyes. Cover the storage container with aluminum foil without damaging the needle tips. Make sure all light is obscured.

4. Collecting embryos for injection

NOTE: The timing of collection depends on what stage of embryos the injection needs to be performed at. With the timing scheme below, the embryos at the time of preparation for injection will be 0-90 min old, which corresponds to cleavage stages¹⁵.

1. On the day of injection, prepare apple juice plates with yeast, as in step 2.2.1. If N rounds of injections are planned, prepare N + 2 plates. Keep these plates at room temperature.
   NOTE: In addition to the N plates to collect embryos for injection, one plate is needed as a pre-collection plate and another one to feed the flies in the cage once collections are completed.
2. Replace the plate on the cage with a fresh yeasted plate (pre-collection plate) and leave it on the cage for 1-2 h.
3. Replace the plate on the cage with a fresh plate (collection plate). Discard the pre-collection plate, as it will contain embryos older than desired. Then, allow the flies to lay eggs for 1.5 h.
   NOTE: As well-fed flies typically lay their eggs shortly after the eggs have been fertilized, a 1.5 h collection time assures that at the end of the collection, most of the embryos on the plate are 0-90 min old, i.e., are in cleavage stages. Female flies that have not been fed fresh yeast since the previous day will tend to retain fertilized eggs for some indeterminate amount of time before laying. Hence, the pre-collection plate may have embryos that were fertilized before the start of collection and thus are older than 60 min, sometimes much older.
4. Replace the plate on the cage with a fresh yeasted plate. Cover the collection plate so that stray flies do not lay eggs on them.

5. Prepare embryos for microinjection

1. Assemble the materials needed.
   1. Attach a piece of double-sided tape to a glass slide. Avoid touching the tape with fingers as that reduces its stickiness.
      NOTE: This slide will be used for removing the chorion and not for imaging.
   2. Using a transfer pipette or a micropipette (p200 or p1000), place a small drop (200 µL or less) of heptane glue roughly in the center of a rectangular coverslip (60 x 25 mm). Allow the heptane to evaporate (this takes less than a min).
      NOTE: This coverslip will be used to mount the embryos for injection and imaging. The dimensions are chosen to fit into an adjustable metal holder on the confocal microscope.
   3. Assemble a desiccation chamber, a sealed chamber (e.g., a Tupperware sandwich box) containing desiccation beads. Only use desiccation beads that have not hydrated.
      NOTE: To ensure the embryos take up the injection solution, the embryo must be slightly desiccated to reduce internal pressure. This is done by placing the coverslip with the dechorionated, air-exposed embryos into the desiccation chamber.
2. Mechanically remove the chorion.
   1. Cover the appropriately aged embryo plate with a thin layer of Halocarbon Oil 27 to turn the eggshell transparent.
      NOTE: Embryos become translucent within tens of seconds¹⁵. If this step does not occur efficiently, the apple juice plates may be too wet and need to be dried off before use.
   2. View the plate under a dissection scope with transillumination (i.e., the light going through the plate into the eyepieces) to confirm the stage of the embryos.
   3. Select embryos in cleavage stages.
      NOTE: Cleavage stage embryos are entirely opaque; the subsequent blastoderm stages can be recognized by a band of transparent cytoplasm all around the opaque center¹⁵. A good introduction on how to recognize various embryonic stages is available on the website (given in reference¹⁶) maintained by the Society of Developmental Biology.
   4. Using fine tweezers, grab an embryo of the desired stage by its dorsal appendages and transfer it onto the prepared glass slide covered with a piece of double-sided tape. Place the embryo on the tape. Minimize the transfer of oil.
   5. As gently as possible, roll the embryo across the surface of the tape by gently nudging the embryo with the side of the tip of the tweezers. Do not poke the embryo directly with the sharp tweezer tips.
      NOTE: If the force applied is too low, the embryo will not roll. If it is too high, it will burst. Finding the appropriate intermediate force requires experience and, thus, this step should be practiced beforehand.
   6. Continue rolling the embryo until the chorion transiently adheres to the tape and cracks.
   7. Once the chorion cracks, keep rolling to separate the embryo (still inside the vitelline membrane) from the chorion as the chorion remains stuck to the tape.
   8. Confirm that the chorion is completely removed by observing the loss of the dorsal appendages from the embryo.
   9. Roll the embryo back onto the chorion, which is less adhesive than the tape. Gently rub the embryo with the tweezers until it attaches to the tweezers for transfer.
   10. Transfer the embryo to the coverslip with the heptane glue and bring it in contact with the glue, which typically detaches it from the tweezers.
   11. Adjust the orientation of the embryo on the coverslip.
       1. Embed the lateral surface of the embryo into the glue.
          NOTE: The surface of the embryo embedded into the glue is the one that will be imaged. Embedding the lateral surface is the simplest, but this can be adjusted depending on personal preference or the biological question to be answered.
       2. Orient the long axis of the embryo perpendicular to the long axis of the coverslip.
       3. If the embryo does not lay in the preferred orientation, clean the tweezers to remove any oil that would detach the embryo from the glue. Then, gently attempt to roll the embryo into position.
   12. Prepare the desired number of embryos for injection in the manner described above.
       NOTE: As time passes after removal of the chorion, the ambient air will begin to desiccate the embryos and thus the effect of step 5.3 will be uneven for the embryos on the coverslip, which were dechorionated at different times. Typically, 1-3 embryos are the most manageable.
3. Desiccate embryos.
   1. Place the coverslip with embryos into the prepared desiccation chamber and seal it.
   2. Allow the embryos to desiccate for 5-12 min.
      NOTE: The timing of this step is dependent on the ambient temperature and humidity and thus needs to be determined empirically.
   3. Remove the coverslip from the chamber and place a drop of Halocarbon Oil 700 onto it, fully covering the embryos, to prevent further desiccation.
   4. Inspect the embryos on a dissecting scope to judge proper desiccation.
   5. If the embryos are just slightly shriveled, but not deflated, proceed to microinjection (step 6).
   6. If the embryos are not sufficiently or overly desiccated, return to step 5.2 and prepare a new batch of embryos as the Halocarbon Oil 700 cannot be removed. If the embryos were overly desiccated, shorten desiccation time for the next batch of embryos by 3 min; if they were under-desiccated, increase desiccation time by 3 min.

6. Microinject embryos

NOTE: Ensure the microinjection setup includes an inverted microscope, a micromanipulator to hold and position the injection needle, and a commercial microinjector to deliver controlled volumes.

1. Power on the microinjector and input the preferred settings.
   NOTE: For this protocol, it is recommended to use linear motion output and the fast setting, but many others will work. The operator should determine personal preference.
2. Load the needle into the micromanipulator. To prevent the needle from being damaged while loading the coverslip containing the embryos, move it out of the way into a safe position.
3. Place the coverslip onto the stage, with embryos on top, toward the needle. Then, carefully move the needle back into position for injection, with its tip still well above the stage.
4. Put the 4x objective into the light path. Using the focus knob on the microscope, bring the embryos into focus.
   NOTE: This will be the focal plane used for injection and will be adjusted only minimally going forward.
5. Using the stage controls, move the embryos horizontally into the center of the field of view.
   1. Pan away from the embryos, keeping them at the edge of the field of view, in preparation for lowering the needle. Stay in the same focal plane to ensure the needle is lowered to the correct position.
6. Lower the needle tip into the correct focal plane and ensure that it is visible in the field of view together with the embryos.
   1. Using the micromanipulator controls and looking from above (not yet through the eyepieces), slowly drop the needle tip into the oil, aiming for the area where the objective points to. As the oil is quite viscous, go slowly to avoid damaging the needle.
   2. Once the needle is visible through the eyepieces, continue using the micromanipulator controls until the needle tip is in focus and at the center of the field of view.
   3. By moving the stage, bring the embryos back to the center of the field of view. Use the stage control, micromanipulator controls, and focus knob to fine-tune the position of the embryo and needle tip relative to each other while both are in focus. Aim to perform injections as close to the coverslip as possible.
7. Perform needle quality control.
   1. To ensure that the needle is functional, dispense some of the injection solution into the Halocarbon Oil 700 surrounding the embryo, visible as a bubble in the oil.
   2. If nothing is flowing out of the needle, gradually increase pressure at the injector. If this does not work, get a new needle.
8. Inject the embryo.
   NOTE: Acceptable injection volumes range from ~0.06-1 pL. The lower limit is set by what can be visualized entering the embryo. The upper limit is set by the trauma the injection exerts on the embryo.
   1. Inject along the lateral edges of the embryo as this is the least invasive.
   2. Using the stage controls, move the embryo toward the needle tip until the latter gently punctures the embryo and enters it.
   3. At the same time, start the flow of injection solution and watch for the appearance of a transient clear spot at the site of the needle tip, which indicates a successful transfer of liquid into the embryo.
   4. Monitor the embryo. If the embryo resists the needle and ruptures (cytoplasm squirts out) upon entry, return to step 5 and increase the desiccation time. If the embryo is flat against the coverslip and appears floppy during the injection, return to step 5, and shorten the desiccation time.
   5. If no dye entered the embryo, confirm that dye can still freely flow from the needle as in step 6.7. If the dye does not flow, replace the needle. If the dye does flow, inject a new embryo, and begin releasing the dye just before the needle punctures the embryo.
      NOTE: Repeated injections of the same embryo are not recommended as it ruptures the previous wound/injection site.
9. Repeat until all the embryos are injected.

7. Image embryos

NOTE: This protocol utilizes a laser scanning confocal microscope.

1. Place coverslip into the metal holder on the confocal microscope so that the embryos are imaged directly from below through the coverslip; above the coverslip, there is only oil and no other barrier.
2. As a quality control step, image first using the epifluorescence function on the confocal microscope, with a 40x objective. Make sure the fluorophore is visible in the embryo before proceeding.
3. If the desired plane of imaging is different from the site of injection, allow enough time for the dye to diffuse to the target area.
   NOTE: For BOPIPY 493/503, this time typically ranges from 30-60 min. As diffusion time depends greatly on the dye, other dyes may require optimizing the site of injection and waiting time.
4. Use different objectives for imaging at different scales. Use a 40x objective to image all of the embryo and a 63x objective to image subsections for smaller scales.
5. For live imaging during the syncytial stages before cellularization, use the following conditions to start with: image size of 512 x 512 pixels, line average of 3, frame rate of 0.1 frames per second (i.e., 1 frame acquired every 10 s). Adjust conditions according to the capabilities of the microscope employed.
   ​NOTE: These conditions allow acquiring ~500+ images from BOPIPY 493/503 and capturing most of the motion.

8. Analyze LD flow by first using FIJI to prepare a time series of images, and then python for PIV analysis

1. Optimize image acquisition.
   1. Perform injections of the desired dye at the appropriate stage. Repeat this step until day-to-day variation is negligible.
   2. Optimize image acquisition settings including magnification, zoom, resolution, and frame rate.
      NOTE: There is a tradeoff between high-quality images (resolution + line averaging) and acquisition speed (frame rate).
2. Prepare data in FIJI.
   1. Acquire a high-quality time series to be analyzed.
   2. Open one frame of the time series to generate a mask with FIJI. Duplicate the image. Convert the Tip of the image to an 8 bit.
   3. Define the boundary of the embryo using the Polygon or Freehand Selection Tool. Use Clear Outside to set pixel values outside the selection to 0. Temizle and then Invert within the selection to set the value of the pixels within the selection to the maximum value (255).
   4. Unselect, then use the Histogram command to ensure that only pixel values of 0 and 255 are present.
   5. Save this new image mask for the specific time series.
   6. Open the time series of interest. Choose which ten frames best capture the time of interest, for example, nuclear cycle 9 at the onset of the cortical contraction. Duplicate the ten frames of interest, forming a substack.
   7. Use the Stack to Images function to generate 10 images to analyze with PIV.
   8. Save the individual files in a manner that preserves their order (SeriesX_1, SeriesX_2, SeriesX_3 …).
3. Use the prepared mask and individual frames for PIV analysis, employing the provided sample python script (Supplemental data) or a custom script generated by the user.
   NOTE: The provided script is based on the python application of OpenPIV¹⁷. It outputs distance in pixels which can be converted using the pixel width and frame rate to generate speeds or velocities.
4. Generate replicates by completing the previous steps for additional embryos and compiling the output values.